CHAPTER 11 BIOLOGICAL MEMBRANES AND TRANSPORT The biological membrane is a lipid bilayer with proteins of various functions (enzymes, transporters) embedded in or associated with the bilayer. The hydrophobic effect stabilizes structures (lipid bilayers and vesicles) in which lipids with some polar and some nonpolar regions can protect their nonpolar regions from interaction with the very polar solvent, water. Membrane proteins are associated with the lipid bilayer more-or-less tightly, and proteins and lipids are both allowed limited lateral motion in the plane of the bilayer. All of the internal membranes of cells are part of an interconnected, functionally specialized, and dynamic endomembrane system. Proteins and lipids synthesized in the endoplasmic reticulum move through the Golgi apparatus, and they are targeted to the membranes of organelles or to the plasma membrane, where they provide the essential properties of these structures. During this membrane trafficking, some proteins are covalently altered, and some lipids are segregated into different organelles or are concentrated in one of the bilayer leaflets. Ra s are functionally specialized regions with unique lipid and protein compositions. Although the lipid bilayer is impermeable to charged or polar solutes, cells of all kinds have many membrane transporters and ion channels that catalyze transmembrane movement of specific solutes. Some transporters merely speed the movement of solutes in the direction that simple diffusion takes them, whereas others use an energy source to move solutes against a concentration gradient. 11.1 The Composition and Architecture of Membranes We begin our discussion of biological membranes by looking at the stable bilayer structures formed spontaneously by phospholipids in water. In biological membranes we see this basic lipid bilayer combined with a wide array of membrane proteins specialized for one of the many roles that membranes play in cells. Each intracellular compartment has membranes with specific protein and lipid content, uniquely disposed across the two leaflets of the bilayer. The intracellular membranes together make up a dynamic endomembrane system that synthesizes and distributes membrane components to create functionally specialized cell compartments. Depending on its function, a protein may form one or several transmembrane segments, or it may adhere to the membrane bilayer through lipids attached to the protein, or it may alternate between membrane-associated and free forms. The Lipid Bilayer Is Stable in Water Glycerophospholipids, sphingolipids, and sterols are virtually insoluble in water. Recall that glycerophospholipids contain two long-chain fatty acids (which are hydrophobic) and a head group consisting of glycerol and one of several polar or charged substituents such as phosphocholine or phosphoethanolamine (see Fig. 10-8). Sphingolipids are constructed from a long-chain alkyl amine (sphingosine), a saturated long-chain fatty acid, and a polar head group that may be as simple as phosphocholine or may be a complex oligosaccharide (see Fig. 10-11). Sterols (cholesterol in animal membranes) have a very nonpolar steroid nucleus of four fused rings, and a polar hydroxyl group at one end of the ring system (see Fig. 10-15). Each type of membrane lipid has its particular effect on membrane structure and function. When mixed with water, these lipids spontaneously form microscopic lipid aggregates, clustering together, with their hydrophobic moieties in contact with each other and their hydrophilic groups in contact with the surrounding water. The clustering reduces the amount of hydrophobic surface exposed to water. This minimizes the number of molecules in the shell of ordered water at the lipid-water interface (see Fig. 2-7), resulting in an increase in entropy. This hydrophobic effect provides the thermodynamic driving force for the formation and maintenance of these clusters of lipid molecules. The term hydrophobic interactions is sometimes used to describe the clustering of hydrophobic molecular surfaces in an aqueous environment, but the molecules are not interacting chemically; they are simply finding the lowest-energy environment by reducing the hydrophobic, or nonpolar, surface area exposed to water. Depending on the precise conditions and the nature of the lipids, several types of lipid aggregate can form when amphipathic lipids are mixed with water (Fig. 11-1). Micelles are spherical structures that contain anywhere from a few dozen to a few thousand amphipathic molecules. These molecules are arranged with their hydrophobic regions aggregated in the interior, where water is excluded, and their hydrophilic head groups at the outer surface, in contact with water. Micelle formation is favored when the cross-sectional area of the head group is greater than that of the acyl side chain(s), as in free fatty acids, lysophospholipids (phospholipids lacking one fatty acid), and many detergents, such as sodium dodecyl sulfate (SDS; p. 88). FIGURE 11-1 Amphipathic lipid aggregates that form in water. (a) In micelles, the hydrophobic chains of the fatty acids are sequestered at the core of the sphere. There is virtually no water in the hydrophobic interior. (b) In an open bilayer, all acyl side chains except those at the edges of the sheet are protected from interaction with water. (c) When a two-dimensional bilayer folds on itself, it forms a closed bilayer, a three- dimensional hollow vesicle (liposome) enclosing an aqueous cavity. A second type of lipid aggregate in water is the bilayer, in which two lipid monolayers (leaflets) form a two-dimensional sheet. Bilayer formation is favored if the cross-sectional areas of the head group and acyl side chain(s) are similar, as in glycerophospholipids and sphingolipids. The hydrophobic portions in each monolayer, excluded from water, make contact with each other. The hydrophilic head groups interact with water at one or the other surface of the bilayer. Because the hydrophobic regions at its edges (Fig. 11-1b) are in contact with water, a bilayer sheet is relatively unstable and spontaneously folds back on itself to form a hollow sphere, called a vesicle or liposome (Fig. 11-1c). The continuous surface of vesicles eliminates exposed hydrophobic regions, allowing bilayers to achieve maximal stability in their aqueous environment. Vesicle formation also creates a separate internal aqueous compartment (the vesicle lumen). It is likely that the antecedents to the first living cells resembled lipid vesicles, their aqueous contents segregated from their surroundings by a lipid bilayer. Studies of lipid bilayers in vitro (Fig. 11-2) show that the hydrocarbon core of the bilayer, made up of the — CH2— and — CH3 of the fatty acyl groups, is about as nonpolar as decane, and about 3 nm (30 Å) thick, roughly the width of two extended fatty acyl chains. Biological membranes, to which we turn next, are 50–80 Å thick, when proteins protruding on either side are included. FIGURE 11-2 Distribution of membrane lipids across an artificial membrane. The membrane, composed of pure phosphatidylcholine in which both fatty acids are oleic acid (18:1Δ9), was studied with x-ray and neutron diffraction. The y axis shows the probability that one of the chemical moieties will be found at the position indicated on the x axis. The phosphatidylcholine molecules are shown at the same scale as the x axis. Fatty acyl chains in the hydrocarbon core are fluid and are about as nonpolar as decane. At either side, a polar region about 15 Å from the center of the bilayer contains the fatty acyl carbonyls. The glycerol, choline, and phosphate that make up the polar head group are at 18–20 Å, outside the hydrocarbon core and in contact with water. [Information from A. Rath and C. M. Deber, Annu. Rev. Biophys. 41:135, 2012, Fig. 5a, based on M. C. Wiener and S. H. White, Biophys. J. 61:434, 1992.] Bilayer Architecture Underlies the Structure and Function of Biological Membranes In the generalized membrane in Figure 11-3, phospholipids form a bilayer in which proteins are embedded, their hydrophobic domains in contact with the fatty acyl chains of membrane lipids. Some proteins protrude from only one side of the membrane; others have domains exposed on both sides. The orientation of both lipids and proteins in the bilayer is asymmetric, giving the membrane structural and functional “sidedness.” The individual lipid and protein units in a membrane form a fluid mosaic with a pattern that, unlike a mosaic of ceramic tile and mortar, is able to change as lipids and proteins move in the plane of the membrane, while maintaining the permeability barrier to polar and charged solutes. FIGURE 11-3 Fluid mosaic model for plasma membrane structure. In this simplified and general model, the fatty acyl chains in the interior of the membrane form a fluid, hydrophobic region. Integral proteins float in this sea of lipid, with their nonpolar amino acid side chains protected from interaction with water. Membranes are not merely passive barriers. They are flexible, self-repairing, and selectively permeable. Their flexibility permits the shape changes that accompany cell growth and movement (such as amoeboid movement). With their ability to break and reseal, two membranes can fuse, as in exocytosis, or a single membrane-enclosed compartment can undergo fission to yield two sealed compartments, as in endocytosis or cell division, without creating gross leaks through cellular surfaces. Their selective permeability allows them to serve as molecular gatekeepers. These membrane functions are accomplished primarily by a broad array of proteins and enzymes in and on membranes. At the cell surface, transporters move specific organic solutes and inorganic ions across the membrane; receptors sense extracellular signals and trigger molecular changes in the cell; ion channels mediate electrical signaling between cells; and adhesion molecules hold neighboring cells together. The relative proportions of protein and lipid vary with the type of membrane, reflecting the diversity of biological roles. For example, certain neurons have a myelin sheath — an extended plasma membrane that wraps around the cell many times and acts as a passive electrical insulator. The myelin sheath consists primarily of lipids (good insulators). In contrast, the plasma membranes of bacteria and the membranes of mitochondria and chloroplasts, the sites of many enzyme-catalyzed processes, contain more protein than lipid (in mass per total mass). Within the cell, membranes organize cellular processes such as the synthesis of lipids and certain proteins and the energy transductions that produce ATP in mitochondria and chloroplasts. Proteins associated with membranes are essentially confined to two-dimensional space, where intermolecular collisions of membrane proteins and lipids are far more probable than in the three-dimensional cytoplasm, so the efficiency of enzyme-catalyzed processes organized within membranes can be vastly increased. The Endomembrane System Is Dynamic and Functionally Differentiated In eukaryotic cells, the endoplasmic reticulum (ER), Golgi apparatus, lysosomes, and various small vesicles that carry lipids and proteins throughout the cell are each surrounded and defined by a single membrane. This endomembrane system (Fig. 11-4) also includes three organelles with double membranes: the nucleus, mitochondrion, and (in plants) chloroplast. (Recall the likely origin of mitochondria and chloroplasts by an endosymbiont mechanism shown in Fig. 1-37.) The convoluted and tubular ER is dynamic and extends throughout the cell, touching the other compartments at specific membrane contact points. Although each compartment is closed, they actively exchange proteins and lipids in several ways. FIGURE 11-4 Trafficking in the endomembrane system of an animal cell. Lipids and proteins move from the site of their synthesis (ER) through the Golgi apparatus to the cell surface (or to organelles such as lysosomes). Mitochondria (and in plants, chloroplasts) are also part of the endomembrane system. Small vesicles bud off the ER, move to and fuse with the cis Golgi apparatus, exit the trans Golgi apparatus as secretory or transport vesicles, and fuse with the plasma membrane or with endosomes, giving rise to lysosomes. Individual lipid molecules can be carried throughout the cell by lipid transfer proteins (LTPs), which act at membrane contact points, shown here in red. Cells have mechanisms to target specific lipids to particular organelles. Each species, each tissue or cell type, and the organelles of each cell type have a characteristic set of membrane lipids. Plasma membranes, for example, are enriched in cholesterol and sphingolipids, but they contain no detectable cardiolipin (Fig. 11-5); mitochondrial membranes are very low in cholesterol and sphingolipids, but they contain most of the cell’s phosphatidylglycerol and cardiolipin, which are synthesized within the mitochondria. Cardiolipin is specifically required for correct assembly of the respiratory complexes of mitochondria. In all but a few cases, the functional significance of these different combinations of lipids is not yet known. FIGURE 11-5 Lipid composition of the plasma and organelle membranes of a rat hepatocyte. The functional specialization of each membrane type is reflected in its unique lipid composition. Cholesterol is prominent in plasma membranes but barely detectable in mitochondrial membranes. Cardiolipin is a major component of the inner mitochondrial membrane but not of the plasma membrane. Phosphatidylserine, phosphatidylinositol, and phosphatidylglycerol are relatively minor components of most membranes but serve critical functions; phosphatidylinositol and its derivatives, for example, are important in signal transductions triggered by hormones. Sphingolipids, phosphatidylcholine, and phosphatidylethanolamine are present in most membranes but in varying proportions. Glycolipids, which are major components of the chloroplast membranes of plants, are virtually absent in animal cells. Most membrane lipids and proteins are synthesized in the ER, and from there they move to their destination organelles or to the plasma membrane. In this process of membrane trafficking, small membrane vesicles bud from the ER, then move to and fuse with the cis Golgi apparatus. As lipids and proteins move across the Golgi apparatus to its trans side, they undergo covalent modifications required for their cellular function. In many cases, these covalent alterations act as molecular ZIP codes, dictating the eventual location of the mature protein. Membrane trafficking achieves striking changes in lipid composition and disposition across the bilayer (Fig. 11-6). Phosphatidylcholine is the principal phospholipid in the lumenal leaflet of the Golgi membrane, but in transport vesicles leaving the trans Golgi apparatus, phosphatidylcholine has largely been replaced by sphingolipids and cholesterol, which, following fusion of the transport vesicles with the plasma membrane, make up the majority of the lipids in the outer leaflet of the cell’s plasma membrane. Plasma membrane lipids are asymmetrically distributed between the two leaflets of the bilayer. Choline- containing lipids (phosphatidylcholine and sphingomyelin) are typically found in the outer (extracellular, or exoplasmic) leaflet, whereas phosphatidylserine, phosphatidylethanolamine, and the phosphatidylinositols are almost exclusively in the inner (cytoplasmic) leaflet. Here the negatively charged serine and inositol phosphate head groups can interact electrostatically with positively charged regions of peripheral or amphitropic membrane proteins (described below). FIGURE 11-6 Change in lipid composition of secretory vesicles with their passage through the Golgi apparatus. Both the lipid composition of the bilayer and the disposition of specific lipids between inner and outer leaflets change remarkably as a result of vesicle trafficking. In the trans Golgi apparatus and plasma membrane, sphingolipids are greatly enriched in the leaflet facing the lumen or extracellular space, and phosphatidylserine, phosphatidylethanolamine, and the inositol phospholipids are almost exclusively in the cytosolic leaflet. The plasma membrane is also enriched with cholesterol. [Information from G. Drin, Annu. Rev. Biochem. 83:51, 2014, Fig. 1.] A second route for redistributing lipids from their site of synthesis to their destination membrane is via lipid transfer proteins (LTPs; Fig. 11-7), which occur universally in various forms. LTPs are soluble in water, but they have a hydrophobic lipid-binding pocket in which they carry a lipid from one membrane to another through the cytosol. Some LTPs are bispecific: they carry, for example, a molecule of cholesterol to the plasma membrane, and they return with a phosphatidylinositol molecule. One class of lipid transfer protein forms a hydrophobic tunnel between two membranes at their contact points. In some cases, ATP is needed to drive the process, supplied by an ATP-binding cassette (ABC) transporter (described below). The rate of lipid movement from one membrane to another in vivo is significantly greater than the rate of vesicle budding and fusion. FIGURE 11-7 Lipid transfer protein (LTP) action. (a) Lipid transfer proteins move lipids and other nonpolar solutes through the very polar cytosol by providing a hydrophobic groove or pocket into which the hydrophobic solute can fit, with its nonpolar regions masked from the surrounding water. (b) Some LTPs form multisubunit structures to generate a hydrophobic groove long enough to connect two separate membranes at contact points (see Fig. 11-5). (c) To move lipid molecules against a concentration gradient (say, from a leaflet poor in phosphatidylserine to one relatively rich in it), the LTP is coupled to an ATP-dependent pump. [Information from L. H. Wong et al., Nat. Rev. Mol. Cell Biol. 20:85, 2019, Fig. 2.] Finally, once phospholipids reach their destination membrane, they can undergo enzymatic remodeling of their fatty acid constituents, a subject discussed in Chapter 21. Membrane Proteins Are Receptors, Transporters, and Enzymes The protein composition of membranes reflects each membrane’s functional specialization. One large group of membrane proteins are the receptors for extracellular signals such as hormones and changes in membrane potential. Hundreds of membrane proteins are transporters that carry specific polar or charged compounds — sugars, amino acids, vitamins, a wide variety of metabolic intermediates, minerals (M g2+, Ca2+, Na+, K+), and trace metal ions (M n2+, Ni2+, Co2+) — across the plasma membrane or between organelles. All transporters and many receptors must span the membrane at least once to play their roles. Some membrane-associated enzymes also span the membrane, but many carry out their catalytic functions on one side or the other of their membrane setting, partially penetrating just one leaflet of the bilayer. The synthesis and delivery to the site of function are more complicated for membrane proteins than for typical cytosolic proteins. Many of the proteins destined for the plasma membrane undergo extensive posttranslational modification as they pass through the ER and the Golgi apparatus. One covalent modification is the glycosylation (attachment of oligosaccharides) of proteins bound for the plasma membrane. In glycophorin, a glycoprotein of the erythrocyte plasma membrane, 60% of the mass consists of complex oligosaccharides covalently attached to specific amino acid residues (Fig. 11-8). For such glycoproteins, Ser, Thr, and Asn residues are the most common points of carbohydrate attachment (see Fig. 7-27), and the carbohydrates are invariably on the outer face of the plasma membrane. Another posttranslational modification of some proteins is the covalent attachment of one or more lipids, which serve as hydrophobic anchors to hold a protein to the membrane or as tags to target the protein to a specific location.
