专题4 物质的跨膜运输

Some mammalian cells, such as nerve and kidney cells, devote up to two-thirds of their total metabolic energy consumption to such transport processes. Cells can also transfer macromolecules and even large particles, such as cell debris, viruses, and bacteria, across their membranes, but the mechanisms involved in most of these cases differ from those used for transferring small molecules, and they are discussed in Chapters 12 and 13.

We begin this chapter by describing some general principles of how small water-soluble molecules traverse cell membranes. We then consider, in turn, the two main classes of membrane proteins that mediate this transmembrane traffic: transporters, which undergo sequential conformational changes to transport specific small molecules across membranes, and channels, which form narrow pores, allowing passive transmembrane movement, primarily of water and small inorganic ions.

Transporters can be coupled to a source of energy to catalyze active transport, which, together with selective passive permeability, creates large differences in the composition of the cytosol compared with that of either the extracellular fluid (Table 11–1) or the fluid within membrane-enclosed organelles. By generating inorganic ion–concentration differences across the lipid bilayer, cell membranes can store potential energy in the form of electrochemical gradients, which drive various transport processes, convey electrical signals in electrically excitable cells, and (in mitochondria, chloroplasts, and bacteria) are harnessed to make most of the cell’s ATP.

We begin this section by describing the permeability properties of an artificial membrane—a synthetic lipid bilayer made solely from lipids without proteins present. We then introduce some of the terms used to describe the various forms of membrane transport and some strategies for characterizing the proteins and processes involved.

Given enough time, virtually any molecule will diffuse down its concentration gradient across a protein-free lipid bilayer. The rate of diffusion, however, varies enormously, depending partly on the size of the molecule but mostly on its relative hydrophobicity (solubility in oil). In general, the smaller the molecule and the more hydrophobic, or nonpolar, it is, the more easily it will diffuse across a lipid bilayer. Small nonpolar molecules, such as O2 and CO2, readily dissolve in lipid bilayers and therefore diffuse rapidly across them.

Small uncharged polar molecules, such as water or urea, also diffuse across a bilayer, albeit much more slowly (Figure 11–1 and see Movie 10.3). By contrast, lipid bilayers are essentially impermeable to charged molecules (ions), no matter how small: the charge and high degree of hydration of such molecules prevent them from entering the hydrocarbon phase of the bilayer (Figure 11–2).

The rate of flow of a solute across the bilayer is directly proportional to the difference in its concentration on the two sides of the membrane. Multiplying this concentration difference (in mol/cm3) by the permeability coefficient (in cm/sec), which is an experimentally determined constant characteristic of each solute, gives the flow of solute in moles per second per square centimeter of bilayer.

Like synthetic lipid bilayers, cell membranes allow small nonpolar molecules to permeate by diffusion. Cell membranes, however, also have to allow the passage of various polar molecules, such as ions, sugars, amino acids, nucleotides, water, and many cell metabolites that cross synthetic lipid bilayers only very slowly. Special membrane transport proteins transfer such solutes across cell membranes. These proteins occur in many forms and in all types of biological membranes. Each protein often transports only a specific molecular species or sometimes a class of molecules (such as ions, sugars, or amino acids).

Early studies found that bacteria with a single-gene mutation were unable to transport a particular class of sugars across their plasma membrane, thereby demonstrating the specificity of membrane transport proteins. We now know that humans with similar mutations suffer from various inherited diseases that hinder the transport of a specific solute or solute class in the kidney, intestine, or other cell type.

All membrane transport proteins that have been studied in detail are multipass transmembrane proteins; that is, their polypeptide chains traverse the lipid bilayer two or more times. By forming a protein-lined pathway across the membrane, these proteins enable specific hydrophilic solutes to cross the membrane without coming into direct contact with the hydrophobic interior of the lipid bilayer.

Transporters and channels are the two major classes of membrane transport proteins (Figure 11–3). Transporters (sometimes called carriers or permeases) bind the specific solute to be transported and undergo a series of conformational changes that alternately expose solute-binding sites on one side of the membrane and then on the other side to transfer the solute across it.

Channels, by contrast, interact much more transiently with the solute to be transported. When opened by conformational changes, channels form continuous pores that extend across the lipid bilayer. The pores allow specific solutes (such as inorganic ions of appropriate size and charge, and in some cases small molecules, including water, glycerol, and ammonia) to pass through them and thereby cross the membrane. Because no stepwise conformational changes are required once a channel is opened, it is not surprising that transport through channels occurs at a much faster rate than transport mediated by transporters. Although water can slowly diffuse across synthetic lipid bilayers, cells use dedicated channel proteins (called water channels, or aquaporins) that greatly increase the permeability of their membranes to water, as we discuss later.

All channels and some transporters allow solutes to cross the membrane only passively (“downhill”), a process called passive transport. In the case of transport of a single uncharged molecule, the difference in the concentration on the two sides of the membrane—its concentration gradient—drives passive transport and determines its direction (Figure 11–4A). If the solute carries a net charge, however, both its concentration gradient and the electrical potential difference across the membrane, the membrane potential, influence its transport. The concentration gradient and the electrical gradient combine to form a net driving force, the electrochemical gradient, for each charged solute (Figure 11–4B). We discuss electrochemical gradients in more detail later and in Chapter 14. In fact, almost all plasma membranes have an electrical potential difference (that is, a voltage) across them, with the inside usually negative with respect to the outside.

In addition to passive transport, cells need to be able to actively pump certain solutes across the membrane “uphill,” against their electrochemical gradients. Such active transport is mediated by transporters whose pumping activity is directional because it is tightly coupled to a source of metabolic energy, such as an ion gradient or ATP hydrolysis, as discussed later. Transmembrane movement of small molecules mediated by transporters can be either active or passive, whereas that mediated by channels is always passive (see Figure 11–4A).

Lipid bilayers are virtually impermeable to most polar molecules. To transport small water-soluble molecules into or out of cells or intracellular membraneenclosed compartments, cell membranes contain various membrane transport proteins, each of which is responsible for transferring a particular solute or class of solutes across the membrane. There are two types of membrane transport proteins—transporters and channels. Both form protein pathways across the lipid bilayer. Whereas transmembrane movement mediated by transporters can be either active or passive, solute flow through channel proteins is always passive. Ion transport across the membrane is influenced by the ion’s concentration gradient and the membrane potential; that is, its electrochemical gradient.

Because cells are mostly water (typically ∼70% by weight), water movement across cell membranes is fundamentally important for life. Cells also contain a high concentration of solutes, including numerous negatively charged organic molecules that are confined inside the cell (the so-called fixed anions) and their accompanying cations that are required for charge balance. This creates an osmotic gradient, which mostly is balanced by an opposite osmotic gradient due to a high concentration of inorganic ions—chiefly Na+ and Cl–—in the extracellular fluid. The small remaining osmotic force tends to “pull” water into the cell, causing it to swell until the forces are balanced.

Because all biological membranes are moderately permeable to water (see Figure 11–2), cell volume equilibrates in minutes or less in response to an osmotic gradient. For most animal cells, however, osmosis has only a minor role in regulating cell volume. This is because most of the cytoplasm is in a gel-like state and resists large changes in its volume in response to changes in osmolarity.

In addition to the direct diffusion of water across the lipid bilayer, some prokaryotic and eukaryotic cells have water channels, or aquaporins, embedded in their plasma membranes to allow water to move more rapidly. Aquaporins are particularly abundant in animal cells that must transport water at high rates, such as the epithelial cells of the kidney or exocrine cells that must transport or secrete large volumes of fluids (Figure 11–19). Water flow is highly regulated in these tissues. In the kidney, hormones such as antidiuretic hormone (vasopressin) regulate the concentration of aquaporin in the plasma membrane.

Aquaporins must solve a problem that is opposite to that facing ion channels. To avoid disrupting ion gradients across membranes, they have to allow the rapid passage of water molecules while completely blocking the passage of ions. The three-dimensional structure of an aquaporin reveals how it achieves this remarkable selectivity. The channels have a narrow pore that allows water molecules to traverse the membrane in single file, following the path of carbonyl oxygens that line one side of the pore (Figure 11–20A and B). Hydrophobic amino acids line the other side of the pore.

The pore is too narrow for any hydrated ion to enter, and the energy cost of dehydrating an ion would be enormous because the hydrophobic wall of the pore cannot interact with a dehydrated ion to compensate for the loss of water. This design readily explains why the aquaporins cannot conduct K+, Na+, Ca2+, or Cl– ions. These channels are also impermeable to H+, which is mainly present in cells as H3O+. These hydronium ions diffuse through water extremely rapidly, using a molecular relay mechanism that requires the making and breaking of hydrogen bonds between adjacent water molecules (Figure 11–20C).

Aquaporins contain two strategically placed asparagines, which bind to the oxygen atom of the central water molecule in the line of water molecules traversing the pore, imposing a bipolarity on the entire single-file column of water molecules (Figure 11–20C and D). Because both valences of this central oxygen are unavailable for hydrogen-bonding, the central water molecule cannot participate in an H+ relay. This makes it impossible for the “making and breaking” sequence of hydrogen bonds (shown in Figure 11–20C) to get past the central asparaginebonded water molecule, and the pore is therefore impermeable to H+.

