Frixos Paraskevas
The discovery of phagocytosis by Elie Metchnikoff marks the birth of immunology (1). As with many other great ideas in the history of science, its origin and evolution can be traced to questions totally unrelated to the subsequent demonstration that phagocytosis plays a major role in defense against intruders in multicellular organisms. Metchnikoff was a zoologist with research in embryology. His early research was concerned with the identification of primordial embryologic structures and functions that could link them to the evolution of species; in other words, he was searching for a link between ontogeny and phylogeny. Digestion was considered one of the most ancient functions—it existed even in unicellular organisms and was also detected among cells of primitive embryologic layers.
These early investigations led to the observations that mobile cells of the primitive digestive tract of the starfish larvae were engulfing foreign material, such as carmine droplets, that Metchnikoff introduced into the larvae. Because carmine had no nutritive value, as he later described the moment, “a new thought suddenly flashed across my brain. It struck me that similar cells might serve in the defense of the organism against intruders.” The now famous Messina experiment provided proof for his new idea. He introduced thorns from his Christmas tangerine tree to starfish larvae, and, after anxiously waiting all night for his results, he observed the next morning that several mobile cells surrounded the thorns. This experiment was followed by observations that, if the spores of the Monospora bicuspidata that had entered the body of Daphniawere quickly engulfed by wandering cells, Daphnia survived; otherwise, the organism died. These results led him to believe that the function of the ameboid cells was important in the defense against infection. The word phagocytosis was suggested to him by Dr. Claus, a professor of zoology at Vienna, defining the function of “devouring cells” (from the Greek phago, eat).
In multicellular organisms, the cells that are involved in phagocytosis have been called professional phagocytes by Rabinovitch. The most widely recognized phagocytes are the polymorphonuclear leukocytes and the mononuclear phagocytes (monocytes and macrophages). A number of mechanisms have been identified by which extracellular material gains entry into the cell (2). The term endocytosis is used to characterize the uptake of macromolecules in solution, and it can be subdivided into clathrin-mediated endocytosis, non–clathrin-mediated endocytosis, and macropinocytosis (3). Clathrin-mediated endocytosis is the best characterized (4). Clathrin consists of three heavy and three light chains (triskelions) forming lattices of hexagons and pentagons. In addition to clathrin, the endocytic vesicles are also coated with adaptors such as adaptor protein 2, can assemble into empty coats or cages, and produce lattice-coated vesicles. The formation of the endocytic vesicle is regulated by a guanosine triphosphatase (GTPase), dynamin. Clathrin-mediated endocytosis is used by certain macromolecules with well-defined receptors present in the clathrin-coated pit or vesicle in the cell membrane, such as receptors for transferrin, low-density lipoprotein, and epidermal growth factor. Non–clathrin-mediated endocytosis is carried out by caveolae and non–clathrin-coated vesicles. Macropinocytosis is a form of endocytosis that results in the formation of large vesicles, usually at the leading edge of the cell membrane where ruffling occurs. It is used by antigen-presenting cells to capture antigen for presentation in the context of major histocompatibility molecules. These various forms of endocytosis involve uptake of fluids, a process that was known as pinocytosis (from the Greek pino, drink). Phagocytosis is endocytosis of large particles (<1 μm). Phagocytosis differs from pinocytosis not only in terms of the size of the material to be ingested but also in its temperature dependence (it does not occur below 18°C) and its requirement of an intact cytoskeletal system.
It is helpful to consider the morphologic aspects of phagocytosis in four stages: (a) recognition, (b) attachment and opsonization, (c) ingestion, and (d) digestion.
Recognition
In order for the particle to be removed by phagocytes, it must express certain ligands that can be recognized by the phagocyte receptors. Such receptors will trigger engulfment and subsequent disposal of the phagocytosed particle. Furthermore, this initial interaction defines the consequences of phagocytosis, i.e., whether it will be followed by inflammatory reaction or not. Recognition depends on the expression of the pathogens of certain ligands that have been called “pathogen-associated molecular patterns” (PAMPs) or, more generally, “molecular patterns” (MPs) (5). A prototype of MP-recognizing receptors is the macrophage mannose receptor (MMR), which recognizes glycoproteins with mannose and fucose residues and belongs to the large family of C-lectins that require calcium for their function. The C-type lectin family includes several members such as DC-SIGN (Dendritic Cell-Specific Intercellular adhesion molecule-3 Grabbing Nonintegrin), DEC-205 (CD205) langerin (a C-type lectin of Langerhans cells), and several others. Several of these receptors may participate concomitantly in the recognition of the ligands on the surface of an infectious agent, thus enlarging the diversity of recognition. The concept of phagocytosis originally involved the removal of infectious agents. In recent years, however, removal of apoptotic cells was shown to be another function of the phagocytic cells. Apoptosis may occur following an insult, such as viral infection, or constitutively as a result of senescence. It appears that phagocytes are able to discriminate between these two mechanisms of death. Only in the former case does a T-cell response follow, and not in the latter. Of course, it may be claimed that it is the viral particles within the dead cell that trigger a T-cell response. Multiple receptors are involved in triggering phagocytosis when they are engaged at the same time, cooperatively or sequentially, perhaps forming a phagocytic synapse, analogous to the T-cell synapse for antigen recognition (5).
However, some of the scavenger receptors such as CD36 or scavenger receptor A (SRA), both of which are involved in removal of apoptotic cells as well as in response to pathogens, are able to discriminate between the two (6).
Attachment: Role of Opsonins
Attachment of foreign particles on the phagocytic surface is facilitated greatly by substances known as opsonins, discovered by Almroth Wright and immortalized in The Doctor’s Dilemma by George Bernard Shaw. The best-known opsonins are immunoglobulin (Ig) G antibodies, complement components (or their activation products), and certain oligosaccharides terminating in mannose or galactose.
The various opsonins bind to the surface of the phagocyte by specific receptors: Fc receptors (FcRs) for the IgG (FcγR), complement receptors (CRs) for complement components, and the mannose receptor for the mannose-terminating oligosaccharides.
Fcγ Receptors
There are three FcγRs: FcγRI (CD64), FcγRII (CD32), and FcγRIII (CD16). Their genes have been mapped on chromosome 1 at q21-23. Many isoforms exist; some are encoded by distinct genes, whereas others are generated as a result of alternative splicing (7,8). Eight genes have been identified so far, three for FcγRI (A, B, and C), three for FcγRII (A, B, and C), and two for FcγRIII (A and B). All FcRs have a ligand-binding polypeptide chain known as α chain, and, in some, the α chain is associated with homo-dimers of the γ chain or heterodimers of the γ and ζ chains (discussed later). The α-chain belongs to the Ig superfamily with conserved extracellular regions, which contain two (FcγRII and FcγRIII) or three (FcγRI) Ig domains. Their cytoplasmic regions are distinct, suggesting that they may be involved in different functions.
One important characteristic feature of their cytoplasmic region is the presence of conserved tyrosine (Y)-containing sequences YXXL (where X is any amino acid and L is leucine) known as ITAM (Immunoreceptor Tyrosine-Based Activation Motif), which is found in components of B-cell and T-cell receptors. The ITAM motif consists of two YXXL sequences separated by nonconserved amino acids and, in its full-length form, contains up to 25 residues (9). This sequence plays a central role in signal transduction of several receptors of the cells of the immune system.
The FcγRI (CD64) is found on monocytes and macrophages and binds monomeric IgG with high affinity (Ka = 5 · 108 mol/L). When present alone, it does not mediate phagocytosis, although it binds IgG-coated sheep erythrocytes. The binding site of the IgG is located in the NH2-terminal region of the CH2 domain of the IgG. IgG1 and IgG3 subclasses bind with higher affinity than IgG4 and IgG2. The α chain of FcγRI lacks the ITAM motif and can mediate phagocytosis even when it totally lacks its cytoplasmic region, provided that it is associated with the γ chain that contains ITAM motifs (10). On monocyte and macrophage cell lines, the FcγRI is associated with the γ chain (11).
