专题5 细胞质基质与内膜系统

cytosol Contents of the main compartment of the cytoplasm, defined as excluding membrane-bounded organelles such as endoplasmic reticulum and mitochondria. (Chapter 12)

The surrounding cytoplasm consists of the cytosol and the cytoplasmic organelles suspended in it. The cytosol constitutes a little more than half the total volume of the cell, and it is the main site of protein synthesis and degradation. It also performs most of the cell’s intermediary metabolism; that is, the many reactions that degrade some small molecules and synthesize others to provide the building blocks for macromolecules (discussed in Chapter 2).

An animal cell contains about 10 billion (1010) protein molecules of perhaps 10,000 kinds, and the synthesis of almost all of them begins in the cytosol, the space of the cytoplasm outside the membrane-enclosed organelles. Each newly synthesized protein is then delivered specifically to the organelle that requires it.

The major process for oxidizing sugars is the sequence of reactions known as glycolysis, which produces ATP without the involvement of molecular oxygen (O2 gas). It occurs in the cytosol of most cells, including many anaerobic microorganisms. Glycolysis probably evolved early in the history of life, before photosynthetic organisms introduced oxygen into the atmosphere.

three main sets of reactions act in series, the products of each being the starting material for the next: glycolysis (which occurs in the cytosol), the citric acid cycle (in the mitochondrial matrix), and oxidative phosphorylation (on the inner mitochondrial membrane).

Nearly all proteins, except a few inside mitochondria and plastids, begin their synthesis on ribosomes in the cytosol. The final location of each protein depends on its amino acid sequence, which can contain one or more sorting signals that direct its delivery to different parts of the cell.

Most proteins on their own do not fold correctly during their synthesis; instead, they require a special class of proteins called molecular chaperones to do so. Molecular chaperones are useful for cells because there are many different folding paths available to an unfolded or partially folded protein. Without these chaperones, some of these pathways would not lead to the correctly folded (and most stable) form; instead, the protein would become “kinetically trapped” in structures that are off-pathway.

molecular chaperone (chaperone) Protein that helps guide the proper folding of other proteins or helps them avoid misfolding. Includes heat-shock proteins. (Chapter 6)

Many molecular chaperones are called heat-shock proteins (designated hsp), because they are synthesized in dramatically increased amounts after a brief exposure of cells to an elevated temperature (for example, 42°C for cells that normally live at 37°C). This increase reflects the operation of a feedback system that responds to an increase in misfolded proteins (such as those produced by elevated temperatures) by boosting the synthesis of the chaperones that help these proteins refold.

There are several major families of molecular chaperones, including the hsp60 and hsp70 proteins, with different family members functioning in different organelles. Thus, as discussed in Chapter 12, mitochondria contain their own hsp60 and hsp70 molecules that are distinct from those that function in the cytosol, and a special hsp70 (called BiP) helps to fold proteins in the endoplasmic reticulum.

Short stretches of hydrophobic amino acids that are abnormally exposed in misfolded proteins (green) trigger the hydrolysis of ATP to ADP, causing hsp70 to close down on its substrate, trapping it in an extended conformation. The rebinding of another molecule of ATP will open the hsp70 “clamp,” releasing the substrate in its extended form. Repeated cycles of hsp70 clamping and release, often involving multiple hsp70 molecules on the same target, help fold the target protein by keeping its hydrophobic regions from aggregating.

An important class is exemplified by the hsp60 proteins. These form a large barrelshaped structure that acts after a protein has been fully synthesized but before it has folded correctly. This type of chaperone, sometimes called a chaperonin, forms an “isolation chamber” for the folding process (Figure 6–84).

These include an hsp90 chaperone that can harness mechanical forces to help proteins fold as part of a collaborative chaperone network (Figure 6–85). The hsp70, hsp60, and hsp90 chaperones often need many cycles of ATP binding and hydrolysis to fold a single polypeptide chain correctly.

Figure 3–65 The marking of proteins by ubiquitin. (A) The three-dimensional structure of ubiquitin, a small protein of 76 amino acids. A family of special enzymes couples its carboxyl end to the amino group of a lysine side chain in a target protein molecule, forming an isopeptide bond.

E1 then passes this activated ubiquitin to one of a set of ubiquitin-conjugating (E2) enzymes, each of which acts in conjunction with a set of accessory (E3) proteins called ubiquitin ligases that select the target proteins to be modified.

Here, the ubiquitin ligase binds to specific degradation signals, called degrons, in protein substrates, thereby helping E2 to form a polyubiquitin chain linked to a lysine of the substrate protein. This polyubiquitin chain on a target protein will then be recognized by a specific receptor in the proteasome, causing the target protein to be rapidly destroyed.

In this case, a series of specialized enzymes recognizes the hydrophobic patch (possibly with the aid of chaperones) and attaches the small protein ubiquitin to a nearby lysine (see Figure 3–65A). Additional ubiquitin molecules are subsequently added to build a polyubiquitin chain. As we saw in Chapter 3, ubiquitin modification of proteins is used for many purposes in the cell. The particular type of ubiquitin linkage that concerns us here is a chain of ubiquitin molecules linked together at lysine 48 (see Figure 3–65B), which is the distinguishing feature of the ubiquitin tag that marks a protein for destruction. This polyubiquitin chain delivers the protein to a protein-destruction machine, a complex protease known as the proteasome.

Proteasomes are very abundant, constituting approximately 1% of the total proteins in cells. They are dispersed throughout the cytosol and the nucleus, but the proteasome also destroys aberrant proteins that have entered the endoplasmic reticulum (ER).

Each end of the cylinder is normally associated with a large protein complex (the 19S cap) that contains a six-subunit protein ring through which target proteins are threaded into the proteasome core, where they are degraded (Figure 6–86).

The entire proteasome, in which the central cylinder (yellow) is supplemented by a 19S cap (blue) at each end. The complex cap selectively binds proteins that are marked by ubiquitin for destruction; if the protein also contains a loosely structured region, the cap uses ATP hydrolysis to further unfold its polypeptide chain and feed it through the narrow channel (see Figure 6–87) into the inner chamber of the 20S cylinder for digestion to short peptides.

The proteasome cap recognizes proteins marked by a polyubiquitin chain (see Figure 3–65B). Most of these proteins also contain an unfolded region, and the cap translocates these proteins into the proteasome core, where they are digested. Before moving through the proteasome cap, the ubiquitin is cleaved from the substrate protein and is recycled.

A crucial property of the proteasome, and one reason for its complexity, is the processivity of its mechanism: in contrast to a “simple” protease that cleaves a substrate’s polypeptide chain just once before dissociating, the proteasome keeps the entire substrate bound until all of it is converted into short peptides.

Yet another function of these proteolytic pathways is to confer short lifetimes on specific normal proteins whose concentrations must change promptly with alterations in the state of a cell. Some of these short-lived proteins are degraded rapidly at all times, while many others are conditionally short-lived; that is, they are metabolically stable under some conditions but become unstable upon a change in the cell’s state. For example, mitotic cyclins are long-lived throughout the cell cycle until their sudden degradation at the end of mitosis, as explained in Chapter 17. How is such a regulated destruction of a protein controlled?

Finally, powerful degradation signals can be created by cleaving a single peptide bond, provided that this cleavage creates a new N-terminus that is recognized by a specific E3 protein as a “destabilizing” N-terminal residue. This E3 protein recognizes only certain amino acids at the N-terminus of a protein; thus, not all protein-cleavage events will lead to degradation of the C-terminal fragment produced.

In humans, nearly 70% of cytosolic proteins are acetylated on their N-terminal residue, and we now know that this modification is recognized by a specific E3 enzyme, which directs the ubiquitylation of the protein and sends it to the proteasome for degradation. Thus, the majority of human proteins carry their own signals for destruction.

As might be expected, there is competition for misfolded proteins between chaperones and the protein degradation machinery. Proteins that are folded quickly escape destruction (at least early in their life before they accumulate damage), whereas those that undergo many rounds of chaperone-assisted folding are more likely to be degraded.

It is estimated that between 1 and 5% of all newly synthesized proteins fail to fold properly and are degraded by the proteasome.

The major intracellular compartments in eukaryotic cells can therefore be categorized into three distinct families: (1) the nucleus and the cytosol, which are topologically equivalent (although functionally distinct) and connected by nuclear pore complexes; (2) all organelles that function in the secretory and endocytic pathways—including the ER, Golgi apparatus, endosomes, lyso somes, and the transport vesicles that move between them—and peroxisomes; (3) the endosymbiont-derived organelles: mitochondria and the plastids (in plants only).