In contrast, most channels in the plasma membrane of animal and plant cells that connect the cytosol to the cell exterior necessarily have narrow, highly selective pores that can open and close rapidly. Because these proteins are concerned specifically with inorganic ion transport, they are referred to as ion channels. For transport efficiency, ion channels have an advantage over transporters, in that they can pass up to 100 million ions through one open channel each second—a rate 105 times greater than even the fastest transporter.

As discussed earlier, however, channels cannot be coupled to an energy source to perform active transport, so the conductance they mediate is always passive (downhill). Thus, the function of ion channels is to allow specific inorganic ions—primarily Na+, K+, Ca2+, or Cl–—to diffuse rapidly down their electrochemical gradients across the lipid bilayer. In this section, we will see that the ability to control ion fluxes through these channels is essential for many cell functions.

Two important properties distinguish ion channels from aqueous pores. First, they show ion selectivity, permitting some inorganic ions to pass, but not others. This suggests that their pores must be narrow enough in places to force permeating ions into intimate contact with the walls of the channel so that only ions of appropriate size and charge can pass. In some cases permeating ions have to shed most or all of their associated water molecules to pass, whereas in other cases hydrated or partially hydrated ions pass through the channel.

Thus, as the ion concentration increases, the flux of the ion through a channel increases proportionally but then levels off (saturates) at a maximum rate. The second important distinction between ion channels and aqueous pores is that ion channels are not continually open. Instead, they are gated, which allows them to open briefly and then close again. Moreover, with prolonged (chemical or electrical) stimulation, most ion channels go into a closed “desensitized,” or “inactivated,” state, in which they are refractory to further opening until the stimulus has been removed, as we discuss later.

In most cases, the gate opens in response to a specific stimulus. As shown in Figure 11–22, the main types of stimuli that are known to cause ion channels to open are a change in the voltage across the membrane (voltage-gated channels), a mechanical stress (mechanically gated channels), or the binding of a ligand (ligand-gated channels). The ligand can be either an extracellular mediator—specifically, a neurotransmitter (transmitter-gated channels)—or an intracellular mediator such as an ion (ion-gated channels) or a nucleotide (nucleotidegated channels).

More than 200 types of ion channels have been identified thus far, and new ones are still being discovered, each characterized by the ions it conducts, the mechanism by which it is gated, and its abundance and localization in the cell and in specific cells. Ion channels are responsible for the electrical excitability of muscle cells, and they mediate most forms of electrical signaling in the nervous system. A single neuron typically contains 10 or more kinds of ion channels, located in different domains of its plasma membrane. But ion channels are not restricted to electrically excitable cells.

Scientists were puzzled by the remarkable ability of ion channels to combine exquisite ion selectivity with a high conductance. K+ leak channels, for example, conduct K+ 10,000-fold faster than Na+, yet the two ions are both featureless spheres and have similar radii (0.133 nm and 0.095 nm, respectively).

We cannot explain the normal K+ selectivity by pore size alone, because Na+ is smaller than K+. Moreover, its high conductance rate is incompatible with the channel having selective, high-affinity K+-binding sites, as the binding of K+ ions to such sites would greatly slow their passage.

The channel is made from four identical transmembrane subunits, which together form a central pore through the membrane. Each subunit contributes two transmembrane α helices, which are tilted outward in the membrane and together form a cone, with its wide end facing the outside of the cell where K+ ions exit from the channel (Figure 11–24). The polypeptide chain that connects the two transmembrane helices forms a short α helix (the pore helix) and a crucial loop that protrudes into the wide section of the cone to form the selectivity filter. The selectivity loops from the four subunits form a short, rigid, narrow pore, which is lined by the carbonyl oxygen atoms of their polypeptide backbones.

The structure of the selectivity filter explains the ion selectivity of the channel. A K+ ion must lose almost all of its bound water molecules to enter the filter, where it interacts instead with the carbonyl oxygens lining the filter; the oxygens are rigidly spaced at the exact distance to accommodate a K+ ion. An Na+ ion, in contrast, cannot enter the filter because the carbonyl oxygens are too far away from the smaller Na+ ion to compensate for the high energy expense associated with the loss of water molecules required for entry (Figure 11–25).

Structural studies of K+ channels and other ion channels have also indicated some general principles of how these channels open and close. The gating involves movement of the helices in the membrane so that they either obstruct or open the path for ion movement.

The structure of a closed K+ channel shows that by tilting the inner helices, the pore constricts like a diaphragm at its cytosolic end (Figure 11–26). Bulky hydrophobic amino acid side chains block the small opening that remains, preventing the entry of ions. Many other ion channels operate on similar principles: the channel’s pore helices are allosterically coupled to sensor domains that in response, say, to ligand binding or altered membrane potential bring about conformational change in the ion-conducting pathway, either opening it or blocking it off.

Ion channels that are permeable to K+ are found in the plasma membrane of almost all cells. An important subset of K+ channels opens even in an unstimulated or “resting” cell, and hence these are called K1 leak channels. Although this term applies to many different K+ channels, depending on the cell type, they serve a common purpose: by making the plasma membrane much more permeable to K+ than to other ions, they have a crucial role in maintaining the membrane potential across all plasma membranes, as we discuss next.

A membrane potential arises when there is a difference in the electrical charge on the two sides of a membrane because of a minute excess of positive ions over negative ones on one side and a minute deficit on the other side. Such charge differences can result both from active electrogenic pumping (see p. 648) and from passive ion diffusion in channels. As we discuss in Chapter 14, electrogenic H+ pumps in the mitochondrial inner membrane generate most of the membrane potential across this membrane. Electrogenic pumps also generate most of the electrical potential across the plasma membrane of plants and fungi. In typical animal cells, however, passive ion movements make the largest contribution to the electrical potential across the plasma membrane.

As explained earlier, because of the action of the Na+-K+ pump, there is little Na+ inside the cell, and other intracellular inorganic cations have to be plentiful enough to balance the charge carried by the cell’s fixed anions—the negatively charged organic molecules that are confined inside the cell. This balancing role is performed largely by K+, which is actively pumped into the cell by the Na+-K+ pump and can also move in or out through the K1 leak channels in the plasma membrane. Because of the presence of these channels, K+ comes almost to equilibrium, where an electrical force exerted by an excess of negative charges attracting K+ into the cell balances the tendency of K+ to leak out down its concentration gradient.

Suppose that initially there is no voltage gradient across the plasma membrane (the membrane potential is zero), but the concentration of K+ is high inside the cell and low outside. K+ will tend to leave the cell through the K+ leak channels, driven solely by its concentration gradient. As K+ begins to move out, each ion leaves behind an unbalanced negative charge, thereby creating an electrical field, or membrane potential, which will tend to oppose the further efflux of K+. The net efflux of K+ halts when the membrane potential reaches a value at which this electrical driving force on K+ exactly balances the effect of its concentration gradient; that is, when the electrochemical gradient for K+ is zero.

The equilibrium condition, in which there is no net flow of ions across the plasma membrane, defines the resting membrane potential for this idealized cell. A simple but very important formula, the Nernst equation, quantifies the equilibrium condition and, as explained in Panel 11–1 (p. 656), makes it possible to calculate the theoretical resting membrane potential if we know the ratio of internal and external ion concentrations. As the plasma membrane of a real cell is not exclusively permeable to K+, however, the actual resting membrane potential is usually not exactly equal to that predicted by the Nernst equation for K+.

The flow of any inorganic ion through a membrane channel is driven by the electrochemical gradient for that ion. This gradient represents the combination of two influences: the voltage gradient and the concentration gradient of the ion across the membrane. When these two influences just balance each other, the electrochemical gradient for the ion is zero, and there is no net flow of the ion through the channel. The voltage gradient (membrane potential) at which this equilibrium is reached is called the equilibrium potential for the ion. It can be calculated from an equation that will be derived below, called the Nernst equation.

When there is a voltage gradient across the membrane, the ions responsible for it—the excess positive ions on one side and the excess negative ions on the other—are concentrated in thin layers on either side of the membrane because of the attraction between positive and negative electric charges. The number of ions that go to form the layer of charge adjacent to the membrane is minute compared with the total number inside the cell. For example, the movement of 6000 Na+ ions across 1 µm2 of membrane will carry sufficient charge to shift the membrane potential by about 100 mV.

Moreover, these movements of charge are generally rapid, taking only a few milliseconds or less. Consider the change in the membrane potential in a real cell after the sudden inactivation of the Na+-K+ pump. A slight drop in the membrane potential may occur immediately. This is because the pump is electrogenic and, when active, makes a small direct contribution to the membrane potential by pumping out three Na+ for every two K+ that it pumps in (see Figure 11–15).