FIGURE 11-8 Transbilayer disposition of glycophorin in an erythrocyte. One hydrophilic domain, containing all the sugar residues, is on the outer surface, and another hydrophilic domain protrudes from the inner face of the membrane. Each red hexagon represents a tetrasaccharide (containing two Neu5Ac (sialic acid), Gal, and GalNAc) O-linked to a Ser or Thr residue; the blue hexagon represents an oligosaccharide N-linked to an Asn residue. The oligosaccharides are much larger than the hexagons used to represent them. A segment of 19 hydrophobic residues (residues 75 to 93) forms an α helix that traverses the membrane bilayer (see Fig. 11-13a). The segment from residues 64 to 74 has some hydrophobic residues and probably penetrates the outer face of the lipid bilayer, as shown. The functional unit of glycophorin is a homodimer; for clarity we show only one of the subunits here. [Information from V. T. Marchesi et al., Annu. Rev. Biochem. 45:667, 1976.] Membrane Proteins Differ in the Nature of Their Association with the Membrane Bilayer Integral membrane proteins are firmly embedded within the lipid bilayer and are removable only by agents such as detergents or organic solvents that overcome hydrophobic interactions (Fig. 11-9). Peripheral membrane proteins associate with the membrane through electrostatic interactions and hydrogen bonding with hydrophilic domains of integral proteins and with the charged head groups of membrane lipids. In the laboratory, they can be released from their membrane association by relatively mild treatments that interfere with electrostatic interactions or that break hydrogen bonds. Amphitropic proteins associate reversibly with membranes and are therefore found both in the membrane and in the cytosol. Their affinity for membranes results in some cases from the protein’s noncovalent interaction with another membrane protein or lipid, and in other cases from the presence of one or more covalently attached lipids (see Fig. 11-16). FIGURE 11-9 Integral, peripheral, and amphitropic proteins. Membrane proteins can be operationally distinguished by the conditions required to release them from the membrane. Integral proteins can only be removed from membranes with harsh treatment such as detergent. Peripheral proteins are more loosely associated. Amphitropic proteins have a reversible association with membranes that depends on a regulatory process, such as reversible palmitoylation or phosphorylation. The firm attachment of integral proteins to membranes is the result of the hydrophobic interaction between the nonpolar core of the lipid bilayer and nonpolar side chains of the protein. Monotopic integral proteins have small hydrophobic domains that interact with only a single leaflet of the membrane (Fig. 11- 10). These domains, which make up as little as a few percent of the protein’s total mass, provide enough hydrophobic surface to hold the proteins firmly to the membrane. Bitopic proteins span the bilayer once, extending on either surface. They have a single hydrophobic sequence somewhere in the molecule. Glycophorin (Fig. 11-8) is bitopic. Polytopic proteins cross the membrane several times. They have multiple hydrophobic sequences, each of which, when in the α -helical conformation, is long enough (about 20 residues) to span the lipid bilayer. (Recall from Worked Example 4-1 that each residue in an α helix adds 1.5 Å to its length.) FIGURE 11-10 Monotopic proteins penetrate only one leaflet. Three typical monotopic membrane proteins have relatively little of their structure (1% to 16%) associated with lipids in one leaflet of the bilayer. They are colored to show surface hydrophobicity, with yellow hydrophobic, blue hydrophilic, and hues of green in between. Each of these proteins catalyzes a reaction that involves a hydrophobic substrate. (a) CPT-II is a mammalian carnitine palmitoyltransferase II; (b) COX2 is a mammalian prostaglandin H2 synthase-1; (c) PlsC is a bacterial acyltransferase (1-acyl-sn-glycerol-6- phosphate acyltransferase). [Data from (a) PDB ID 2H4T, Y. S. Hsiao et al., Biochem. Biophys. Res. Commun. 346:974, 2006; (b) PDB ID 1CQE, D. Picot et al., Nature 367:243, 1994; (c) PDB ID 5KYM, R. M. Robertson et al., Nat. Struct. Mol. Biol. 24:666, 2017.] One of the best-studied polytopic proteins, bacteriorhodopsin, has seven very hydrophobic internal sequences and spans the lipid bilayer seven times. Bacteriorhodopsin is a light-driven proton pump densely packed in regular arrays in the purple membrane of the bacterium Halobacterium salinarum. X-ray crystallography and cryo-electron microscopy reveal a structure in which each of the seven α -helical segments crosses the lipid bilayer, and all are connected by nonhelical loops at the inner and outer faces of the membrane (Fig. 11-11). In the amino acid sequence of bacteriorhodopsin, seven segments of about 20 hydrophobic residues can be identified, each forming an α helix that spans the bilayer. The seven helices are clustered together and oriented not quite perpendicular to the bilayer plane, a motif that (as we shall see in Chapter 12) is very common in membrane proteins involved in signal reception. The hydrophobic interaction keeps the nonpolar amino acid residues firmly anchored among the fatty acyl groups of the membrane lipids in the membrane’s lipid core. FIGURE 11-11 Bacteriorhodopsin, a membrane-spanning protein. The single polypeptide chain folds into seven hydrophobic α helices, each of which traverses the lipid bilayer roughly perpendicular to the plane of the membrane. The seven transmembrane helices are clustered, and the space around and between them is filled with the acyl chains of membrane lipids. The light-absorbing pigment retinal (see Fig. 10-20) is buried deep within the membrane in contact with several of the helical segments (not shown). The helices are colored to correspond with the hydropathy plot in Figure 11- 13. [Data from PDB ID 2AT9, K. Mitsuoka et al., J. Mol. Biol. 286:861, 1999.] Many membrane proteins co-crystalize with phospholipid molecules, which are presumed to be positioned in the native membranes as they are in the protein crystals. Many of these phospholipid molecules lie on the protein surface, their head groups interacting with polar amino acid residues at the inner and outer membrane–water interfaces and their side chains associated with nonpolar residues. Other phospholipids are found at the interfaces between monomers of multisubunit membrane proteins, where they form a “grease seal” (Fig. 11-12). FIGURE 11-12 Lipid annuli associated with an integral protein. The crystal structure of sheep aquaporin, a transmembrane water channel, includes a shell of phospholipids positioned with their head groups (blue) at the expected positions on the inner and outer membrane surfaces and their hydrophobic acyl chains (gold) intimately associated with the surface of the protein exposed to the bilayer. The lipid forms a “grease seal” around the protein, which is depicted by a dark blue surface representation. [Data from PDB ID 2B6O, T. Gonen et al., Nature 438:633, 2005.] The Topology of an Integral Membrane Protein Can O en Be Predicted from Its Sequence Determination of the location of a membrane protein relative to the bilayer — that is, its topology — is generally much more difficult than determining its amino acid sequence, either directly or by gene sequencing. The amino acid sequences of many thousands of membrane proteins are known, but fewer three- dimensional structures have been established by x-ray crystallography or cryo-electron microscopy. The presence of unbroken sequences of more than 20 hydrophobic residues in a membrane protein is commonly taken as evidence that these sequences traverse the lipid bilayer, acting as hydrophobic anchors or forming transmembrane channels. All known integral proteins have at least one such sequence. Application of this logic to entire genomic sequences leads to the conclusion that in many organisms, 20% to 30% of all proteins are integral proteins. What can we predict about the secondary structure of the membrane-spanning portions of integral proteins? At 1.5 Å (0.15 nm) per amino acid residue, an α -helical sequence of 20 to 25 residues is just long enough to span the thickness (30 Å) of the lipid bilayer. A polypeptide chain surrounded by lipids, having no water molecules with which to hydrogen-bond, will tend to form α helices or β sheets in which intrachain hydrogen bonding is maximized. If the side chains of all amino acids in a helix are nonpolar, the helix is further stabilized in the surrounding lipids by the hydrophobic effect. Several simple methods of analyzing amino acid sequences yield reasonably accurate predictions of secondary structure for transmembrane proteins. The relative polarity of each amino acid has been determined experimentally by measuring the free- energy change accompanying the movement of that amino acid side chain from a hydrophobic environment into water. This free energy of transfer, which can be expressed as a hydropathy index (see Table 3-1), ranges from highly exergonic for charged or polar residues to highly endergonic for amino acids with aromatic or aliphatic hydrocarbon side chains. The overall hydropathy index (hydrophobicity) of a sequence of amino acids is estimated by summing the free energies of transfer for the residues in the sequence. To scan a polypeptide sequence for potential membrane-spanning segments, an investigator calculates the hydropathy index for successive segments (called windows) of a given size, from 7 to 20 residues. For a window of seven residues, for example, the average indices for residues 1 to 7, 2 to 8, 3 to 9, and so on, are plotted as in Figure 11-13 (plotted for the middle residue in each window — residue 4 for residues 1 to 7, for example). A region with more than 20 residues of high hydropathy index is presumed to be a transmembrane segment. When the sequences of membrane proteins of known three-dimensional structure are scanned using simple online bioinformatics tools, we find a reasonably good correspondence between predicted and known membrane-spanning segments. Hydropathy analysis predicts a single hydrophobic helix for glycophorin (Fig. 11-13a) and seven transmembrane segments for bacteriorhodopsin (Fig. 11-13b) — in agreement with structures known from x-ray crystallography. FIGURE 11-13 Hydropathy plots. Average hydropathy index (see Table 3-1) is plotted against residue number for two integral membrane proteins. The hydropathy index for each amino acid residue in a sequence of defined length, or “window,” is used to calculate the average hydropathy for that window. The horizontal axis shows the residue number in the middle of the window. (a) Glycophorin from human erythrocytes has a single hydrophobic sequence between residues 75 and 93 (yellow); compare this with Figure 11-8. (b) Bacteriorhodopsin, known from independent physical studies to have seven transmembrane helices (see Fig. 11-11), has seven hydrophobic regions. Note, however, that the hydropathy plot is ambiguous in the region of segments 6 and 7. X-ray crystallography has confirmed that this region has two transmembrane segments. Not all integral proteins are composed of transmembrane α helices. Another structural motif common in bacterial and mitochondrial membrane proteins is the β barrel (see Fig. 4- 16b), in which 20 or more transmembrane segments form β sheets that line a cylinder (Fig. 11-14). The same factors that favor α helix formation in the hydrophobic interior of a lipid bilayer also stabilize β barrels: when no water molecules are available to hydrogen-bond with the carbonyl oxygen and nitrogen of the peptide bond, maximal intrachain hydrogen bonding gives the most stable conformation. Planar β sheets do not maximize these interactions and are generally not found in the membrane interior; β barrels allow all possible hydrogen bonds and are common among membrane proteins. Porins, proteins that allow certain polar solutes to cross the outer membrane of gram- negative bacteria such as E. coli, have many-stranded β barrels lining the polar transmembrane passage. The outer membranes of mitochondria and chloroplasts also contain a variety of β barrels, as do the outer membranes of the modern bacteria descended from those believed to have evolved to mitochondria and chloroplasts.
FIGURE 11-14 Polytopic integral proteins with β -barrel structure. Three proteins of the E. coli outer membrane are shown, viewed in the plane of the membrane. FepA, involved in iron uptake, has 22 membrane-spanning β strands. Outer membrane phospholipase A, or OmpLA, is a 12-stranded β barrel that exists as a dimer in the membrane. Maltoporin, a maltose transporter, is a trimer; each monomer consists of 16 β strands. [Data from FepA, PDB ID 1FEP, S. K. Buchanan et al., Nat. Struct. Biol. 6:56, 1999; OmpLA, PDB ID 1QD5, H. J. Snijder et al., Nature 401:717, 1999; maltoporin, PDB ID 1MAL, T. Schirmer et al., Science 267:512, 1995.] A polypeptide is more extended in the β conformation than in an α helix; just seven to nine residues of β conformation are needed to span a membrane. Recall that in the β conformation, alternating side chains project above and below the sheet (see Fig. 4-5). In β strands of membrane proteins, every second residue in the membrane-spanning segment is hydrophobic and interacts with the lipid bilayer; aromatic side chains are commonly found at the lipid-protein interface. The other residues may or may not be hydrophilic. A further remarkable feature of many transmembrane proteins of known structure is the presence of Tyr and Trp residues at the interface between lipid and water (Fig. 11-15). The side chains of these residues seem to serve as membrane interface anchors, able to interact simultaneously with the central lipid phase and the aqueous phases on either side of the membrane. Another generalization about amino acid location relative to the bilayer is described by the positive-inside rule: the positively charged Lys and Arg residues in the extramembrane loop of membrane proteins occur more commonly on the cytoplasmic face of plasma membranes. FIGURE 11-15 Tyr and Trp residues of integral proteins clustering at the water-lipid interface. The detailed structures of these five integral proteins are known from crystallographic studies. The K+ channel is from the bacterium Streptomyces lividans (see Fig. 11-45); maltoporin, OmpLA, OmpX, and phosphoporin E are proteins of the outer membrane of E. coli. Residues of Tyr and Trp are found predominantly where the nonpolar region of acyl chains meets the polar head-group region. Charged residues (Lys, Arg, Glu, Asp) are found almost exclusively in the aqueous phases. [Data from K+ channel, PDB ID 1BL8, D. A. Doyle et al., Science 280:69, 1998; maltoporin, PDB ID 1AF6, Y. F. Wang et al., J. Mol. Biol. 272:56, 1997; OmpLA, PDB ID 1QD5, H. J. Snijder et al., Nature 401:717, 1999; OmpX, PDB ID 1QJ9, J. Vogt and G. E. Schulz, Structure 7:1301, 1999; phosphoporin E, PDB ID 1PHO, S. W. Cowan et al., Nature 358:727, 1992.] Covalently Attached Lipids Anchor or Direct Some Membrane Proteins Some membrane proteins are covalently linked to one or more lipids, which may be of several types: long-chain fatty acids, isoprenoids, sterols, or glycosylated derivatives of phosphatidylinositol (GPIs; Fig. 11-16). The attached lipid provides a hydrophobic anchor that inserts into the lipid bilayer and holds the protein at the membrane surface. The strength of the hydrophobic interaction between a bilayer and a single hydrocarbon chain linked to a protein is barely enough to anchor the protein securely, but many proteins have more than one attached lipid moiety. Furthermore, other interactions, such as ionic attractions between positively charged Lys residues in the protein and negatively charged lipid head groups, can add to the anchoring effect of a covalently bound lipid. For example, the plasma membrane protein MARCKS (myristoylated alanine-rich C-kinase substrate), which interacts with actin filaments in the process of cell motility, has a covalently attached myristoyl moiety, but it also contains the sequence which adds to the protein’s affinity for the membrane. Three clusters of positively charged Lys and Arg residues (screened blue) interact with the negatively charged head group of phosphatidylinositol 4,5-bisphosphate (PIP2) on the cytoplasmic face of the plasma membrane; five aromatic residues (screened yellow) insert into the lipid bilayer. When the head-group phosphates of PIP2 are enzymatically removed, MARCKS loses its hold on the plasma membrane and dissociates. The reversible nature of the association of MARCKS with the membrane makes it an amphitropic protein. FIGURE 11-16 Lipid-linked membrane proteins. Covalently attached lipids anchor membrane proteins to the lipid bilayer. A palmitoyl group is shown attached by thioester linkage to a Cys residue; an N-myristoyl group is generally attached to an amino- terminal Gly residue, typically of a protein that also has a hydrophobic transmembrane segment; the farnesyl and geranylgeranyl groups attached to carboxyl-terminal Cys residues are isoprenoids of 15 and 20 carbons, respectively. The carboxyl-terminal Cys residue is invariably methylated. Glycosyl phosphatidylinositol (GPI) anchors are derivatives of phosphatidylinositol in which the inositol bears a short oligosaccharide covalently joined to the carboxyl-terminal residue of a protein through phosphoethanolamine. GPI-anchored proteins are always on the extracellular face of the plasma membrane. Farnesylated and palmitoylated membrane proteins are found on the inner surface of the plasma membrane, and myristoylated proteins have domains both inside and outside the plasma membrane. For some lipid-linked proteins, the lipid attachment process is reversible, so the protein is amphitropic: membrane-bound when lipid-linked, soluble when not. Beyond merely anchoring a protein to the membrane, the attached lipid may have a more specific role. In the plasma membrane, GPI-anchored proteins are exclusively on the outer face and are clustered in certain regions, as discussed later in the chapter (p. 380), whereas other types of lipid-linked proteins (with farnesyl or geranylgeranyl groups attached; Fig. 11-16) are exclusively on the inner face. In polarized epithelial cells (such as intestinal epithelial cells), in which apical and basal surfaces have different roles, GPI-anchored proteins are directed specifically to the apical surface. Attachment of a specific lipid to a newly synthesized membrane protein therefore has a targeting function, directing the protein to its correct cellular location. SUMMARY 11.1 The Composition and Architecture of Membranes Biological phospholipids and sterols spontaneously form a lipid bilayer to protect their hydrophobic hydrocarbon chains from energetically unfavorable interaction with water. In the fluid mosaic model, a lipid bilayer is the basic structural unit, and proteins associate with or span the bilayer. Biological membranes are flexible, self-repairing, and selectively permeable. They regulate the traffic of small molecules into and out of the cell and among the organelles, and they provide a scaffold on which proteins assemble into catalytic and structural aggregates that function in two-dimensional space. Proteins and lipids are trafficked through the dynamic endomembrane system of a eukaryote from their point of synthesis to their appropriate cellular locations. Small vesicles and soluble transporter proteins ensure that each membrane leaflet has its unique complement of lipids and proteins to achieve its specialized function. Cells have hundreds of membrane transporters that carry polar solutes and ions in and out of cells and throughout the endomembrane system. The plasma membrane contains protein receptors that sense signals from outside the cell and carry their message into the cell. Many enzymes are associated with membranes, where they interact with each other in essentially two-dimensional space. Integral proteins are embedded within membranes, their nonpolar amino acid side chains stabilized by contact with the lipid bilayer. Peripheral proteins associate with membranes through electrostatic interactions and hydrogen bonding with membrane phospholipids and integral proteins. Amphitropic proteins associate reversibly with the membrane. Many integral membrane proteins span the lipid bilayer several times, with hydrophobic sequences of about 20 amino acid residues forming transmembrane α helices. Multistranded β barrels are also common in integral proteins of bacterial and mitochondrial membranes. A hydropathy plot of the amino acid sequence identifies segments that are likely to cross the lipid bilayer as helices or barrels. Covalent attachment of a hydrophobic molecule such as a fatty acid serves to anchor some membrane proteins to the bilayer. 