First, the cell now has a set of internal membranes that are derived from an ancestral prokaryotic plasma membrane. These internal membranes enclose interior spaces that are said to be topologically equivalent to each other and to the exterior of the cell (Figure 12–4), because they can communicate with one another, in the sense that molecules can get from one to the other without having to cross a membrane. We shall see that this topological relationship holds for all of the organelles involved in the secretory and endocytic pathways, including the ER, Golgi apparatus, endosomes, lysosomes, and peroxisomes.

Figure 12–4 Topologically equivalent compartments in the secretory and endocytic pathways in a eukaryotic cell. Topologically equivalent spaces are shown in red. (A) Molecules can be carried from one compartment to another topologically equivalent compartment by transport vesicles that bud from one and fuse with the other.

On the way to lysosomes, endocytosed material must first pass through a series of organelles called endosomes. As we will see, the ER, Golgi apparatus, lysosomes, endosomes, and plasma membrane are linked by the cell’s major pathways of membrane traffic.

Their common origin from a primordial intracellular compartment explains why all of these compartments can exchange material with each other through vesicular transport. The nucleus was formerly the cytosol in the ancient archaeon, explaining why the cytosol and nucleus are topologically equivalent compartments that can intermix during mitosis.

In protein translocation, transmembrane protein translocators directly transport specific proteins from the cytosol into a space that is topologically distinct: either the other side of a membrane or within the lipid bilayer in the case of integral membrane proteins. The transported protein molecule usually must unfold to snake through the translocator. The initial transport of selected proteins from the cytosol into the ER lumen, the ER membrane, or mitochondria occurs in this way.

Sorting signals are usually composed of amino acid side chains in a protein and come in two general varieties: a linear sequence of amino acids (called a signal sequence) or a specific three-dimensional arrangement of amino acids (called a signal patch).

Proteins destined for mitochondria have signal sequences of yet another type, in which positively charged amino acids alternate with hydrophobic ones. The signal for protein import into the nucleus is composed primarily of positively charged amino acids. Finally, many proteins destined for peroxisomes have a signal sequence of three characteristic amino acids at their C-terminus.

Placing the N-terminal ER signal sequence at the beginning of a cytosolic protein, for example, redirects the protein to the ER; removing or mutating the signal sequence of an ER protein causes its retention in the cytosol. Signal sequences are therefore both necessary and sufficient for protein targeting.

Sorting signals are recognized by complementary sorting receptors that guide proteins to their appropriate destination, where the receptors unload their cargo. The receptors function catalytically: after completing one round of targeting, they return to their point of origin to be reused.

The membrane of the endoplasmic reticulum (ER) typically constitutes more than half of the total membrane of an average animal cell (see Table 12–2). The ER is organized into a netlike labyrinth of branching tubules and flattened sacs that extends throughout the cytosol (Figure 12–14 and Movie 12.2). The tubules and sacs interconnect, and their membrane is continuous with the outer nuclear membrane. This membrane system encloses a single internal space, called the ER lumen, which is continuous with the space between the inner and outer nuclear membranes. The ER often occupies more than 10% of the total cell volume (see Table 12–1).

The ER has a central role in the biosynthesis of both lipids and proteins, and the ER lumen stores intracellular Ca2+ that is mobilized in many cell signaling responses (discussed in Chapter 15). The ER membrane is the site of production of many of the transmembrane proteins and lipids of the cell’s organelles, includ ing the ER itself, the Golgi apparatus, lysosomes, endosomes, secretory vesicles, peroxisomes, and the plasma membrane.

While the various functions of the ER are essential to every cell, their relative importance varies greatly between individual cell types. To meet different func tional demands, distinct regions of the ER become highly specialized. Functional specialization entails dramatic changes in the proportional abundance of dif ferent parts of the ER. These changes are observed as characteristically different types of ER membrane in different types of cells.

The most visually remarkable specializations are the rough ER and smooth ER (Figure 12–15). The rough appearance is due to the abundance of ribosomes engaged in protein synthesis bound to the surface of this part of the ER. By contrast, regions of smooth ER lack ribosomes and are dedicated to other ER functions such as the biosynthesis and metabolism of lipids. All cells have both rough and smooth ER, but their relative abundance can vary enormously in specialized cells.

Most secreted proteins are synthesized by the ribosomes that stud the sur face of the rough ER. Thus, cells specialized to secrete vast amounts of protein are packed with an abundance of rough ER. For example, exocrine cells of the pancreas secrete their own weight in digestive enzymes every day, explaining why the rough ER makes up 60% of these cells’ membranes (see Table 12–2). Similarly, antibody-secreting plasma cells and insulin-secreting β cells also contain a mark edly expanded rough ER.

In contrast to rough ER, functions for the smooth ER are more diverse and can become highly specialized. A type of smooth ER found in all cells is called transitional ER, from which transport vesicles carrying newly synthesized proteins and lipids bud off for transport to the Golgi apparatus. In certain specialized cells, the smooth ER has additional functions that warrant its expansion. For example, cells that synthesize steroid hormones contain prominent smooth ER to accom modate the enzymes that make cholesterol and modify it to form a variety of steroid hormones (see Figure 12–15B).

To initiate their journey along the secretory pathway, proteins that have entered the ER and are destined for the Golgi apparatus or beyond are first packaged into COPII-coated transport vesicles. These vesicles bud from specialized regions of the ER called ER exit sites, whose membrane lacks bound ribosomes. Most animal cells have ER exit sites dispersed throughout the ER network.

To study the functions and biochemistry of the ER, it is necessary to isolate it. This may seem to be a hopeless task because the ER is intricately interleaved with other components of the cytoplasm. Fortunately, when tissues or cells are disrupted by homogenization, the ER breaks into fragments, which reseal to form small (∼100–200 nm in diameter) closed vesicles called microsomes (Figure 12–17).

To the biochemist, microsomes represent small authentic ver sions of the ER, still capable of protein translocation, protein glycosylation (discussed later), Ca2+ uptake and release, and lipid synthesis. Rough microsomes, derived from rough ER, contain ribosomes on their outside surface and enclose a small part of the ER lumen. Smooth microsomes, which lack ribosomes, are derived from vesiculated fragments of the smooth ER, plasma membrane, Golgi apparatus, endosomes, and mitochondria.

The ribosomes attached to rough microsomes make them denser than smooth microsomes. As a result, scientists use equilibrium density centrifugation to separate the rough and smooth micro somes (Figure 12–17).

In the key experiment, the mRNA encoding a secreted protein was added to cytosol extracted from cells. In this cell-free reaction, ribosomes in the cytosol translated the mRNA into a protein that was slightly larger than the normal secreted protein (Figure 12–18). When the reaction was repeated in the presence of microsomes derived from the rough ER, a protein of the correct size was pro duced and located inside the microsomes (Figure 12–18).

Finally, the smooth ER can be specialized in regions that make intimate con tacts with other organelles, most notably the mitochondria, plastids, endosomes, and the plasma membrane (Figure 12–16). These organelle contact sites are enriched for proteins involved in communication or transport of key metabolites between the juxtaposed membranes. For example, the transport of lipids from their site of synthesis in the ER to the mitochondrion is thought to occur at ER–mitochondria contact sites.

Mitochondria and plastids do not communicate with the ER by vesicular trans port, so they require different mechanisms to import many of their lipids from the ER for growth. Carrier proteins in the cytosol called lipid transfer proteins ferry individual lipid molecules between membranes, functioning much like fatty acid binding proteins that shepherd fatty acids through the cytosol (see Figure 12–38). In many cases, lipid transfer proteins function at organelle con tact sites where the originating and destination membranes are held within ∼10–30 nm of each other by specific junction complexes.

Different lipid trans fer proteins shuttle phosphatidylcholine and phosphatidylserine from the ER to mitochondria at contact sites. Disruption of the junctional complexes or the lipid transfer proteins impairs lipid import into mitochondria and causes their dysfunction. The extensive network of the ER participates in contact sites with most other cellular organelles (Figure 12–41).

Proteins enter the ER lumen as unfolded polypeptides. They must therefore fold and assemble into their correct three-dimensional structures just as newly made proteins in the cytosol must fold (discussed in Chapter 3). To meet this demand, the lumen of the ER contains a high concentration of resident chaperones and other protein-folding catalysts. These ER resident proteins contain an ER retention signal of four amino acids at their C-terminus that is responsible for retaining the protein in the ER (see Figure 12–13; discussed in Chapter 13, p. 768).

Soluble ER resident proteins, such as BiP, also contain a short ER retrieval signal at their C-terminal end, but it is different: it consists of a Lys-Asp-Glu-Leu or a similar sequence. If this signal (called the KDEL sequence) is removed from BiP by genetic engineering, the protein is slowly secreted from the cell. If the signal is transferred to a protein that is normally secreted, the protein is now efficiently returned to the ER, where it accumulates.