However, switching off the pump does not abolish the major component of the resting potential, which is generated by the K+ equilibrium mechanism just described. This component of the membrane potential persists as long as the Na+ concentration inside the cell stays low and the K+ ion concentration high—typically for many minutes. But the plasma membrane is somewhat permeable to all small ions, including Na+. Therefore, without the Na+-K+ pump, the ion gradients set up by the pump will eventually run down, and the membrane potential established by diffusion through the K+ leak channels will fall as well. As Na+ enters, the cell eventually comes to a new resting state where Na+, K+, and Cl– are all at equilibrium across the membrane.

A small flow of inorganic ions through an ion channel carries sufficient charge to cause a large change in the membrane potential. The ions that give rise to the membrane potential lie in a thin (<1 nm) surface layer close to the membrane, held there by their electrical attraction to their oppositely charged counterparts (counterions) on the other side of the membrane. For a typical cell, 1 microcoulomb of charge (6 × 1012 monovalent ions) per square centimeter of membrane, transferred from one side of the membrane to the other side, changes the membrane potential by roughly 1 V. This means, for example, that in a spherical cell of diameter 10 μm, the number of K+ ions that have to flow out to alter the membrane potential by 100 mV is only about 1/100,000 of the total number of K+ ions in the cytosol. This amount is so minute that the intracellular K+ concentration remains virtually unchanged.

The resting potential of an animal cell varies between –20 mV and –120 mV, depending on the organism and cell type. Although the K+ gradient always has a major influence on this potential, the gradients of other ions (and the disequilibrating effects of ion pumps) also have a significant effect: the more permeable the membrane for a given ion, the more strongly the membrane potential tends to be driven toward the equilibrium value for that ion. Consequently, changes in a membrane’s relative permeability to different ions can cause significant changes in the membrane potential. This is one of the key principles relating the electrical excitability of cells to the activities of ion channels.

All organisms, from single-cell bacteria to multicellular animals and plants, must sense and respond to mechanical forces both in their external environment (such as sound, touch, pressure, shear forces, and gravity) and in their internal environment (such as osmotic pressure and membrane bending). Numerous proteins are known to be capable of responding to such mechanical forces, and a large subset of those proteins has been identified as possible mechanosensitive channels, but very few of the candidate proteins have been shown directly to be mechanically activated ion channels.

One reason for this dearth in our knowledge is that most such channels are extremely rare. Auditory hair cells in the human cochlea, for example, contain extraordinarily sensitive mechanically gated ion channels, but each of the approximately 15,000 individual hair cells is thought to have a total of only 50–100 of them (Movie 11.9 and Movie 11.10).

Three identical copies of either of these proteins can assemble into a mechanosensitive Piezo ion channel that lies in the plasma membranes of numerous types of animal cells (Figure 11–27). Each of the three subunits of a Piezo channel contains a large arm built from 36 transmembrane helices. The arms extend outward from a central hub that contains the channel’s ion-conducting pore. The particular packing of the wing helices deforms the membrane into a dome. In this resting state, the Piezo channel is closed. When the membrane becomes stretched by mechanically pushing on the plasma membrane elsewhere, the resulting membrane tension pulls the deformation flat, which opens the channel’s central pore.

Mechanical gating has been demonstrated using biophysical techniques in which force is exerted on pure lipid bilayers containing the bacterial mechanosensitive channels; for example, by applying suction with a micropipette. Such measurements demonstrate that the cell has several different channels that open at different levels of pressure. The mechanosensitive channel of small conductance, called the MscS channel, opens at low and moderate pressures (Figure 11–28). It is composed of seven identical subunits, which in the open state form a pore about 1.3 nm in diameter—just big enough to pass ions and small molecules.

Patch-clamp recording, developed in the 1970s and 1980s, revolutionized the study of ion channels and made it possible to examine transport through a single channel in a small patch of membrane covering the mouth of a micropipette (Figure 11–36). With this simple but power ful technique, one can study the detailed properties of ion channels in all sorts of cell types. This work led to the discovery that even cells that are not electrically excitable usually have a variety of ion channels in their plasma membrane. Many of these cells, such as yeasts, are too small to be investigated by the traditional electrophysiologist’s method of impalement with an intracellular microelectrode.

Patch-clamp recording indicates that individual ion channels open in an allor-nothing fashion. For example, a voltage-gated Na+ channel opens and closes stochastically with a voltage-dependent probability, but when open, the channel always has the same large conductance, allowing more than 1000 ions to pass per millisecond (Figure 11–37). Therefore, the aggregate current crossing the membrane of an entire cell does not indicate the degree to which a typical individual channel is open but rather the total number of channels in its membrane that are open at any one time.

Some simple physical principles allow us to refine our understanding of voltagegating from the perspective of a single Na+ channel. The interior of the resting neuron or muscle cell is at an electrical potential about 40–100 mV more negative than the external medium. Although this potential difference seems small, it exists across a plasma membrane only about 5 nm thick, so that the resulting voltage gradient is about 100,000 V/cm. Charged proteins in the membrane such as Na+ channels are thus subjected to a very large electrical field that can profoundly affect their conformation.

The process by which a transporter transfers a solute molecule across the lipid bilayer resembles an enzyme–substrate reaction, and in many ways transporters behave like enzymes. In contrast to ordinary enzyme–substrate reactions, however, the transporter does not modify the transported solute but instead delivers it unchanged to the other side of the membrane.

Each type of transporter has one or more specific binding sites for its solute (substrate). It transfers the solute across the lipid bilayer by undergoing reversible conformational changes that alternately expose the solute-binding site first on one side of the membrane and then on the other side—but never on both sides at the same time. The transition occurs through an intermediate state in which the solute is inaccessible, or occluded, from either side of the membrane (Figure 11–5).

When the transporter is saturated (that is, when all solute-binding sites are occupied), the rate of transport is maximal. This rate, referred to as Vmax (V for velocity), is characteristic of the specific carrier. Vmax is the maximal rate at which the carrier can flip between its conformational states. In addition, each transporter has a characteristic affinity for its solute, reflected in the Km of the reaction, which is equal to the concentration of solute when the transport rate is half its maximum value (Figure 11–6).

As with enzymes, the binding of solute can be blocked by either competitive inhibitors (which compete for the same binding site and may or may not be transported) or noncompetitive inhibitors (which bind elsewhere and alter the structure of the transporter). As we discuss shortly, conceptually it requires only a relatively minor modification of the mechanism shown in Figure 11–5 to link a transporter to a source of energy in order to pump a solute uphill against its electrochemical gradient. Cells carry out such active transport in three main ways (Figure 11–7): 1. Coupled transporters harness the energy stored in concentration gradients to couple the uphill transport of one solute across the membrane to the downhill transport of another.

3. Light- or redox-driven pumps, which are known in bacteria, archaea, mitochondria, and chloroplasts, couple uphill transport to an input of energy from light, as with bacteriorhodopsin and photosystem II (discussed in Chapters 10 and 14, respectively), or from a redox reaction, as with cytochrome c oxidase (discussed in Chapter 14). Comparisons of amino acid sequences and three-dimensional structures suggest that, in many cases, there are strong similarities in structure between transporters that mediate active transport and those that mediate passive transport. Some bacterial transporters, for example, that use the energy stored in the H+ gradient across the plasma membrane to drive the active uptake of various sugars are structurally similar to the transporters that mediate passive glucose transport into most animal cells. There is thus a clear evolutionary relationship between various transporters.

Given the cell’s essential need to transport small metabolites across membranes, it comes as no surprise that the superfamily of transporters is a large and ancient one. We begin our discussion of active membrane transport by considering a class of coupled transporters that are driven by ion-concentration gradients. These proteins have a crucial role in the transport of small metabolites across membranes in all cells. We then discuss ATP-driven pumps, including the Na+-K+ pump that is found in the plasma membrane of most animal cells. Examples of the third class of active transport—light- or redox-driven pumps—are discussed in Chapter 14.

Some transporters simply facilitate the passive movement of a single solute from one side of the membrane to the other at a rate determined by their Vmax and Km; they are called uniporters. Others function as coupled transporters, in which the transfer of one solute strictly depends on the transport of a second. Coupled transport involves either the intimately coupled transfer of a second solute in the same direction, performed by symporters (also called co-transporters), or the transfer of a second solute in the opposite direction, performed by antiporters (also called exchangers) (Figure 11–8).

The tight coupling between the transfer of two solutes allows the coupled transporters to harvest the energy stored in the electrochemical gradient of one solute, typically an inorganic ion, to transport the other. In this way, the free energy released during the movement of an inorganic ion or H+ down an electrochemical gradient is used as the driving force to pump other solutes uphill, against their electrochemical gradient. This strategy can work in either direction; some coupled transporters function as symporters, others as antiporters.

In the plasma membrane of animal cells, Na+ is the usual co-transported ion because its electrochemical gradient provides a large driving force for the active transport of a second molecule. Such ion-driven coupled transporters are said to mediate secondary active transport. The Na+ that enters the cell during coupled transport is subsequently pumped out by an ATP-driven Na+-K+ pump in the plasma membrane (as we discuss later), which, by exchanging K+ for Na+, maintains the Na+ gradient, indirectly driving the coupled transport.