The FcγRII (CD32) is a low-affinity receptor and does not bind monomeric IgG. Its expression on neutrophils is low (10,000 to 40,000 per cell) and shows subclass specificity for IgG dimers (IgG3 > IgG1 = IgG2 ≫ IgG4). The CH2 and CH3 domains of the IgG may be involved in the binding. FcγRIIA is expressed on neutrophils, monocytes, and macrophages, whereas the FcγRIIB is expressed only on B lymphocytes (12,13). The FcγRIIA isoform contains two YXXL sequences (i.e., one complete ITAM) and effectively induces phagocytosis, whereas the FcγRIIB contains only a single cytoplasmic YXXL sequence and does not induce a phagocytic signal unless a second YXXL sequence is inserted by transfection (14). The position of the two sequences and the number of amino acids separating them play a role in phagocytosis.
The FcγRIIIA (CD16) is also a low-affinity receptor and consists of an α chain (the ligand binding) and a homodimer of γ chains or a heterodimer of γ and ζ subunits (15,16). The γ subunit is important not only for the induction of the phagocytic signal but also for the surface expression of FcγRIII. The FcγRIIIB is found on neutrophils and is anchored to the outer leaflet of the plasma membrane by a glycosylphosphatidylinositol group, whereas the FcγRIIIA is a transmembrane protein with a cytoplasmic domain and is expressed on natural killer cells (17). The most critical difference between the two FcγRIII proteins is located in amino acid 203, where a serine in FcγRIIIB determines the glycosylphosphatidylinositol anchor, whereas a phenylalanine in the FcγRIIIA directs for the trans-membrane form.
|
Table 13.1 Fcγ Receptor (FcγR) Expression |
||||||||||||||||||||||||||||||||||||||||||||||||||||||||
|
||||||||||||||||||||||||||||||||||||||||||||||||||||||||
The expression of FcγR can be constitutive or inducible by cytokines. The resting neutrophils do not express FcγRI but express between 10,000 and 40,000 copies of FcγRII and 100,000 to 300,000 copies of FcγRIII (18).
Soluble forms have been demonstrated for all three FcγRs, which are generated by several mechanisms. Serum contains high levels of FcγRIIIB, probably released from polymorphonuclear cells by serine protease activity (19). Other mechanisms involve alternative splicing (FcγRIIA) or stop codons (FcγRI). It becomes apparent that FcRs in general exert a wide range of biologic functions within the network of immunologic regulation and function (20). The expression of the various forms of FcγR and their functions are summarized in Tables 13.1 and 13.2, respectively.
Complement Receptors
Certain complement components and their activation-induced fragments bind to receptors present on phagocytes and enhance phagocytosis. The best studied are the C3b receptor (CR1; CD35), the C3d receptor (CR2; CD21), and the receptor for the inactive fragments of C3b, that is, iC3b (CR3; CD11b and CR4; CD11c). Under certain conditions, some of them mediate ingestion of complement-coated particles. Opsonization by C3 is necessary for phagocytosis to occur and although by itself does not induce ingestion, it enhances IgG-mediated phagocytosis by 20-fold. CRs, however, can promote both attachment and ingestion in macrophages activated under the influence of T-cell–derived lymphokines, such as macrophage colony-stimulating factor and interleukin (IL)-4 (21). The ability of C3b receptors to promote phagocytosis relates to their capacity to be redistributed on the surface of the phagocyte, and lymphokines may do precisely this (22). This lateral mobility of the CRs may permit their engagement with the intracellular machinery that is involved in engulfment, and is induced by soluble (23) or immobilized immune complexes (24).
|
Table 13.2 Fcγ Receptor (FcγR) Function |
||||||||||||||||||||||||||||||
|
||||||||||||||||||||||||||||||
A receptor for C1q also mediates phagocytosis (24), and of the three cell-surface C1q-binding proteins, two (60 kDa and 126 kDa) bind to the “collagenlike” region of C1q (cC1qR), whereas the third (33 kDa) binds to the globular heads (gC1qR). The 60-kDa cC1qR is structurally related to the Ca-binding protein, calreticulin (25), and binds also to mannose-binding lectin and pulmonary surfactant protein A (26), all of which enhance phagocytosis (27). Mannose-binding lectin and pulmonary surfactant protein A have primary structures similar to C1q, that is, they possess collagenous and globular domains.
The mannose-binding lectin, more commonly known as the mannose receptor, is a C-type lectin expressed by macrophages and dendritic cells (28) and is involved in uptake of mycobacteria and other pathogens as a result of binding to the lipoglycan, lipoarabinomannan (29,30). Phagocytosed pathogens are directed to the endocytic pathway, and the lipoarabinomannan antigens associated with these pathogens are presented to T cells in the context of CD1b molecules (31) (see Chapter 17), linking innate and adaptive immunity.
Macrophages also express high-affinity receptors for lipopolysaccharides, the lipopolysaccharide receptor (CD14) (32). This receptor enhances ingestion of Gram-negative bacteria (33) and triggers release of cytokines, especially tumor necrosis factor-α.
Opsonization of pathogens (coated with antibody or complement) markedly facilitates attachment and subsequent ingestion and is particularly important for encapsulated bacteria, such as pneumococci, that resist phagocytosis. Capsules are usually very hydrophilic and prevent the approach of the phagocyte. Opsonization with antibody or complement, or both combined, results in bacterial clearance by several logs more than without opsonization or opsonization with either opsonin alone. It is interesting that the subclasses of IgG that bind strongly to the FcγR also fix complement well. Nonencapsulated bacteria can be ingested without opsonization through receptors that bind latex particles and zymosan.
Ingestion
After attachment, the phagocyte extends pseudopodia that encircle the particle and eventually enclose it within a vesicle known as aphagosome. Phagocytosis stimulated by one particle does not induce ingestion of other particles attached to the same phagocyte that by themselves would not be phagocytosed (34). In other words, the response of the membrane to the phagocytic stimulus is segmental, not generalized. This segmental response engages the receptors of the phagocyte with the respective ligand on the particle. Successive ligand–receptor engagements lead to the extension of broad lamellipodia of the phagocytic membrane over the particle, forming a phagocytic cup (Fig. 13.1). Eventually, the advancing ends of the lamellipodia fuse and enclose the particle within the phagosome. This sequential and circumferential ligand–receptor interaction has been called the zipper hypothesis of phagocytosis (35) (Fig. 13.2). The validity of the zipper mechanism is shown by the inability of macrophages to phagocytose lymphocytes when their surface Ig is capped by anti-Ig antibodies. Lymphocytes treated with anti-Ig antibodies can be phagocytosed if their entire surface is coated with the anti-Ig antibody (36) (Fig. 13.3). However, if capping of surface Ig is allowed to take place (i.e., the Ig–anti-Ig complexes have moved to one pole of the cell), the phagocyte is firmly attached to the cap, but pseudopodia do not extend to the remaining surface of the cell.