Unlike the retrieval signals on ER membrane proteins, which can interact directly with the COPI coat, soluble ER resident proteins must bind to specialized receptor proteins such as the KDEL receptor—a multipass transmembrane protein that binds to the KDEL sequence and packages any protein displaying it into COPI-coated retrograde transport vesicles (Figure 13–26). The KDEL receptor accomplishes this task by cycling between the ER and the Golgi apparatus, selectively binding proteins with the KDEL sequence in the Golgi apparatus and releasing them in the ER.

The protein BiP, a member of the hsp70 family of chaperone proteins, is a major component of the ER folding machinery. We have already discussed how BiP pulls proteins post-translationally into the ER through the Sec61 ER translo cator.

Like other chaperones (discussed in Chapter 6), BiP recognizes incorrectly folded proteins, as well as protein subunits that have not yet assembled into their final oligomeric complexes. It does so by binding to exposed hydrophobic amino acid sequences that would normally be buried in the interior of correctly folded or assembled polypeptide chains. The bound BiP both prevents the protein from aggregating and helps keep it in the ER (and thus out of the Golgi apparatus and later parts of the secretory pathway).

BiP hydrolyzes ATP to shuttle between highand low-affinity polypeptide-binding states. In this way, BiP periodically lets go of its substrate proteins to allow them an opportunity to fold, and then re-binds them if folding is not yet achieved.

The ER resident protein protein disulfide isomerase (PDI) catalyzes the oxida tion of free sulfhydryl (SH) groups on cysteines to form disulfide (S}S) bonds (Figure 12–31).

Disulfide bonds stabilize the folded state of a protein, enabling it to better withstand a harsh, variable, and chaperone-free extracellular environment. Because proteins often contain multiple cysteines, they sometimes pair incorrectly. PDI resolves this problem by rearranging the disulfide bonds in a protein until it is correctly folded. This is possible because PDI enzymes are capa ble of operating in reverse to reduce incorrectly paired disulfides of immature proteins.

The ER lumen contains multiple members of the PDI family, some of which are specialized for reducing disulfide bonds to fully unfold misfolded pro teins that need to be translocated back to the cytosol for degradation (discussed later). All PDI enzymes are therefore oxidoreductases that can catalyze either the formation or breakage of disulfide bonds in their client proteins. The formation of disulfide bonds relies on maintaining an oxidizing environment in the ER lumen. Disulfide bonds form only very rarely in domains exposed to the cytosol because of the reducing environment there.

It has long been debated why glycosylation is such a common modification of pro teins that enter the ER. One particularly puzzling observation has been that some proteins require N-linked glycosylation for proper folding in the ER, yet the precise location of the oligosaccharides attached to the protein’s surface does not seem to matter. A clue to the role of glycosylation in protein folding came from studies of two ER chaperone proteins, which are called calnexin and calreticulin because they require Ca2+ for their activities.

Like other chaperones, they prevent incompletely folded proteins from irreversibly aggregating. Both calnexin and calreticulin also promote the association of incompletely folded proteins with another ER chaper one, which binds to cysteines that have not yet formed disulfide bonds.

How do calnexin and calreticulin distinguish properly folded from incom pletely folded proteins? The answer lies in the structure of the oligosaccharide attached to the protein. Shortly after a newly made protein acquires an N-linked precursor oligosaccharide, ER glucosidases rapidly remove two glucoses, leaving behind a single terminal glucose. This singly glucosylated oligosaccharide is rec ognized by calnexin and calreticulin, ensuring that all newly made (and hence, likely to be not yet folded) glycoproteins bind to one of these chaperones.

This last glucose is removed over time, leaving a de-glucosylated glycoprotein that no lon ger binds to calnexin or calreticulin. If the glycoprotein is folded, it can leave the ER. However, yet another ER enzyme, a glucosyl transferase, re-adds the terminal glucose selectively to glycoproteins that have not yet folded completely. The ter minal glucose then causes re-association of the unfolded protein with calnexin or calreticulin.

The covalent addition of oligosaccharides to proteins is one of the major bio synthetic functions of the ER. About half of the soluble and membrane-bound proteins that are processed in the ER—including those destined for transport to the Golgi apparatus, lysosomes, plasma membrane, or extracellular space—are glycoproteins that are modified in this way. Some proteins in the cytosol and nucleus are also glycosylated, but not with large oligosaccharides: they instead carry a much simpler sugar modification, in which a single N-acetylglucosamine group is added to a serine or threonine of the protein.

During the most common form of protein glycosylation in the ER, a preformed precursor oligosaccharide (containing 14 sugars composed of 2 N- acetylglucosamines, 9 mannoses, and 3 glucoses) is transferred as a complete unit to proteins. Because this oligosaccharide is transferred to the side-chain NH2 group of an asparagine in the protein, it is said to be N-linked, or asparagine-linked (Figure 12–32A). A special lipid molecule called dolichol (see Panel 2–5, pp. 102–103) anchors the precursor oligosaccharide in the ER membrane. The pre cursor oligosaccharide is transferred to the target asparagine in a single enzymatic step by an oligosaccharyl transferase.

The precursor oligosaccharide is built up sugar by sugar on the membranebound dolichol lipid. The sugars are first activated in the cytosol by the formation of nucleotide (UDP or GDP)-sugar intermediates, which then donate their sugar first to the dolichol lipid and then to the partially assembled oligosaccharide tree in an orderly sequence. Partway through this process, the lipid-linked oligosac charide is flipped, with the help of a transporter, from the cytosolic to the lumenal side of the ER membrane (Figure 12–33).

The N-linked oligosaccharides are by far the most common oligosaccharides, being found on 90% of all glycoproteins. Less frequently, oligosaccharides are linked to the hydroxyl group on the side chain of a serine, threonine, hydroxylysine, or hydroxyproline amino acid. The first sugar of these O-linked oligosaccharides is added in the ER. N-linked and O-linked oligosaccharides undergo extensive pro cessing, modification, and extension in the Golgi apparatus (Chapter 13), producing the diversity of oligosaccharide structures observed on mature glycoproteins.

The ER membrane is the site of synthesis of nearly all of the cell’s major classes of lipids, including both phospholipids and cholesterol, required for the production of new cell membranes. The major phospholipid made is phosphatidylcholine, which can be formed in three steps from choline, two fatty acids, and glycerol phosphate (Figure 12–38). Each step is catalyzed by enzymes in the ER membrane, which have their active sites facing the cytosol, where all of the required metabolites are found. Thus, phospholipid synthesis occurs exclu sively in the cytosolic leaflet of the ER membrane.

Phosphatidic acid is suf ficiently water-insoluble to remain in the lipid bilayer; it cannot be extracted from the bilayer by the fatty acid binding proteins. It is therefore this first step that enlarges the ER lipid bilayer. The later steps determine the head group of a newly formed lipid molecule and therefore the chemical nature of the bilayer, but they do not result in net membrane growth.

Because phospholipid synthesis takes place in the cytosolic leaflet of the ER lipid bilayer, there needs to be a mechanism that transfers some of the newly formed phospholipid molecules to the lumenal leaflet of the bilayer. In synthetic lipid bilayers, lipids do not “flip-flop” in this way (see Figure 10–10). In the ER, however, phospholipids equilibrate across the membrane within minutes, which is almost 100,000 times faster than can be accounted for by spontaneous “flipflop.”

As discussed in Chapter 13, the plasma membrane and the membranes of the Golgi apparatus, lysosomes, and endosomes all form part of a membrane system that communicates with the ER by means of transport vesicles. A large propor tion of the lipids that compose the membranes of these organelles is acquired via the membranes delivered by transport vesicles.

About half the total area of membrane in a eukaryotic cell encloses the laby rinthine spaces of the endoplasmic reticulum (ER). Most soluble and integral membrane proteins destined for the cell exterior or for other organelles are initially assembled at the ER. These proteins are transported into the ER as they are synthesized by ribosomes. The distinctive appearance in electron micro graphs of ribosomes studding the surface of these regions of the ER is the reason they are termed the rough ER. The ER also produces most of the lipid and sterols for the rest of the cell and functions as a store for Ca2+ ions. These regions of the ER typically lack bound ribosomes and are called smooth ER.

The main cell type in the liver, the hepatocyte, also has expanded amounts of smooth ER (see Table 12–2) serving two separate purposes. The hepatocyte is the principal site of production of lipoprotein particles, which carry lipids via the bloodstream to other parts of the body. The enzymes that synthesize the lipid components of the particles are enriched in the membrane of the smooth ER.

In addition, these membranes contain enzymes that catalyze a series of reactions to detoxify drugs and various harmful compounds produced by metabolism. The most extensively studied of these detoxification reactions are carried out by the cytochrome P450 family of enzymes. They catalyze a series of reactions in which water-insoluble drugs or metabolites that would otherwise accumulate to toxic levels in cell membranes are rendered sufficiently water soluble to leave the cell and be excreted in the urine or bile.