Such ATP-driven pumps are therefore said to mediate primary active transport because in these the free energy of ATP hydrolysis is used to directly drive the transport of a solute against its electrochemical gradient. The energy stored in the gradient is then used to fuel the secondary active transport processes.

Because the Na+ tends to move into the cell down its electrochemical gradient, the sugar or amino acid is, in a sense, “dragged” into the cell with it. The greater the electrochemical gradient for Na+, the more solute is transported into the cell (Figure 11–9). Neurotransmitters (released by nerve cells to signal at synapses—as we discuss later) are taken up again by Na+ symporters after their release. This both terminates their signaling to postsynaptic cells and recycles them for reuse. These neurotransmitter transporters are important drug targets: stimulants, such as cocaine and antidepressants, inhibit them and thereby prolong signaling by the neurotransmitters because they are not cleared efficiently.

Despite their great variety, transporters share structural features that can explain how they function and how they evolved. Transporters are typically built from bundles of 10 or more α helices that span the membrane. Solute- and ionbinding sites are located midway through the membrane, where some helices are broken or distorted and amino acid side chains and polypeptide backbone atoms form ion- and solute-binding sites. In the inward-open and outward-open conformations, these binding sites are accessible by passageways from one side of the membrane but not the other.

In switching between the two conformations, the transporter protein transiently adopts an occluded conformation, in which both passageways are closed; this prevents the driving ion and the transported solute from crossing the membrane unaccompanied, which would deplete the cell’s energy store to no purpose. Because only transporters with both types of binding sites appropriately filled change their conformation, tight coupling between ion and solute transport is ensured. Like enzymes, transporters can work in the reverse direction if ion and solute gradients are appropriately adjusted experimentally. This chemical symmetry is mirrored in their physical structure. Protein structural analyses have revealed that many transporters are built from inverted repeats: the packing of the transmembrane α helices in one half of the helix bundle is structurally similar to the packing in the other half, but the two halves are inverted in the membrane relative to each other.

It is thought that the two halves evolved by gene duplication of a smaller ancestor protein. Some other types of important membrane transport proteins are also built from inverted repeats. Examples even include channel proteins such as the aquaporin water channel (discussed later) and the Sec61 channel through which nascent polypeptides move into the endoplasmic reticulum (discussed in Chapter 12). It is thought that these channels evolved from coupled transporters in which the gating functions were lost, allowing them to open toward both sides of the membrane simultaneously to provide a continuous path across the membrane.

In bacteria, yeasts, and plants, as well as in many membrane-enclosed organelles of animal cells, most ion-driven active transport systems depend on H+ rather than Na+ gradients, reflecting the predominance of H+ pumps in these membranes. An electrochemical H+ gradient across the bacterial plasma membrane, for example, drives the inward active transport of many sugars and amino acids.

Most proteins operate optimally at a particular pH. Lysosomal enzymes, for example, function best at the low pH (∼5) found in lysosomes, whereas cytosolic enzymes function best at the close-to-neutral pH (∼7.2) found in the cytosol. It is therefore crucial that cells control the pH of their intracellular compartments. Most cells have one or more types of Na+-driven antiporters in their plasma membrane that help to maintain the cytosolic pH at about 7.2. These transporters use the energy stored in the Na+ gradient to pump out excess H+, which either leaks in or is produced in the cell by acid-forming reactions. Two mechanisms are used: either H+ is directly transported out of the cell or HCO3– is brought into the cell to neutralize H+ in the cytosol (according to the reaction HCO3– + H+ → H2O + CO2).

The Na+-driven Cl––HCO3– exchanger is twice as effective as the Na+–H+ exchanger: it pumps out one H+ and neutralizes another for each Na+ that enters the cell. If HCO3– is available, as is usually the case, this antiporter is the most important transporter regulating the cytosolic pH. The pH inside the cell regulates both exchangers; when the pH in the cytosol falls, both exchangers sense the change and increase their activity.

An Na+-independent Cl––HCO3– exchanger in the membrane of red blood cells (called band 3 protein—see Figure 10–38) facilitates the quick discharge of CO2 (as HCO3–) as the cells pass through capillaries in the lung. The intracellular pH is not entirely regulated by transporters in the plasma membrane: ATP-driven H+ pumps are used to control the pH of many intracellular compartments. As discussed in Chapter 13, H+ pumps maintain the low pH in lysosomes, as well as in endosomes and secretory vesicles. These H+ pumps use the energy of ATP hydrolysis to pump H+ into these acidic organelles from the cytosol.

An advantage of using electrochemical H+ gradients to power intracellular transport events is that they can dissipate and regenerate quickly, thus affording more opportunity to switch transport reactions on and off. To create an electrochemical H+ gradient of similar energy to that of the Na+ gradient at the plasma membrane requires the movement of far fewer H+. This is because the H+ concentration is many orders of magnitude smaller (0.1 μM at pH 7) than that of Na+ and K+ (∼100 mM).

In epithelial cells, such as those that absorb nutrients from the gut, transporters are distributed nonuniformly between the apical and basolateral plasma membranes and thereby contribute to the transcellular transport of absorbed solutes. By the actions of the transporters in these cells, solutes are moved across the epithelial-cell layer into the extracellular fluid from where they pass into the blood.

As shown in Figure 11–11, Na+-linked symporters located in the apical (absorptive) domain of the plasma membrane actively transport nutrients into the cell, building up substantial concentration gradients for these solutes across the plasma membrane. Uniporters in the basal and lateral (basolateral) domain allow the nutrients to leave the cell passively down these concentration gradients to enter the bloodstream for use in the rest of the body.

In many of these epithelial cells, the plasma membrane area is greatly increased by the formation of thousands of microvilli, which extend as thin, fingerlike projections from the apical surface of each cell. Such microvilli can increase the total absorptive area of a cell as much as 25-fold, thereby enhancing its transport capabilities. As we have seen, ion gradients have a crucial role in driving many essential transport processes in cells. Ion pumps that use the energy of ATP hydrolysis establish and maintain these gradients, as we discuss next.

ATP-driven pumps are often called transport ATPases because they hydrolyze ATP to ADP and phosphate and use the energy released to pump ions or other solutes across a membrane. There are three principal classes of ATP-driven pumps (Figure 11–12), and representatives of each are found in all prokaryotic and eukaryotic cells.

1. P-type pumps are structurally and functionally related multipass transmembrane proteins. They are called “P-type” because they phosphorylate themselves during the pumping cycle. This class includes many of the ion pumps that are responsible for setting up and maintaining gradients of Na+, K+, H+, and Ca2+ across cell membranes.

3. V-type pumps are turbine-like protein machines constructed from multiple different subunits. The V-type proton pump transfers H+ into organelles, such as lysosomes, synaptic vesicles, and plant or yeast vacuoles (V = vacuolar), to acidify the interior of these organelles (see Figures 13–46 and 13–47).

Structurally related to the V-type pumps is a distinct subclass of F-type ATPases, more commonly called ATP synthases because they normally work in reverse: instead of using ATP hydrolysis to drive H+ transport, they use the H+ gradient across the membrane to drive the synthesis of ATP from ADP and phosphate (see Figure 14–31).

ATP synthases are found in the plasma membrane of bacteria, the inner membrane of mitochondria, and the thylakoid membrane of chloroplasts. The H+ gradient is generated either during the electron-transport steps of oxidative phosphorylation (in aerobic bacteria and mitochondria), during photosynthesis (in chloroplasts), or by the light-driven H+ pump (bacteriorhodopsin) in Halobacterium. We discuss some of these proteins in detail in Chapter 14. For the remainder of this section, we focus on P-type pumps and ABC transporters.

Eukaryotic cells maintain very low concentrations of free Ca2+ in their cytosol (∼10–7 M) in the face of a very much higher extracellular Ca2+ concentration (∼10–3 M). Therefore, even a small influx of Ca2+ significantly increases the concentration of free Ca2+ in the cytosol, and the flow of Ca2+ down its steep concentration gradient in response to extracellular signals is one means of transmitting these signals rapidly across the plasma membrane (discussed in Chapter 15).

It is thus important that the cell maintains a steep Ca2+ gradient across its plasma membrane. Ca2+ transporters that actively pump Ca2+ out of the cytosol help maintain the gradient. One of these is a P-type Ca2+ ATPase; the other is an antiporter (called an Na1–Ca21 exchanger) that is driven by the Na+ electrochemical gradient (discussed in Chapter 15).

The Ca21 pump, or Ca21 ATPase, in the sarcoplasmic reticulum (SR) membrane of skeletal muscle cells is a well-understood P-type transport ATPase. The SR is a specialized type of endoplasmic reticulum that forms a network of tubular sacs in the muscle-cell cytoplasm, and it serves as an intracellular store of Ca2+. When an action potential depolarizes the muscle-cell plasma membrane, Ca2+ is released into the cytosol from the SR through Ca21-release channels, stimulating the muscle to contract (discussed in Chapters 15 and 16).