Signal Transduction by Phagocytic Receptors
Cross-linking of the Fc receptors initiates a series of well-orchestrated and highly complex series of molecular interactions, resulting in the extension of pseudopodia, engulfment of the particle, which then is drawn to the cell interior for digestion (37).
|
|
|
Figure 13.1. Phagocytosis of a red blood cell. Scanning electron micrograph of phagocytosis of an immunoglobulin G–coated red blood cell. Lamellipodia extending from the macrophage over the red cell form a “cup” that engulfs the red cell. (From Kay MM. Hodgkin’s disease: a war between T-lymphocytes and transformed macrophages? Recent Results Cancer Res 1976;56:111–121, with permission.) |
|
|
|
Figure 13.2. The “zipper” mechanism of phagocytosis. A phagocyte establishes contact with an immunoglobulin G (IgG)–opsonized particle through interaction of the Fcγ receptor (FcR) of the phagocyte with the IgG opsonin. Signals generated by this interaction lead to actin polymerization and the formation of pseudopodia. New contacts between the FcγR on the advancing pseudopodia and the IgG opsonin lead to further advancement of the pseudopodia and additional contacts resulting in progressive encircling of the opsonized particle and the formation of the phagosome. This stepwise binding of the ligands (opsonins) to their respective receptors is known as the zipper mechanism of phagocytosis. Partially opsonized particles are not ingested (Fig. 13.3). |
|
|
|
Figure 13.3. Phagocytosis of opsonized and partially opsonized lymphocytes. A: Mouse lymphocytes were treated with rabbit immunoglobulin IgG anti-mouse Ig followed by sheep IgG antirabbit-F(ab′)2 antibody. The peroxidase reaction shows that these lymphocytes have surface Ig distributed around the entire surface of the cell. As shown by electron microscopy (a) and phase microscopy (b), these lymphocytes are phagocytosed by the macrophages. B: Lymphocytes were first allowed to cap their surface Ig and then were treated with the sheep antirabbit-F(ab′)2antibody. The macrophage is attached to the lymphocyte only in the area of the cap, whereas pseudopodia are unable to extend beyond the cap over the lymphocyte surface that has lost the Ig–anti-Ig complexes. Peroxidase-labeled antibodies are seen in vacuoles (asterisks) and tubular invaginations of the membrane (arrows). (From Griffin FM Jr, Griffin JA, Silverstein SC. Studies on the mechanism of phagocytosis. II. The interaction of macrophages with anti-immunoglobulin IgG-coated bone marrow-derived lymphocytes. J Exp Med 1976;144:788, with permission.) |
Six members of the Src family have been identified in phagocytes: Fgr, Fyn, Hck, Lyn, Ycs, and Src. Some of these are associated with specific receptors. These kinases are important for the early phosphorylation events after FcγR engagement (38). There is a high degree of redundancy among these kinases, because significant interference of phagocytosis requires inactivation of several members (39).
Upon stimulation with IgG-coated particles, these kinases are activated and are redistributed to actin-rich phagocytic caps (Fig. 13.4, part 1). The ITAMs of the FcRs are phosphorylated by Src kinases (40) and become docking sites for Syk kinase, which has a critical role in phagocytosis because Syk-/- macrophages cannot internalize IgG-opsonized particles (41).
Syk phosphorylates the p85 regulatory subunit of phosphatidyl inositol 3-kinase (PI-3K), which in turn activates the catalytic subunit p110β (42). Lipid products from PI3-K activate some isoforms of PKC that may be needed for recruitment of signaling molecules, bearing plecstrin homology domains, such as Vav, PLCγ, etc. (43). PI-3K in neutrophils and macrophages regulates membrane extension for pseudopod formation mediated by myosin-X (44,45), as well as activation of the extracellular signal-regulated kinase, which stimulates phospholipase A2 for arachidonic acid, which acts as second messenger in phagocytosis.
Ca2+ concentration increases around the phagocytic cup, but its role remains controversial. The increase seems to be due to Ca2+ release from the phagosome into the cytosol (46). Ca2+ is important for triggering actin depolymerization through gelsolin activation, and its mobilization is independent of inositol 1,4,5-triphosphate in neutrophils, indicating that the increase of Ca2+ around the forming phagosome does not come from intracellular stores.
Phospholipase Cγ (47) has also been shown to have a role in phagocytosis in some cells.
Rac and Cdc42 regulate actin assembly and internalization of FcγR-coated particles (48,49). Rac1, Rac2, and Cdc42 are subfamilies of the RhoGTPases, and cycle between GDP-bound (inactive) and GTP-bound (active) forms (50,51). They regulate several leukocyte functions such as cell motility, movements of cellular vesicles, chemotaxis, and the uptake of particles during phagocytosis. All these functions depend on dynamic and rapid reorganization of the actin cytoskeleton (52). The Arp 2/3 complex, an actin nucleator (see later) localizes to phagosomes formed by FcγR or complement and is required for both types of phagocytosis.
Cdc42 is restricted to the leading edge of the cell, whereas Rac1 is active throughout the phagocytic cup (53) and Rac2 is an integral component of the NADPH oxidase (54) (see later). The main phagocytic receptor for particles opsonized by the complement fragment C3bi is the αMβ2integrin, also known as complement receptor 3 (CR3) or CD11b/CD18 (55). Three C-terminal residues of this integrin are necessary for recruitment of Rho at the phagosome, and 16 amino acids of the membrane proximal region are needed for Rho activation (56).
Multiple subfamilies of the RhoGTPases are activated by members of the Vav family of guanine nucleotide exchange factors (GEFs), i.e., Vav1, Vav2, and Vav3. Activation involves the exchange of GDP for GTP (57). Vav1 is found predominantly in hematopoietic cells, whereas Vav2 and Vav3 are more broadly expressed. The Vav proteins are activated by immune receptors, integrins, growth factor receptors, etc. Activation involves phosphorylation of tyrosine residues in the amino terminus (57a).
The Wiskott-Aldrich syndrome protein (WASP) directly binds Cdc42 and Rac and is recruited to the phagocytic cup (58,59) (see later).
The signaling for inhibition of phagocytosis is regulated through inhibitory receptors, which express immunoreceptor tyrosine-based inhibitory motifs (ITIM). Ligation of inhibitory receptors results in phosphorylation of the ITIMs by Src kinases, which become docking sites for protein tyrosine phosphatases SHP-1 and SHP-2. The phosphatases dephosphorylate activating receptors, blocking downstream signaling, and the activation is arrested (60). The inhibitory receptor, FcγRIIB, recruits the inositol phosphatases SHIP-1 and SHIP-2, which act on PI(3,4,5)P3(PIP3) and break it down to PI(3,4)P2 (PIP2), thus arresting the downstream signaling of PI3K. Recruitment of SHIP-phosphatases is determined by the amino acid in the Y+2 position within the ITIMs (61) and by inadequate phosphorylation level (62). Overexpression of SHIP in macrophages results in inhibition of FcγRIIB-mediated phagocytosis (63), and colligation of FcγRIIA and FcγRIIB reduces phagocytosis in monocytes (64).
Enclosure of the Phagocytic Vacuole
It is believed that the receptor interaction is solely sufficient to trigger phagocytosis. However, mechanical parameters of the target particle can affect the outcome. When presented with particles of identical chemical properties but of different rigidities, macrophages show a preference to engulf rigid objects (65). Constitutively active Rac1 stimulates phagocytosis of soft particles.