A Ca2+ pump transports Ca2+ from the cyto sol into the ER lumen. A high concentration of Ca2+-binding proteins in the ER facilitates Ca2+ storage. In some cell types, specific regions of the ER are special ized for Ca2+ storage. Muscle cells have an abundant, modified smooth ER called the sarcoplasmic reticulum. The release and reuptake of Ca2+ by the sarcoplasmic reticulum trigger myofibril contraction and relaxation, respectively, during each round of muscle contraction (discussed in Chapter 16).

Despite all the help from chaperones, many protein molecules translocated into the ER fail to achieve their properly folded or oligomeric state. Such pro teins are exported from the ER back into the cytosol, where they are degraded in proteasomes (discussed in Chapter 6). In many ways, the mechanism of such retrotranslocation is similar to other post-translational modes of translocation.

Selecting proteins from the ER for degradation is a challenging process: misfolded proteins or unassembled protein subunits should be degraded, but folding intermediates of newly made proteins should not. Help in making this distinction comes from the N-linked oligosaccharides, which serve as timers that measure how long a protein has spent in the ER. The slow trimming of a particular mannose on the core oligosaccharide tree by an enzyme (a mannosi dase) in the ER creates a new oligosaccharide structure that ER-lumenal lectins of the retrotranslocation apparatus recognize. A protein that folds and exits from the ER faster than the mannosidase can remove its target mannose therefore escapes degradation.

Multiple translocator complexes move different proteins from the ER mem brane or lumen into the cytosol. Translocator complexes always contain an E3 ubiquitin ligase enzyme (Chapter 6), which attaches polyubiquitin tags to the unfolded proteins as they emerge into the cytosol, marking them for destruc tion. Fueled by the energy derived from ATP hydrolysis, a hexameric ATPase of the family of AAA-ATPases (see Figure 6–88) pulls the unfolded protein through the translocator into the cytosol. An N-glycanase removes en bloc any oligosac charide chains attached to the retrotranslocated protein. Guided by its ubiquitin tag, the de-glycosylated polypeptide is rapidly fed into proteasomes, where it is degraded (Figure 12–35).

Sometimes, however, there are drawbacks to the stringent quality-control mechanism. The predominant mutations that cause cystic fibrosis, a common inherited disease, result in the production of a slightly misfolded form of a plasma membrane protein important for Cl– transport. Although the mutant protein would function almost normally if it reached the plasma membrane, it is retained in the ER and then is degraded by cytosolic proteasomes. This devastating disease thus results not because the mutation inactivates the protein but because potentially active protein is discarded before it reaches the plasma membrane.

Cells carefully monitor the amount of misfolded protein in various compart ments. An accumulation of misfolded proteins in the cytosol, for example, triggers a heat-shock response (discussed in Chapter 6), which stimulates the transcrip tion of genes encoding cytosolic chaperones that help to refold the proteins.

The stimulated genes code for ER chaperones, the machinery for protein retrotranslocation and degradation, fac tors for protein transport out of the ER, and factors for expansion of the ER. This multipronged response operates by coupling the detection of misfolded proteins in the ER lumen to the production of transcription regulatory proteins that enter the nucleus to tune the transcription of hundreds of genes. How do misfolded proteins in the ER signal to the nucleus? There are three parallel pathways that execute the unfolded protein response (Figure 12–36).

The relative importance of each of these three pathways in the unfolded protein response differs in different cell types, enabling each cell type to tailor the unfolded protein response to its particular needs. The signaling pathways that execute the unfolded protein response are used during normal physiological conditions to adjust ER capacity to closely match demand for the ER.

The first pathway, which was initially discovered in yeast cells, is conserved in all eukaryotic cells and is particularly remarkable. Misfolded proteins in the ER cause IRE1, a transmembrane protein kinase in the ER, to oligomerize and phosphorylate itself. This mechanism of activation is similar to how some cell-surface receptor kinases in the plasma membrane are activated (discussed in Chapter 15). Oligomeric and phosphorylated IRE1 enables its cytosolic endoribonuclease domain to remove an intron from a specific cytosolic mRNA molecule.

IRE1 accomplishes this task by cleaving the mRNA at two positions that are then joined together by an RNA ligase. The mRNA produced by this splicing reaction is translated to produce an active transcription regulatory pro tein that increases expression of a subset of the genes of the unfolded protein response (Figure 12–37). The regulated splicing of a cytosolic mRNA by IRE1 is a unique exception to the rule that all mRNA splicing occurs in the nucleus and is catalyzed by the spliceosome.

During normal conditions, IRE1 is maintained in an inactive state by its association with the ER-lumenal chaperone BiP. Elevated levels of misfolded proteins activate IRE1 by a combination of two mechanisms. First, BiP dissociates from IRE1 to bind and protect misfolded proteins from aggregation. Second, misfolded proteins bind to the lumenal domain of IRE1 facilitating the formation of IRE1 oligomers. The oligomerized IRE1 phosphorylates itself on the cytosolic side, activating its ribonuclease domain.

In another example, IRE1 is activated when B cells begin differentiating into antibody-secreting plasma cells. IRE1 activation dramatically expands the ER content of the cell in preparation for the very high levels of immunoglobulins that will soon be assembled there.

Misfolded proteins also activate a second transmembrane kinase in the ER, PERK. The target of activated PERK is a translation initiation protein whose phosphorylation has two consequences. First, translation of new proteins is reduced throughout the cell, thereby reducing the load of proteins that need to be folded in the ER. Second, some proteins are preferentially translated when translation initiation factors are scarce, and one of these is a transcription regu lator that helps activate the transcription of the genes that execute the unfolded protein response.

For example, insulin production increases substantially in pancreatic β cells in response to eating a meal. The elevated demand for the processing capacity of the ER, where insulin is initially assembled, partially acti vates PERK so cells can adjust insulin synthesis rates to avoid overburdening the ER.

The unfolded protein response ultimately increases the production of proteins that improve protein processing in the ER and reduce the burden of misfolded proteins. As homeostasis is restored, the activities of IRE1, PERK, and ATF6 abate. If homeostasis cannot be restored, persistently active signaling from the ER, particularly via PERK, activates genes that initiate apoptosis. In multicellular organisms, it is often less detrimental to eliminate a persistently dysfunctional cell than risk its aberrant interactions with neighboring cells.

Because it could be selectively visualized by silver stains, the Golgi apparatus was one of the first organelles described by early light microscopists. It consists of a collection of flattened, membrane-enclosed compartments called cisternae, that somewhat resemble a stack of pita breads. Each Golgi stack typically consists of four to six cisternae (Figure 13–27), although some unicellular flagellates can have more than 20.

In animal cells, tubular connections between corresponding cisternae link many stacks, thus forming a single complex, which is usually located near the cell nucleus and close to the centrosome (Figure 13–28A). This localization depends on microtubules. If microtubules are experimentally depolymerized, the Golgi apparatus reorganizes into individual stacks that are found throughout the cytoplasm, adjacent to ER exit sites.

In this section, we focus mainly on the Golgi apparatus (also called the Golgi complex). It is a major site of carbohydrate synthesis, as well as a sorting and dispatching station for products delivered to it from the ER. The cell makes many polysaccharides in the Golgi apparatus, including the pectin and hemicellulose of the cell wall in plants and most of the glycosaminoglycans of the extracellular matrix in animals (discussed in Chapter 19).

Each Golgi stack has two distinct faces: a cis face (or entry face) and a trans face (or exit face). Both cis and trans faces are closely associated with special compartments, each composed of a network of interconnected tubular and cisternal structures: the cis Golgi network (CGN) and the trans Golgi network (TGN), respectively. The CGN is a collection of fused vesicular tubular clusters arriving from the ER. Proteins and lipids enter the cis Golgi network and exit from the trans Golgi network, bound for the cell surface or another compartment.

Both networks are important for protein sorting: proteins entering the CGN can either move onward in the Golgi apparatus or be returned to the ER. Similarly, proteins exiting from the TGN move onward and are sorted according to their next destination: endosomes, secretory vesicles, or the cell surface. They also can be returned to an earlier compartment. Some membrane proteins are retained in the part of the Golgi apparatus where they function.

After arrival in the CGN, proteins enter the first of the Golgi processing compartments (the cis Golgi cisterna). They then move to the next compartment (the medial cisterna) and finally to the trans cisterna, where glycosylation is completed. The lumen of the trans cisterna is thought to be continuous with the TGN, the place where proteins are segregated into different transport packages and dispatched to their next destinations.