The Ca2+ pump, which accounts for about 90% of the membrane protein of the SR, moves Ca2+ from the cytosol back into the SR. The endoplasmic reticulum of non-muscle cells also stores Ca2+ using a closely homologous Ca2+ pump and Ca2+-release channels. Enzymatic studies and analyses of the three-dimensional structures of transport intermediates of the SR Ca2+ pump and related pumps have revealed the molecular mechanism of P-type transport ATPases in great detail. They all have similar structures, containing 10 transmembrane α helices connected to three cytosolic domains (Figure 11–13).

In the Ca2+ pump, amino acid side chains protruding from the transmembrane helices form two centrally positioned binding sites for Ca2+. As shown in Figure 11–14, in the pump’s ATP-bound nonphosphorylated state, these binding sites are accessible only from the cytosolic side of the SR membrane. Ca2+ binding triggers a series of conformational changes that close the passageway to the cytosol and activate a phosphotransfer reaction in which the terminal phosphate of the ATP is transferred to an aspartate that is conserved among all P-type ATPases.

The ADP then dissociates and is replaced with a fresh ATP, causing another conformational change that opens a passageway to the SR lumen through which the two Ca2+ ions exit. They are replaced by two H+ ions and water molecules that stabilize the empty Ca2+-binding sites and close the passageway to the SR lumen, switching the pump to the occluded conformation. Hydrolysis of the labile phosphoryl–aspartate bond opens the passageway to the cytosol.

The concentration of K+ is typically 10–30 times higher inside cells than outside, whereas the reverse is true of Na+ (see Table 11–1, p. 638). An Na1-K1 pump, or Na1-K1 ATPase, found in the plasma membrane of virtually all animal cells, maintains these concentration differences.

Like the Ca2+ pump, the Na+-K+ pump belongs to the family of P-type ATPases and operates as an ATP-driven antiporter, actively pumping Na+ out of the cell against its steep electrochemical gradient and pumping K+ in (Figure 11–15). We mentioned earlier that the Na+ gradient produced by the Na+-K+ pump drives the transport of most nutrients into animal cells and also has a crucial role in regulating cytosolic pH.

Because the Na+-K+ pump drives three positively charged ions out of the cell for every two it pumps in, it is electrogenic: it drives a net electric current across the membrane, tending to create an electrical potential, with the cell’s inside being negative relative to the outside. This electrogenic effect of the pump, however, seldom directly contributes more than 10% to the membrane potential. The remaining 90%, as we discuss later, depends only indirectly on the Na+-K+ pump.

The last type of transport ATPase that we discuss here is the family of ABC transporters, so named because each member contains two highly conserved ATPase domains, or ATP-binding “cassettes,” on the cytosolic side of the membrane. ATP binding brings together the two ATPase domains, and ATP hydrolysis leads to their dissociation (Figure 11–16). These movements of the cytosolic domains are transmitted to the transmembrane segments, driving cycles of conformational changes that alternately expose solute-binding sites on one side of the membrane and then on the other side, as we have seen for other transporters.

In this way, ABC transporters harvest the energy released upon ATP binding and hydrolysis to drive transport of solutes across the bilayer. The transport is directional toward the inside or toward the outside, depending on the particular conformational change in the solute-binding site that is linked to ATP hydrolysis (see Figure 11–16). ABC transporters constitute the largest family of membrane transport proteins and are of great clinical importance. The first of these proteins to be characterized was found in bacteria. We have already mentioned that the plasma membranes of all bacteria contain transporters that use the H+ gradient across the membrane to actively transport a variety of nutrients into the cell. In addition, bacteria use ABC transporters to import certain small molecules.

In E. coli, 78 genes (an amazing 5% of the bacterium’s genes) encode ABC transporters, and animal genomes encode an even larger number. Although each transporter is thought to be specific for a particular molecule or class of molecules, the variety of substrates transported by this superfamily is great and includes inorganic ions, amino acids, monosaccharides and polysaccharides, peptides, lipids, drugs, and, in some cases, even proteins that can be larger than the transporter itself.

The first eukaryotic ABC transporters identified were discovered because they pump hydrophobic drugs out of the cytosol. One of these transporters is the multidrug resistance (MDR) protein, also called P-glycoprotein. It is present at elevated levels in many human cancer cells and makes the cells simultaneously resistant to a variety of chemically unrelated cytotoxic drugs that are widely used in cancer chemotherapy.

Treatment with any one of these drugs can result in the selective survival and overgrowth of those cancer cells that express an especially large amount of the MDR transporter. These cells pump drugs out of the cell very efficiently and are therefore relatively resistant to the drugs’ toxic effects (Movie 11.5). Selection for cancer cells with resistance to one drug can thereby lead to resistance to a wide variety of anticancer drugs. Some studies indicate that up to 40% of human cancers develop multidrug resistance, making it a major hurdle in the battle against cancer.

A related and equally sinister phenomenon occurs in the protist Plasmodium falciparum, which causes malaria. More than 200 million people are infected worldwide with this parasite, which remains a major cause of human death, killing almost a million people every year. The development of resistance to the antimalarial drug chloroquine has hampered the control of malaria. The resistant P. falciparum have amplified a gene encoding an ABC transporter that pumps out the chloroquine.

In most vertebrate cells, an ABC transporter in the endoplasmic reticulum (ER) membrane (named transporter associated with antigen processing, or TAP transporter) actively pumps a wide variety of peptides from the cytosol into the ER lumen. These peptides are produced by protein degradation in proteasomes (discussed in Chapter 6). They are carried from the ER to the cell surface, where they are displayed for scrutiny by cytotoxic T lymphocytes, which kill the cell if the peptides are derived from a virus or other microorganism lurking in the cytosol of an infected cell (discussed in Chapter 24).

Yet another member of the ABC transporter family is the cystic fibrosis transmembrane conductance regulator (CFTR) protein, which was discovered through studies of the common genetic disease cystic fibrosis. This disease is caused by a mutation in the gene encoding CFTR, a Cl– transport protein in the plasma membrane of epithelial cells. CFTR regulates ion concentrations in the extracellular fluid, especially in the lung. One in 27 Caucasians carries a gene encoding a mutant form of this protein; in 1 in 2900, both copies of the gene are mutated, causing the disease.

In contrast to other ABC transporters, ATP binding and hydrolysis in the CFTR protein do not drive the transport process. Instead, they control the opening and closing of a continuous channel, which provides a passive conduit for Cl– to move down its electrochemical gradient. Thus, some ABC proteins can function as transporters and others as gated channels. Even though we treat transporters and channels as distinct in this chapter, the border is not absolute as this example illustrates. Indeed, the structure of some other Cl– channels reveals that they also resemble transporters more than they resemble most other ion channels.

Transporters bind specific solutes and transfer them across the lipid bilayer by undergoing conformational changes that alternately expose the solute-binding site on one side of the membrane and then on the other side. Some transporters move a single solute “downhill,” whereas others can act as pumps to move a solute “uphill” against its electrochemical gradient, by using energy provided by ATP hydrolysis, by a downhill flow of another solute (such as Na1 or H1), or by light to drive the requisite series of conformational changes in an orderly manner.

Transporters belong to a small number of protein families. Each family evolved from a common ancestral protein, and its members all operate by a similar mechanism. The family of P-type transport ATPases, which includes Ca21 and Na1-K1 pumps, is an important example; each of these ATPases sequentially phosphorylates and dephosphorylates itself during the pumping cycle.

The superfamily of ABC transporters is the largest family of membrane transport proteins and is especially important clinically. It includes proteins that are responsible for drug resistance in both cancer cells and cells infected with malaria-causing parasites and for pumping pathogen-derived peptides into the ER for cytotoxic lymphocytes to reorganize on the surface of infected cells, and mutations in an ABC transporter cause cystic fibrosis.

Transport begins when a special coat of proteins is assembled on a region of the cytosolic face of a membrane compartment. The coat is used to collect specific cargo components from the membrane and compartment lumen for delivery to another compartment. The coat, with the help of additional proteins, shapes the membrane into a transport vesicle that buds from the originating compartment. These vesicles selectively dock at the appropriate destination membrane and then fuse with it to deliver their cargo.

Most transport vesicles form from specialized, coated regions of membranes. They bud off as coated vesicles, which have a distinctive cage of proteins covering their cytosolic surface. Before the vesicles fuse with a target membrane, they shed their coat so that the membrane surfaces of the vesicle and destination compartment can interact directly and fuse.

First, an inner coat layer concentrates specific membrane proteins in a specialized patch, which then gives rise to the vesicle membrane. In this way, the inner layer selects the appropriate membrane molecules for transport. Second, an outer coat layer assembles into a curved, basketlike lattice that deforms the membrane patch and thereby shapes the vesicle.

There are four well-characterized types of coated vesicles, distinguished by their major coat proteins: clathrin-coated, COPI-coated, COPII-coated, and retromer-coated (Figure 13–4). Each type is used for different transport steps (Figure 13–5). Clathrin-coated vesicles mediate transport originating from the Golgi apparatus, endosome, and the plasma membrane. COPI-coated and COPII-coated vesicles mediate transport originating from the Golgi cisternae and the ER, respectively.