Macrophages are able to ingest a number of particles with a total surface larger than the cell itself without net loss of surface, and even with surface gains (66). Phagocytes must, therefore, supply new membrane from sources within the cell to be able to meet the needs for the enclosure of the phagosome (67). Membranous pieces are supplied from an endosomal compartment recruited by phospholipase A2 (68). Focal exocytosis at sites of phagocytosis is necessary along with actin rearrangements to induce proper pseudopodial extension (69). Such addition of endomembranes is now considered a partner to cytoskeletal actin rearrangements for several cellular processes (i.e., extension of lamellipodia in phagocytosis, cell spreading, and chemotaxis) (70). Actin reorganization and membrane insertions by exocytosis are regulated by different mechanisms. Inhibition of PI3K plays no role in actin polymerization, yet it blocks completion of the phagosome, indicative of mechanisms other than rearrangements of actin cytoskeleton (44,71). The detection of endosomal markers on the plasma membrane during phagocytosis is accepted as evidence of membrane insertions during phagosome formation (79).
|
|
|
Figure 13.4. From stimulus to response: signaling pathways in phagocytosis. A particle opsonized by immunoglobulin G binds to Fcγ receptor (stimulus). Two types of responses are triggered by the stimulus: response 1 (1), formation of the phagosome through actin polymerization and extension of pseudopodia, which ingest the particle; and response 2 (2), the respiratory burst through activation of nicotinamide adenine dinucleotide phosphate (NADPH) oxidase. Response 1 involves SYK and other kinases (phosphatidyl-inositol 3-kinase, etc.), generating a second wave of downstream signaling molecules (phosphoinositides, activation of guanosine triphosphatases, etc.), which focus on the Wiskott-Aldrich syndrome protein (WASP), a master activator that links the process to actin polymerization through Arp2/3 (see text for details). Response 2 consists of the assembly of the NADPH oxidase in the membrane of the forming phagosome from several individual proteins, some present in the membrane and others translocated from the cytosol (see text for details). Response 1 will enclose the invader within the phagosome, to be killed by response 2 and eventually withdrawn in the interior of the cell to be disintegrated in the endosomes and lysosomes. Arp2/3, actin-related proteins 2 and 3; ITAM, immunoreceptor tyrosine-based activation motif; PIP2, phosphatidylinositol 4,5-bisphosphate. |
Most intracellular fusion events are determined by a specific protein machinery, which includes soluble factors, such as the N-ethylmaleimide-sensitive factor (NSF) and the soluble NSF-attachment protein, SNAP, as well as membrane complexes such as SNAP-attachment protein receptor, SNARE (Soluble N-ethylmaleimide-sensitive factor Attachment protein Receptor) NSF, is an adenosine triphosphatase (ATPase) important for SNARE function. The membranes to be fused need to bear specific SNAREs, known as vesicle or v-SNAREs and target or t-SNAREs. Particular v- and t-SNAREs dictate the specificity of intracellular molecular fusion events (73). Inactivation of two SNARE proteins, VAMP2 (vesicle-associated membrane protein 2) and VAMP3, reduces efficiency of phagocytosis. Exocytosis of endosomal membranes is a focal event and occurs at the sites of phagocytosis.
Another group of GTPases, the Rabs, contributes to vesicle exocytosis and interact with SNARE proteins (74).
Two other proteins, dynaminII and amphiphysin IIm, which are associated with endocytic vesicles, also participate in phagocytosis (75,76). DynaminII is a GTPase and, through its SH3 (Src homology 3) domain, interacts with amphiphysin IIm recruited to clathrin-coated pits. It also contributes to pseudopod extension.
In summary, several proteins, following distinct pathways, contribute to pseudopod extension, i.e., PI3-K, dynaminII, amphiphysin, and myosin X.
Once the phagosome is formed, it is pulled to the interior of the cell by actin filaments and myosins. The Rac, Cdc42, and Vav proteins coordinate F-actin accumulation and its polymerization at the phagosome while it is still at its nascent state. Their recruitment depends on activities between the ITAM motif of the FcR and Src and Syk kinases (77). There are several myosin isoforms (78), which are localized around phagosomes in macrophages, and their ATPase activity moves the phagosome along actin filaments. As the phagosome is internalized, it matures, i.e., changes the composition of its membrane. During maturation, the phagosomes progressively acquire a variety of proteins that are characteristic of endosomes and lysosomes (79,80,81), whereas other proteins acquired during invagination are recycled back to the plasma membrane.
Early in the maturation process, an ATPase acidifies the phagosomal vacuole. During maturation, phagosomes and endosomes do not coalesce, but membrane components and luminal contents are exchanged by momentary fusion followed by fission (“kiss and run” model) (82). Multiple fusion and fission events eventually change the composition of their membrane and content. Others believe that early and late endosomes and lysosomes simply exchange components by means of carrier vesicles (3).
Phagosomes move toward the cell interior with the help of microtubules, and during this journey, they have multiple contacts with endocytic organelles with which they fuse (Fig. 13.5). Eventually, their membrane is remodeled and comes to resemble a lysosomal membrane. This change is characterized by loss of small GTPases and acquisition of lamp proteins. The Rab family of proteins consists of ras-like GTP-binding proteins that serve as key regulators in endocytic trafficking. Rab proteins may, therefore, be used as markers for the endosomal vesicles. Rab5 is present on early endosome, whereas Rab7 is a late endosome–specific marker. The Rab5 member of the family, which is detected in the phagosome and normally regulates fusion of early endosomes, suggests that the phagosome fuses at an early stage with endocytic vesicles. The evidence is certainly more convincing that the phagosome fuses with late endosomes and eventually transforms to terminal lysosomes.
The relationship of late endosome to lysosome is dynamic. Molecules (i.e., receptors) that will be recycled are sorted out in the early endosome and are directed back to the membrane through the recycling vesicles, whereas other endocytosed material is delivered by carrier vesicles to late endosomes. The late endosomes are of lighter density than lysosomes, and although lysosomes eventually contain all internalized substances, they probably fuse with late endosomes on more than one occasion, indicating that an equilibrium likely exists between late endosomes and lysosomes.
This suggests that the lysosomal enzymes that are transported by the MPRs are delivered to the acidic endosomal compartment rather than to lysosomes, and the MPRs are then recycled back to the Golgi, plasma membrane, or both. The mannose receptor is considered an endocytic scavenger molecule that binds and internalizes glycoconjugate ligands, lyososomal enzymes, pituitary hormones, and other molecules that may be damaging to the organism if they accumulate in large concentrations (83).
The mannose receptor is not, strictly speaking, a phagocytic receptor, but it may trigger endocytosis in association with a partner (84).
|
|
|
Figure 13.5. The journey of the phagosome along the endocytic pathway to the cell interior. The membrane of the phagosome in the process of being born (phagosome nascendi) has the same composition as the cell membrane and contains the phagocytosed particle. The intracellular movement of the phagosome and other endocytic vesicles involves the microtubules. During this journey, the membrane composition of the phagosome changes as it bounces and partially fuses with other endocytic vesicles such as the early (EE) and late (LE) endosomes. Eventually, the altered phagosome fuses with lysosomal vesicles, which are in a dynamic, continuous exchange cycle with the LEs. The fusion between endocytic vesicles is regulated by the Rab family of guanosine triphosphate–binding proteins. Some of them are highly restricted to a certain stage in the endosomal pathway, such as Rab 4 and Rab 5 in the EEs and Rab 7 and Rab 9 in the LEs. The Rab proteins are not present on the lysosome. At the stage of LE and later, other proteins, such as the lysosomal-associated membrane proteins (Lamp), are characteristic markers of these vesicles. Marcks, myristoylated alanine-rich C-kinase substrate; MPR, mannose 6-phosphate receptor. |
Actin and Partners: An Incredible Machine
Extension of pseudopods and locomotion of cells is achieved by the assembly of actin filaments and their branching to form networks.
Actin is an ATP-binding protein that exists as a globular monomer (G-actin) and as polymerized filaments (F-actin). In electron micrographs, actin filaments have a characteristic arrowhead appearance with a pointed end and a barbed end. The barbed end has a higher affinity for actin monomers than the pointed end.
Each actin monomer has a bound ATP, which, after polymerization, is hydrolyzed to adenosine diphosphate (ADP), releasing inorganic phosphate. Monomers released from polymerized actin carry ADP, and, to polymerize again, ADP must be exchanged with ATP because ATP-actin has a higher affinity for the barbed ends of F-actin.