11.2 Membrane Dynamics One remarkable feature of all biological membranes is their plasticity — their ability to change shape without losing their integrity and becoming leaky. The bases for this property are the noncovalent interactions among lipids in the bilayer and the mobility allowed to individual lipids because they are not covalently anchored to one another. We turn now to the dynamics of membranes: the motions that occur and the transient structures that are allowed by these motions. Acyl Groups in the Bilayer Interior Are Ordered to Varying Degrees Although the lipid bilayer structure is stable, its individual phospholipid molecules have much freedom of motion (Fig. 11- 17), depending on the temperature and the lipid composition. Below normal physiological temperatures, the lipids in a bilayer form a gel-like liquid-ordered (Lo) state, in which all types of motion of individual lipid molecules are strongly constrained (Fig. 11-17a). Above physiological temperatures, individual hydrocarbon chains of fatty acids are in constant motion produced by rotation about the carbon–carbon bonds of the long acyl side chains and by lateral diffusion of individual lipid molecules in the plane of the bilayer. This is the liquid- disordered (Ld) state (Fig. 11-17b). In the transition from the Lo state to the Ld state, the general shape and dimensions of the bilayer are maintained; what changes is the degree of motion (lateral and rotational) allowed to individual lipid molecules. FIGURE 11-17 Two extreme states of bilayer lipids. (a) In the liquid- ordered (Lo) state, polar head groups are uniformly arrayed at the surface, and the acyl chains are nearly motionless and packed with regular geometry. (b) In the liquid-disordered (Ld) state, or fluid state, acyl chains undergo much thermal motion and have no regular organization. The state of membrane lipids in biological membranes is maintained somewhere between these extremes. [Information from H. Heller et al., J. Phys. Chem. 97:8343, 1993.] At temperatures in the physiological range for a mammal (about 20 to 40 °C), long-chain saturated fatty acids (such as 16:0 and 18:0) tend to pack into an Lo gel phase, but the kinks in unsaturated fatty acids interfere with packing, favoring the Ld state (see Fig. 10-1). Shorter-chain fatty acyl groups are more mobile than longer-chain fatty acyl groups and thus also favor the Ld state. The sterol content of a membrane, which varies greatly with organism and organelle, is another important determinant of lipid state. Sterols (such as cholesterol) have paradoxical effects on bilayer fluidity: they interact with phospholipids containing unsaturated fatty acyl chains, compacting them and constraining their motion in bilayers. In contrast, their association with sphingolipids and phospholipids having long, saturated fatty acyl chains tends to make a bilayer fluid that would otherwise, without cholesterol, adopt the Lo state. In biological membranes composed of a variety of phospholipids and sphingolipids, cholesterol tends to associate with sphingolipids and to form regions in the Lo state surrounded by cholesterol-poor regions in the Ld state (see the discussion of membrane ra s below). Transbilayer Movement of Lipids Requires Catalysis At physiological temperatures, lateral diffusion in the plane of the bilayer is very rapid, but the movement of a lipid molecule from one leaflet of the bilayer to the other occurs very slowly, if at all, in most membranes (Fig. 11-18). Transbilayer — or “flip-flop” — movement requires that a polar or charged head group leave its aqueous environment and move into the hydrophobic interior of the bilayer, a process with a large, positive free-energy change. There are, however, situations in which such movement is essential. For example, in the ER, membrane glycerophospholipids are synthesized on the cytosolic face, whereas sphingolipids are synthesized or modified on the lumenal surface. To get from their site of synthesis to their eventual point of deposition, these lipids must undergo flip-flop diffusion. FIGURE 11-18 Motion of single phospholipids in a bilayer. (a) Lateral diffusion within a leaflet is very rapid, but (b) uncatalyzed movement from one leaflet to the other is very slow. (c) Three types of phospholipid translocators in the plasma membrane. PE is phosphatidylethanolamine; PS is phosphatidylserine. Membrane proteins called flippases, floppases, and scramblases (Fig. 11-18c) facilitate the transbilayer movement (translocation) of individual lipid molecules. Like enzymes, these translocators act by providing a path that is energetically more favorable and therefore much faster than the uncatalyzed movement. The combination of asymmetric biosynthesis of membrane lipids, no uncatalyzed flip-flop diffusion, and the presence of selective, energy-dependent lipid translocators accounts for the transbilayer asymmetry in lipid composition discussed in Section 11.1. Flippases catalyze translocation of the aminophospholipids phosphatidylethanolamine and phosphatidylserine from the extracellular to the cytoplasmic leaflet of the plasma membrane, contributing to the asymmetric distribution of phospholipids: phosphatidylethanolamine and phosphatidylserine primarily in the cytoplasmic leaflet, and the sphingolipids and phosphatidylcholine in the outer leaflet. Keeping phosphatidylserine out of the extracellular leaflet is important: its exposure on the outer surface triggers apoptosis (programmed cell death; see Chapter 12) and engulfment by macrophages that carry phosphatidylserine receptors. Flippases also act in the ER, where they move newly synthesized phospholipids from their site of synthesis in the cytosolic leaflet to the lumenal leaflet. Flippases consume about one ATP per molecule of phospholipid translocated, and they are structurally and functionally related to the P-type ATPases (active transporters) described in Section 11.3. There are three other types of lipid-translocating activities: floppases, scramblases, and phosphatidylinositol transfer proteins. Floppases move plasma membrane phospholipids and sterols from the cytoplasmic leaflet to the extracellular leaflet and, like flippases, are ATP-dependent. Floppases are members of the ABC transporter family, described on page 395; all ABC transporters actively transport hydrophobic substrates outward across the plasma membrane. Each floppase specializes in movement of specific lipids: cholesterol, phosphatidylcholine, sphingomyelin, and phosphatidylserine. Scramblases are proteins that move any membrane phospholipid across the bilayer down its concentration gradient (from the leaflet where it has a higher concentration to the leaflet where it has a lower concentration); their activity is not dependent on ATP, although some require Ca2+. Scramblase activity leads to controlled randomization of the head-group composition on the two faces of the bilayer. The activity of some scramblases rises sharply with an increase in cytosolic Ca2+ concentration, which may result from cell activation, cell injury, or apoptosis. Finally, a group of proteins that act primarily to move phosphatidylinositol lipids across lipid bilayers, the phosphatidylinositol transfer proteins, are believed to have important roles in lipid signaling and membrane trafficking. Lipids and Proteins Diffuse Laterally in the Bilayer Individual lipid molecules can move laterally in the plane of the membrane by changing places with neighboring lipid molecules; that is, they undergo Brownian movement within the bilayer (Fig. 11-18a). This rapid lateral diffusion in the plane of the bilayer tends to randomize the positions of individual molecules in a few seconds. Lateral diffusion can be shown experimentally by attaching fluorescent probes to the head groups of lipids and using fluorescence microscopy to follow the probes over time (Fig. 11- 19). In one technique, a small region (5 μm 2) of a cell surface with fluorescence-tagged lipids is bleached by intense laser radiation so that the irradiated patch no longer fluoresces when viewed with less-intense (nonbleaching) light in the fluorescence microscope. However, within milliseconds, the region recovers its fluorescence as unbleached lipid molecules diffuse into the bleached patch and bleached lipid molecules diffuse away from it. The rate of fluorescence recovery a er photobleaching, or FRAP, is a measure of the rate of lateral diffusion of the lipids. Using the FRAP technique, researchers have shown that some membrane lipids diffuse laterally at rates of up to 1 μ m/s. At this rate, a lipid molecule could move from one end of a eukaryotic cell to the other in a few seconds. FIGURE 11-19 Measurement of lateral diffusion rates of lipids by fluorescence recovery a er photobleaching (FRAP). Lipids in the outer leaflet of the plasma membrane are labeled by reaction with a membrane- impermeant fluorescent probe (red) so that the surface is uniformly labeled when viewed with a fluorescence microscope. A small area is bleached with a laser, then recovers its fluorescence. With time, unbleached lipid molecules diffuse into the bleached region, and it again becomes fluorescent. The FRAP method can also be used to measure lateral diffusion of membrane proteins labeled with a fluorescent tag. Another technique, single particle tracking, has allowed researchers to follow the movement of a single lipid molecule in the plasma membrane on a much shorter time scale. Results from these studies confirm that lipid molecules diffuse laterally with rapidity within small, discrete regions of the cell surface but that movement from one such region to a nearby region (“hop diffusion”) is rarer; membrane lipids behave as though corralled by fences that they can occasionally cross by hop diffusion (Fig. 11-20). FIGURE 11-20 Hop diffusion of individual lipid molecules. The motion of a single fluorescently labeled lipid molecule in a cell surface is recorded on video by fluorescence microscopy, with a time resolution of 25 µs (equivalent to 40,000 frames/s). The track shown here represents a molecule followed for 56 ms (2,250 frames); the trace begins in the purple area and continues through blue, green, and orange. The pattern of movement indicates rapid diffusion within a confined region (about 250 nm in diameter, shown by a single color), with occasional hops into an adjoining region. This finding suggests that the lipids are corralled by molecular fences that they occasionally jump. [Data from Takahiro Fujiwara, Ken Ritchie, Hideji Murakoshi, Ken Jacobson, and Akihiro Kusumi.] Like membrane lipids, many membrane proteins are free to diffuse laterally in the plane of the bilayer and are in constant motion, as shown by the FRAP technique with fluorescence- tagged plasma membrane proteins. Some membrane proteins associate to form large aggregates (“patches”) on the surface of a cell or an organelle in which individual protein molecules do not move relative to one another; for example, acetylcholine receptors form dense, near-crystalline patches on neuronal plasma membranes at synapses. Other membrane proteins are anchored to internal structures that prevent their free diffusion. In the erythrocyte membrane, both glycophorin and the chloride- bicarbonate exchanger (p. 389) are tethered to spectrin, a filamentous cytoskeletal protein (Fig. 11-21). One possible explanation for the pattern of lateral diffusion of lipid molecules shown in Figure 11-20 is that membrane proteins immobilized by their association with spectrin form the “fences” that define the regions within which relatively unrestricted lipid motion can occur. FIGURE 11-21 Restricted motion of the erythrocyte chloride-bicarbonate exchanger and glycophorin. The proteins span the membrane and are tethered to spectrin, a cytoskeletal protein, by another protein, ankyrin, limiting their lateral mobility. Ankyrin is anchored in the membrane by a covalently bound palmitoyl side chain (see Fig. 11-16). Spectrin, a long, filamentous protein, is cross-linked at junctional complexes containing actin. A network of cross-linked spectrin molecules attached to the cytoplasmic face of the plasma membrane stabilizes the membrane, making it resistant to deformation. This network of anchored membrane proteins may form the “corral” suggested by the experiment shown in Figure 11-20; the lipid tracks shown here are confined to different regions defined by the tethered membrane proteins. Occasionally a lipid molecule (green track) jumps from one corral to another (blue track), then another (red track). Sphingolipids and Cholesterol Cluster Together in Membrane Ra s We have seen that diffusion of membrane lipids from one bilayer leaflet to the other does not occur unless catalyzed and that the different lipid species of the plasma membrane are asymmetrically distributed in the two leaflets of the bilayer (Fig. 11-6). Even within a single leaflet, the lipid distribution is not uniform. Glycosphingolipids (cerebrosides and gangliosides), which typically contain long-chain saturated fatty acids, form transient clusters in the outer leaflet; these clusters largely exclude glycerophospholipids, which typically contain one unsaturated fatty acyl group and a shorter saturated acyl group. The long, saturated acyl groups of sphingolipids can form more compact, more stable associations with the long ring system of cholesterol than can the shorter, o en unsaturated, chains of phospholipids. The cholesterol-sphingolipid microdomains of the plasma membrane make the bilayer slightly thicker and more ordered (less fluid) than neighboring regions, which are rich in phospholipids. These microdomains are more difficult to dissolve with nonionic detergents; they behave like liquid-ordered sphingolipid ra s adri on an ocean of liquid-disordered phospholipids (Fig. 11-22). Proteins with relatively short hydrophobic helical sections (19 to 20 residues) cannot span the thicker bilayer in ra s, and thus they tend to be excluded. Proteins with longer hydrophobic helices (24 to 25 residues) segregate into the thicker bilayer regions of ra s, where the entire length of the helix is stabilized by the hydrophobic effect. FIGURE 11-22 Membrane microdomains (ra s). Stable associations of sphingolipids and cholesterol in the outer leaflet produce a microdomain, slightly thicker than other membrane regions, that is enriched with specific types of membrane proteins. GPI-anchored proteins are prominent in the outer leaflet of these ra s, and proteins with one or several covalently attached long-chain acyl groups are common in the inner leaflet. Inwardly curved ra s called caveolae are especially enriched in proteins called caveolins (see Fig. 11-23). Lipid ra s are remarkably enriched in two classes of integral proteins, with two specific types of covalently attached lipids. The integral proteins of one class have two long-chain saturated fatty acids (two palmitoyl groups, or a palmitoyl and a myristoyl group) covalently attached through Cys residues. Caveolin is one such protein; there are many others. Those of the second class, the GPI-anchored proteins, have a glycosyl phosphatidylinositol on their carboxyl-terminal residue (Fig. 11-16). Presumably, these lipid anchors, like the long, saturated acyl chains of sphingolipids, form more stable associations with the cholesterol and long acyl groups in ra s than they form with the surrounding phospholipids. (It is notable that other lipid-linked proteins, those with covalently attached isoprenyl groups such as farnesyl, are not preferentially associated with the outer leaflet of sphingolipid ra s; see Fig. 11-22.) The “ra ” and “sea” domains of the plasma membrane are not rigidly separated; membrane proteins can move into and out of lipid ra s in a fraction of a second. But in the shorter time scale (microseconds) more relevant to many membrane-mediated biochemical processes, many of these proteins reside primarily in a ra . The approximate fraction of the cell surface occupied by ra s can be as high as 50% in some cases; the ra s cover half of the ocean. Indirect measurements in cultured fibroblasts suggest a diameter of roughly 50 nm for an individual ra , which corresponds to a patch containing a few thousand sphingolipids and perhaps 10 to 50 membrane proteins. Because most cells express more than 50 different kinds of plasma membrane proteins, it is likely that a single ra contains only a subset of membrane proteins and that this segregation of membrane proteins is functionally significant. Their presence in a single ra would hugely increase the likelihood of their collision. Certain membrane receptors and signaling proteins, for example, seem to be segregated together in membrane ra s. Experiments show that signaling through these proteins can be disrupted by manipulations that deplete the plasma membrane of cholesterol and destroy lipid ra s. Plasma membranes of many cells have specialized ra s called caveolae (“little caves”), which can represent about half of the total area of the plasma membrane (Fig. 11-23a). Closely associated with caveolae is caveolin, an integral protein with two globular domains connected by a hairpin-shaped hydrophobic domain that binds the protein to the cytoplasmic leaflet of the plasma membrane (Fig. 11-23b). Three palmitoyl groups attached to the carboxyl-terminal globular domain further anchor the protein to the membrane. Caveolins form dimers and associate with cholesterol-rich regions in the membrane. The presence of caveolin dimers forces the associated lipid bilayer to curve inward, forming caveolae. Caveolae involve both leaflets of the bilayer — the cytoplasmic leaflet, from which the caveolin globular domains project, and the extracellular leaflet, a cholesterol and sphingolipid ra with associated GPI-anchored proteins. Caveolae are implicated in a variety of cellular functions, including membrane trafficking within cells and the transduction of external signals into cellular responses.
FIGURE 11-23 Caveolin forces inward curvature of a membrane. (a) Caveolae are small invaginations in the plasma membrane. (b) Cartoon showing the location and role of a caveolin dimer in causing inward membrane curvature. Each caveolin monomer has a central hydrophobic domain and three long-chain acyl groups (red), which hold the molecule to the inside of the plasma membrane. When several caveolin dimers are concentrated in a small region (a ra ), they force a curvature in the lipid bilayer, forming a caveola. Cholesterol molecules in the bilayer are shown in orange. (c) Flattening of caveolae allows the plasma membrane to expand in response to various stresses. [Information from R. G. Parton, Nat. Rev. Mol. Cell Biol. 8:185–194, 2007.] Caveolae may also provide a means of expanding the cell surface. The lipid bilayer itself is not elastic, but if existing caveolae lose their associated caveolin as the result of a regulatory signal, the caveolae flatten into the plasma membrane (Fig. 11-23c). The effect is to add surface area, allowing the cell to expand without bursting in response to osmotic or other stress. Membrane Curvature and Fusion Are Central to Many Biological Processes Caveolins are not unique in their ability to induce curvature in membranes. Changes of curvature are central to one of the most remarkable features of biological membranes: their ability to undergo fusion with other membranes without losing their continuity. Within the eukaryotic endomembrane system, the membranous compartments constantly reorganize. Vesicles bud from the ER to carry newly synthesized lipids and proteins to other organelles and to the plasma membrane. Exocytosis, endocytosis, cell division, fusion of egg and sperm cells, and entry of a membrane-enveloped virus into a host cell all involve a membrane reorganization that requires the fusion of two membrane segments without loss of continuity (Fig. 11-24). Most of these processes begin with a local increase in membrane curvature. FIGURE 11-24 Membrane fusion. The fusion of two membranes is central to a variety of cellular processes involving organelles and the plasma membrane. Cardiolipin, present primarily in mitochondrial membranes of eukaryotes, but also a major component of the membrane lipids of bacteria, can create or recognize membrane curvature. It is cone-shaped — its head group is small relative to its four fatty acyl chains — so it can act as a wedge in the monolayer, contracting that monolayer relative to the other. Membrane curvature is the result. In E. coli, cardiolipin is highly localized at the two poles of the rod-shaped cell, where its curvature is sharpest. It seems very likely that other membrane lipids will be found to influence local curvature of the bilayer. A protein that is intrinsically curved may force a bilayer to curve by binding to it (Fig. 11-25); the binding energy provides the driving force for the increase in bilayer curvature. Alternatively, multiple subunits of a scaffold protein may assemble into curved supramolecular complexes and stabilize curves that spontaneously form in the bilayer. For example, a superfamily of proteins containing BAR domains (named for the first three members of the family that were identified: BIN1, amphiphysin, and RVS167) can assemble into a crescent-shaped scaffold that binds to the membrane surface, forcing or favoring membrane curvature. BAR domains consist of coiled coils that form long, thin, curved dimers with a positively charged concave surface that tends to form ionic interactions with the negatively charged head groups of membrane lipids PIP2 and PIP3. The enzymatic formation of these inositol lipids can tag a plasma membrane area for creation of inward curvature by a BAR protein (Fig. 11- 25). Some of these BAR proteins also have an amphipathic helix (having one polar face and one hydrophobic face; see Fig. 11-32) that inserts like a wedge into one leaflet of the bilayer, expanding its area relative to the other leaflet and thereby forcing curvature. Such a protein might also serve as a detector of preexisting membrane curvature due to differences in lipid composition of two leaflets of a bilayer.
FIGURE 11-25 Three models for protein-induced curvature of membranes. [Information from (a, b) B. Qualmann et al., EMBO J. 30:3501, 2011, Fig. 1; (c) B. J. Peter et al., Science 303:495, 2004, Fig. 1A.] Septins make up a family of GTP-binding proteins (14 genes in humans) that polymerize at curved regions of the plasma membrane and participate in cellular processes such as cell division, exocytosis, phagocytosis, and apoptosis. All septins have an amphipathic helix that can submerge its hydrophobic side in one leaflet of the bilayer, forcing one leaflet of the bilayer to expand laterally, either causing or sensing local curvature in the membranes. Studies of cells with mutations in this helix show its biological importance in vesicle trafficking and neurotransmitter release. Specific fusion of two membranes requires mediation by fusion proteins. Fusion proteins ensure (1) that the two membranes recognize each other; (2) that their surfaces become closely apposed, which requires removal of the water molecules normally associated with the polar head groups of lipids; (3) that their bilayer structures become locally disrupted, resulting in fusion of the outer leaflets of the two membranes (hemifusion); and (4) that their bilayers fuse to form a single continuous bilayer. The fusion occurring in receptor-mediated endocytosis, or regulated secretion, also requires (5) that the process is triggered at the appropriate time or in response to a specific signal. Fusion proteins mediate these events, bringing about specific recognition and a transient local distortion of the bilayer structure that favors membrane fusion. (Note that these fusion proteins are unrelated to the products encoded by two fused genes, also called fusion proteins, discussed in Chapter 9.) A well-studied example of membrane fusion occurs at synapses, when intracellular (neuronal) vesicles loaded with neurotransmitter fuse with the plasma membrane. Yeast cells provide another experimentally accessible system in which vesicles fuse with the plasma membrane, releasing their secretion products. Both processes involve a family of proteins called SNAREs (snap receptors; Fig. 11-26). SNAREs in the cytoplasmic face of the intracellular vesicle are called v-SNAREs (v for vesicle); those in the target membrane with which the vesicle fuses (the plasma membrane during exocytosis) are t- SNAREs (t for target). The protein NSF (N-ethylmaleimide sensitive factor) regulates the interactions among SNAREs. During fusion, a v-SNARE and a t-SNARE bind to each other and undergo a structural change that produces a bundle of long, thin rods made up of helices from both SNAREs and two helices from the protein SNAP25 (Fig. 11-26). The two SNAREs initially interact at their ends, then zip up into the bundle of helices. This structural change pulls the two membranes into contact and initiates the fusion of their lipid bilayers. An alternative designation of SNARE types is based on structural features of the proteins: R-SNAREs have an Arg residue critical to their function, and Q-SNAREs have a critical Gln residue. Typically, R-SNAREs act as v-SNAREs, and Q-SNAREs act as t-SNAREs.