Proteins are modified in successive stages as they move from cisterna to cisterna across the stack, so that the stack forms a multistage processing unit. Investigators discovered the functional differences between the cis, medial, and trans subdivisions of the Golgi apparatus by localizing the enzymes involved in processing N-linked oligosaccharides in distinct regions of the organelle, both by physical fractionation of the organelle and by labeling the enzymes in electron microscope sections with antibodies (Figure 13–29).

A series of electron micrographs shows the Golgi apparatus (A) unstained, (B) stained with osmium, which preferentially labels the cisternae of the cis compartment, and (C and D) stained to reveal the location of specific enzymes. Nucleoside diphosphatase is found in the trans Golgi cisternae (C), while acid phosphatase is found in the trans Golgi network (D). Note that usually more than one cisterna is stained. The enzymes are therefore thought to be highly enriched rather than precisely localized to a specific cisterna.

The localization of each processing step shown was determined by a combination of techniques, including biochemical subfractionation of the Golgi apparatus membranes and electron microscopy after staining with antibodies specific for some of the processing enzymes. Processing enzymes are not restricted to a particular cisterna; instead, their distribution is graded across the stack, such that early-acting enzymes are present mostly in the cis Golgi cisternae and later-acting enzymes are mostly in the trans Golgi cisternae.

Whereas the ER lumen is full of soluble resident proteins and enzymes, the resident proteins in the Golgi apparatus are all membrane bound. All of the Golgi glycosidases and glycosyl transferases, for example, are single-pass transmembrane proteins, many of which are organized in multienzyme complexes.

The unique architecture of the Golgi apparatus depends on both the microtubule cytoskeleton, as already mentioned, and cytoplasmic Golgi matrix proteins. The Golgi reassembly and stacking proteins (called GRASPs) form a scaffold between adjacent cisternae and give the Golgi stack its structural integrity. Other matrix proteins, called golgins, form long tethers composed of stiff coiled-coil domains with interspersed hinge regions. Golgins form a forest of tentacles that can extend 100–400 nm from the surface of the Golgi stack.

Because transport vesicles arriving from different locations have their characteristic Rab proteins on them, golgins are thought to function as tethers that initially select which part of the Golgi stack a transport vesicle engages (Figure 13–37).

When the cell prepares to divide, mitotic protein kinases phosphorylate the Golgi matrix proteins, causing the Golgi apparatus to fragment and disperse throughout the cytosol. The Golgi fragments are then distributed evenly to the two daughter cells, where the matrix proteins are dephosphorylated, leading to the reassembly of the Golgi stack. Similarly, during apoptosis, proteolytic cleavage of golgins by caspases (discussed in Chapter 18) leads to fragmentation of the Golgi apparatus as the cell self-destructs.

During the most common form of protein glycosylation in the ER, a preformed precursor oligosaccharide (containing 14 sugars composed of 2 N- acetylglucosamines, 9 mannoses, and 3 glucoses) is transferred as a complete unit to proteins. Because this oligosaccharide is transferred to the side-chain NH2 group of an asparagine in the protein, it is said to be N-linked, or asparagine-linked (Figure 12–32A). A special lipid molecule called dolichol (see Panel 2–5, pp. 102–103) anchors the precursor oligosaccharide in the ER membrane.

Almost as soon as a polypeptide chain enters the ER lumen, it is glycosylated on target asparagine amino acids. The precursor oligosaccharide (shown in color) is attached only to asparagine side chains in the sequences Asn-X-Ser and Asn-X-Thr (where X is any amino acid except proline). These sequences occur much less frequently in glycoproteins than in nonglycosylated cytosolic proteins.

Some proteins in the cytosol and nucleus are also glycosylated, but not with large oligosaccharides: they instead carry a much simpler sugar modification, in which a single N-acetylglucosamine group is added to a serine or threonine of the protein.

Two broad classes of N-linked oligosaccharides, the complex oligosaccharides and the high-mannose oligosaccharides, are attached to mammalian glycoproteins. Sometimes, both types are attached (in different places) to the same polypeptide chain. Complex oligosaccharides are generated when the original N-linked oligosaccharide added in the ER is trimmed and further sugars are added; by contrast, high-mannose oligosaccharides are trimmed but have no new sugars added to them in the Golgi apparatus (Figure 13–31).

Both complex oligosaccharides and high-mannose oligosaccharides share a common core region derived by trimming the original N-linked oligosaccharide added in the ER (see Figure 12–32) and typically containing two N-acetylglucosamines (GlcNAc) and three mannoses (Man). The three amino acids constitute the sequence recognized by the oligosaccharyl transferase enzyme that adds the initial oligosaccharide to the protein. Asn, asparagine; Ser, serine; Thr, threonine; X, any amino acid, except proline.

Step 2: Golgi mannosidase I removes three more mannoses. Step 3: N-acetylglucosamine transferase I then adds an N-acetylglucosamine. Step 4: Golgi mannosidase II then removes two additional mannoses. This yields the final core of three mannoses that is present in a complex oligosaccharide. At this stage, the bond between the two N-acetylglucosamines in the core becomes resistant to attack by a highly specific endoglycosidase (Endo H). Because all later structures in the pathway are also Endo H–resistant, treatment with this enzyme is widely used to distinguish complex from high-mannose oligosaccharides.

Whether a given oligosaccharide remains high-mannose or is processed depends largely on its position in the protein. If the oligosaccharide is accessible to the processing enzymes in the Golgi apparatus, it is likely to be converted to a complex form; if it is inaccessible because its sugars are tightly held to the protein’s surface, it is likely to remain in a high-mannose form.

This O-linked glycosylation (Figure 13–33), like the extension of N-linked oligosaccharide chains, is catalyzed by a series of glycosyl transferase enzymes that use the sugar nucleotides in the lumen of the Golgi apparatus to add sugars to a protein one at a time. Usually, N-acetylgalactosamine is added first, followed by a variable number of additional sugars, ranging from just a few to 10 or more.

The Golgi apparatus confers the heaviest O-linked glycosylation of all on mucins, the glycoproteins in mucus secretions, and on proteoglycan core proteins, which it modifies to produce proteoglycans. As discussed in Chapter 19, this process involves the polymerization of one or more glycosaminoglycan chains (long, unbranched polymers composed of repeating disaccharide units; see Figure 19–35) onto serines on a core protein. Many proteoglycans are secreted and become components of the extracellular matrix, while others remain anchored to the extracellular face of the plasma membrane.

The sugars incorporated into glycosaminoglycans are heavily sulfated in the Golgi apparatus immediately after these polymers are made, thus adding a significant portion of their characteristically large negative charge. Some tyrosines in proteins also become sulfated shortly before they exit from the Golgi apparatus.

The N-linked oligosaccharides are by far the most common oligosaccharides, being found on 90% of all glycoproteins. Less frequently, oligosaccharides are linked to the hydroxyl group on the side chain of a serine, threonine, hydroxylysine, or hydroxyproline amino acid. The first sugar of these O-linked oligosaccharides is added in the ER. N-linked and O-linked oligosaccharides undergo extensive pro cessing, modification, and extension in the Golgi apparatus (Chapter 13), producing the diversity of oligosaccharide structures observed on mature glycoproteins.

Note that, as a biosynthetic organelle, the Golgi apparatus differs from the ER: all sugars in the Golgi are assembled inside the lumen from sugar nucleotides, whereas in the ER, the N-linked precursor oligosaccharide is assembled partly in the cytosol and partly in the lumen, and most lumenal reactions use dolichol-linked sugars as their substrates (see Figure 12–33).

There is an important difference between the construction of an oligosaccharide and the synthesis of other macromolecules such as DNA, RNA, and protein. Whereas nucleic acids and proteins are copied from a template in a repeated series of identical steps using the same enzyme or set of enzymes, complex carbohydrates require a different enzyme at each step. The product of each enzyme is recognized as the exclusive substrate for the next enzyme in the series.

N-linked glycosylation promotes protein folding in two ways. First, it has a direct role in making folding intermediates more soluble, thereby preventing their aggregation. Second, the sequential modifications of the N-linked oligosaccharide establish a “glyco-code” that marks the progression of protein folding. This glyco-code is used by chaperones and lectins in the ER to guide protein folding and degradation (discussed in Chapter 12) and by other lectins that guide ER-to-Golgi transport.

The recognition of sugar chains by lectins in the extracellular space is important in many developmental processes and in cell–cell recognition: selectins, for example, are transmembrane lectins that function in cell–cell adhesion during blood-cell migration, as discussed in Chapter 19.

In order to function, the Golgi apparatus must maintain its polarized multicisternal structure while facilitating the transit of a large number of diverse molecules. It is likely that multiple mechanisms are used to transport cargo molecules through the Golgi cisternae while efficiently retaining Golgi resident proteins.