The assembly of actin filaments is a highly complex process and, at the beginning, must involve reactions between actin monomers de novo, but later, when a filament is formed, monomers are added to the filaments for extension or branching. This process is called nucleation, which refers to “initiation of new actin filaments by assembly from monomers” (85). Elongation, on the other hand, is the addition of new monomers to barbed ends of the filament (Fig. 13.4, part 1). Recently, the protein complex Arp2/3 has been shown to be the universal actin nucleation and organizer machine, conserved across the species (86,87). Arp2/3 consists of seven subunits, two actin-related proteins (Arp), Arp 2 and Arp 3, and five other proteins (ARPC1 through 5). The crystal structure of Arp2/3 has recently been solved (88) and shows that the Arp2/3 complex is a flat ellipsoid with dimensions 150 Å long, 140 Å wide, and 70 to 100 Å thick. The overall shape is in the form of a C-shaped clamp formed by the ARPC2 and ARPC4 subunits, while the Arp2 and Arp3 sit in the center. The other ARPC subunits reinforce and decorate the edges of the clamp. Activation of Arp2/3 complex by the WASP family of proteins brings Arp2 into proximity with Arp3 for nucleation (see later). For efficient performance, during the forward extension of lamellepodia, the cytoskeleton growth at the leading edge needs to be restricted by Arp 2/3-mediated actin polymerization mostly within a two-dimensional plane. The addition of new Arp 2/3 complexes to maintain proper overall ultrastructural organization of actin networks is still poorly understood (89).
ATP-bound actin polymerizes into filaments while hydrolysis and release of ATP destabilizes the filament, leading to depolymerization and recycling of actin subunits. The Arp 2/3 subunits hydrolyze ATP, resulting in debranching of actin filaments followed by remodeling, suggesting that ATP hydrolysis is critical for endocytic internalization (90).
Purified Arp2/3 has little activity (91) and requires activation by nucleation-promoting factors. Nucleation-promoting factors are members of the WASP family of proteins (92,93). WASP has a modular structure consisting of several structurally and functionally distinct domains (Fig. 13.6). At the C-terminal end of the protein is the VCA region, also referred to as WA. It consists of three domains: W or V (verprolin homology), C (cofilin homology or central or “connecting”), and A (acidic). The W is also known as WH2 (WASP homology 2 domain), and binds monomeric actin, competing with profilin and thymosin-β4.
The A and C domains (at the N terminus of VCA), on the other hand, bind to the Arp2/3 complex. The polyproline region (P), rich in ProXXPro motifs (X = any amino acid), binds SH3 domain–containing proteins (i.e., Grb2, Fyn, BTK), as well as the actin-binding protein, profilin, and the GTPase Cdc-42, linking WASP to the microtubules.
On the N-terminal side of the polyproline region is the GTPase-binding domain (GBD), which is the target for the small GTPases, Cdc42 and Rac. It is followed by a short lysine-rich basic region, which may be the actual binding site of PIP2 (94,95). The N-terminal domain, WH1, binds WIP (WASP-interacting protein), a member of the proline-rich actin-binding family of proteins. WIP has been implicated in regulation of actin cytoskeleton (96). It contains a WH2 actin monomer–binding domain, a profilin-binding domain, and a WASP-binding domain in its carboxy terminus. It regulates N-WASP–mediated polymerization and filopodium formation and stabilizes actin filaments. N-WASP has two WH2 domains, and Scar (suppressor of G-protein–coupled cyclic AMP receptor), also known as WAVE (WASP family verpolin-homologous proteins), lacks GBD and WH1 domains. WAVE proteins are composed of multiple conserved domains and their C-terminal domain; VCA (verpolin/cofilim/acidic) binds both monomeric actin and the Arp 2/3 complex (97). There are three members in this group (WAVE-1 to -3). WAVE proteins act as anchors for kinases, such as Abl, PKA (protein kinase A), and PI-3k, and as a result of the assembly of large signaling complexes, coordinate actin polymerization and branching by Arp 2/3. In conclusion, the C and A domains contribute to the binding of the Arp2/3 complex but not the WH2 domain. The binding sites for actin and for the Arp2/3 complex are close together or even overlapping. The WASP is a highly conserved protein in evolution, from yeast to humans, and coordinates the complex processes that extend pseudopodia from the cell body for survival, i.e., feeding or defense.
|
|
|
Figure 13.6. The structure of the Wiskott-Aldrich syndrome protein (WASP) and its ligands. WASP consists of several domains that mediate different functions (see text for details). Arp2/3, actin-related proteins 2 and 3; BR, basic region; GBD, guanosine triphosphatase–binding domain; PIP2, phosphatidylinositol 4,5-bisphosphate; SH3, Src homology 3; VCA, verpolin homology, cofilin homology, acidic domains; WH1, WASP homology domain 1; WH2, WASP homology domain 2; WIP, WASP-interacting protein. |
Activation of Arp2/3 by the Wiskott-Aldrich Syndrome Protein (WASP)
The crucial role of WASPs depends on their C-terminal WA (VCA) region, which binds and activates the Arp2/3 complex. WA binds simultaneously to G-actin and to Arp2/3 complex, with the actin filament enhancing the affinity of WASP for the Arp2/3 complex. The WASP dissociates rapidly from the complex after nucleation (98).
WASP mediates the “presentation” of G-actin to Arp2/3, followed by its activation. As shown by constructs containing individual domains of the VCA region, the A domain confers high-affinity binding to Arp2/3 but not activation, whereas the V domain is indispensable for activation and, at the same time, enhances G-actin binding (99). Thus, the V, C, and A domains act cooperatively in Arp2/3-mediated function for actin polymerization (100,101).
The ARPC2 and ARPC1 subunits anchor the complex to the side of the “mother” filament at a 70-degree angle. WASPs stabilize the conformation by linking ARPC1 and ARPC3 subunits.
Activity of WASPs is markedly enhanced by GTP-Cdc42, PIP2, and GRB2. Each of these factors individually has a weak effect, and it is likely that their action is additive or even synergistic (102,103). For example, GTP-Cdc42 is unable to activate WASP by itself, but it augments PIP2stimulation. Electron micrographs show that the Arp2/3 complex contacts three successive subunits of the “mother” filament, and the Arp2/3 form the first two subunits of the branching or “daughter” filament (104).
Actin Polymerization in Phagocytosis
Nucleation could take place either on the side of a filament (“branching nucleation”) (104,105,106) or at its barbed ends (107). Strong evidence by a variety of techniques (108,109,110) shows that a complex network of F-actin grows like a tree, with branches always extending from the sides of a filament (Fig. 13.7). As shown in Figure 13.8, gold-labeled Arp2/3 binds to the side of a pre-existing branch or mother filament (104). Elongation of the daughter filaments then takes place by the addition of new actin monomers with their pointed ends fitting to the barbed ends of the previous units (Fig. 13.7). Thus, the branches of the advancing actin network push forward by their free barbed ends. This model of actin polymerization is supported also by the fact that Arp2/3 initiates nucleation even from mother filaments that have their barbed ends capped (111). Finally, the protein cortactin, which binds only to the sides of the F-actin filaments, mediates nucleation by Arp2/3 complex (112). Cortactin has an N-terminal acidic domain and five to six repeats in the central part of the molecule, which have the F-actin–binding site (113). Cortactin acts as a mediator between WASP and the Arp2/3 complex. They both bind simultaneously to the complex and synergistically contribute to Arp2/3 activation, despite the fact that they compete for a common binding site on Arp2/3 (114). Cortactin binds to the Arp3 subunit of the complex and mediates the binding of the complex to F-actin, whereas WASP activates the complex. Cortactin thus promotes and stabilizes the formation of new daughter actin filaments (115).