FIGURE 11-26 Membrane fusion during neurotransmitter release at a synapse. The secretory vesicle membrane contains the v-SNARE synaptobrevin (red). The target (plasma) membrane contains the t-SNAREs syntaxin (blue) and SNAP25 (violet). When a local increase in [Ca2+] signals release of neurotransmitter, the v-SNARE, SNAP25, and t-SNARE interact, forming a coiled bundle of four α helices, pulling the two membranes together and disrupting the bilayer locally. This leads first to hemifusion, joining the outer leaflets of the two membranes, then to complete membrane fusion and neurotransmitter release. When fusion is complete, the SNARE complex is disassembled. [Information from Y. A. Chen and R. H. Scheller, Nat. Rev. Mol. Cell Biol. 2:98, 2001.] The complex of SNAREs and SNAP25 is the target of several powerful neurotoxins. Clostridium botulinum toxin is a bacterial protease that cleaves specific bonds in SNARE proteins, preventing neurotransmission and causing paralysis and death. Because of its very high specificity for these proteins, purified botulinum toxin has served as a powerful tool for dissecting the mechanism of neurotransmitter release in vivo and in vitro. Used in small amounts, botulinum toxin (Botox) is used in medicine to treat disorders of eye and neck muscles, as well as cosmetically for the removal of skin wrinkles. Tetanus toxin, produced by the bacterium Clostridium tetani, also is a protease with high specificity for SNARE proteins. It causes painful muscle spasms and rigidity of voluntary muscles — hence the characteristic symptom “lockjaw.” Integral Proteins of the Plasma Membrane Are Involved in Surface Adhesion, Signaling, and Other Cellular Processes Several families of integral proteins in the plasma membrane provide specific points of attachment between cells or between a cell and proteins of the extracellular matrix. Integrins are surface adhesion proteins that mediate a cell’s interaction with the extracellular matrix and with other cells, including some pathogens. Integrins also carry signals in both directions across the plasma membrane, integrating information about the extracellular and intracellular environments. All integrins are heterodimeric proteins composed of two unlike subunits, α and β , each anchored to the plasma membrane by a single transmembrane helix. The large extracellular domains of the α and β subunits combine to form a specific binding site for extracellular proteins such as collagen and fibronectin, which contain a common determinant of integrin binding, the sequence Arg–Gly–Asp (RGD). Other plasma membrane proteins involved in surface adhesion are the cadherins, which undergo homophilic (“with the same kind”) interactions with identical cadherins in an adjacent cell. Selectins have extracellular domains that, in the presence of Ca2+ , bind specific polysaccharides on the surface of an adjacent cell. Selectins are present primarily in the various types of blood cells and in the endothelial cells that line blood vessels (see Fig. 7-29). They are an essential part of the blood-clotting process. SUMMARY 11.2 Membrane Dynamics Lipids in a biological membrane can exist in liquid-ordered or liquid-disordered states. Membrane fluidity is affected by temperature, fatty acid composition, and sterol content. Flip-flop diffusion of lipids between the inner and outer leaflets of a membrane occurs only when the diffusion is specifically catalyzed by flippases, floppases, scramblases, or PI transporters. Proteins and lipids can diffuse laterally within the plane of the membrane, but this mobility is limited by interactions of membrane proteins with internal cytoskeletal structures and interactions of lipids with lipid ra s. One class of lipid ra s is enriched for sphingolipids and cholesterol with a subset of membrane proteins that are GPI- linked or attached to several long-chain fatty acyl moieties. Caveolins are integral proteins that associate with the inner leaflet of the plasma membrane, forcing it to curve inward to form caveolae, which are involved in membrane transport, signaling, and the expansion of plasma membranes. Proteins containing BAR domains cause local membrane curvature and mediate the fusion of two membranes, which accompanies processes such as endocytosis, exocytosis, and viral invasion. Septins are proteins that sense or cause membrane curvature. Membrane fusion depends on SNARE proteins, which draw two membranes close together and favor fusion. Integrins, cadherins, and selectins are transmembrane proteins of the plasma membrane that act both to attach cells to each other and to carry messages between the extracellular matrix and the cytoplasm. 11.3 Solute Transport across Membranes Every living cell must acquire from its surroundings the raw materials for biosynthesis and for energy production, and must release the byproducts of metabolism to its environment; both processes require that small compounds or inorganic ions cross the plasma membrane. Within the eukaryotic cell, different compartments have different concentrations of ions and of metabolic intermediates and products, and these, too, must move across intracellular membranes in tightly regulated processes. A few nonpolar compounds can dissolve in the lipid bilayer and cross a membrane unassisted, but for any polar compound or ion, a specific membrane protein carrier is essential. Approximately 2,000 genes in the human genome encode proteins that function in transporting solutes across membranes. In some cases, a membrane protein simply facilitates the diffusion of a solute down its concentration gradient; but transport can also occur against a gradient of concentration, electrical potential, or both, and in these cases, as we shall see, the transport process requires energy. Ions may also diffuse across membranes via ion channels formed by proteins, or they may be carried across by ionophores, small molecules that mask the charge of ions and allow them to diffuse through the lipid bilayer. Figure 11-27 summarizes the various types of transport mechanisms discussed in this section. FIGURE 11-27 Summary of transporter types. Some types (ionophores, ion channels, and passive transporters) simply speed transmembrane movement of solutes (S) down their electrochemical gradients, whereas others (active transporters) can pump solutes against a gradient, using ATP or a gradient of a second solute to provide the energy. Transport May Be Passive or Active When two aqueous compartments containing unequal concentrations of a soluble compound or ion are separated by a permeable divider (membrane), the solute moves by simple diffusion from the region of higher concentration, through the membrane, to the region of lower concentration, until the two compartments have equal solute concentrations (Fig. 11-28a). When ions of opposite charge are separated by a permeable membrane, there is a transmembrane electrical gradient, a membrane potential, Vm (expressed in millivolts). This membrane potential produces a force opposing ion movements that increase Vm and driving ion movements that reduce Vm (Fig. 11-28b). Thus, the direction in which a charged solute tends to move spontaneously across a membrane depends on both the chemical gradient (the difference in solute concentration) and the electrical gradient (Vm) across the membrane. Together these two factors are referred to as the electrochemical gradient or electrochemical potential. This behavior of solutes is in accord with the second law of thermodynamics: molecules tend to spontaneously assume the distribution of greatest randomness and lowest energy. FIGURE 11-28 Movement of solutes across a permeable membrane. (a) Net movement of an electrically neutral solute is toward the side of lower solute concentration until equilibrium is achieved. The solute concentrations on the le and right sides of the membrane, as shown here, are designated C1 and C2. The rate of transmembrane solute movement (indicated by the arrows) is proportional to the concentration ratio. (b) Net movement of an electrically charged solute is dictated by a combination of the electrical potential (Vm) and the ratio of chemical concentrations (C2/C1) across the membrane; net ion movement continues until this electrochemical potential reaches zero. Membrane proteins that act by increasing the rate of solute movement across membranes are called transporters or carriers. Transporters are of two general types: passive and active. Passive transporters simply facilitate movement down a concentration gradient, increasing the transport rate. This process is called passive transport or facilitated diffusion. Active transporters (sometimes called pumps) can move substrates across a membrane against a concentration gradient or an electrical potential, a process called active transport. Primary active transporters use energy provided directly by a chemical reaction; secondary active transporters couple uphill transport of one substrate with downhill transport of another. Transporters and Ion Channels Share Some Structural Properties but Have Different Mechanisms To pass through a lipid bilayer, a polar or charged solute must first give up its interactions with the water molecules in its hydration shell, then diffuse about 3 nm (30 Å) through a substance (lipid) in which it is poorly soluble (Fig. 11-29a). The energy used to strip away the hydration shell and to move the polar compound from water into lipid, then through the lipid bilayer, is regained as the compound leaves the membrane on the other side and is rehydrated. However, the intermediate stage of transmembrane passage is a high-energy state comparable to the transition state in an enzyme-catalyzed chemical reaction. In both cases, an activation barrier must be overcome to reach the intermediate stage (Fig. 11-29; compare with Fig. 6-3). The energy of activation (ΔG‡) for translocation of a polar solute across the bilayer is so large that pure lipid bilayers are virtually impermeable to polar and charged species on time scales relevant to cell growth and division. FIGURE 11-29 Energy changes accompanying passage of a hydrophilic solute through the lipid bilayer of a biological membrane. (a) In simple diffusion, removal of the hydration shell is highly endergonic, and the energy of activation (ΔG‡) for diffusion through the bilayer is very high. (b) A transporter protein reduces the ΔG‡ for transmembrane diffusion of the solute. It does this by forming noncovalent interactions with the dehydrated solute to replace the hydrogen bonding with water and by providing a hydrophilic transmembrane pathway. Membrane proteins lower the activation energy for transport of polar compounds and ions by providing an alternative path across the membrane for specific solutes. Lowering the activation energy greatly increases the rate of transmembrane movement (recall p. 182). Transporters are not enzymes in the usual sense; their “substrates” are moved from one compartment to another but are not chemically altered. Like enzymes, however, transporters bind their substrates with stereochemical specificity through multiple weak, noncovalent interactions. The negative free-energy change associated with these weak interactions, ΔGbinding, counterbalances the positive free-energy change that accompanies loss of the water of hydration from the substrate, ΔGdehydration, thereby lowering ΔG‡ for transmembrane passage (Fig. 11-29b). Transporter proteins span the lipid bilayer several times, forming a transmembrane pathway lined with hydrophilic amino acid side chains. The pathway provides an alternative route for a specific substrate to move across the lipid bilayer without its having to dissolve in the bilayer, further lowering ΔG‡ for transmembrane diffusion. The result is an orders-of-magnitude increase in the substrate’s rate of passage across the membrane. Ion channels use a different mechanism than transporters to move inorganic ions across membranes. Ion channels speed the passage of ions across membranes by providing an aqueous path across the membrane through which inorganic ions can diffuse at very high rates. Most ion channels have a “gate” (Fig. 11-30a) regulated by a biological signal. When the gate is open, ions move across the membrane, through the channel, in the direction dictated by the ion’s charge and the electrochemical gradient. Movement occurs at rates approaching the limit of unhindered diffusion (tens of millions of ions per second per channel — much higher than typical transporter rates). Ion channels typically show some specificity for an ion, but they are not saturable with their ion substrate. Flow through a channel stops either when the gating mechanism is closed (again, by a biological signal) or when there is no longer an electrochemical gradient providing the driving force for the movement. In contrast, transporters, which bind their “substrates” with high stereospecificity, catalyze transport at rates well below the limits of free diffusion, and they are saturable in the same sense as are enzymes: there is some substrate concentration above which further increases will not produce a greater rate of transport. Transporters have a gate on either side of the membrane, and the two gates are never open at the same time (Fig. 11-30b). FIGURE 11-30 Differences between channels and transporters. (a) Ion channels have a transmembrane pore that is either open or closed, depending on the position of the single gate. When the gate is open, ions move through at a rate limited only by the maximum rate of diffusion. (b) Transporters have two gates, and both are never open at the same time. Movement of a substrate (an ion or a small molecule) through the membrane is therefore limited by the time needed for one gate to open and close (on one side of the membrane) and the second gate to open. Rates of movement through ion channels can be orders of magnitude greater than rates through transporters, but channels simply allow the ion to flow down the electrochemical gradient, whereas active transporters (pumps) can move a substrate against its concentration gradient. [Information from D. C. Gadsby, Nat. Rev. Mol. Cell Biol. 10:344, 2009, Fig. 1.] Both transporters and ion channels constitute large families of proteins, defined not only by their primary sequences but also by their secondary structures. We next consider some well-studied representatives of the main transporter and channel families. You will also encounter some of these in Chapter 12 when we discuss transmembrane signaling, and some in later chapters in the context of the metabolic pathways in which they participate. The Glucose Transporter of Erythrocytes Mediates Passive Transport Energy-yielding metabolism in erythrocytes depends on a constant supply of glucose from the blood plasma, where the glucose concentration is maintained at about 4.5 to 5 mM. Glucose enters the erythrocyte by passive transport via a specific glucose transporter called GLUT1, at a rate about 50,000 times greater than it could cross the membrane unassisted. The process of glucose transport can be described by analogy with an enzymatic reaction in which the “substrate” is glucose outside the cell (Sout), the “product” is glucose inside the cell (Sin), and the “enzyme” is the transporter, T. When the initial rate of glucose uptake is measured as a function of external glucose concentration (Fig. 11-31), the resulting plot is hyperbolic: at high external glucose concentrations, the rate of uptake approaches Vmax. Formally, such a transport process can be described by the set of equations in which k1, k−1, and so forth are the forward and reverse rate constants for each step; T 1 is the transporter conformation in which the glucose-binding site faces outward (in contact with blood plasma), and T 2 is the conformation in which it faces inward. Given that every step in this sequence is reversible, the transporter is, in principle, equally able to move glucose into or out of the cell. As for enzyme assays, this transporter assay measures the initial rate of uptake, when the product concentration (glucose concentration inside the cell) is zero, while the substrate concentration (glucose on the outside) is varied. In the living cell, GLUT1 accelerates the movement of glucose down its concentration gradient, which normally means into the cell. Glucose that enters a cell is generally metabolized immediately, and the intracellular glucose concentration is thereby kept low relative to its concentration in the blood. FIGURE 11-31 Kinetics of glucose transport into erythrocytes. (a) The initial rate of glucose entry into an erythrocyte, V0, depends on the initial concentration of glucose on the outside, [S]out. (b) Double-reciprocal plot of the data in (a). The kinetics of passive transport is analogous to the kinetics of an enzyme-catalyzed reaction. (Compare these plots with Fig. 6-12 and 6- 14.) Kt is analogous to Km, the Michaelis constant. The rate equations for glucose transport can be derived exactly as for enzyme-catalyzed reactions (Chapter 6), yielding an expression analogous to the Michaelis-Menten equation V0= (11-1) in which V0 is the initial velocity of accumulation of glucose inside the cell when its concentration in the surrounding medium is [S]out, and Kt (Ktransport) is a constant analogous to the Michaelis constant, a combination of rate constants that is characteristic of each transport system. This equation describes the initial velocity, the rate observed when [S]in = 0. As is the case for enzyme-catalyzed reactions, the slope-intercept form of the equation describes a linear plot of 1/V0 against 1/[S]out, from which we can obtain values of Kt and Vmax (Fig. 11-31b). When [S]out = Kt, the rate of uptake is ½Vmax; the transport process is half-saturated. The concentration of glucose in blood, as noted above, is 4.5 to 5 mM, which is close enough to the Kt to ensure that GLUT1 is half-saturated with substrate and operates near one-half Vmax. Vmax[S]out Kt+ [S]out Because no chemical bonds are made or broken in the conversion of Sout to Sin, neither “substrate” nor “product” is intrinsically more stable, and the process of entry is therefore fully reversible. As [S]in approaches [S]out, the rates of entry and exit become equal. Such a system is therefore incapable of accumulating glucose within a cell at concentrations above that in the surrounding medium; it simply equilibrates glucose on the two sides of the membrane much faster than would occur in the absence of a specific transporter. GLUT1 is specific for D-glucose, with a Kt of about 6 mM. For the close analogs D-mannose and D- galactose, which differ only in the position of one hydroxyl group, the values of Kt are 20 mM and 30 mM, respectively, and for L- glucose, Kt exceeds 3,000 mM. Thus, GLUT1 shows the three hallmarks of passive transport: high rates of diffusion down a concentration gradient, saturability, and stereospecificity. GLUT1 is an integral protein with 12 hydrophobic segments, each forming a membrane-spanning helix (Fig. 11-32a). The helices that line the transmembrane path for glucose are amphipathic; for each helix, the residues along one side are predominantly nonpolar, and those on the other side are mainly polar. This amphipathic structure is evident in a helical wheel diagram (Fig. 11-32b). A cluster of amphipathic helices are arranged so that their polar sides face each other and line a hydrophilic pore through which glucose can pass (Fig. 11-32c), while their hydrophobic sides interact with the surrounding membrane lipids such that the hydrophobic effect stabilizes the entire transporter structure. FIGURE 11-32 Membrane topology of the glucose transporter GLUT1. (a) Transmembrane helices are represented here as oblique (angled) rows of three or four amino acid residues, each row depicting one turn of the α helix. Of the 12 helices, 9 contain three or more polar or charged residues (blue or red), o en separated by several hydrophobic residues (yellow). (b) A helical wheel diagram shows the distribution of polar and nonpolar residues on the surface of a helical segment. The helix is diagrammed as though observed along its axis from the amino terminus. Adjacent residues in the linear sequence are connected, and each residue is placed around the wheel in the position it occupies in the helix; recall that 3.6 residues are required to make one complete turn of the α helix. In this example, the polar residues (blue) are on one side of the helix, and the hydrophobic residues (yellow) are on the other. This is, by definition, an amphipathic helix. (c) Side-by-side association of amphipathic helices, each with its polar face oriented toward the central cavity, produces a transmembrane channel lined with polar (and charged) residues, available for interaction with glucose. [Information from (a, c) M. Mueckler, Eur. J. Biochem. 