A different way for cargo to move through the Golgi apparatus involves the cisternal maturation mechanism. According to this view, new cis cisternae continually form as vesicular tubular clusters arrive from the ER and fuse with transport vesicles containing Golgi resident proteins and enzymes. As the cargo within a cis cisterna is modified, the enzymes leave in transport vesicles that will fuse with newly arriving vesicular tubular clusters. At the same time, the cisterna accepts transport vesicles containing enzymes from later Golgi cisternae, converting it into a medial cisterna.

(A) In the vesicle transport mechanism, Golgi cisternae are static compartments, which contain a characteristic complement of resident enzymes. The passing of molecules from cis to trans through the Golgi is accomplished by forward-moving transport vesicles, which bud from one cisterna and fuse with the next in a cisto-trans direction. (B) In the cisternal maturation mechanism, each Golgi cisterna matures as it migrates outward through the stack.

It is likely that aspects of both mechanisms are used to varying degrees depending on the type of cell and the nature of cargo molecules that need to be transported. A stable core of long-lasting cisternae might exist in the center of each Golgi cisterna, while regions at the rim may undergo continual maturation, perhaps utilizing Rab cascades that change their identity.

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.

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.

In animal cells they carry a unique marker in the form of mannose 6-phosphate (M6P) groups, which are added exclusively to the N-linked oligosaccharides of these soluble lysosomal enzymes as they pass through the lumen of the cis Golgi network (Figure 13–39). Transmembrane M6P receptor proteins, which are present in the TGN, recognize the M6P groups and bind to the lysosomal hydrolases on the lumenal side of the membrane and to adaptor proteins in assembling clathrin coats on the cytosolic side.

In this way, the receptors help package the hydrolases into clathrin-coated vesicles that bud from the TGN and deliver their contents to early endosomes. The M6P receptor protein binds to M6P at pH 6.5–6.7 in the TGN lumen and releases it at pH 6, which is the pH in the lumen of endosomes.

Cells that are specialized for secreting some of their products rapidly on demand concentrate and store these products in secretory vesicles (often called densecore secretory granules because they have dense cores when viewed in the electron microscope). 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. Clumps of aggregated, electrondense material can be detected by electron microscopy in the lumen of the TGN. The signals that direct secretory proteins into such aggregates are not well defined and may be quite diverse.

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.

Immature secretory vesicles also fuse with one another, and the lumens become progressively more acidic from the increasing concentration of V-type ATPases in the vesicle membrane. Acidification of the lumen further condenses the secretory protein aggregate within a vesicle whose excess membrane has now been retrieved back to the TGN.

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.

Many protein hormones and small neuropeptides, as well as many secreted hydrolytic enzymes, are synthesized as inactive precursors. Proteolysis is necessary to liberate the active molecules from these precursor proteins. The cleavages occur in the secretory vesicles and sometimes in the extracellular fluid after secretion. Additionally, many of the precursor proteins exit the ER with an N-terminal propeptide that is cleaved off only later in the secretory pathway to yield the mature protein.

In other cases, peptide signaling molecules are made as polyproteins that contain multiple copies of the same amino acid sequence. In still more complex cases, a variety of peptide signaling molecules are synthesized as parts of a single polyprotein that acts as a precursor for multiple end products, which are individually cleaved from the initial polypeptide chain. The same polyprotein may be processed in various ways to produce different peptides in different cell types (Figure 13–44).

The initial cleavages are made by proteases that cut next to pairs of positively charged amino acids (Lys-Arg, Lys-Lys, Arg-Lys, or Arg-Arg pairs). Trimming reactions then produce the final secreted products. Different cell types produce different concentrations of individual processing enzymes, so that the same prohormone precursor is cleaved to produce different peptide hormones.

In addition, for secreted hydrolytic enzymes—or any other protein whose activity could be harmful inside the cell that makes it—delaying activation of the protein until it reaches a secretory vesicle or until after it has been secreted has a clear advantage: the delay prevents the protein from acting prematurely inside the cell in which it is synthesized.

Proteins destined for different domains travel together from the ER until they reach the TGN, where they are separated and dispatched in secretory or transport vesicles to the appropriate plasma membrane domain (Figure 13–49). These routes are known as the direct pathways for polarized secretion because cargo destined for the apical and basolateral domains is delivered there directly.

The apical plasma membrane of most epithelial cells is greatly enriched in glycosphingolipids, which help protect this exposed surface from damage; for example, from the digestive enzymes and low pH in sites such as the gut or stomach, respectively. Similarly, plasma membrane proteins that are linked to the lipid bilayer by a glycosylphosphatidylinositol (GPI) anchor (see Figure 12–30) are found predominantly in the apical plasma membrane. If recombinant DNA techniques are used to attach a GPI anchor to a protein that would normally be delivered to the basolateral surface, the protein is delivered to the apical surface instead.

In this route, both apical and basolateral cargo travel together in transport vesicles from the TGN to the basolateral membrane. Membrane proteins that do not belong in that region of the plasma membrane are retrieved by endocytosis and are transported via early endosomes to the correct region. Membrane proteins destined for delivery to the basolateral membrane contain sorting signals in their cytosolic tail.

The lysosome is a terminal destination for the degradation of proteins, microorganisms, dead cells, and other materials ingested by endocytosis and phagocytosis. In addition, proteins, old organelles, and other components in the cytosol can also be degraded in lysosomes through a process termed autophagy.

Lysosomes are membrane-enclosed organelles filled with soluble hydrolytic enzymes that digest macromolecules. Lysosomes contain about 40 types of hydrolytic enzymes, including proteases, nucleases, glycosidases, lipases, phospholipases, phosphatases, and sulfatases. All are acid hydrolases; that is, hydrolases that work best at acidic pH. For optimal activity, they need to be activated by proteolytic cleavage, which also requires an acid environment. The lysosome provides this acidity, maintaining an interior pH of about 4.5–5.0.

Most of the lysosome membrane proteins, for example, are highly glycosylated, which helps to protect them from the lysosome proteases in the lumen. Transport proteins in the lysosome membrane carry the final products of the digestion of macromolecules—such as amino acids, sugars, and nucleotides—to the cytosol, where the cell can either reuse or excrete them. A vacuolar H1 ATPase in the lysosome membrane uses the energy of ATP hydrolysis to pump H+ into the lysosome, thereby maintaining the lumen at its acidic pH (Figure 13

The lysosome H+ pump belongs to the family of V-type ATPases and has a similar architecture to the mitochondrial and chloroplast ATP synthases (F-type ATPases), which convert the energy stored in H+ gradients into ATP (see Figure 11–12). By contrast to these enzymes, however, the vacuolar H+ ATPase exclusively works in reverse, pumping H+ into the organelle. Similar or identical V-type ATPases acidify all endocytic and exocytic organelles, including lysosomes, endosomes, some compartments of the Golgi apparatus, and many transport and secretory vesicles.

(3) Lysosome proteins, including lumenal hydrolases and a membrane-embedded V-type ATPase, are delivered from the TGN to the maturing endosome. (4) The V-type ATPase pumps H+ from the cytosol into the endosome lumen and further acidifies the organelle. Crucially, the increasing acidity that accompanies maturation renders lysosomal hydrolases increasingly more active, influencing many receptor–ligand interactions, thereby controlling receptor loading and unloading.

They contain unique membrane proteins and a wide variety of soluble hydrolytic enzymes that operate best at pH 5, which is the internal pH of lysosomes. An ATPdriven H1 pump in the lysosomal membrane maintains this low pH.

In animal cells they carry a unique marker in the form of mannose 6-phosphate (M6P) groups, which are added exclusively to the N-linked oligosaccharides of these soluble lysosomal enzymes as they pass through the lumen of the cis Golgi network (Figure 13–39). Transmembrane M6P receptor proteins, which are present in the TGN, recognize the M6P groups and bind to the lysosomal hydrolases on the lumenal side of the membrane and to adaptor proteins in assembling clathrin coats on the cytosolic side.

Because all glycoproteins leave the ER with identical N-linked oligosaccharide chains, the signal for adding the M6P units to oligosaccharides must reside somewhere in the polypeptide chain of each hydrolase. Genetic engineering experiments have revealed that the recognition signal is a cluster of neighboring amino acids on each protein’s surface, known as a signal patch (Figure 13–41). Because most lysosomal hydrolases contain multiple oligosaccharides, they acquire many M6P groups, providing a high-affinity signal for the M6P receptor.

Sorting signals are usually composed of amino acid side chains in a protein and come in two general varieties: a linear sequence of amino acids (called a signal sequence) or a specific three-dimensional arrangement of amino acids (called a signal patch). Sorting signals for protein translocation into organelles are linear signal sequences, while examples of linear signals and signal patches are known for nuclear and vesicular transport.