|
|
|
Figure 13.7. Organization of actin polymerization. Actin-branching polymerization by actin-related proteins 2 and 3 (Arp2/3). Arp2/3 binds to the sides of pre-existing (“mother”) filaments and adds new G-actin monomers by side branching. The “push” of the actin filaments comes from the barbed ends of the new, elongated “daughter” filaments. |
WASP is subject to autoinhibition and exists in an inactive form. The “inactive conformation” results from an intramolecular interaction between the hydrophobic core of GBD and the C-terminal WA effector domain, which brings the occlusion of the WASP WA domain. Binding of GTP-Cdc42 to GBD disrupts the hydrophobic core, an action enhanced by PIP2. Change in the conformation “opens” the C-terminal VCA domain for binding of the Arp2/3 complex (102,116).
|
|
|
Figure 13.8. Localization of the actin-related proteins 2 and 3 (Arp2/3) complex at branching points of actin filaments. Electron micrographs of actin filaments. The Arp2/3 complex is localized at the branching points (arrow). The complex was identified by a specific antibody, the binding of which was detected by a secondary gold-conjugated antiimmunoglobulin antibody. (From Svitkina TM, Borisy GG. Arp2/3 complex and actin depolymerizing factor/cofilin in dendritic organization and treadmilling of actin filament array in lamellipodia. J Cell Biol 1999;145:1009–1026, with permission.) |
|
|
|
Figure 13.9. Phagocytosis of yeast particles by polymorphonuclear leukocytes. A human neutrophil extends long pseudopodia that encircle two yeast particles. HE, hyaline ectoplasm. (From Stossel TP. Phagocytosis (second of three parts). N Engl J Med 1974;290:744, with permission.) |
The leading edge of a moving phagocyte is known as the lamellipodium. It extends forward and seems to pull the rest of the cell body along with it. Pseudopodia project from the lamellipodium and attach closely to the particle to be ingested. The lamellipodium and pseudopodia contain no cytoplasmic organelles and, as a result, have a glassy appearance (Fig. 13.9). Both, however, are rich in actin filaments. Actin constitutes 10% of the total protein in macrophages and is present in two forms, G-actin (globular or monomeric) and F-actin (filamentous or polymeric).
Actin polymerization during phagocytosis has been well documented (117,118), as has the role of the WASP family of proteins in the mechanism of polymerization through activation of the Arp2/3 complex (119,120,121). Monocytes and macrophages from patients with Wiskott-Aldrich syndrome (WAS) have reduced capacity for phagocytosis (122). WAS is a rare X-linked, recessive immunodeficiency disease. Patients suffer from recurrent pyogenic and opportunistic infections, eczema, and thrombocytopenia with small platelets. WAS is due to a wide spectrum of gene mutations that result in the expression of no WASP or a truncated protein lacking the crucial C-terminal VCA or WA region. In an X-linked thrombocytopenia form or attenuated WAS, the WASP gene has missense mutations in the N-terminal region (123). Finally, an X-linked neutropenia form or myelodysplasia has been detected as a result of mutations of part of the gene encoding the GBD domain. Mice with targeted disruption of the GBD domain share some of the features (i.e., lack of T-cell activation) but not others (124). WASP is essential for activation of NF-AT. In WASP-/- mice the nuclear translocation of NF-AT and Erk (Extra-cellular regulated kinase) are impaired (125). These defects may underlie defective IL-2 production by T cells in Wiskott-Aldrich syndrome.
Fragments of Arp2/3 containing the WA domain almost completely inhibit phagocytosis, a finding that emphasizes the importance of Arp2/3 in the process (126). The Arp2/3 complex, WASP, and N-WASP localize to the surface of phagosomes during FcγR- and CR3-mediated phagocytosis (122,126).
An intriguing function of actin polymerization is related to the movement of various endosomal vesicles. Actin polymerization propels them to their destinations by a “rocketing” mechanism (127). The Arp2/3 complex and N-WASP are localized to the surface of moving endosomes and lysosomes and to the surface of motile macropinosomes (128,129). A similar function of actin polymerization has been demonstrated with the motility of intracellular pathogens (130). Such pathogens are associated with actin “comet tails,” which contain many of the components of lamellipodia. Structurally, however, the filaments in lamellipodia are branched as a result of dendritic nucleation, whereas the “comet tails” are short, randomly oriented filaments with some dendritic nucleation (131). Thus, it is evident that actin polymerization and dendritic nucleation with an expanding F-actin network provide all the mechanical force necessary for “pushing” lamellipodia around the particle and completing the enclosure of the phagosome (132).
Actin-Binding Proteins
Profilin and Cofilin
Profilin and thymosin-β4 bind monomeric actin and modify its properties for polymerization.
Thymosin-β4 is a peptide of 43 residues that competes with profilin (125 to 139 residues) for binding to actin (133). Thymosin-β4 binds more strongly to ATP-actin and is a sequestering protein.
Profilin has a higher affinity for ATP-actin monomers than for the barbed ends of actin in filaments because its binding site is buried in the filament structure. As a result, profilin maintains a pool of free ATP-actin ready for filament elongation.
In cells with both proteins, profilin serves as a carrier between the thymosin-β4–actin pool and the barbed ends of actin filaments (134).
At least two F-actin pools exist in vertebrate cells, one relatively stable, which is often associated with tropomyosin (an actin filament–binding protein with an α-helical coiled-coil structure), and one more dynamic, usually found in the leading edge of motile cells.
Recently, it has been shown that the proteins that regulate the high turnover of F-actin filaments belong to the family of ADF-cofilin (ADF, actin depolymerizing factor; cofilin, forming cofilamentous structure with actin) (135,136). There are approximately 30 members in the family with conserved actin-binding regions. A second homologous region outside the actin-binding site suggests that they bind to another protein (137). ADF and cofilin are usually cytoplasmic proteins, but, under stress, they accumulate in the nucleus accompanied by actin, which forms actin rods (135). Nuclear localization, which is mediated by a specific sequence, has been detected in many types of cells as inclusion bodies that frequently have a fibrillar or lattice appearance. Actin and cofilin have been detected in these inclusions.
The three-dimensional structure of yeast cofilin shows that it consists of a β sheet with four strands surrounded by four α helices. This structure has been called the ADF homology domain. The ADF homology domain shares similarities with the gelsolin family. Gelsolin has six repeats of this domain (138).
Cofilin interacts with actin through its N-terminal region (139), an interaction inhibited by PIP2. The structure of the cofilin–actin filament complex shows that cofilin makes contacts on the outside of the filament with both the pointed and barbed ends (140).
In general, ADF/cofilin binds to the actin-ADP subunits of the F-actin filament, increasing the off-rate from the pointed end and thus depolymerizing F-actin and severing actin filaments (118); in doing so, it increases the number of barbed ends. When actin monomers dissociate from the pointed ends of the filament, they have ADP nucleotide, and, to be reused, ADP must be exchanged with ATP. However, ADP/cofilin prevents the exchange, limiting the reuse of actin monomers released from filaments (141).
Profilin, on the other hand, strongly increases the nucleotide exchange, even in the presence of ADF/cofilin (142), and it appears that profilin and ADF/cofilin regulate recycling of actin-ADP and F-actin filament turnover based on their opposite effects on nucleotide exchange. Filament turnover is rapid during extension of pseudopods and cell movement. Lamellipodia move up to 1 μm/s (370 subunits/s). The mechanisms that regulate the high rate of depolymerization, and of severing when it is required, are not well understood. Polymerization or filament extension requires free barbed ends, which can be generated by various mechanisms such as uncapping and severing.