219:713, 1994.] Structural studies of mammalian GLUT1 and other GLUT transporters suggest that the protein cycles through a series of conformational changes, interconverting a form (T 1) with its glucose-binding site accessible only from the extracellular side, through a form in which the bound glucose is sequestered and inaccessible from either side, to a form (T 2) with the glucose- binding site open only to the intracellular side (Fig. 11-33). FIGURE 11-33 Model of glucose transport into erythrocytes by GLUT1. (a) The transporter exists in two extreme conformations: T 1, with the glucose- binding site exposed on the outer surface of the plasma membrane, and T 2, with the binding site exposed on the inner surface. Glucose transport occurs in four steps. Glucose in blood plasma binds to a stereospecific site on T 1; this lowers the activation energy for a conformational change from glucoseout • T 1 to glucosein • T 2, effecting transmembrane passage of the glucose. Glucose is released from T 2 into the cytoplasm, and the transporter returns to the T 1 conformation, ready to transport another glucose molecule. Between the forms T 1 and T 2, there is an intermediate form (not shown here) in which glucose is sequestered within the transporter, with access to neither side. The structures of (b) human GLUT3 in the T 1 conformation and (c) human GLUT1 in the T 2 conformation, determined by x-ray crystallography, support the model shown in (a). [Data from (b) PDB ID 4ZWC, D. Deng et al., Nature 526:391, 2015; (c) PDB ID 4PYP, D. Deng et al., Nature 510:121, 2014.] Twelve passive glucose transporters are encoded in the human genome, each with its unique kinetic properties, patterns of tissue distribution, and function (Table 11-1). GLUT1, in addition to supplying glucose to erythrocytes, also transports glucose across the blood-brain barrier, supplying the glucose that is essential for normal brain metabolism. The very rare individuals with defects in GLUT1 have a variety of brain-related symptoms, including seizures, movement and language disorders, and developmental delays. Standard care for such individuals includes a ketogenic diet, which provides the ketones that can serve as an alternative energy source for the brain. In the liver, GLUT2 transports glucose out of hepatocytes when liver glycogen is broken down to replenish blood glucose. GLUT2 has a large Kt(≥ 17 mM ) and can therefore respond to increased levels of intracellular glucose (produced by glycogen breakdown) by increasing outward transport. Skeletal and heart muscle and adipose tissue have yet another glucose transporter, GLUT4 (Kt= 5 mM ), which is distinguished by its response to insulin: its activity increases when insulin signals a high blood glucose concentration, thus increasing the rate of glucose uptake into muscle and adipose tissue. Box 11-1 describes the effect of insulin on this transporter. TABLE 11-1 Glucose Transporters in Humans Transporter Tissue(s) where expressed Kt (mM) Role/characteristics GLUT1 Erythrocytes, blood- brain barrier, placenta, 3 Basal glucose uptake; defective in De Vivo disease a most tissues at a low level GLUT2 Liver, pancreatic islets, intestine, kidney 17 In liver and kidney, removal of excess glucose from blood; in pancreas, regulation of insulin release GLUT3 Brain (neuron), testis (sperm) 1.4 Basal glucose uptake; high turnover number GLUT4 Muscle, fat, heart 5 Activity increased by insulin GLUT5 Intestine (primarily), testis, kidney 6 Primarily fructose transport GLUT6 Spleen, leukocytes, brain > 5 Possibly no transporter function GLUT7 Small intestine, colon, testis, prostate 0.3 — GLUT8 Testis, sperm acrosome ~2 — GLUT9 Liver, kidney, intestine, lung, placenta 0.6 Urate and glucose transporter in liver, kidney GLUT10 Heart, lung, brain, liver, muscle, pancreas, placenta, kidney 0.3 Glucose and galactose transporter GLUT11 Heart, skeletal muscle 0.16 Glucose and fructose transporter GLUT12 Skeletal muscle, heart, prostate, placenta — — Information on localization from M. Mueckler and B. Thorens, Mol. Aspects Med. 34:121, 2013. Kt values for glucose from R. Augustin, IUBMB Life 62:315, 2010. b c Dash indicates role uncertain. Km for fructose. Km for 2-deoxyglucose. BOX 11-1 MEDICINE Defective Glucose Transport in Diabetes When ingestion of a carbohydrate-rich meal causes blood glucose to exceed the usual concentration between meals (about 5 m ), excess glucose is taken up by the myocytes of cardiac and skeletal muscle (which store it as glycogen) and by adipocytes (which convert it to triacylglycerols). Glucose uptake into myocytes and adipocytes is mediated by the glucose transporter GLUT4. Between meals, some GLUT4 is present in the plasma membrane, but most (90%) is sequestered in the membranes of small intracellular vesicles (Fig. 1). Insulin is released from the pancreas in response to high blood glucose, and it triggers, within minutes, the movement of these vesicles to the plasma membrane. The vesicles fuse with the membrane, bringing most of the GLUT4 molecules to the cell surface (see Fig. 12-23) and increasing the rate of glucose uptake 15-fold or more. When blood glucose levels return to normal, insulin release slows and most GLUT4 molecules are removed from the plasma membrane by endocytosis and stored in vesicles. a b c FIGURE 1 Transport of glucose into a myocyte by GLUT4 is regulated by insulin. [Information from F. E. Lienhard et al., Sci. Am. 266 (January):86, 1992.] In type 1 (insulin-dependent) diabetes mellitus, the inability to release insulin (and thus to mobilize glucose transporters) results in low rates of glucose uptake into muscle and adipose tissue. One consequence is a prolonged period of high blood glucose a er a carbohydrate-rich meal. This condition is the basis for the glucose tolerance test used to diagnose diabetes (Chapter 23). The Chloride-Bicarbonate Exchanger Catalyzes Electroneutral Cotransport of Anions across the Plasma Membrane The erythrocyte contains another passive transport system, an anion exchanger that is essential in CO2 transport to the lungs from tissues such as skeletal muscle and liver. Waste CO2 released from respiring tissues into the blood plasma enters the erythrocyte, where it is converted to bicarbonate HCO− 3 by the enzyme carbonic anhydrase. (Recall that HCO− 3 is the primary buffer of blood pH; see Fig. 2-20.) The HCO− 3 reenters the blood plasma for transport to the lungs (Fig. 11-34). Because HCO− 3 is much more soluble in blood plasma than is CO2, this roundabout route increases the capacity of the blood to carry carbon dioxide from the tissues to the lungs. In the lungs, HCO− 3 reenters the erythrocyte and is converted to CO2, which is eventually released into the lung space and exhaled. To be effective, this shuttle requires very rapid movement of HCO− 3 across the erythrocyte membrane. As described in Chapter 5 (pp. 160–161), there is a second mechanism for moving CO2 from tissue to lung, involving reversible binding of CO2 to hemoglobin.) FIGURE 11-34 Chloride-bicarbonate exchanger of the erythrocyte membrane. This cotransport system allows the entry and exit of HCO−3 without changing the membrane potential. Its role is to increase the CO2- carrying capacity of the blood. The top half of the figure illustrates the events that take place in respiring tissues; the bottom half illustrates the events in the lungs. The chloride-bicarbonate exchanger, also called the anion exchange (AE) protein, increases the rate of HCO− 3 transport across the erythrocyte membrane more than a millionfold. Like the glucose transporter, it is a dimeric integral protein that spans the membrane 14 times. This protein mediates the simultaneous movement of two anions: for every HCO− 3 ion that moves in one direction, one Cl− ion moves in the opposite direction, with no net transfer of charge: the exchange is electroneutral. The coupling of Cl− and HCO− 3 movements is obligatory; in the absence of chloride, bicarbonate transport stops. In this respect, the anion exchanger is typical of those systems, called cotransport systems, that simultaneously carry two solutes across a membrane (Fig. 11-35). When, as in this case, the two substrates move in opposite directions, the process is antiport. In symport, two substrates are moved simultaneously in the same direction. Transporters that carry only one substrate, such as the erythrocyte glucose transporter, are known as uniport systems. FIGURE 11-35 Three general classes of transport systems. Transporters differ in the number of solutes (substrates) transported and the direction in which each solute moves. Examples of all three types of transporter are discussed in the text. Note that this classification tells us nothing about whether these are energy-requiring (active transport) or energy- independent (passive transport) processes. Active Transport Results in Solute Movement against a Concentration or Electrochemical Gradient In passive transport, the transported species always moves down its electrochemical gradient and is not accumulated above the equilibrium concentration. Active transport, by contrast, results in the accumulation of a solute above the equilibrium point. Active transport is essential when cells function in an environment in which key substrates are present outside the cell only at very low concentrations. For example, the bacterium E. coli can grow in a medium containing only 1 μ M Pi (inorganic phosphate), but the cell must maintain internal Pi levels in the millimolar range. Worked Example 11-2 describes another such situation, which requires cells to pump Ca2+ outward across the plasma membrane. Active transport is thermodynamically unfavorable (endergonic) and takes place only when coupled, directly or indirectly, to an exergonic process such as the absorption of sunlight, an oxidation reaction, the breakdown of ATP, or the concomitant flow of some other chemical species down its electrochemical gradient. In primary active transport, solute accumulation is coupled directly to an exergonic chemical reaction, such as conversion of ATP to AD P + Pi (Fig. 11-36). Secondary active transport occurs when endergonic (uphill) transport of one solute is coupled to the exergonic (downhill) flow of a different solute that was originally pumped uphill by primary active transport. FIGURE 11-36 Two types of active transport. (a) In primary active transport, the energy released by ATP hydrolysis drives solute (S1) movement against an electrochemical gradient. (b) In secondary active transport, a gradient of an ion (designated S1; o en Na+) has been established by primary active transport. Movement of S1 down its electrochemical gradient now provides the energy to drive cotransport of a second solute, S2, against its electrochemical gradient. The amount of energy needed for the transport of a solute against a gradient can be calculated from the initial concentration gradient. The general equation for the free-energy change in the chemical process that converts substrate (S) to product (P) is ΔG = ΔG′°+RT ln ([P]/[S]) (11-2) where Δ G′° is the standard free-energy change, R is the gas constant (8.315 J/mol • K), and T is the absolute temperature. When the “reaction” is simply transport of a solute from a region where its concentration is C1 to a region where its concentration is C2, no bonds are made or broken and Δ G′° is zero. The free- energy change for transport, ΔGt, is then ΔGt= RT ln (C2/C1) (11-3) If there is, say, a 10-fold difference in concentration between two compartments, the cost of moving 1 mol of an uncharged solute at 25 °C uphill across a membrane separating the compartments is ΔGt = (8.315 J /mol∙K)(298 K) ln (10/1)= 5,700 J /mol = 5.7 kJ /mol Equation 11-3 holds for all uncharged solutes. WORKED EXAMPLE 11-1 Energy Cost of Pumping an Uncharged Solute Calculate the energy cost (free-energy change) of pumping an uncharged solute against a 104-fold concentration gradient at 25 °C. SOLUTION: Begin with Equation 11-3. Substitute 1.0× 104 for (C2/C1), 8.315 J/mol • K for R, and 298 K for T: ΔGt = RT ln (C2/C1) = (8.315 J /mol∙K)(298 K)(1.0× 104) = 23 kJ /mol When the solute is an ion, its movement without an accompanying counterion results in the endergonic separation of positive and negative charges, producing an electrical potential; such a transport process is said to be electrogenic. The energetic cost of moving an ion depends on the electrochemical potential (Fig. 11-25), the sum of the chemical and electrical gradients: ΔGt= RT ln (C2/C1)+ ZF Δψ (11-4) where Z is the charge on the ion, F is the Faraday constant (96,480 J/V • mol), and Δψ is the transmembrane electrical potential (in volts). Eukaryotic cells typically have plasma membrane potentials of about 0.05 V (with the inside negative relative to the outside), so the second term on the right side of Equation 11-4 can make a significant contribution to the total free-energy change for transporting an ion. Most cells maintain more than a 10-fold difference in ion concentrations across their plasma or intracellular membranes, and for many cells and tissues active transport is therefore a major energy-consuming process. WORKED EXAMPLE 11-2 Energy Cost of Pumping a Charged Solute Calculate the energy cost (free-energy change) of pumping Ca2+ from the cytosol, where its concentration is about 1.0× 10−7 M, to the extracellular fluid, where its concentration is about 1.0 mM. Assume a temperature of 37 °C (body temperature in a mammal) and a standard transmembrane potential of 50 mV (inside negative) for the plasma membrane. SOLUTION: This is a case in which energy must be expended to counter two forces acting on the ion being transported: the membrane potential and the concentration difference across the membrane. These forces are expressed in the two terms on the right side of Equation 11-4: ΔGt= RT ln (C2/C1)+ ZF Δψ in which the first term describes the chemical gradient and the second describes the electrical potential. In Equation 11-4, substitute 8.315 J/mol • K for R, 310 K for T, 1.0× 10−3 for C2, 1.0× 10−7 for C1, + 2 (the charge on a Ca2+ ion) for Z, 96,500 J/V • mol for F, and 0.050 V for Δψ . Note that the transmembrane potential is 50 mV (inside negative), so the change in potential when an ion moves from inside to outside is 50 mV. ΔGt = RT ln (C2/C1)+ ZF Δψ = (8.315 J /mol∙K) (310 K) ln + 2(96,500 J /V ∙mol) (0.050 V) = 33 kJ /mol The mechanism of active transport is of fundamental importance in biology. As we shall see in Chapters 19 and 20, ATP is formed in mitochondria and chloroplasts by a mechanism that is essentially ATP-driven ion transport operating in reverse. The energy made available by the spontaneous flow of protons across a membrane is calculable from Equation 11-4; remember that ΔG for flow down an electrochemical gradient has a negative value, and ΔG for transport of ions against an electrochemical gradient has a positive value. P-Type ATPases Undergo Phosphorylation during Their (1.0× 10−3) (1.0× 10−7) Catalytic Cycles The family of active transporters called P-type ATPases are cation transporters that are reversibly phosphorylated by ATP (thus the name P-type) as part of the transport cycle. Phosphorylation forces a conformational change that is central to movement of the cation across the membrane. The human genome encodes at least 70 P-type ATPases that share similarities in amino acid sequence and topology, especially near the Asp residue that undergoes phosphorylation. All are integral proteins with 8 or 10 predicted membrane-spanning regions in a single polypeptide, and all are sensitive to inhibition by the transition-state analog vanadate, which mimics phosphate when under nucleophilic attack by a water molecule. The P-type ATPases are widespread in eukaryotes and bacteria. The Na+K+ ATPase of animal cells (an antiporter for Na+ and K+ ions) and the plasma membrane H+ ATPase of plants and fungi set the transmembrane electrochemical potential in cells by establishing ion gradients across the plasma membrane. These gradients provide the driving force for secondary active transport and are also the basis for electrical signaling in neurons. In animal tissues, the sarcoplasmic/endoplasmic reticulum Ca2+ ATPase (SERCA) pump and the plasma membrane Ca2+ ATPase pump, which are uniporters for Ca2+ ions, together maintain the cytosolic level of Ca2+ below 1 µM. The SERCA pump moves Ca2+ from the cytosol into the lumen of the sarcoplasmic reticulum. Parietal cells in the lining of the mammalian stomach have a P- type ATPase that pumps H+ and K+ out of the cells and into the stomach, thereby acidifying the stomach contents. Lipid flippases, as we noted earlier, are structurally and functionally related to P-type transporters. Bacteria and eukaryotes use P-type ATPases to pump toxic heavy metal ions such as Cd2+ and Cu2+ out of cells. All P-type pumps have similar structures (Fig. 11-37) and similar mechanisms. The mechanism postulated for P-type ATPases takes into account the large conformational changes and the phosphorylation-dephosphorylation of the critical Asp residue in the P (phosphorylation) domain that is known to occur during a catalytic cycle. For the SERCA pump (Fig. 11-38), each catalytic cycle moves two Ca2+ ions across the membrane and converts an ATP to ADP and Pi. The role of ATP binding and phosphoryl transfer to the enzyme is to bring about the interconversion of two conformations, E1 and E2, of the transporter. In the E1 conformation, the two Ca2+-binding sites are exposed on the cytosolic side of the ER or sarcoplasmic reticulum and bind Ca2+ with high affinity. ATP binding and Asp phosphorylation drive a conformational change from E1 to E2 that exposes the Ca2+- binding sites to the lumen and greatly reduces their affinity for Ca2+, which releases the Ca2+ ions into the lumen. By this mechanism, the energy released by hydrolysis of ATP during one phosphorylation-dephosphorylation cycle drives Ca2+ across the membrane against a large electrochemical gradient. FIGURE 11-37 The general structure of the P-type ATPases. (a) P-type ATPases have three cytoplasmic domains (A, N, and P) and two transmembrane domains (T and S) consisting of multiple helices. The N (nucleotide-binding) domain binds ATP and M g2+ and has protein kinase activity that phosphorylates a specific Asp residue in the P (phosphorylation) domain of all P-type ATPases. The A (actuator) domain has protein phosphatase activity and removes the phosphoryl group from the Asp residue with each catalytic cycle of the pump. A transport (T) domain with six transmembrane helices includes the ion-transporting structure, and four more transmembrane helices make up the support (S) domain, which provides physical support to the transport domain and may have other specialized functions in certain P-type ATPases. The binding sites for the ions to be transported are near the middle of the membrane, 40 to 50 Å from the phosphorylated Asp residue — thus, Asp phosphorylation-dephosphorylation does not directly affect ion binding. The A domain communicates movements of the N and P domains to the ion-binding sites. (b) A ribbon representation of the Ca2+ ATPase (SERCA pump). ATP binds to the N domain, and the Ca2+ ions to be transported bind to the T domain. (c) Other P-type ATPases have domain structures, and presumably mechanisms, like those of the SERCA pump; shown here are Na+K+ ATPase, the plasma membrane H+ ATPase, and the gastric H+K+ ATPase. [(a) Information from M. Bublitz et al., Curr. Opin. Struct. Biol. 20:431, 2010, Fig. 1. Data from (b) PDB ID 1SU4, C. Toyoshima et al., Nature 405:647, 2000; (c) Na+K+ ATPase, PDB ID 3KDP, J. Preben Morth et al., Nature 450:1043, 2007; H+ ATPase, PDB ID 3B8C, B. P. Pedersen et al., Nature 450:1111, 2007; H+K+ ATPase, PDB ID 3IXZ, K. Abe et al., EMBO J. 28:1637, 2009, and PDB ID 3B8E, J. Preben Morth et al., Nature 450:1043, 2007.] FIGURE 11-38 Postulated mechanism of the SERCA pump. The transport cycle begins with the protein in the E1 conformation, with the Ca2+-binding sites facing the cytosol. Two Ca2+ ions bind, then ATP binds to the transporter and phosphorylates Asp351, forming E1-P. Phosphorylation favors the second conformation, E2-P, in which the Ca2+- binding sites, now with a reduced affinity for Ca2+, are accessible on the other side of the membrane (the lumen or extracellular space), and the released Ca2+ diffuses away. ADP is released, and E2-P is dephosphorylated. The A domain resets and the protein returns to the E1 conformation for another round of transport. [Information from W. Kühlbrandt, Nat. Rev. Mol. Cell Biol. 5:282, 2004.] A variation on this basic mechanism is seen in the Na+K+ ATPase of the plasma membrane. This cotransporter couples phosphorylation-dephosphorylation of the critical Asp residue to the simultaneous movement of both Na+ and K+ against their electrochemical gradients. The Na+K+ ATPase is responsible for maintaining low Na+ and high K+ concentrations in the cell relative to the extracellular fluid (Fig. 11-39). For each molecule of ATP converted to ADP and Pi, the transporter moves two K+ ions inward and three Na+ ions outward across the plasma membrane. Cotransport is therefore electrogenic, creating a net separation of charge across the membrane; in animals, this produces the membrane potential of –50 to –70 mV (inside negative relative to outside) that is characteristic of most cells and is essential to the conduction of action potentials in neurons. The central role of the Na+K+ ATPase is reflected in the energy invested in this single reaction: about 25% of the total energy consumption of a human at rest. FIGURE 11-39 Role of the Na+K+ ATPase in animal cells. This active transport system is primarily responsible for setting and maintaining the intracellular concentrations of Na+ and K+ in animal cells and for generating the membrane potential. It does this by moving three Na+ ions out of the cell for every two K+ ions it moves in. The electrical potential across the plasma membrane is central to electrical signaling in neurons, and the gradient of Na+ is used to drive the uphill cotransport of solutes in many cell types. V-Type and F-Type ATPases Are ATP- Driven Proton Pumps V-type ATPases, a class of proton-transporting ATPases, are responsible for acidifying intracellular compartments in many organisms (thus, V for vacuolar). Proton pumps of this type maintain the vacuoles of fungi and higher plants at a pH between 3 and 6, well below that of the surrounding cytosol (pH 7.5). V- type ATPases are also responsible for the acidification of lysosomes, endosomes, the Golgi complex, and secretory vesicles in animal cells. All V-type ATPases have a similar complex structure, with an integral (transmembrane) domain (Vo) that serves as a proton channel and a peripheral domain (V1) that contains the ATP-binding site and the ATPase activity (Fig. 11- 40a). The structure is similar to that of the well-characterized F- type ATPases.