There is, however, much more variety in coated vesicles and their functions than this short list suggests. As we discuss shortly, there are several types of clathrin-coated vesicles, each specialized for a different transport step, and the COPI-coated and COPII-coated vesicles may be similarly diverse. We discuss clathrin-coated vesicles first, as they provide a good example of how vesicles form.

The major protein component of clathrin-coated vesicles is clathrin, which forms the outer layer of the coat. Clathrin is composed of a large subunit (the heavy chain) and a small subunit (the light chain). Three heavy chains and three light chains assemble into a three-legged structure called a triskelion (Figure 13–6A and B). Clathrin triskelions assemble into a basketlike framework of hexagons and pentagons on the cytosolic surface of membranes. Clathrin assembly induces the formation of coated buds (called coated pits when on the plasma membrane), which eventually pinch off to become clathrin-coated vesicles (Figure 13–7).

Adaptor proteins, another major coat component in clathrin-coated vesicles, form a discrete inner layer of the coat, positioned between the clathrin cage and the cytosolic face of the membrane. They bind to various transmembrane protein cargoes and transmembrane receptors that capture soluble cargo molecules inside the vesicle—so-called cargo receptors. Adaptor proteins also bind to clathrin and recruit it to the membrane surface where it assembles and bends the membrane. In this way, the specific set of transmembrane and soluble cargoes selected by adaptor proteins is packaged into a newly formed clathrin-coated transport vesicle (Figure 13–8).

The assembly of adaptor proteins on the membrane is tightly controlled, in part by the cooperative interaction of the adaptor proteins with the membrane, transmembrane cargoes, and other components of the coat. The adaptor protein AP2 serves as a well-understood example. When it binds to a specific phosphorylated phosphatidylinositol lipid (a phosphoinositide), AP2 acquires a different conformation that exposes binding sites for cargo receptors in the membrane. The simultaneous binding to the cargo receptors and lipid head groups greatly enhances the binding of AP2 to the membrane (Figure 13–9).

Upon binding, AP2 induces membrane curvature, which makes the binding of additional AP2 proteins in its proximity more likely. The cooperative assembly of the AP2 coat layer then is further amplified by clathrin binding, which leads to the formation and budding of a transport vesicle.

There are several types of adaptor proteins. The best characterized, like AP2, have four different protein subunits; others are single-chain proteins. Many of the adaptor proteins bind to phosphoinositides. As we will see next, different types of phosphoinositides are located in different membrane compartments, serving as one of the molecular markers of that compartment’s identity. Each type of adaptor protein is specific for transmembrane cargoes and cargo receptors that share a particular amino acid sequence motif displayed on the cytosolic side of the membrane. Because different adaptor proteins have different specificities for both the type of phosphoinositide and the sequence motif they recognize, each type of adaptor protein directs assembly of a clathrin-coated vesicle only at particular membranes.

Many proteins involved at different steps in vesicle transport contain domains that bind with high specificity to the head groups of particular PIPs, distinguishing one phosphorylated form from another (see Figure 13–10). Local control of the PI and PIP kinases and PIP phosphatases can therefore be used to rapidly control the binding of proteins to a membrane or membrane domain. The production of a particular type of PIP recruits proteins containing matching PIP-binding domains. The PIP-binding proteins then help regulate vesicle formation and other steps in the control of vesicle traffic (Figure 13–11).

Although vesicle-budding is similar at various locations in the cell, each cell membrane poses its own special challenges. The plasma membrane, for example, is comparatively flat and stiff, owing to its cholesterol-rich lipid composition and underlying actin-rich cortex. Thus, the forces generated by clathrin coat assembly alone are not sufficient to shape and pinch off a vesicle from the plasma membrane. Other membrane-bending and force-generating proteins participate at every stage of the process.

Some of these proteins also contain amphiphilic helices that induce membrane curvature after being inserted as wedges into the cytoplasmic leaflet of the membrane. The curved membrane generated by BAR-domain proteins is thought to help AP2 nucleate the formation of a clathrin-coated bud. Other BAR-domain proteins are important in shaping the neck of a budding vesicle, where stabilization of sharp membrane bends is essential. These BAR-domain proteins, together with the clathrin machinery they help nucleate, stimulate the local assembly of actin filaments (Figure 13–13). The growing filaments push on the membrane surrounding the budding vesicle and further help propel it away from the membrane.

As a clathrin-coated bud grows, soluble cytoplasmic proteins, including dynamin, assemble at the neck of the bud and ultimately pinch off the membrane to release the fully formed clathrin-coated vesicle (Figure 13–14). Dynamin contains a phosphoinositide-binding domain, which tethers the protein to the membrane, and a GTPase domain, which regulates the rate at which vesicles pinch off from the membrane. The pinching-off process brings the two noncytosolic leaflets of the membrane at the bud neck into close proximity and seals off the forming vesicle (see Figure 13–2). To perform this task, dynamin assembles in a ring around the neck, then undergoes a conformational change when it hydrolyzes its bound GTP. This constricts the dynamin ring together with the underlying membrane at the bud neck.

Once released from the membrane, the vesicle rapidly loses its clathrin coat because factors that are co-packaged into a clathrin-coated vesicle initiate reactions that lead to coat disassembly. A phosphoinositide phosphatase in the vesicle depletes the phosphoinositide that binds to the adaptor proteins of the coat. In addition, auxilin, another vesicle protein, activates the ATPase of an hsp70 chaperone protein (see Figure 6–80) that uses the energy of ATP hydrolysis to peel off the clathrin coat. The release of the coat, however, must not happen prematurely, so additional control mechanisms must somehow prevent the clathrin from being removed before it has formed a complete vesicle.

The routes that lead inward from the cell surface start with the process of endocytosis, by which cells take up plasma membrane components, fluid, solutes, macromolecules, and particulate substances. Endocytosed cargo includes receptor–ligand complexes, a spectrum of nutrients and their carriers, extracellular matrix components, cell debris, bacteria, viruses, and, in specialized cases, even other cells. Through endocytosis, the cell regulates the composition of its plasma membrane in response to changing extracellular conditions. In endocytosis, the material to be ingested is progressively enclosed by a small portion of the plasma membrane, which first invaginates and then pinches off to form an endocytic vesicle containing the ingested substance or particle.

Most eukaryotic cells constantly form endocytic vesicles, a process called pinocytosis (“cell drinking”); in addition, some specialized cells contain dedicated pathways that take up large particles on demand, a process called phagocytosis (“cell eating”). Endocytic vesicles form at the plasma membrane by multiple mechanisms that differ in both the molecular machinery used and how that machinery is regulated. Once generated at the plasma membrane, most endocytic vesicles fuse with a common receiving compartment, the early endosome, where internalized cargo is sorted: some cargo molecules are returned to the plasma membrane, either directly or via a recycling endosome, and others remain as the early endosome changes into a late endosome by a process termed endosome maturation (Figure 13–50).

Virtually all eukaryotic cells continually ingest portions of their plasma membrane in the form of small pinocytic (endocytic) vesicles. The rate at which plasma membrane is internalized in this process of pinocytosis varies between cell types, but it is usually surprisingly high. A macrophage, for example, ingests 25% of its own volume of fluid each hour. This means it must ingest 3% of its plasma membrane each minute, or 100% in about half an hour. Fibroblasts endocytose at a somewhat lower rate (1% of their plasma membrane per minute), whereas some amoebae ingest their plasma membrane even more rapidly.

The endocytic part of the cycle often begins at clathrin-coated pits. These specialized regions typically occupy about 2% of the total plasma membrane area. The lifetime of a clathrin-coated pit is short: within a minute or so of being formed, it invaginates into the cell and pinches off to form a clathrin-coated vesicle (Figure 13–51). About 2500 clathrin-coated vesicles pinch off from the plasma membrane of a cultured fibroblast every minute. The coated vesicles are even more transient than the coated pits: within seconds of being formed, they shed their coat and fuse with early endosomes.

Caveolae, sometimes seen in the electron microscope as deeply invaginated flasks, are present in the plasma membrane of most vertebrate cell types (Figure 13–52). The major structural proteins in caveolae are caveolins, a family of unusual integral membrane proteins that each insert a hydrophobic loop into the membrane from the cytosolic side but do not extend across the membrane. On their cytosolic side, caveolins are bound to large protein complexes of cavin proteins, which are thought to stabilize the membrane curvature. Caveolae are especially rich in cholesterol, glycosphingolipids, and glycosylphosphatidylinositol (GPI)-anchored membrane proteins and might represent a type of lipid raft in the plasma membrane (see Figure 10–13).

In contrast to clathrin-coated and COPI-coated or COPII-coated vesicles, caveolae are usually static structures that can serve as a reservoir of additional plasma membrane. It is thought that cells subjected to dynamic changes in shear forces, such as the endothelial cells of arteries, exploit this reservoir to provide their plasma membranes greater resilience to stretch. This is accomplished by the rapid disassembly of the cavin protein scaffold in response to mechanical force, thereby allowing the underlying membrane to temporarily increase the surface area of the cell. The ability to rapidly change membrane surface area is thought to be important for accommodating dynamic changes in blood flow to different parts of the brain.