Gelsolin
Gelsolin is the most abundant protein that caps filament ends, and it regulates cytoskeleton remodeling in response to Ca2+ and PIP2 binding (143). Severing is the breaking of noncovalent bonds between two actin molecules within a filament. After severing, gelsolin remains attached to the barbed ends of the filament as a cap (capping). Gelsolin is the most effective severing protein identified to date (144). Through actin cytoskeleton reorganization, it plays a role in cell motility, phagocytosis, chemotaxis, and wound healing (145). Animals of mixed-strain background that are deficient in gelsolin have no gross pathology, but inbred strains are not viable at perinatal stages (146).
Gelsolin consists of six structurally similar domains (G1 through G6), evolved by gene duplications from one prototypical domain (142). Significant structural similarities exist between the domains in each triplex (G1 through G3 and G4 through G6). The actin-binding site is masked and is revealed in response to Ca2+ increase, which releases contacts between domains and exposes actin-binding sites in G2, G1, and G4domains. There are up to eight Ca2+-binding sites of two types. Type 1 sites are shared between gelsolin and actin and govern the strength of interactions between them. Type 2 sites have the potential for causing movement between the domains. The structure of the gelsolin–F-actin complex has been solved and provides insights into the mechanism of severing (147,148). Ca2+ activation triggers binding of G2 to F-actin, and this brings the G1 into a position to disrupt actin—actin interaction. The second disruption needed to sever the monomer is contributed by G6, which brings G4 domain for a pincer action with G1 to remove the actin monomer. There are at least two G-actin–binding sites in domains 1, 2, and 4 and one PIP2-binding site in domain 2. Lowering Ca2+ concentration may not induce gelsolin uncapping after severing. PIP2 is the only known agent that inhibits gelsolin severing and dissociates gelsolin from actin in vitro (149).
In platelets, removal of gelsolin contributes to actin polymerization (150) and to filament growth. Severing is the major mechanism for generating free barbed ends in platelets (151), and the thrombin receptor–activating peptide stimulates Arp2/3 activation for all actin-dependent morphologic changes (152).
Regulatory Mechanisms: Downstream Pathways
Calcium
One of the first signals observed in response to FcγR activation of phagocytosis is the increase of cytosolic Ca2+ (153). The role of Ca2+ in FcR-mediated phagocytosis remains controversial, but, for CR3-mediated phagocytosis in neutrophils, it plays no role. In macrophages, neither phagocytosis nor phagosome–lysosome fusion is Ca2+-dependent (154). Ca2+, however, is important in some aspects of actin network reorganization and in the function of gelsolin.
Protein Kinase C Family
These serine/threonine kinases are activated by the phospholipase product diacylglycerol, as well as by Ca2+. Members of the family have been identified in the phagosome (PKC-α, -β, -γ, and -δ) during FcγR-mediated phagocytosis, and pleckstrin, a major substrate for PKC phosphorylation, is recruited in macrophage phagosomes (155). Other substrates for PKC are the proteins MARCKS (myristoylated alanine-rich C-kinase substrate), which mediate cross-linking of actin filaments. In mice deficient in MARCKS, however, the FcγR- and C3-mediated phagocytosis is normal (156), indicating that simple localization around the phagosome is not necessarily indicative of functional participation.
Phosphoinositide Kinases
Phosphoinositides bind to pleckstrin homology domains, which are detected in well over 100 different proteins. The majority of these proteins are found in membranes and have been implicated in a variety of membrane functions (157,158).
Phosphoinositides are major regulators of various aspects of the phagocytic process (159,160) and are produced from the function of PI3K. PI3K, however, is not localized in the phagosome (161) as a result of displacement by its own product, PIP3 (162). PI3K is activated by FcγRs and is required for phagocytosis by macrophages—not in the initial stages of phagosome formation but mainly for its closure (163), when actin reorganization is important. Different sizes of particles seem to require different handling by the phagocytic machinery. Among the PI3K classes, PI3K class I is involved for phagocytosis of particles larger than 3 μm but not for their subsequent maturation (164). In contrast, PI3K class III (which forms PIP3) is needed for phagosome maturation (164,165). PIP2 is an important regulator of WASPs and of gelsolin (see earlier).
Guanosine Triphosphatases
Small GTP-binding proteins are key regulators of actin cytoskeleton (166,167). Assembly of actin beneath an opsonized particle requires the small GTPases, Cdc42, and Rac1, which control the cytoskeletal rearrangements during particle uptake (118,168,169). Cdc42 and Rac1 control different steps in the phagocytic process (i.e., pseudopod formation for Cdc42 and phagosome closure for Rac1) (169). In contrast, actin polymerization mediated by CR3 depends on the activity of Rho A, whereas Cdc42 and Rac1 are not involved (170). GTPases act on WASP family members, which activate the Arp2/3 complex (171). GTP-Cdc42 and PIP2 act on the inactive form of WASP as a result of autoinhibition and restore its active conformation (172,173,174). The transient PIP2accumulation at the phagocytic cups is part of the activation signal for WASP, together with Cdc42 recruitment. Rac1 stimulates gelsolin dissociation, thus preventing depolymerization of filaments and promoting the closure of the phagosome (175).
Ezrin, Radixin, and Moesin Family
The gap between cytoskeleton and membrane may be filled, in some systems, by a family of three proteins, ezrin, radixin, and moesin, known as the ERM family (176). These proteins act as linkers between plasma membrane and actin (177). Their N terminal binds to some membrane proteins, such as CD44, and their C terminal binds to actin. This intermolecular cross-linking requires activation, which is probably mediated by the Rho protein. Normally, the ERM proteins have a head-to-tail association that is released after activation (178).
The cytoskeleton is linked to the extracellular matrix focal adhesions (FA) that assemble at the lamellipodium base (179). They contain transmembrane integrins that link the exracellular matrix to F-actin via structural and signaling proteins. Interaction with talin, which is a cytosolic protein recruited under the particle attachment at F-actin sites, facilitates FcR-mediated phagocytosis (180).
Digestion
Microbicidal Mechanisms
Microbicidal mechanisms used by the phagocytes can be divided into (a) oxygen-independent and (b) oxygen-dependent mechanisms. Oxygen-dependent mechanisms may be subdivided into (a) myeloperoxidase (MPO)-mediated (halide radicals–generated) and (b) MPO-independent.
Oxygen-Independent Mechanisms
Detailed description of neutrophil and macrophage granule content and their role in digestion of phagocytosed material is provided in Chapters 9and 12, respectively.
The neutrophil primary or azurophilic granules contain several bacteriocidal proteins (i.e., lysozyme, lactoferrin, cathepsin G, and elastase) and small peptides known as defensins (181). Defensins are peptides of 29 to 35 amino acids, including six invariant cysteines forming intramolecular disulfide bonds, which stabilize the cyclic structure of the peptide with a triple-stranded β-sheet configuration. They kill microorganisms and mammalian cells, even those resistant to tumor necrosis factor-α, by inserting themselves into the cell membrane, which they permeabilize. The mechanism of insertion involves interaction with negatively charged molecules, probably membrane lipids.
The secondary (or specific) granules contain also lysozyme and collagenase. The lysozyme is a cationic enzyme that hydrolyzes glycosidic bonds and has bactericidal properties for some bacteria.
Oxygen-Dependent Microbicidal Activity
During particle ingestion, phagocytes show a two- to threefold increase in O2 consumption, which is insensitive to cyanide, a two- to tenfold increase of glucose oxidation via the hexose monophosphate shunt, and the generation of hydrogen peroxide. This combination of metabolic activities came to be known as respiratory burst (182) (Fig. 13.4, part 2). Respiratory burst was actually discovered in 1933 (183).