FIGURE 11-40 Two proton pumps with similar structures. (a) The VoV1 H+ ATPase uses ATP to pump protons into vacuoles and lysosomes, creating their low internal pH. It has an integral (membrane-embedded) domain, Vo, that includes multiple identical c subunits, and a peripheral domain that projects into the cytosol and contains the ATP- hydrolyzing sites, on three identical B subunits (purple). (b) The FoF1 ATPase/ATP synthase of mitochondria has an integral domain, Fo, with multiple copies of the c subunit, and a peripheral domain, F1, consisting of three α subunits, three β subunits, and a central sha joined to the integral domain. Fo and Vo provide transmembrane channels through which protons are pumped as ATP is hydrolyzed on the β subunits of F1 (B subunits of V1). An ATP-driven proton transporter also can catalyze ATP synthesis (red arrows) as protons flow down their electrochemical gradient. This is the central reaction in the processes of oxidative phosphorylation and photophosphorylation. F-type ATPase transporters catalyze the uphill transmembrane passage of protons, driven by ATP hydrolysis. The “F-type” designation derives from the identification of these ATPases as energy-coupling factors. The Fo integral membrane protein complex (Fig. 11-40b; subscript “o” denotes its inhibition by the drug oligomycin) provides a transmembrane pathway for protons, and the peripheral protein F1 (subscript “1” indicating that this was the first of several factors isolated from mitochondria) uses the energy of ATP to drive protons uphill (into a region of higher H+ concentration). The FoF1 organization of proton-pumping transporters must have developed very early in evolution. Bacteria such as E. coli use an FoF1 ATPase complex in their plasma membrane to pump protons outward, and archaea have a closely homologous proton pump, the AoA1 ATPase. Like all enzymes, F-type ATPases catalyze their reactions in both directions. Therefore, a sufficiently large proton gradient can supply the energy to drive the reverse reaction, ATP synthesis (Fig. 11-40b). When functioning in this direction, the F-type ATPases are more appropriately named ATP synthases. ATP synthases are central to ATP production in mitochondria during oxidative phosphorylation and in chloroplasts during photophosphorylation, as well as in bacteria and archaea. The proton gradient needed to drive ATP synthesis is produced by other types of proton pumps powered by substrate oxidation or sunlight. We provide a detailed description of these processes in Chapters 19 and 20. ABC Transporters Use ATP to Drive the Active Transport of a Wide Variety of Substrates ABC transporters constitute a large family of ATP-driven transporters that pump amino acids, peptides, proteins, metal ions, various lipids, bile salts, and many hydrophobic compounds, including drugs, across a membrane against a concentration gradient. Many ABC transporters are located in the plasma membrane, but some are also found in the ER and in the membranes of mitochondria and lysosomes. All members of this family have two ATP-binding domains (“cassettes”) that give the family its name — ATP-binding cassette transporters — and two transmembrane domains, each containing six transmembrane helices. In some cases, all these domains are in a single, long polypeptide; other ABC transporters have two subunits, each contributing a nucleotide-binding domain (NBD) and a domain with six transmembrane helices. The transport mechanism is believed to involve two forms of the transporter, one with its substrate-binding site facing the outside of the cell, the other open to substrate on the inside (Fig. 11-41). Substrates move across the membrane when the two forms interconvert, driven by ATP hydrolysis. The NBDs of all ABC proteins are similar in sequence and presumably in three-dimensional structure. They constitute the conserved molecular motor that can be coupled to a wide variety of transmembrane domains, each capable of pumping one specific substrate across a membrane. When coupled this way, the ATP-driven motor moves solutes against a concentration gradient, with a stoichiometry of about one ATP hydrolyzed per molecule of substrate transported. FIGURE 11-41 ABC transporters. The protein has two homologous halves, each with a six-helix transmembrane domain (TMD; blue), and a cytoplasmic nucleotide-binding domain (NBD; red). In the mechanism proposed for the coupling of ATP hydrolysis to transport, with ATP bound to the NBD sites, substrate binds to the transporter on the cytoplasmic side. Upon substrate binding and ATP hydrolysis to ADP, a conformational change exposes the substrate to the outside surface and lowers the transporter’s affinity for its substrate; substrate diffuses away from the transporter into the extracellular space. This mechanism for the coupling of ATP hydrolysis to transport is based on structures of a number of ABC transporters crystallized under different conditions. Compare this process with the model of glucose transport in Figure 11-33. The human genome contains at least 48 genes that encode ABC transporters; a number of these are presented in Table 11-2. Some of these transporters have very high specificity for a single substrate; others are more promiscuous, able to transport drugs that cells presumably did not encounter during their evolution. Many ABC transporters are involved in maintaining the composition of the lipid bilayer, such as the floppases that move membrane lipids from one leaflet of the bilayer to the other. Many others are needed to move sterols, sterol derivatives, and fatty acids into the bloodstream for transport throughout the body. For example, the cellular machinery for exporting excess cholesterol includes an ABC transporter (see Fig. 21-47). Mutations in the genes that encode some of these proteins contribute to genetic diseases, including liver failure, retinal degeneration, and Tangier disease. The cystic fibrosis transmembrane conductance regulator protein (CFTR) of the plasma membrane is an interesting case of an ABC protein that is an ion channel (for Cl−), regulated by ATP hydrolysis, but without the pumping function characteristic of an active transporter (Box 11-2). TABLE 11-2 Some ABC Transporters in Humans Gene(s) Role/characteristics Text reference ABCA1 Reverse cholesterol transport; defect causes Tangier disease Fig. 21-47 ABCA4 Only in visual receptors, recycling of all-trans-retinal Fig. 12-19 ABCB1 Multidrug resistance P-glycoprotein 1; transport across blood-brain barrier — ABCB4 Multidrug resistance; transport of phosphatidylcholine in bile — ABCB11 Transports bile salts out of hepatocytes Fig. 17-1 ABCC6 Sulfonylurea receptor; targeted by the drug glipizide in type 2 diabetes Fig. 23-27 ABCG2 Breast cancer resistance protein (BCRP); major exporter of anticancer drugs p. 396 ABCC7 CFTR (Cl− channel); defect causes cystic fibrosis Box 11-2 BOX 11-2 MEDICINE A Defective Ion Channel in Cystic Fibrosis Cystic fibrosis (CF) is a serious hereditary disease. In the United States, the frequency of CF is about 1 in 3,200 live births, and 1% to 4% (depending on ethnicity) are carriers, having one defective copy of the gene and one normal copy. Only individuals with two defective copies show the severe symptoms of the disease: obstruction of the gastrointestinal and respiratory tracts, commonly leading to bacterial infection of the airways. The defective gene underlying CF was discovered in 1989. It encodes a membrane protein called cystic fibrosis transmembrane conductance regulator, or CFTR. This protein has two segments, each containing six transmembrane helices, two nucleotide-binding domains (NBDs), and a regulatory region that connects them (Fig. 1). CFTR is therefore very similar to other ABC transporter proteins, except that it functions as an ion channel (for Cl−), not as a pump. The channel conducts Cl− across the plasma membrane when both NBDs have bound ATP, and it closes when the ATP on one of the NBDs is broken down to ADP and Pi. The Cl− channel is further regulated by phosphorylation of several Ser residues in the regulatory domain, catalyzed by cAMP-dependent protein kinase. When the regulatory domain is not phosphorylated, the Cl− channel is closed. FIGURE 1 Three states of the CFTR protein. The protein has two segments, each with six transmembrane helices, and three functionally significant domains extend from the cytoplasmic surface: NBD 1 and NBD 2 (green) are nucleotide-binding domains that bind ATP, and the regulatory R domain (blue) is the site of phosphorylation by cAMP-dependent protein kinase. When this R domain is phosphorylated but no ATP is bound to the NBDs (le ), the channel is closed. The binding of ATP opens the channel (middle) until the bound ATP is hydrolyzed. When the R domain is unphosphorylated (right), it binds the NBD domains and prevents ATP binding and channel opening. CFTR is a typical ABC transporter in all but two respects: most ABC transporters lack the regulatory domain, and CFTR acts as an ion channel (for Cl−), not as a typical transporter. The mutation responsible for CF in up to 90% of cases results in deletion of a Phe residue at position 508 (a mutation denoted F508del). The mutant protein folds incorrectly, causing it to be degraded in proteasomes. As a result, Cl− movement is reduced across the plasma membranes of epithelial cells that line the airways, digestive tract, exocrine glands (pancreas, sweat glands), bile ducts, and vas deferens. Less-common mutations, such as G551D (Gly551 changed to Asp), lead to production of CFTR that is correctly folded and inserted into the membrane but is defective in Cl− transfer. Diminished export of Cl− in individuals with CF is accompanied by diminished export of water from cells, causing the mucus on cell surfaces to become dehydrated, thick, and excessively sticky. In normal circumstances, cilia on the epithelial cells lining the inner surface of the lungs constantly sweep away bacteria that settle in this mucus, but the thick mucus in individuals with CF hinders this process, providing a haven in the lungs for pathogenic bacteria. Frequent infections by bacteria such as Staphylococcus aureus and Pseudomonas aeruginosa cause progressive damage to the lungs and reduce respiratory efficiency, eventually resulting in death due to inadequate lung function. Advances in therapy have raised the average life expectancy for people who have CF from just 10 years in 1960 to more than 40 years today. CFTR potentiators such as ivaca or (VX-770) increase the function of the mutant G551D protein that is properly folded and in place in the plasma membrane. For individuals with the folding defect, F508del, CFTR correctors improve the processing and delivery of the mutant protein to the cell surface; a combination of potentiator and corrector drugs is more effective than the corrector drug alone for these patients (Fig. 2). In 2019, clinicals trials of a combination of three drugs (ivaca or, tezaca or, and elexaca or), acting as both potentiators and correctors, showed dramatic improvements in individuals with the most common mutation (F508del). FIGURE 2 (a) The CFTR mutation G551D (replacement of Gly551 with Asp) results in a protein that is inserted into the membrane correctly but is defective as a Cl− channel. Addition of the potentiator drug VX-770 (ivaca or) restores partial function to the Cl− channel. (b) The more common mutation F508del (deletion of Phe508) prevents proper folding of CFTR, causing it to be degraded in proteasomes. In the presence of a corrector drug, folding and membrane insertion can take place; addition of the potentiator drug results in partial restoration of Cl− channel activity. The channel is unstable and is degraded over time. [Information from J. P. Clancy, Sci. Transl. Med. 6:1, 2014.] One human ABC transporter with very broad substrate specificity is the multidrug transporter (MDR1), encoded by the ABCB1 gene. MDR1 in the placental membrane and in the blood-brain barrier ejects toxic compounds that would damage the fetus or the brain. But it is also responsible for the striking resistance of certain tumors to some generally effective antitumor drugs. For example, MDR1 pumps the chemotherapeutic drugs doxorubicin and vinblastine out of cells, thus preventing their accumulation within a tumor and blocking their therapeutic effects. Overexpression of MDR1 is o en associated with treatment failure in cancers of the liver, kidney, and colon. A related ABC transporter, BCRP (breast cancer resistance protein, encoded by the ABCG2 gene), is overexpressed in breast cancer cells, also conferring resistance to anticancer drugs. Highly selective inhibitors of these multidrug transporters are expected to enhance the effectiveness of antitumor drugs and are the objects of current drug discovery and design. ABC transporters are also present in simpler animals and in plants and microorganisms. Yeast has 31 genes that encode ABC transporters, Drosophila has 56, and E. coli has 80, representing 2% of its entire genome. ABC transporters that are used by E. coli and other bacteria to import essentials such as vitamin B12 are the presumed evolutionary precursors of the MDRs of animal cells. The presence of ABC transporters that confer antibiotic resistance in pathogenic microbes (Pseudomonas aeruginosa, Staphylococcus aureus, Candida albicans, Neisseria gonorrhoeae, and Plasmodium falciparum) is a serious public health concern and makes these transporters attractive targets for drug design. Ion Gradients Provide the Energy for Secondary Active Transport The ion gradients formed by primary transport of Na+ or H+ can, in turn, provide the driving force for cotransport of other solutes. Many cell types have transport systems that couple the spontaneous, downhill flow of these ions to the simultaneous uphill pumping of another ion, sugar, or amino acid. In intestinal epithelial cells, glucose and certain amino acids are accumulated by symport with Na+, down the Na+ gradient established by the Na+K+ ATPase of the plasma membrane (Fig. 11-42). The apical surface of the intestinal epithelial cell (the surface that faces the intestinal contents) is covered with microvilli — long, thin projections of the plasma membrane that greatly increase the surface area exposed to the intestinal contents. The Na+-glucose symporter in the apical plasma membrane takes up glucose from the intestine in a process driven by the downhill flow of Na+: 2Na+out+ glucoseout → 2Na+ in+ glucosein The energy required for this process comes from two sources: the greater concentration of Na+ outside than inside the cell (the chemical potential) and the membrane (electrical) potential, which is inside negative and therefore draws Na+ inward. The strong thermodynamic tendency for Na+ to move into the cell provides the energy needed for the transport of glucose into the cell, against its concentration gradient. An ion gradient created and sustained by energy-dependent ion pumping serves as the potential energy for cotransport of another species against its concentration gradient. FIGURE 11-42 Glucose transport in intestinal epithelial cells. Glucose is cotransported with Na+ across the apical plasma membrane into the epithelial cell. It moves through the cell to the basal surface, where it passes into the blood via GLUT2, a passive glucose uniporter. The Na+K+ ATPase continues to pump Na+ outward to maintain the Na+ gradient that drives glucose uptake. WORKED EXAMPLE 11-3 Energetics of Pumping by Symport Calculate the maximum ratio that can be achieved by the plasma membrane Na+-glucose symporter of an epithelial cell when [Na+]in is 12 mM, [Na+]out is 145 mM, the membrane potential is −50 mV (inside negative), and the temperature is 37 °C. [glucose]in [glucose]out SOLUTION: Using Equation 11-4 (p. 392), we can calculate the energy inherent in an electrochemical Na+ gradient — that is, the cost of moving one Na+ ion up this gradient: ΔGt= RT ln + ZF Δψ We then substitute standard values for R, T, and F; the given values for [Na+] (expressed as molar concentrations); +1 for Z (because Na+ has a positive charge); and 0.050 V for Δψ . Note that the membrane potential is −50 mV (inside negative), so the change in potential when an ion moves from inside to outside is 50 mV. ΔGt = (8.315 J /mol∙K)(310 K) ln +1(96,500 J /V ∙mol)(0.050 V) = 11.2 kJ /mol When Na+ reenters the cell, it releases the electrochemical potential created by pumping it out; ΔG for reentry is −11.2 kJ/mol of Na+. This is the potential energy per mole of Na+ that is available to pump glucose. Given that two Na+ ions pass down their electrochemical gradient and into the cell for each glucose [Na+]out [Na+]in (1.45× 10−1) (1.2× 10−2) carried in by symport, the energy available to pump 1 mol of glucose is 2× 11.2 kJ /mol= 22.4 kJ /mol. We can now calculate the maximum concentration ratio of glucose that can be achieved by this pump (from Eqn 11-3, p. 392): ΔGt= RT ln Rearranging, then substituting the values of ΔGt, R, and T, gives ln = = = 8.69 = e8.69 = 5.94× 103 Thus, the cotransporter can pump glucose inward until its concentration inside the epithelial cell is about 6,000 times the concentration outside (in the intestine). (This is the maximum theoretical ratio, assuming a perfectly efficient coupling of Na+ reentry and glucose uptake.) As glucose molecules are pumped from the intestine into the epithelial cell at the apical surface, glucose is simultaneously moved from the cell into the blood by passive transport through a glucose transporter (GLUT2) in the basal surface (Fig. 11-42). The crucial role of Na+ in symport and antiport systems such as this [glucose]in [glucose]out [glucose]in [glucose]out ΔGt RT 22.4 kJ /mol (8.315 J /mol∙K)(310 K) [glucose]in [glucose]out requires the continued outward pumping of Na+ to maintain the transmembrane Na+ gradient. In the kidney, a different Na+-glucose symporter (SGLT2) is the target of drugs used to treat type 2 diabetes. Gliflozins are specific inhibitors of this Na+-glucose symporter. They lower blood glucose by inhibiting glucose reabsorption in the kidney, thus preventing the damaging effects of elevated blood glucose. Glucose not reabsorbed in the kidney is cleared in the urine. This class of drugs, taken orally in combination with diet and exercise, lowers blood glucose significantly in individuals with type 2 diabetes. Because of the essential role of ion gradients in active transport and energy conservation, compounds that collapse ion gradients across cellular membranes are effective poisons, and those that are specific for infectious microorganisms can serve as antibiotics. One such substance is valinomycin, a small cyclic peptide that neutralizes the K+ charge by surrounding the ion with six carbonyl oxygens (Fig. 11-43). The hydrophobic peptide then acts as a shuttle, carrying K+ across the membrane down its concentration gradient and deflating that gradient. Compounds that shuttle ions across membranes in this way are called ionophores (“ion bearers”). Both valinomycin and monensin (a Na+-carrying ionophore) are antibiotics; they kill microbial cells by disrupting secondary transport processes and energy- conserving reactions. Monensin is widely used as an antifungal and antiparasitic agent.