Two other endocytosis pathways are known, neither of which uses clathrin. Macropinocytosis is a process whereby the plasma membrane protrudes from the cell and engulfs a portion of the surrounding extracellular fluid into a macropinosome. This is a nonselective process for bringing fluid into the cell under certain conditions. In phagocytosis, the plasma membrane is directed to wrap around the particle to be engulfed until it fuses with itself, resulting in an enclosed phagosome inside the cell.

Because ligands are selectively captured by receptors, receptor-mediated endocytosis provides a selective concentrating mechanism that increases the efficiency of internalization of particular ligands more than a hundredfold. In this way, even minor components of the extracellular fluid can be efficiently taken up in large amounts. A particularly well-understood and physiologically important example is the process that mammalian cells use to import cholesterol. Many animal cells take up cholesterol through receptor-mediated endocytosis and, in this way, acquire most of the cholesterol they require to make new membrane. If the uptake is blocked, cholesterol accumulates in the blood and can contribute to the formation in blood vessel (artery) walls of atherosclerotic plaques, deposits of lipid and fibrous tissue that can cause strokes and heart attacks by blocking arterial blood flow. In fact, it was a study of humans with a strong genetic predisposition for atherosclerosis that first revealed the mechanism of receptor-mediated endocytosis.

Most cholesterol is transported in the blood as cholesterol esters in the form of lipid–protein particles known as low-density lipoproteins (LDLs) that, architecturally, resemble lipid droplets bearing a core of triacylglycerol, free cholesterol, and cholesterol esters. The droplet is stabilized by a single molecule of apolipoprotein B, a very large protein that wraps around the LDL particle (Figure 13–53). When a cell needs cholesterol for membrane synthesis, it makes transmembrane receptor proteins for LDL at the ER and transports them to the plasma membrane. Once in the plasma membrane, the LDL receptor diffuses until an endocytosis signal in its cytoplasmic tail binds the adaptor protein AP2 after AP2’s conformation has been locally unlocked by binding to PI(4,5)P2 on the plasma membrane.

AP2 then recruits clathrin to initiate endocytosis. Because coated pits constantly pinch off to form coated vesicles, any LDL particles bound to LDL receptors in the coated pits are rapidly internalized in coated vesicles. After shedding their clathrin coats, the vesicles deliver their contents to early endosomes. Once the LDL and LDL receptors encounter the low pH in early endosomes, LDL is released from its receptor and is delivered via late endosomes to lysosomes. There, the cholesterol esters in the LDL particles are hydrolyzed to free cholesterol, which is now available to the cell for new membrane synthesis (Movie 13.7).

If too much free cholesterol accumulates in a cell, the cell simultaneously shuts off endogenous cholesterol synthesis (Figure 12–64) and reduces exogenous cholesterol intake by shutting off the synthesis of LDL receptors. This regulated pathway for cholesterol uptake is disrupted in individuals who inherit defective genes encoding LDL receptors. The resulting high levels of blood cholesterol predispose these individuals to develop atherosclerosis prematurely, and many would die at an early age of heart attacks resulting from coronary artery disease if they were not treated with drugs such as statins that lower the level of blood cholesterol.

In some cases, the receptor is lacking altogether. In others, the receptors are defective—in either the extracellular binding site for LDL or the intracellular binding site for the AP2 adaptor protein in clathrin-coated pits. In the latter case, normal numbers of LDL receptors are present, but they fail to become localized in clathrin-coated pits. Although LDL binds to the surface of these mutant cells, it is not internalized, directly demonstrating the importance of clathrin-coated pits for the receptor-mediated endocytosis of cholesterol.

Many of these receptors, like the LDL receptor, enter coated pits irrespective of whether they have bound their specific ligands. Others enter preferentially when bound to a specific ligand, suggesting that a ligand-induced conformational change is required for them to activate the signal sequence that guides them into the pits. Because most plasma membrane proteins fail to become concentrated in clathrin-coated pits, the pits serve as molecular filters, preferentially collecting certain plasma membrane proteins (receptors) over others.

Early endosomes are the main sorting stations in the endocytic pathway, just as the cis and trans Golgi networks serve this function in the secretory pathway. In the mildly acidic environment of the early endosome, many internalized receptor proteins change their conformation and release their ligand, as already discussed for the M6P receptors. Those endocytosed ligands that dissociate from their receptors in the early endosome are usually destined for delivery to lysosomes, where they are either degraded and recycled into building blocks or utilized directly by the cell (such as the cholesterol just discussed).

The recycling transport vesicles bud from long, narrow tubules that extend from the early endosomes (Figure 13–55). It is likely that the geometry of these tubules helps the sorting process: because tubules have a large membrane area enclosing a small volume, membrane proteins become enriched over soluble proteins. The transport vesicles return the LDL receptor directly to the plasma membrane. The transferrin receptor follows a similar recycling pathway as the LDL receptor, but unlike the LDL receptor it also recycles its ligand. Transferrin is a soluble protein that carries iron in the blood.

Cell-surface transferrin receptors deliver transferrin with its bound iron to early endosomes by receptor-mediated endocytosis. The low pH in the endosome induces transferrin to release its bound iron, but the iron-free transferrin itself (called apotransferrin) remains bound to its receptor. The receptor–apotransferrin complex enters the tubular extensions of the early endosome and from there is recycled back to the plasma membrane. When the apotransferrin returns to the neutral pH of the extracellular fluid, it dissociates from the receptor and is thereby freed to pick up more iron and begin the cycle again. Thus, transferrin shuttles back and forth between the extracellular fluid and early endosomes, avoiding lysosomes and delivering iron to the cell interior, as needed for cells to grow and proliferate.

Cells can regulate the release of membrane proteins from recycling endosomes, thus adjusting the flux of proteins through the transcytotic pathway according to need. This regulation, the mechanism of which is uncertain, allows recycling endosomes to play an important part in adjusting the concentration of specific plasma membrane proteins. Fat cells and muscle cells, for example, contain large intracellular pools of the glucose transporters that are responsible for the uptake of glucose across the plasma membrane. These membrane transport proteins are stored in specialized recycling endosomes until the hormone insulin stimulates the cell to increase its rate of glucose uptake. In response to the insulin signal, transport vesicles rapidly bud from the recycling endosome and deliver large numbers of glucose transporters to the plasma membrane, thereby greatly increasing the rate of glucose uptake into the cell (Figure 13–57).

EGF is a small, extracellular signal protein that stimulates epidermal and various other cells to divide. Unlike LDL receptors, EGF receptors accumulate in clathrin-coated pits only after binding their ligand, and most do not recycle but are degraded in lysosomes, along with the ingested EGF. EGF binding therefore first activates intracellular signaling pathways and then leads to a decrease in the concentration of EGF receptors on the cell surface, a process called receptor down-regulation, that reduces the cell’s subsequent sensitivity to EGF (see Figure 15–21).

Receptor down-regulation is highly regulated. The activated receptors are first covalently modified on the cytosolic face with the small protein ubiquitin. Unlike polyubiquitylation, which adds a chain of ubiquitins that typically targets a protein for degradation in proteasomes (discussed in Chapter 6), ubiquitin tagging for sorting into the clathrin-dependent endocytic pathway adds just one or a few single ubiquitin molecules to the protein—a process called monoubiquitylation or multiubiquitylation, respectively. Ubiquitin-binding proteins recognize the attached ubiquitin and help direct the modified receptors into clathrin-coated pits.

The ubiquitylated receptor does not get recycled back to the plasma membrane from the early endosome. Instead, it remains there as the endosome matures. During the maturation process, which we discuss next, the ubiquitin tag is used to selectively sort the receptor and its bound ligand into intralumenal vesicles. Receptor signaling is terminated when the receptor is sequestered into intralumenal vesicles, which ultimately are degraded in lysosomes. In this way, addition of ubiquitin blocks receptor recycling to the plasma membrane and directs the receptors into the degradation pathway.

Macropinocytosis was among the first types of endocytosis to be described because it is visible by light microscopy, where cells can be seen taking up the surrounding fluid into large vesicles called macropinosomes (Figure 13–68). In most cell types, macropinocytosis does not operate continually but rather is induced for a limited time in response to cell-surface receptor activation by specific cargoes, including growth factors, integrin ligands, apoptotic-cell remnants, and some viruses. These ligands activate a complex signaling pathway, resulting in a change in actin dynamics and the formation of cell-surface protrusions, called ruffles (discussed in Chapter 16).

Macropinosomes form when the protruding ends of ruffles fuse with each other or the cell membrane, thereby trapping a portion of extracellular content. Macropinocytosis is a dedicated degradative pathway: macropinosomes acidify and then fuse with late endosomes or endolysosomes, without recycling their cargo back to the plasma membrane. Micropinocytosis is stimulated by activation of the oncogene Ras. Induction of macropinocytosis can increase the bulk fluid uptake of a cell by up to tenfold. Cancer cells that contain constitutively active Ras (see Chapter 15) use enhanced micropinocytosis to obtain increased nutrients from the surrounding environment in order to support their rapid growth and division.