The M6P receptor protein binds to M6P at pH 6.5–6.7 in the TGN lumen and releases it at pH 6, which is the pH in the lumen of endosomes. Thus, after the receptor is delivered, the lysosomal hydrolases dissociate from the M6P receptors, which are retrieved into transport vesicles that bud from endosomes. These vesicles are coated with retromer, a coat protein complex specialized for endosome-to-TGN transport, which returns the receptors to the TGN for reuse (Figure 13

Not all the hydrolase molecules that are tagged with M6P get to lysosomes. Some escape the normal packaging process in the trans Golgi network and are transported by the constitutive secretory pathway to the cell surface, where they are secreted into the extracellular fluid. Some M6P receptors, however, also take a detour to the plasma membrane, where they recapture the escaped lysosomal hydrolases and return them by receptor-mediated endocytosis (discussed later) to lysosomes via early and late endosomes.

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). This conversion process changes the protein composition of the endosome membrane, patches of which invaginate and become incorporated within the organelles as intralumenal vesicles, while the endosome itself moves from the cell periphery to a location close to the nucleus. As an endosome matures, it ceases to recycle material to the plasma membrane.

Early endosomes are mainly derived from incoming endocytic vesicles that fuse with one another (Movie 13.8). Typically, an early endosome receives incoming vesicles for about 10 minutes before beginning its maturation into a late endosome. Early endosomes have tubular and vacuolar domains (see Figure 13–55). Most of the membrane surface is in the tubules, and most of the volume is in the vacuolar domain. Many changes occur during the maturation process.

(1) The endosome changes shape and location as the tubular domains are mostly recycled back to the plasma membrane, the vacuolar domains are thoroughly modified, and the endosome is moved by motors along microtubules toward the nucleus. (2) Rab proteins drive changes in phosphoinositide lipids and fusion machinery (SNAREs and tethers) on the cytosolic face of the endosome membrane to change the functional characteristics of the organelle.

Most of these events occur gradually but eventually lead to a complete transformation of the endosome into an early endolysosome. In addition to committing selected cargo for degradation, the maturation process is important for lysosome maintenance. The continual delivery of lysosome components from the TGN to maturing endosomes ensures a steady supply of new lysosome proteins. The endocytosed materials mix in early endosomes with newly arrived acid hydrolases.

As endosomes mature, patches of their membrane invaginate into the endosome lumen and pinch off to form intralumenal vesicles. Because of their appearance in the electron microscope, such maturing endosomes are also called multivesicular bodies (Figure 13

Lysosomes are found in all eukaryotic cells. They were initially discovered by the biochemical fractionation of cell extracts; only later were they seen clearly in the electron microscope. Although extraordinarily diverse in shape and size, staining them with specific antibodies shows they are members of a single family of organelles. They can also be identified by histochemical techniques that reveal which organelles contain acid hydrolase (Figure 13

Late endosomes containing material received from the plasma membrane by endocytosis and containing newly synthesized lysosomal hydrolases fuse with preexisting lysosomes to form structures that are referred to as endolysosomes, which then fuse with one another (Figure 13–64). When the majority of the endocytosed material within an endolysosome has been digested so that only resistant or slowly digestible residues remain, these organelles become “classical” lysosomes. These are relatively dense, round, and small, but they can enter the cycle again by fusing with late endosomes or endolysosomes. Thus, there is no real distinction between endolysosomes and lysosomes: they are the same except that they are in different stages of a maturation cycle. For this reason, lysosomes are sometimes viewed as a heterogeneous collection of distinct organelles, the common feature of which is a high content of hydrolytic enzymes.

Lysosomes are the end product of endosome maturation. Cell-surface proteins that are endocytosed into endosomes are degraded in lysosomes unless they are retrieved back to the plasma membrane.

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.

Genetic defects that affect one or more of the lysosomal hydrolases cause a number of human lysosomal storage diseases. The defects result in an accumulation of undigested substrates in lysosomes, with severe pathological consequences, most often in the nervous system. In most cases, there is a mutation in a structural gene that codes for an individual lysosomal hydrolase. This occurs in Hurler’s disease, for example, in which the enzyme required for the breakdown of certain types of glycosaminoglycan chains is defective or missing.

The most severe form of lysosomal storage disease, however, is a very rare inherited metabolic disorder called inclusion-cell disease (I-cell disease). In this condition, almost all of the hydrolytic enzymes are missing from the lysosomes of many cell types, and their undigested substrates accumulate in these lysosomes, which consequently form large inclusions in the cells. The consequent pathology is complex, affecting all organ systems, skeletal integrity, and mental development; individuals rarely live beyond 6 or 7 years.

In individuals with I-cell disease, all the hydrolases missing from lysosomes are found in the blood: because they fail to sort properly in the Golgi apparatus, they are secreted rather than transported to lysosomes. The mis-sorting has been traced to a defective or missing GlcNAc phosphotransferase. Because lysosomal enzymes are not phosphorylated in the cis Golgi network, the M6P receptors do not segregate them into the appropriate transport vesicles in the TGN. Instead, the lysosomal hydrolases are carried to the cell surface and secreted.

In I-cell disease, the lysosomes in some cell types, such as hepatocytes, contain a normal complement of lysosomal enzymes, implying that there is another pathway for directing hydrolases to lysosomes that is used by some cell types but not others. Alternative sorting receptors function in these M6P-independent pathways. Similarly, an M6P-independent pathway in all cells sorts the membrane proteins of lysosomes from the TGN for transport to late endosomes, and those proteins are therefore normal in I-cell disease.

Lysosomes are meeting places where several streams of intracellular traffic converge. A route that leads outward from the ER via the Golgi apparatus delivers most of the lysosome’s digestive enzymes, as we discussed earlier. In addition, at least three paths from the cell surface and extracellular space feed substances into lysosomes for digestion, while a fourth route called autophagy originates in the cytoplasm and is used to digest intracellular macromolecules and organelles.

We have already discussed how macromolecules taken up from plasma membrane and extracellular fluid by endocytosis can reside in endosomes until they mature and fuse with lysosomes. A second pathway called macropinocytosis specializes in the nonspecific uptake of fluids, membrane, and particles attached to the plasma membrane. A third pathway found in phagocytic cells, such as macrophages and neutrophils in vertebrates, is dedicated to the engulfment, or phagocytosis, of large particles and microorganisms to form phagosomes. In contrast to these routes originating from the plasma membrane, autophagy is used to digest cytosol, worn-out organelles, and microbes that invade the cytosol.

All eukaryotic cells can carry out a process called autophagy, or “self-eating.” During autophagy, a portion of the cytoplasm is engulfed into a membrane structure called the autophagosome that subsequently fuses with the lysosome where the autophagosome’s contents are degraded (Figure 13–71). Autophagy can be either nonselective or selective. In nonselective autophagy, a bulk portion of cytoplasm is sequestered in autophagosomes. In selective autophagy, autophagosomes tightly enclose specific cargo and mostly exclude the surrounding cytosol.

The autophagosome assembles by the fusion of small vesicles of unknown origin. The process begins when a phosphoinositide lipid kinase complex (termed the ATG1 complex) locally produces PI(3)P (see Figure 13–10) to mark a potential membrane site for the recruitment of several autophagy-related factors. These factors catalyze the covalent attachment of the membrane lipid phosphatidylethanolamine to a ubiquitin-like protein called ATG8. The ATG8-marked vesicles undergo homotypic fusion with each other and heterotypic fusion with vesicles containing ATG9. This results in expansion of a membrane whose identity is provided by ATG8 and other autophagy-related proteins.

For reasons that are not known, the growing membrane structure formed by vesicle fusion is not spherical. Instead, it is a flattened disc (similar to a Golgi apparatus cisterna) that curls into a cup-shaped structure. Fusion of the lips of this cup encloses the contents inside a compartment that is now surrounded by two membranes. The outer membrane of this newly formed autophagosome then fuses with the lysosome in a SNARE-mediated process. The inner membrane and its enclosed cargo are released into the lysosome where they are degraded by the acid hydrolases.

The activity of the ATG1 kinase complex that initiates autophagosome formation is tightly regulated. Most of the time, it is kept inactive because of phosphorylation by another protein kinase called the mTOR complex 1 (discussed in Chapter 15). The activity of mTOR complex 1 is dependent on the availability of certain amino acids generated by the recycling of proteins in the lysosome. When these amino acids become limiting, mTOR complex 1 activity is reduced, relieving its inhibition of the ATG1 complex. Activation of the ATG1 complex initiates nonselective autophagy to degrade bulk cytosol in lysosomes, thereby generating amino acids that activate the mTOR complex 1.

Starvation-induced nonselective autophagy is particularly important in mammals in the hours immediately after birth. During this transition period, the constant nutrient supply from the womb is abruptly lost, and feeding by mouth has not yet started.