FIGURE 11-43 Valinomycin, a peptide ionophore that binds K+. The central K+ ion is surrounded by inward-facing polar and charged amino acid side chains, and the outside surface is covered with nonpolar side chains that make the whole structure hydrophobic enough to diffuse through the lipid bilayer, carrying K+ down its concentration gradient. The resulting dissipation of the transmembrane ion gradient kills microbial cells, making valinomycin a potent antibiotic. [Information from P. Barak and E. A. Nater, http://virtual-museum.soils.wisc.edu, and K. Neupert-Laves and M. Dobler, Helv. Chim. Acta 58:432, 1975.] Aquaporins Form Hydrophilic Transmembrane Channels for the Passage of Water A family of integral membrane proteins, the aquaporins (AQPs), provide channels for rapid movement of water molecules across all plasma membranes. Aquaporins are found in all organisms, and multiple aquaporin genes are generally present, encoding similar but not identical proteins. Eleven aquaporins are known in mammals, each with a specific location and role (Table 11-3). The exocrine glands that produce sweat, saliva, and tears secrete water through aquaporins. Erythrocytes, which swell or shrink rapidly in response to abrupt changes in extracellular osmolarity as blood travels through the renal medulla, have a high density of aquaporin in their plasma membrane (2× 105 copies of AQP1 per cell). Seven aquaporins play roles in urine production and water retention in the nephron (the functional unit of the kidney). Each renal AQP has a specific location in the nephron, and each has specific properties and regulatory features. For example, AQP2 in the epithelial cells of the renal collecting duct is regulated by vasopressin (also called antidiuretic hormone): more water is reabsorbed from the duct into the kidney tissues when the vasopressin level is high. Mutant mice with no AQP2 gene have greater urine output (polyuria) and more dilute urine, the result of the proximal tubule becoming less permeable to water. In humans, genetically defective AQPs are known to be responsible for a variety of diseases. TABLE 11-3 Permeability Characteristics and Predominant Distribution of Known Mammalian Aquaporins Aquaporin Permeant (permeability) Tissue distribution Primary subcellular distribution AQP0 Water (low) Lens Plasma membrane AQP1 Water (high) Erythrocyte, kidney, lung, vascular endothelium, brain, eye Plasma membrane AQP2 Water (high) Kidney, vas deferens Apical plasma membrane, intracellular vesicles AQP3 Water (high), glycerol (high), urea (moderate) Kidney, skin, lung, eye, colon Basolateral plasma membrane AQP4 Water (high) Brain, muscle, kidney, lung, stomach, small intestine Basolateral plasma membrane AQP5 Water (high) Salivary gland, lacrimal gland, sweat gland, lung, cornea Apical plasma membrane AQP6 Water (low), anions (NO−3 > Cl−) Kidney Intracellular vesicles AQP7 Water (high), glycerol (high), urea (high) Adipose tissue, kidney, testis Plasma membrane AQP8 Water (high) Testis, kidney, liver, pancreas, small intestine, colon Plasma membrane, intracellular vesicles AQP9 Water (low), glycerol (high), Liver, leukocyte, brain, testis Plasma membrane a b urea (high) AQP10 Water (low), glycerol (high), urea (high) Small intestine Intracellular vesicles Information from L. S. King et al., Nat. Rev. Mol. Cell Biol. 5:688, 2004. The apical plasma membrane faces the lumen of the gland or tissue; the basolateral plasma membrane is along the sides and base of the cell, not facing the lumen of the gland or tissue. AQP8 might also be permeated by urea. Water molecules flow through an AQP1 channel at a rate of about 109 s−1. For comparison, the highest known turnover number for an enzyme is that for catalase, 4× 107s−1, and many enzymes have turnover numbers between 1 s−1 and 104 s−1 (see Table 6-7). The low activation energy for passage of water through aquaporin channels (ΔG‡< 15 kJ /mol) suggests that water moves through the channels in a continuous stream, in the direction dictated by the osmotic gradient. (For a discussion of osmosis, see p. 52.) Aquaporins do not allow passage of protons (hydronium ions, H3O+), which would collapse membrane electrochemical gradients. Ion-Selective Channels Allow Rapid Movement of Ions across Membranes Ion-selective channels — first recognized in neurons and now known to be present in the plasma membranes of all cells, as well as in the intracellular membranes of eukaryotes — provide another mechanism for moving inorganic ions across a b membranes. Ion channels, together with ion pumps such as the Na+K+ ATPase, determine a plasma membrane’s permeability to specific ions and regulate the cytosolic concentration of ions and the membrane potential. In neurons, very rapid changes in the activity of ion channels cause the changes in membrane potential (action potentials) that carry signals from one end of a neuron to the other. In myocytes, rapid opening of Ca2+ channels in the sarcoplasmic reticulum releases the Ca2+ that triggers muscle contraction. We discuss the signaling functions of ion channels in Chapter 12. Here we describe the structural basis for ion-channel function, using as our example a well-studied K+ channel. Ion channels are distinct from ion transporters in at least three ways. First, the rate of flux through channels can be orders of magnitude greater than the turnover number for a transporter — 107 to 108 ions/s for an ion channel, approaching the theoretical maximum for unrestricted diffusion. By contrast, the turnover rate of the Na+K+ ATPase is about 100 s−1. Second, ion channels are not saturable: rates do not approach a maximum at high substrate concentration. Third, they are gated in response to some type of cellular event. In ligand-gated channels (which are generally oligomeric), binding of an extracellular or intracellular small molecule forces an allosteric transition in the protein, which opens or closes the channel. In voltage-gated ion channels, a change in transmembrane electrical potential (Vm) causes a charged protein domain to move relative to the membrane, opening or closing the channel. Both types of gating can be very fast. A channel typically opens in a fraction of a millisecond and may remain open for only milliseconds, making these molecular devices effective for very fast signal transmission in the nervous system. Because a single ion channel typically remains open for only a few milliseconds, monitoring this process is beyond the limit of most biochemical measurements. Ion fluxes must therefore be measured electrically, either as changes in Vm (in the millivolt range) or as electric current I (in the microampere or picoampere range), using microelectrodes and appropriate amplifiers. In patch-clamping, very small currents are measured through a tiny region of the membrane surface containing only one or a few ion- channel molecules (Fig. 11-44). The researcher can measure the size and duration of the current that flows during one opening of an ion channel and can determine how o en a channel opens and how that frequency is affected by membrane potential, regulatory ligands, toxins, and other agents. Patch-clamp studies have revealed that as many as 104 ions can move through a single ion channel in 1 ms. Such an ion flux represents a huge amplification of the initial signal, which may be just one or two signaling molecules (neurotransmitters, for example). FIGURE 11-44 Electrical measurements of ion-channel function. The “activity” of an ion channel is estimated by measuring the flow of ions through it, using the patch-clamp technique. A finely drawn-out pipette (micropipette) is pressed against the cell surface, and negative pressure in the pipette forms a pressure seal between pipette and membrane. As the pipette is pulled away from the cell, it pulls off a tiny patch of membrane (which may contain one or a few ion channels). A er placing the pipette and attached patch in an aqueous solution, the researcher can measure channel activity as the electric current that flows between the contents of the pipette and the aqueous solution. In practice, a circuit is set up that “clamps” the transmembrane potential at a given value and measures the current that must flow to maintain this voltage. With highly sensitive detectors, researchers can measure the current flowing through a single ion channel, typically a few picoamperes. The trace shows the current through a single acetylcholine receptor channel as a function of time (in milliseconds), revealing how fast the channel opens and closes, how frequently it opens, and how long it stays open. Downward deflection represents channel opening. Clamping the Vm at different values permits determination of the effect of membrane potential on these parameters of channel function. [Information from V. Witzemann et al., Proc. Natl. Acad. Sci. USA 93:13,286, 1996.] The Structure of a K+ Channel Reveals the Basis for Its Specificity The structure of a potassium channel from the bacterium Streptomyces lividans provides important insight into the way ion channels work. This bacterial ion channel is related in sequence to all other known K+ channels and serves as the prototype for such channels, including the voltage-gated K+ channel of neurons. Among the members of this protein family, the similarities in sequence are greatest in the “pore region,” which contains the ion selectivity filter that allows K+ (radius 1.33 Å) to pass 104 times more readily than Na+ (radius 0.95 Å) — at a rate (about 108 ions/s) approaching the theoretical limit for unrestricted diffusion. The K+ channel consists of four identical subunits that span the membrane and form a cone within a cone surrounding the ion channel, with the wide end of the double cone facing the extracellular space (Fig. 11-45a). Each subunit has two transmembrane α helices and a third, shorter helix that contributes to the pore region. The outer cone is formed by one of the transmembrane helices of each subunit. The inner cone, formed by the other four transmembrane helices, surrounds the ion channel and cradles the ion selectivity filter. Viewed perpendicular to the plane of the membrane, the central channel is seen to be just wide enough to accommodate an unhydrated metal ion such as potassium (Fig. 11-45b). FIGURE 11-45 The K+ channel of Streptomyces lividans. (a) Viewed in the plane of the membrane, the channel consists of eight transmembrane helices (two from each of four identical subunits), forming a cone with its wide end toward the extracellular space. The inner helices of the cone (lighter colored) line the transmembrane channel, and the outer helices interact with the lipid bilayer. Short segments of each subunit converge in the open end of the cone to make a selectivity filter. (b) This view, perpendicular to the plane of the membrane, shows the four subunits arranged around a central channel just wide enough for a single K+ ion to pass. (c) Diagram of a K+ channel in cross section, showing the structural features critical to function. K+ ions go through the channel in pairs, first in sites 1 and 3, then in sites 2 and 4. Carbonyl oxygens (red) of the peptide backbone in the selectivity filter protrude into the channel, interacting with and stabilizing the K+ ions that are passing through. [Data from (a, b) PDB ID 1BL8, D. A. Doyle et al., Science 280:69, 1998; (c) G. Yellen, Nature 419:35, 2002, and PDB ID 1J95, M. Zhou et al., Nature 411:657, 2001.] Both the ion specificity and the high flux through the channel are understandable from what we know of the channel’s structure (Fig. 11-45c). At the inner and outer plasma membrane surfaces, the entryways to the channel have several negatively charged amino acid residues, which presumably increase the local concentration of cations such as K+ and Na+. The ion path through the membrane begins (on the inner surface) as a wide, water-filled channel in which the ion can retain its hydration sphere. Further stabilization is provided by the short helices in the pore region of each subunit, with the partial negative charges of their electric dipoles pointed at K+ in the channel. About two- thirds of the way through the membrane, this channel narrows in the region of the selectivity filter, forcing the ion to give up its hydrating water molecules. Carbonyl oxygen atoms in the backbone of the selectivity filter replace the water molecules in the hydration sphere, forming a series of perfect coordination shells through which the K+ moves. This favorable interaction with the filter is not possible for Na+, which is too small to make contact with all the potential oxygen ligands. The preferential stabilization of K+ is the basis for the ion selectivity of the filter, and mutations that change residues in this part of the protein eliminate the channel’s ion selectivity. The K+-binding sites of the filter are flexible enough to collapse to fit any Na+ that enters the channel, and this conformational change closes the channel. There are four potential K+-binding sites along the selectivity filter, each composed of an oxygen “cage” that provides ligands for the K+ ions (Fig. 11-45c). In the crystal structure, two K+ ions are visible within the selectivity filter, about 7.5 Å apart, and two water molecules occupy the unfilled positions. K+ ions pass through the filter in single file; their mutual electrostatic repulsion probably just balances the interaction of each ion with the selectivity filter and keeps them moving. Movement of the two K+ ions is concerted: first they occupy positions 1 and 3, then they hop to positions 2 and 4. The energetic difference between these two configurations (1, 3 and 2, 4) is very small; energetically, the selectivity pore is not a series of hills and valleys but a flat surface, which is ideal for rapid ion movement through the channel. The structure of the channel seems to have been optimized during evolution to give maximal flow rates and high specificity. SUMMARY 11.3 Solute Transport across Membranes Some transporters simply facilitate passive diffusion of a solute across the membrane, from a higher concentration to a lower concentration. Others transport solutes against an electrochemical gradient; this requires a source of metabolic energy. Transporters move solutes across a membrane one or a few at a time, providing a binding site on each side of the membrane. The binding sites alternate between being accessible from the outside and from the inside. Ion channels provide a path across the membrane, which is either open or closed. When open, the channel allows the movement of large numbers of solute ion across the membrane at nearly the speed of unhindered diffusion. A family of glucose transporters in humans includes the passive transporter GLUT1, which is saturated at normal levels of glucose in the blood. GLUT1 facilitates movement of glucose from blood into erythrocytes. The chloride-bicarbonate exchanger of erythrocytes exchanges one Cl− ion for one HCO− 3 ion across the erythrocyte plasma membrane, mediating the uptake of CO2 in the tissues and its release in the lungs. Active transporters use energy to pump solutes against an electrochemical gradient. P-type ATPases, including the Na+K+ ATPase of the plasma membrane and the Ca2+ transporters of the sarcoplasmic/endoplasmic reticulum, couple phosphorylation and dephosphorylation of the transporter to alternate exposure of solute binding sites on the inside and the outside of the membrane. In animal cells, the Na+K+ ATPase maintains the differences in cytosolic and extracellular concentrations of Na+ and K+, and the resulting Na+ gradient is used as the energy source for a variety of secondary active transport processes. V-type and F-type ATPases are active transporters that couple ATP cleavage to the uphill transport of H+ ions. The same mechanism, working in reverse, allows the synthesis of ATP, driven by movement of protons down their electrochemical gradient. ABC transporters carry a variety of substrates (including many drugs) out of cells, using ATP as the energy source. The ATP-using domain is conserved in many ABC transporters, and it is coupled with various transmembrane domains that give substrate specificity. Some active cotransporters use the energy in an ion gradient generated catabolically to move a solute uphill. The Na+-glucose cotransporter of the kidney and intestine is such a transporter. Water moves across membranes through aquaporins. Some aquaporins are regulated; some also transport glycerol or urea. Ion channels provide hydrophilic pores through which select ions can diffuse, moving down their electrical or chemical concentration gradients. Ion channels are unsaturable, have very high flux rates, and are highly specific for one ion. Structural studies of K+ channels reveal the mechanism that allows great discrimination between K+ and other ions like Na+. The polar transmembrane passage precisely fits the K+ ion, but allows neither larger ions nor smaller ions to pass. Chapter Review KEY TERMS Terms in bold are defined in the glossary. micelle bilayer vesicle fluid mosaic model lipid transfer protein integral proteins peripheral proteins amphitropic proteins monotopic bitopic polytopic hydropathy index β barrel porin positive-inside rule GPI-anchored protein flippases floppases scramblases FRAP microdomains ra s caveolae caveolin BAR domain septins fusion protein v-SNAREs t-SNAREs integrin selectin simple diffusion membrane potential (Vm) electrochemical gradient electrochemical potential transporters passive transport active transport ion channels Kt(Ktransport) amphipathic electroneutral cotransport antiport symport uniport electrogenic P-type ATPases SERCA pump Na+K+ ATPase V-type ATPases F-type ATPases ATP synthase ABC transporters multidrug transporters Na+-glucose symporter ionophore aquaporins (AQPs) ligand-gated channel voltage-gated ion channel patch-clamping PROBLEMS 1. Determining the Cross-Sectional Area of a Lipid Molecule When phospholipids are layered gently onto the surface of water, they orient at the air-water interface with their head groups in the water and their hydrophobic tails in the air. The experimental apparatus shown can be used to progressively reduce the surface area available to a layer of lipids. By measuring the force necessary to push the lipids together, we can determine when the molecules are packed tightly in a continuous monolayer; as that area is approached, the force needed to further reduce the surface area increases sharply, as shown in the graph. How would you use this apparatus to determine the average area occupied by a single lipid molecule in the monolayer? 2. Properties of Lipids and Lipid Bilayers Lipid bilayers form when phospholipids are suspended in water. The edges of these sheets close upon each other and undergo self-sealing to form vesicles (liposomes). a. What properties of lipids are responsible for this property of bilayers? Explain. b. What are the consequences of this property for the structure of biological membranes? 3. Length of a Fatty Acid Molecule The carbon–carbon bond distance for single-bonded carbons, such as those in a saturated fatty acyl chain, is about 1.5 Å. Estimate to one significant figure the length of a single molecule of palmitate in its fully extended form. If two molecules of palmitate were placed end to end, how would their total length compare with the thickness of the lipid bilayer in a biological membrane? 4. Membrane Proteins What are the three main categories of membrane proteins, and how are they distinguished experimentally? 5. Location of a Membrane Protein Treatment of disrupted erythrocyte membranes with a concentrated salt solution released an unknown membrane protein, X. Proteolytic enzymes cleaved X into fragments. In additional experiments, intact erythrocytes were treated with proteolytic enzymes, washed, then disrupted. Extraction of membrane components yielded intact X. What do these observations indicate about the location of X in the plasma membrane? Do the properties of X resemble those of an integral membrane protein or a peripheral membrane protein? 6. Predicting Membrane Protein Topology from Sequence You have cloned the gene for a human erythrocyte protein, which you suspect is a membrane protein. You deduce the amino acid sequence of the protein from the nucleotide sequence of the gene. From this sequence alone, how would you evaluate the possibility that the protein is an integral protein? Suppose the protein proves to be an integral protein with one transmembrane segment. Suggest biochemical or chemical experiments that might allow you to determine whether the protein is oriented with the amino terminus on the outside of the cell or on the inside of the cell. 7. Surface Density of a Membrane Protein E. coli can be induced to make about 10,000 copies of the lactose transporter (Mr 31,000) per cell. Assume that E. coli is a cylinder 1 µm in diameter and 2 µm long. The diameter of the lactose transporter is approximately 6 nm. What fraction of the plasma membrane surface is occupied by the lactose transporter molecules? Explain how you arrived at this conclusion. 8. Molecular Species in the Plasma Membrane The plasma membrane of E. coli is about 75% protein and 25% phospholipid by weight. How many molecules of membrane lipid are present for each molecule of membrane protein? Assume an average protein Mr of 50,000 and an average phospholipid Mr of 750. What more would you need to know to estimate the fraction of the membrane surface that is covered by lipids? 9. Temperature Dependence of Lateral Diffusion The experiment described in Figure 11-19 was performed at 37 °C. If the experiment were carried out at 10 °C, what effect would you expect on the rate of diffusion? Why? 10. Membrane Self-Sealing Cellular membranes are self- sealing — if they are punctured or disrupted mechanically, they quickly and automatically reseal. What properties of membranes are responsible for this important feature? 11. Lipid Melting Temperatures Membrane lipids in tissue samples obtained from different parts of a reindeer’s leg have different fatty acid compositions. Membrane lipids from tissue near the hooves contain a larger proportion of unsaturated fatty acids than those from tissue in the upper leg. What is the significance of this observation? 12. Flip-Flop Diffusion What is the physical explanation for the very slow movement of membrane phospholipids from one leaflet of a biological membrane to the other? What factors influence this rate? 13. Bilayer Asymmetry The inner leaflet (monolayer) of the human erythrocyte membrane consists predominantly of phosphatidylethanolamine and phosphatidylserine. The outer leaflet consists predominantly of phosphatidylcholine and sphingomyelin. Although the phospholipid components of the membrane can diffuse in the fluid bilayer, this sidedness is preserved at all times. How? 14. Scramblase and Flippase Explain the difference between the scramblase enzymes and flippase enzymes based on the membranes with which they are associated, the symmetry of these membranes, and their energy requirements. 