Phagocytosis is a special form of endocytosis in which a cell uses large endocytic vesicles called phagosomes to ingest large particles such as microorganisms and dead cells. Phagocytosis is distinct, both in purpose and mechanism, from macropinocytosis, which we discussed earlier. In protozoa, phagocytosis is a form of feeding: large particles taken up into phagosomes end up in lysosomes, and the products of the subsequent digestive processes pass into the cytosol to be used as food. However, few cells in multicellular organisms are able to ingest such large particles efficiently. In the gut of animals, for example, extracellular processes break down food particles, and cells import the small products of hydrolysis.

In mammals, two important classes of white blood cells that act as professional phagocytes are macrophages and neutrophils (Movie 13.9). These cells develop from hemopoietic stem cells (discussed in Chapter 22), and they ingest invading microorganisms to defend us against infection. Macrophages also have an important role in scavenging senescent cells and cells that have died by apoptosis (discussed in Chapter 18). In quantitative terms, the clearance of senescent and dead cells is by far the most important: our macrophages, for example, phagocytose more than 1011 senescent red blood cells in each of us every day.

The diameter of a phagosome is determined by the size of its ingested particles, and those particles can be almost as large as the phagocytic cell itself (Figure 13–69). Phagosomes fuse with lysosomes, and the ingested material is then degraded. Indigestible substances remain in the lysosomes, forming residual bodies that can be excreted from cells by exocytosis Some of the internalized plasma membrane components never reach the lysosome, because they are retrieved from the phagosome in transport vesicles and returned to the plasma membrane.

Phagocytosis is a cargo-triggered process. That is, it requires the activation of cell-surface receptors that transmit signals to the cell interior. Thus, to be phagocytosed, particles must first bind to the surface of the phagocyte (although not all particles that bind are ingested). Phagocytes have a variety of cell-surface receptors that are functionally linked to the phagocytic machinery of the cell. The best-characterized triggers of phagocytosis are antibodies, which protect us by binding to the surface of infectious microorganisms (pathogens) and initiating a series of events that culminate in the invader being phagocytosed.

Localized actin polymerization, initiated by Rho family GTPases and their activating Rho GEFs (discussed in Chapters 15 and 16), shapes the pseudopods. The activated Rho GTPases switch on the kinase activity of local PI kinases to produce PI(4,5)P2 in the membrane (see Figure 13–11), which stimulates actin polymerization. To seal off the phagosome and complete the engulfment, actin is depolymerized by a PI 3-kinase that converts the PI(4,5)P2 to PI(3,4,5)P3, which is required for closure of the phagosome and may also contribute to reshaping the actin network to help drive the invagination of the forming phagosome (Figure 13–70). In this way, the ordered generation and consumption of specific phosphoinositides guides sequential steps in phagosome formation.

After transiting the Golgi cisternae, cargo molecules that arrive at the trans Golgi network (TGN) are sorted and packaged into transport vesicles that depart for different destinations. Transport vesicles destined for the cell surface normally leave the TGN in a steady stream as irregularly shaped tubules. The membrane proteins and the lipids in these vesicles provide new components for the cell’s plasma membrane, while the soluble proteins inside the vesicles are secreted to the extracellular space. The fusion of the vesicles with the plasma membrane is called exocytosis. This is the route, for example, by which cells secrete most of the proteoglycans and glycoproteins of the extracellular matrix, as discussed in Chapter 19. All cells require this constitutive secretory pathway, which operates continually (Movie 13.5).

This is the regulated secretory pathway, found mainly in cells specialized for secreting products rapidly on demand—such as hormones, neurotransmitters, or digestive enzymes. The third major destination from the TGN is endosomes. Hydrolases that function in the lumen of lysosomes use this pathway to first arrive at endosomes, which progressively mature into lysosomes (discussed later). The sorting mechanism at the TGN for lysosomal hydrolase proteins is especially well understood and provides an example of how cargo molecules in the TGN are segregated among different types of transport vesicles. In this section, we consider the role of the Golgi apparatus in sorting proteins between these three pathways and compare the mechanisms of constitutive and regulated secretion.

A cell capable of regulated secretion must separate at least three classes of proteins before they leave the TGN—those destined for lysosomes (via endosomes), those destined for secretory vesicles, and those destined for immediate delivery to the cell surface (Figure 13–38). Specific signals are needed to direct secretory proteins into secretory vesicles and lysosomal proteins into different specialized transport vesicles. The nonselective constitutive secretory pathway transports most other proteins directly to the cell surface. Because entry into this pathway does not require a particular signal, it is also called the default pathway.

As we discussed (see Figure 13–38), secretory vesicles form from the TGN, and they release their contents to the cell exterior by exocytosis in response to specific signals. The secreted product can be either a small molecule (such as histamine or a neuropeptide) or a protein (such as a hormone or digestive enzyme). Proteins destined for secretory vesicles (called secretory proteins) are packaged into appropriate vesicles in the TGN by a mechanism that involves the selective aggregation of the secretory proteins.

Initially, the membrane of the secretory vesicles that leave the TGN is only loosely wrapped around the clusters of aggregated secretory proteins. Morphologically, these immature secretory vesicles resemble dilated trans Golgi cisternae that have pinched off from the Golgi stack. As immature secretory vesicles mature, clathrin-coated transport vesicles bud from them and go back to the TGN (Figure 13–42). This recycling process not only returns Golgi components to the Golgi apparatus, but also serves to concentrate the contents of secretory vesicles. The sum of all the retrieval pathways during the transit of a secretory protein from the ER through the Golgi cisternae to a mature secretory vesicle results in a 200- to 400-fold increase in net concentration.

Microtubules also guide transport vesicles to the cell surface for constitutive exocytosis. Whereas transport vesicles containing materials for constitutive release fuse with the plasma membrane once they arrive there, secretory vesicles in the regulated pathway wait at the membrane until the cell receives a signal for the vesicles to secrete their cargo. The signal can be an electrical nerve impulse (an action potential) or an extracellular signal molecule, such as a hormone. In either case, it leads to a transient increase in the concentration of free Ca2+ in the cytosol, which is the trigger for secretory vesicle fusion.

When a secretory vesicle fuses with the plasma membrane, its contents are discharged from the cell by exocytosis and its membrane becomes part of the plasma membrane. Although this should increase the surface area of the plasma membrane, it does so only transiently, because an equivalent amount of membrane is removed from the surface by endocytosis almost as fast as it is added by exocytosis, a process reminiscent of the endocytic–exocytic cycle discussed later. The proteins of the secretory vesicle membrane that are endocytosed from the plasma membrane are either recycled or shuttled to lysosomes for degradation through mechanisms discussed later.

The amount of secretory vesicle membrane that is temporarily added to the plasma membrane can be enormous: in a pancreatic acinar cell discharging digestive enzymes for delivery to the gut lumen, about 900 μm2 of vesicle membrane is inserted into the apical plasma membrane (whose area is only 30 μm2) when the cell is stimulated to secrete. Control of membrane traffic thus has a major role in maintaining the composition of the various membranes of the cell.

To maintain each membrane-enclosed compartment in the secretory and endocytic pathways at a constant size, the balance between the outward and inward flows of membrane needs to be precisely regulated. For cells to grow, however, the forward flow needs to be greater than the retrograde flow, so that the membrane can increase in area. For cells to maintain a constant size, the forward and retrograde flows must be equal. We still know very little about the mechanisms that coordinate these flows.

An important task of regulated exocytosis is to deliver more membrane to enlarge the surface area of a cell’s plasma membrane when such a need arises. A spectacular example is the plasma membrane expansion that occurs during the cellularization process in a fly embryo, which initially is a syncytium—a single cell containing about 6000 nuclei surrounded by a single plasma membrane (see Figure 21–14). Within tens of minutes, the embryo is converted into the same number of cells. This process of cellularization requires a vast amount of new plasma membrane, which is added by a carefully orchestrated fusion of cytoplasmic vesicles, eventually forming the plasma membranes that enclose the separate cells. Similar vesicle fusion events are required to enlarge the plasma membrane when other animal cells or plant cells divide during cytokinesis (discussed in Chapter 17).

In addition to providing an emergency barrier against leaks, the patch reduces membrane tension over the wounded area, allowing the bilayer to flow back together to restore continuity and seal the puncture. The fusion and exocytosis of vesicles that mediated membrane repair is triggered by the sudden increase of Ca2+, which is abundant in the extracellular space and rushes into the cell as soon as the plasma membrane is punctured. Figure 13–48 shows four examples in which regulated exocytosis leads to plasma membrane expansion.

Cells ingest fluid, molecules, and particles by endocytosis, in which localized regions of the plasma membrane invaginate and pinch off to form endocytic vesicles. In most cells, endocytosis internalizes a large fraction of the plasma membrane every hour. The cells remain the same size because most of the plasma membrane components (proteins and lipids) that are endocytosed are continually returned to the cell surface by exocytosis. This large-scale endocytic–exocytic cycle is mediated largely by clathrin-coated pits and vesicles, but clathrin-independent endocytic pathways also contribute.