Selective autophagy can be used to degrade bacteria and viruses that invade the cytosol, as well as damaged proteins, protein aggregates, and damaged whole organelles. The range of cargoes degraded by autophagy explains why dysregulated autophagy contributes to diseases ranging from infectious disorders to neurodegeneration and cancer.

Selective autophagy mediates the degradation of invading microbes, damaged or otherwise unwanted organelles, and protein complexes or aggregates that are too large for degradation by the proteasome. The steps of autophagosome formation and its fusion with lysosomes are the same as in nonselective autophagy. Selective autophagy differs only at the very early stages when the cargo destined for degradation is recruited to the concave surface of the forming autophagosome membrane. This recruitment is mediated by specialized receptor proteins that recognize the cargo and have a binding site for the autophagosome-specific protein ATG8.

In most cases, autophagy receptors recognize their cargo via a mark that is acquired only when the cargo is destined for degradation. The most commonly used mark is ubiquitin. For example, bacteria that escape from phagosomes and invade the cytosol are recognized by cytosolic proteins that ubiquitylate proteins on the bacterial surface. Several cargo receptors recognize the ubiquitin and other bacteria-specific proteins and recruit ATG8-containing vesicles. These vesicles fuse together, and the forming autophagosome effectively zippers around the cargo to engulf it without trapping bulk cytosol in the process (Figure 13

Damaged mitochondria cannot maintain the gradient, so protein import is blocked. As a consequence, a protein kinase called Pink1, which is normally imported into mitochondria, is instead retained on the mitochondrial surface where it recruits the ubiquitin ligase Parkin from the cytosol. Parkin ubiquitylates mitochondrial outer membrane proteins, which serves as a mark for a ubiquitin-dependent cargo receptor for autophagy. Mutations in Pink1 or Parkin cause a form of early-onset Parkinson’s disease, a degenerative disorder of the central nervous system.

Peroxisomes are major sites of oxygen utilization and are found in virtually all eukaryotic cells. They contain oxidative enzymes, such as catalase and urate oxidase, at such high concentrations that, in some cells, the peroxisomes stand out in electron micrographs because of the presence of a crystalloid protein core (Figure 12

Peroxisomes are so named because they usually contain one or more enzymes that use molecular oxygen to remove hydrogen atoms from specific organic sub strates (designated here as R) in an oxidation reaction that produces hydrogen peroxide (H2O2):

Catalase uses the H2O2 generated by other enzymes in the organelle to oxidize a variety of substrates—including formic acid, formaldehyde, and alcohol—by the “peroxidation” reaction: H2O2 + R′H2 → R′ + 2H2O. This type of oxidation reaction is particularly important in liver and kidney cells, where the peroxisomes detoxify various harmful molecules that enter the bloodstream. About 25% of the ethanol we drink is oxidized to acetaldehyde in this way.

A major function of the oxidation reactions performed in peroxisomes is the breakdown of fatty acid molecules. The process, called b oxidation, shortens the alkyl chains of fatty acids sequentially in blocks of two carbon atoms at a time, thereby converting the fatty acids to acetyl CoA. The peroxisomes then export the acetyl CoA to the cytosol for use in biosynthetic reactions. In mammalian cells, β oxidation occurs in both mitochondria and peroxisomes; in fungi and plant cells, however, this essential reaction occurs exclusively in peroxisomes.

An essential biosynthetic function of animal peroxisomes is to catalyze the first reactions in the formation of plasmalogens. This abundant class of phospho lipids is found in all human cells but is particularly enriched in brain, where it is a major constituent of myelin (Figure 12

The other type of peroxisome is present in germinating seeds, where it converts the fatty acids stored in seed lipids into the sugars needed for the growth of the young plant. Because this conversion of fats to sugars is accomplished by a series of reactions known as the glyoxylate cycle, these per oxisomes are also called glyoxysomes. In the glyoxylate cycle, two molecules of acetyl CoA produced by fatty acid breakdown in the peroxisome are used to make succinic acid, which then leaves the peroxisome and is converted into glucose in the cytosol. The glyoxylate cycle does not occur in animal cells, and animals are therefore unable to convert fats into carbohydrates.

Yeasts grown on sugar, for example, have a few small peroxisomes. But when some yeasts are grown on methanol, numerous large peroxisomes are formed that oxidize methanol; and when grown on fatty acids, they develop numerous large peroxisomes that break down fatty acids to acetyl CoA by β oxidation.

Proteins destined for mitochondria have signal sequences of yet another type, in which positively charged amino acids alternate with hydrophobic ones. The signal for protein import into the nucleus is composed primarily of positively charged amino acids. Finally, many proteins destined for peroxisomes have a signal sequence of three characteristic amino acids at their C-terminus. A sorting signal for any particular destination needs to be sufficiently distinctive from all other sequences to permit its selective recognition by the appropriate sorting receptor.

The proteins that compose peroxisomes are delivered by two different routes (Figure 12–46). In the first route, some of the integral membrane proteins of the peroxisomal membrane are first inserted into the ER using the ER-resident Sec61 protein translocator. These peroxisome-destined proteins are then pack aged into specialized peroxisomal precursor vesicles.

In the second route, peroxisomal proteins can be imported into preexisting peroxisomes directly from the cytosol. A specific sequence of three amino acids (Ser-Lys-Leu) located at the C-terminus of many peroxisomal proteins functions as an import signal (see Figure 12–13). Other peroxisomal proteins contain a slightly longer and partially hydrophobic signal sequence near the N-terminus. If either sequence is attached to a cytosolic pro tein, the protein is imported into peroxisomes.

Peroxisomal protein import is driven by ATP hydrolysis and utilizes a col lection of proteins, called peroxins, that catalyze the import cycle. C-terminal peroxisomal sorting signals are recognized by the peroxin Pex5 in the cytosol. This import receptor accompanies its cargo all the way into a protein translocator in the peroxisomal membrane. After cargo release inside the peroxisome, Pex5 is recycled back to the cytosol. This recycling step requires modification of Pex5 with ubiquitin, which is used as a handle by an ATPase complex composed of Pex1 and Pex6. The Pex1–Pex6 complex harnesses the energy of ATP hydroly sis to release Pex5 from peroxisomes so it can pick up the next cargo molecule.

N-terminal peroxisomal signal sequences are recognized by the peroxin Pex7. The Pex7–cargo complex, together with additional accessory peroxins, appear to participate in an import cycle similar to that mediated by Pex5. The protein translocator in the peroxisomal membrane is composed of at least six different peroxins. Unlike protein translocators in the ER, the peroxi somal translocator can transport fully folded and even oligomeric proteins across the membrane.

Peroxisomal precursor vesicles bud from the ER. At least two peroxisomal membrane proteins, Pex3 and Pex15, follow this route. The machinery that drives the budding reaction and that selects only peroxisomal proteins for packaging into these vesicles depends on Pex19 and other cytosolic proteins that are still unknown. Peroxisomal precursor vesicles may then fuse with one another or with preexisting peroxisomes.

The importance of protein import into peroxisomes is demonstrated by the inherited human disease Zellweger syndrome. Mutations in any of a dozen differ ent peroxins, the most common being Pex1, cause an impairment in peroxisomal protein import. These individuals, whose cells contain “empty” peroxisomes, accumulate very-long-chain and branched-chain fatty acids that are normally broken down in peroxisomes. Furthermore, they are deficient in plasmalogens. These metabolic impairments cause severe abnormalities in the brain, liver, and kidneys of individuals, and they die soon after birth.

The evolutionary origin of peroxisomes is not firmly established, but they are generally thought to represent a specialized offshoot of the membrane system that composes the secretory and endocytic pathways. One hypothesis is that peroxisomes are a vestige of an ancient organelle that performed all the oxygen metabolism in the primitive ancestors of eukaryotic cells.

The major intracellular compartments in eukaryotic cells can therefore be categorized into three distinct families: (1) the nucleus and the cytosol, which are topologically equivalent (although functionally distinct) and connected by nuclear pore complexes; (2) all organelles that function in the secretory and endocytic pathways—including the ER, Golgi apparatus, endosomes, lyso somes, and the transport vesicles that move between them—and peroxisomes; (3) the endosymbiont-derived organelles: mitochondria and the plastids (in plants only).

Lysosomes contain digestive enzymes that degrade defunct intracellular organelles, as well as macro molecules and particles taken in from outside the cell by endocytosis. On the way to lysosomes, endocytosed material must first pass through a series of organelles called endosomes.

Finally, peroxisomes are small vesicular compartments that contain enzymes used in various oxidative reactions. On average, the membrane-enclosed compartments together occupy nearly half the volume of a cell (Table 12