15. Membrane Permeability At pH 7, tryptophan crosses a lipid bilayer at about one-thousandth the rate of indole, a closely related compound: Suggest an explanation for this observation. 16. Glucose Transporters A cell biologist working with cultured cells from intestinal epithelium finds that the cells take up glucose from the growth medium 10 times faster when the glucose concentration is 5 mM than when it is 0.2 mM. She also finds that glucose uptake requires Na+ in the growth medium. What can you say about the glucose transporter in these cells? 17. Use of the Helical Wheel Diagram A helical wheel is a two-dimensional representation of a helix, a view along its central axis. Use the helical wheel diagram shown here to determine the distribution of amino acid residues in a helical segment with the sequence –Val–Asp–Arg–Val–Phe–Ser–Asn– Val–Cys–Thr–His–Leu–Lys–Thr–Leu–Gln–Asp–Lys– What can you say about the surface properties of this helix? How would you expect the helix to be oriented in the tertiary structure of an integral membrane protein? 18. Synthesis of Gastric Juice: Energetics Parietal cells acidify gastric juice (pH 1.5) by pumping HCl from blood plasma (pH 7.4) into the stomach. How much ATP (in moles) is required to pump a mole of protons (H+) against this concentration gradient? The free-energy change for ATP hydrolysis under cellular conditions is about –58 kJ/mol. Ignore the effects of the transmembrane electrical potential. 19. Electrogenic Transporters A single-cell organism, Paramecium, is large enough to allow the insertion of a microelectrode, permitting the measurement of the electrical potential between the inside of the cell and the surrounding medium (the membrane potential). The measured membrane potential is −50 mV (inside negative) in a living cell. What would happen if you added valinomycin to the surrounding medium, which contains K+ and Na+? 20. Energetics of the Na+K+ ATPase For a typical vertebrate cell with a membrane potential of −0.070 V (inside negative), what is the free-energy change for transporting 1 mol of Na+ from the cell into the blood at 37 °C? Assume the Na+ concentration is 12 mM inside the cell and 145 mM in blood plasma. 21. Action of Ouabain on Kidney Tissue Ouabain specifically inhibits the Na+K+ ATPase activity of animal tissues but is not known to inhibit any other enzyme. When ouabain is added to thin slices of living kidney tissue, it inhibits oxygen consumption by 66%. Why? What does this observation tell us about the use of respiratory energy by kidney tissue? 22. Digoxin to Inhibit Na+K+ ATPase The Na+Ca2+ exchanger expressed in cardiac myocytes is a bidirectional antiporter protein that removes calcium from the cytoplasm by exchanging it with sodium. Cardiac myocytes also express the Na+K+ ATPase. Suppose that a Na+K+ ATPase inhibitor (digoxin) is added to cardiac myocytes. Using your knowledge of the relative concentrations of ions (intracellular versus extracellular) and the important role of the Na+K+ ATPase in maintaining the electrochemical gradient, what change would you expect in the intracellular [Ca2+]? Why? 23. Energetics of Symport Suppose you determined experimentally that a cellular transport system for glucose, driven by symport of Na+, could accumulate glucose to concentrations 25 times greater than in the external medium, while the external [Na+] was only 10 times greater than the intracellular [Na+]. Would this violate the laws of thermodynamics? If not, how could you explain this observation? 24. Labeling the Lactose Transporter A bacterial lactose transporter, which is highly specific for lactose, contains a Cys residue that is essential to its transport activity. Covalent reaction of N-ethylmaleimide (NEM) with this Cys residue irreversibly inactivates the transporter. A high concentration of lactose in the medium prevents inactivation by NEM, presumably by sterically protecting the Cys residue, which is in or near the lactose-binding site. You know nothing else about the transporter protein. Suggest an experiment that might allow you to determine the Mr of this Cys-containing transporter polypeptide. 25. Transport Types You have just discovered a new L- alanine transporter in liver cells (hepatocytes). Poisoning hepatocytes with cyanide (which blocks ATP synthesis) reduces alanine transport by 90%. Tenfold reduction in extracellular [Na+] has no immediate effect on alanine transport. How would you use these observations to decide whether the alanine transporter is passive or active, primary or secondary? 26. Intestinal Uptake of Leucine You are studying the uptake of L-leucine by epithelial cells of the mouse intestine. Measurements of the rates of uptake of L-leucine and several of its analogs, with and without Na+ in the assay buffer, yield the results given in the table. What can you conclude about the properties and mechanism of the leucine transporter? Would you expect L-leucine uptake to be inhibited by ouabain (see problem 21)? Substrate Uptake in presence of Na+ Uptake in absence of Na+ Vmax Kt(mM) Vmax Kt(mM) -Leucine 420 0.24 23 0.2 -Leucine 310 4.7 5 4.7 -Valine 225 0.31 19 0.31 27. Ion Channel Selectivity Potassium channels consist of four subunits that form a channel just wide enough for K+ ions to pass through. Although Na+ ions are smaller (Mr 23, radius 0.95 Å) than K+ ions (Mr 39, radius 1.33 Å), the potassium channels in the bacterium Streptomyces lividans transport 104 times more K+ ions than Na+ ions. What prevents Na+ ions from passing through potassium channels? 28. Effect of an Ionophore on Active Transport Consider the leucine transporter described in Problem 26. Would Vmax and/or Kt change if you added a Na+ ionophore to the assay solution containing Na+? Explain. BIOCHEMISTRY ONLINE 29. Predicting Membrane Protein Topology I Online bioinformatics tools make hydropathy analysis easy if you know the amino acid sequence of a protein. At the Protein Data Bank (www.rcsb.org), the Protein Feature View displays additional information about a protein gleaned from other databases, such as UniProt and SCOP2. A simple graphical view of a hydropathy plot created using a window of 15 residues shows hydrophobic regions in red and hydrophilic regions in blue. a. Looking only at the displayed hydropathy plots in the Protein Feature View, what predictions would you make about the membrane topology of these proteins: glycophorin A (PDB ID 1AFO), myoglobin (PDB ID 1MBO), and aquaporin (PDB ID 2B6O)? b. Now, refine your information using the ProtScale tools at the ExPASy bioinformatics resource portal. Each of the PDB Protein Feature Views was created with a UniProt Knowledgebase ID. For glycophorin A, the UniProtKB ID is P02724; for myoglobin, P02185; and for aquaporin, Q6J8I9. Go to the ExPASy portal (http://web.expasy.org/protscale) and select the Kyte & Doolittle hydropathy analysis option, with a window of 7 amino acids. Enter the UniProtKB ID for aquaporin (Q6J8I9, which you can also get from the PDB’s Protein Feature View page), then select the option to analyze the complete chain (residues 1 to 263). Use the default values for the other options and click Submit to get a hydropathy plot. Save a GIF image of this plot. Now repeat the analysis using a window of 15 amino acids. Compare the results for the 7-residue and 15-residue window analyses. Which window size gives you a better signal-to-noise ratio? c. Under what circumstances would it be important to use a narrower window? 30. Predicting Membrane Protein Topology II The epinephrine receptor in animal cells is an integral membrane protein (Mr 64,000) that is believed to have seven transmembrane α -helical regions. a. Show that a protein of this size is capable of spanning the membrane seven times. b. Given the amino acid sequence of this protein, how would you predict which regions of the protein form the membrane-spanning helices? c. Go to the Protein Data Bank (www.rcsb.org). Use the PDB identifier 1DEP to retrieve the data page for a portion of the β -adrenergic receptor (one type of epinephrine receptor) isolated from turkey. Using JSmol to explore the structure, predict whether this portion of the receptor is located within the membrane or at the membrane surface. Explain your answer. Now use the Protein Feature View to see the hydrophobicity analysis of the sequence. Does this support your answer? d. Retrieve the data for a portion of another receptor, the acetylcholine receptor of neurons and myocytes, using the PDB identifier 1A11. As in (c), predict where this portion of the receptor is located and explain your answer. DATA ANALYSIS PROBLEM 31. The Fluid Mosaic Model of Biological Membrane Structure Figure 11-3 shows the currently accepted fluid mosaic model of biological membrane structure. This model was presented in detail in a review article by S. J. Singer in 1971. In the article, Singer presented the three models of membrane structure that had been proposed up to that time: A. The Davson-Danielli-Robertson Model. This was the most widely accepted model in 1971, when Singer’s review was published. In this model, the phospholipids are arranged as a bilayer. Proteins are found on both surfaces of the bilayer, attached to it by ionic interactions between the charged head groups of the phospholipids and charged groups of the proteins. Crucially, there is no protein in the interior of the bilayer. B. The Benson Lipoprotein Subunit Model. Here the proteins are globular and the membrane is a protein-lipid mixture. The hydrophobic tails of the lipids are embedded in the hydrophobic parts of the proteins. The lipid head groups are exposed to the solvent. There is no lipid bilayer. C. The Lipid–Globular Protein Mosaic Model. This is the model shown in Figure 11-3. The lipids form a bilayer and proteins are embedded in it, some extending through the bilayer and others not. Proteins are anchored in the bilayer by interactions between the hydrophobic tails of the lipids and hydrophobic portions of the protein. For the data given below, consider how each piece of information aligns with each of the three models of membrane structure. Which model(s) are supported, which are not supported, and what reservations do you have about the data or their interpretation? Explain your reasoning. a. When cells were fixed, stained with osmium tetroxide, and examined in the electron microscope, the membranes showed a “railroad track” appearance, with two dark-staining lines separated by a light space. b. The thickness of membranes in cells fixed and stained in the same way was found to be 5 to 9 nm. The thickness of a “naked” phospholipid bilayer, without proteins, was 4 to 4.5 nm. The thickness of a single monolayer of proteins was about 1 nm. c. Singer wrote in his article: “The average amino acid composition of membrane proteins is not distinguishable from that of soluble proteins. In particular, a substantial fraction of the residues is hydrophobic” (p. 165). d. As described in Problems 1 and 2 of this chapter, researchers had extracted membranes from cells, extracted the lipids, and compared the area of the lipid monolayer with the area of the original cell membrane. The interpretation of the results was complicated by the issue illustrated in the graph of Problem 1: the area of the monolayer depended on how hard it was pushed. With very light pressures, the ratio of monolayer area to cell membrane area was about 2.0. At higher pressures — thought to be more like those found in cells — the ratio was substantially lower. e. Circular dichroism spectroscopy uses changes in polarization of UV light to make inferences about protein secondary structure (see Fig. 4-9). On average, this technique showed that membrane proteins have a large amount of α helix and little or no β sheet. This finding was consistent with most membrane proteins having a globular structure. f. Phospholipase C is an enzyme that removes the polar head group (including the phosphate) from phospholipids. In several studies, treatment of intact membranes with phospholipase C removed about 70% of the head groups without disrupting the “railroad track” structure of the membrane. g. Singer described in his article a study in which “a glycoprotein of molecular weight about 31,000 in human red blood cell membranes is cleaved by tryptic treatment of the membranes into soluble glycopeptides of about 10,000 molecular weight, while the remaining portions are quite hydrophobic” (p. 199). Trypsin treatment did not cause gross changes in the membranes, which remained intact. Singer’s review also included many more studies in this area. In the end, though, the data available in 1971 did not conclusively prove Model C was correct. As more data have accumulated, this model of membrane structure has been accepted by the scientific community. Reference Singer, S.J. 1971. The molecular organization of biological membranes. In Structure and Function of Biological Membranes (L.I. Rothfield, ed.), pp. 145–222. New York: Academic Press.
Stems are from the chapter Problems section; correct choices are drawn from Abbreviated Solutions to Problems (Appendix B) in the same edition.
1. Determining the Cross-Sectional Area of a Lipid Molecule When phospholipids are layered gently onto the surface of water, they orient at the air-water interface with their head groups in the water and their hydrophobic tails in the air. The experimental apparatus shown can be used to progressively reduce the surface area available to a layer of lipids. By measuring the force necessary to push the lipids together, we can determine when the molecules are packed tightly in a continuous monolayer; as that area is approached, the force needed to further reduce the surface area increases sharply, as shown in the graph. How would you use this apparatus to determine the average area occupied by a single lipid molecule in the monolayer?
2. Properties of Lipids and Lipid Bilayers Lipid bilayers form when phospholipids are suspended in water. The edges of these sheets close upon each other and undergo self-sealing to form vesicles (liposomes). a. What properties of lipids are responsible for this property of bilayers? Explain. b. What are the consequences of this property for the structure of biological membranes?
3. Length of a Fatty Acid Molecule The carbon–carbon bond distance for single-bonded carbons, such as those in a saturated fatty acyl chain, is about 1.5 Å. Estimate to one significant figure the length of a single molecule of palmitate in its fully extended form. If two molecules of palmitate were placed end to end, how would their total length compare with the thickness of the lipid bilayer in a biological membrane?
4. Membrane Proteins What are the three main categories of membrane proteins, and how are they distinguished experimentally?
5. Location of a Membrane Protein Treatment of disrupted erythrocyte membranes with a concentrated salt solution released an unknown membrane protein, X. Proteolytic enzymes cleaved X into fragments. In additional experiments, intact erythrocytes were treated with proteolytic enzymes, washed, then disrupted. Extraction of membrane components yielded intact X. What do these observations indicate about the location of X in the plasma membrane? Do the properties of X resemble those of an integral membrane protein or a peripheral membrane protein?
6. Predicting Membrane Protein Topology from Sequence You have cloned the gene for a human erythrocyte protein, which you suspect is a membrane protein. You deduce the amino acid sequence of the protein from the nucleotide sequence of the gene. From this sequence alone, how would you evaluate the possibility that the protein is an integral protein? Suppose the protein proves to be an integral protein with one transmembrane segment. Suggest biochemical or chemical experiments that might allow you to determine whether the protein is oriented with the amino terminus on the outside of the cell or on the inside of the cell.
7. Surface Density of a Membrane Protein E. coli can be induced to make about 10,000 copies of the lactose transporter (Mr 31,000) per cell. Assume that E. coli is a cylinder 1 µm in diameter and 2 µm long. The diameter of the lactose transporter is approximately 6 nm. What fraction of the plasma membrane surface is occupied by the lactose transporter molecules? Explain how you arrived at this conclusion.
8. Molecular Species in the Plasma Membrane The plasma membrane of E. coli is about 75% protein and 25% phospholipid by weight. How many molecules of membrane lipid are present for each molecule of membrane protein? Assume an average protein Mr of 50,000 and an average phospholipid Mr of 750. What more would you need to know to estimate the fraction of the membrane surface that is covered by lipids?
9. Temperature Dependence of Lateral Diffusion The experiment described in Figure 11-19 was performed at 37 °C. If the experiment were carried out at 10 °C, what effect would you expect on the rate of diffusion? Why?
10. Membrane Self-Sealing Cellular membranes are self- sealing — if they are punctured or disrupted mechanically, they quickly and automatically reseal. What properties of membranes are responsible for this important feature?
11. Lipid Melting Temperatures Membrane lipids in tissue samples obtained from different parts of a reindeer’s leg have different fatty acid compositions. Membrane lipids from tissue near the hooves contain a larger proportion of unsaturated fatty acids than those from tissue in the upper leg. What is the significance of this observation?
12. Flip-Flop Diffusion What is the physical explanation for the very slow movement of membrane phospholipids from one leaflet of a biological membrane to the other? What factors influence this rate?
13. Bilayer Asymmetry The inner leaflet (monolayer) of the human erythrocyte membrane consists predominantly of phosphatidylethanolamine and phosphatidylserine. The outer leaflet consists predominantly of phosphatidylcholine and sphingomyelin. Although the phospholipid components of the membrane can diffuse in the fluid bilayer, this sidedness is preserved at all times. How?
14. Scramblase and Flippase Explain the difference between the scramblase enzymes and flippase enzymes based on the membranes with which they are associated, the symmetry of these membranes, and their energy requirements.
15. Membrane Permeability At pH 7, tryptophan crosses a lipid bilayer at about one-thousandth the rate of indole, a closely related compound: Suggest an explanation for this observation.
16. Glucose Transporters A cell biologist working with cultured cells from intestinal epithelium finds that the cells take up glucose from the growth medium 10 times faster when the glucose concentration is 5 mM than when it is 0.2 mM. She also finds that glucose uptake requires Na+ in the growth medium. What can you say about the glucose transporter in these cells?
17. Use of the Helical Wheel Diagram A helical wheel is a two-dimensional representation of a helix, a view along its central axis. Use the helical wheel diagram shown here to determine the distribution of amino acid residues in a helical segment with the sequence –Val–Asp–Arg–Val–Phe–Ser–Asn– Val–Cys–Thr–His–Leu–Lys–Thr–Leu–Gln–Asp–Lys– What can you say about the surface properties of this helix? How would you expect the helix to be oriented in the tertiary structure of an integral membrane protein?
18. Synthesis of Gastric Juice: Energetics Parietal cells acidify gastric juice (pH 1.5) by pumping HCl from blood plasma (pH 7.4) into the stomach. How much ATP (in moles) is required to pump a mole of protons (H+) against this concentration gradient? The free-energy change for ATP hydrolysis under cellular conditions is about –58 kJ/mol. Ignore the effects of the transmembrane electrical potential.
19. Electrogenic Transporters A single-cell organism, Paramecium, is large enough to allow the insertion of a microelectrode, permitting the measurement of the electrical potential between the inside of the cell and the surrounding medium (the membrane potential). The measured membrane potential is −50 mV (inside negative) in a living cell. What would happen if you added valinomycin to the surrounding medium, which contains K+ and Na+?
20. Energetics of the Na+K+ ATPase For a typical vertebrate cell with a membrane potential of −0.070 V (inside negative), what is the free-energy change for transporting 1 mol of Na+ from the cell into the blood at 37 °C? Assume the Na+ concentration is 12 mM inside the cell and 145 mM in blood plasma.
21. Action of Ouabain on Kidney Tissue Ouabain specifically inhibits the Na+K+ ATPase activity of animal tissues but is not known to inhibit any other enzyme. When ouabain is added to thin slices of living kidney tissue, it inhibits oxygen consumption by 66%. Why? What does this observation tell us about the use of respiratory energy by kidney tissue?
22. Digoxin to Inhibit Na+K+ ATPase The Na+Ca2+ exchanger expressed in cardiac myocytes is a bidirectional antiporter protein that removes calcium from the cytoplasm by exchanging it with sodium. Cardiac myocytes also express the Na+K+ ATPase. Suppose that a Na+K+ ATPase inhibitor (digoxin) is added to cardiac myocytes. Using your knowledge of the relative concentrations of ions (intracellular versus extracellular) and the important role of the Na+K+ ATPase in maintaining the electrochemical gradient, what change would you expect in the intracellular [Ca2+]? Why?
23. Energetics of Symport Suppose you determined experimentally that a cellular transport system for glucose, driven by symport of Na+, could accumulate glucose to concentrations 25 times greater than in the external medium, while the external [Na+] was only 10 times greater than the intracellular [Na+]. Would this violate the laws of thermodynamics? If not, how could you explain this observation?
24. Labeling the Lactose Transporter A bacterial lactose transporter, which is highly specific for lactose, contains a Cys residue that is essential to its transport activity. Covalent reaction of N-ethylmaleimide (NEM) with this Cys residue irreversibly inactivates the transporter. A high concentration of lactose in the medium prevents inactivation by NEM, presumably by sterically protecting the Cys residue, which is in or near the lactose-binding site. You know nothing else about the transporter protein. Suggest an experiment that might allow you to determine the Mr of this Cys-containing transporter polypeptide.
25. Transport Types You have just discovered a new L- alanine transporter in liver cells (hepatocytes). Poisoning hepatocytes with cyanide (which blocks ATP synthesis) reduces alanine transport by 90%. Tenfold reduction in extracellular [Na+] has no immediate effect on alanine transport. How would you use these observations to decide whether the alanine transporter is passive or active, primary or secondary?