Myeloperoxidase-Independent Systems
The burst of respiratory activity results from the activation of an enzyme known as nicotinamide adenine dinucleotide phosphate (NADPH) oxidase, a key enzyme present only in professional phagocytes and in much smaller quantities in B cells. NADPH oxidase transfers electrons from NADPH in the cytoplasm across the wall of the phagosome to its interior, forming superoxide (184).
When oxygen accepts one electron, it is reduced to superoxide anion, O2-, according to the reaction shown in Equation 1.
where NADP+ is the oxidized form of nicotinamide adenine dinucleotide phosphate.
H2O2 is produced by oxidation-reduction of two superoxide radicals (O2-), a reaction known as dismutation, catalyzed by the enzyme superoxide dismutase according to the reaction shown in Equation 2.
Alternatively, H2O2 can be produced by reduction of O2 by the transfer of two electrons catalyzed by the NADPH oxidase. It is possible, however, in conditions in which H2O2 is detected in the absence of detectable O2-, that the superoxide may not be diffusing out of the space of the enzyme (184).
Superoxide and H2O2 are not used by the phagocyte for bacterial killing because H2O2 is weakly microbicidal and superoxide is innocuous. They are, however, used as intermediates to produce oxidizing radicals or in reaction with halides to generate hypochlorous acid (HOCl) or hypochlorite mediated by MPO (185).
Other O2 radicals are generated by interaction of O2- and H2O2, such as the hydroxyl radical (OH·). This reaction is slow and may not take place directly but through the catalytic action of trace metals such as iron (Equation 3, known as the Haber-Weiss reaction).
Singlet oxygen (1O2) is another O2 intermediate that is generated as a result of the shifting of one of the two unpaired electrons to another orbit of energy and a change in its direction of spin. 1O2dissipates its excess energy quickly by thermal decay or light emission (like chemiluminescence, CL), and it can be produced by a number of mechanisms such as spontaneous dismutation of two superoxide anions (Equation 4).
Myeloperoxidase-Dependent Oxidants
The MPO system produces potent antimicrobial reactive oxygen radicals that are effective against bacteria, fungi, viruses, and mycoplasma. MPOs are enzymes that catalyze the oxidation of a number of substances termed hydrogen or electron donors. The components of the system are MPO, H2O2, and halide. The products of the reactions are also toxic to tissues and cells of the host.
H2O2 reacts with a halide, a reaction mediated by MPO that is present in the primary or azurophilic granules of the neutrophils and in the lysosomes of the juvenile monocytes but is not present in macrophages. This reaction generates hypochlorous acid as shown in Equation 5,
or hypochlorite (OCl-) as shown in Equation 6.
HOCl attacks the phagocytosed microorganisms and causes damage to their cell membranes. Of the halides, chlorine is present in leukocytes at a high concentration, but iodine is considerably more effective on a molar basis. The halide oxidants are quickly detoxified by reacting with low-molecular-weight amines to yield chloramines (i.e., taurine), present in high concentrations in neutrophils.
The acceleration of the hexose monophosphate shunt is a result of the production of NADP+ and the regeneration of NADPH by the action of glucose 6-phosphate dehydrogenase (Fig. 13.10). However, NADP+ increases also from the glutathione (GSH) peroxidase/GSH reductase system, which reduces H2O2 to H2O (as shown in Equations 7 and 8).
where GSSG is oxidized GSH.
NADPH is regenerated through the metabolism of the glucose (186).
Not only microorganisms but also the phagocyte and surrounding tissues are susceptible to damage (187). Phagocytes apply different mechanisms to detoxify the oxidant radicals, such as the superoxide dismutase (converts O2- to H2O2), catalase (converts H2O2 to H2O), GSH reductase (converts H2O2 to H2O), amines (convert HOCl to chloramines), and so forth. The oxidants have been implicated in the pathogenesis of a number of clinical conditions such as chronic lung disease.
The NADPH oxidase is the enzyme that catalyzes the generation of O2-derived oxidants during phagocytosis or upon stimulation by a variety of substances such as phorbol myristate acetate, N-formylated oligopeptides of bacterial origin that are secreted or released from dead organisms, complement-derived anaphylatoxin C5a, and IL-8. The activity of the enzyme depends on a number of components, which are brought together immediately after stimulation. The active enzyme is associated exclusively with the plasma membrane.
|
|
|
Figure 13.10. The production of oxygen radicals. The NADPH oxidase is assembled from five subunits (two localized on the membrane of a secretory vesicle and three residing within the cytoplasm). The assembly is powered by signals emanating from the triggering of the Fc receptor. Phosphorylation of p47phox in the cytoplasm may be an early event that triggers translocation of all three cytosolic subunits to join the other two on the membrane. G-6P, glucose 6-phosphate; G-6PD, glucose 6-phosphate dehydrogenase; GPX, glutathione peroxidase; GR, glutathione reductase; GSH, glutathione; GSSG, oxidized glutathione; Heme, Fe-protoporphyrin IX; MPO, myeloperoxidase; NADP+, oxidized form of nicotinamide adenine dinucleotide phosphate; 6PGD, 6-phosphogluconate dehydrogenase. |
Nicotinamide Adenine Dinucleotide Phosphate Reduced Oxidase
There are five protein members of the active enzyme, known as the phox (phagocytic oxidase) family of proteins. Two members, gp91phox and p22phox, reside in the membrane and three, p67phox, p47phox, and p40phox, in the cytoplasm. New homologs of the gp91phox subunit have recently been discovered and form a family designated as the Nox family of NADPH oxidases (188). It has seven members, Nox1 through Nox5 and Duox1 and Duox2, with the original gp91phox being Nox2. The members of this family are expressed in different tissues, Nox2 in myeloid cells, Nox1 in the colon, Nox3 in the vestibule (of the ear), Nox4 in the kidney, and Nox5 in the spleen and testis.
The Duox members (Dual oxidases) have been detected in cells of the mucosal surfaces, salivary glands, bronchi, trachea, and thyroid (188a). They perform a potential host defense role, generating H2O2 for lactoperoxidase, which oxidizes thiocyanate and iodide into antimicrobial compounds, while in the thyroid the iodide is oxidized by thyroperoxidase into reactive iodide species.
Structure
Membrane Components: Cytochrome b558
Two of the subunits of the oxidase, the gp91phox (or Nox2), and the p22phox, are located in the membranes of secretory vesicles and specific granules and form a heterodimer known as cytochrome b558 (Fig. 13.10) (189). The heterodimer contains two heme prosthetic “groups,” both residing within the gp91phox (190) subunit, which are coordinated by histidine residues (191). One heme is toward the outer face of the membrane (His115-His222) and the second is in the inner face of the membrane (His101 and His209) (189). The carboxy terminal of gp91phoxhas flavin- and NADPH-binding sites, homologous to the ferredoxin-NADPH+ reductase of the flavoenzyme family (192,193). In almost all patients with mutations in b558, the neutrophils lack both subunits regardless of which subunit is affected by the mutation (194), i.e., the expression of one subunit affects the expression of the other. In patients deficient in p22phox, the fully glycosylated gp91phox subunit is not detected, but only a partially processed precursor (65 kDa) with high-mannose carbohydrate side chains (195). Expression of the mature gp91phox polypeptide with fully processed oligosaccharide side chains was restored, however, by expression of a recombinant p22phox. Inhibition of heme biosynthesis results in decrease of b558 expression with marked decrease of p22phox and of mature gp91phox(196), indicating that the heme prosthetic groups contribute to the assembly and stability of the heterodimer.
The gene for gp91phox is located on chromosome X p21.1. The protein contains 570 amino acids, with four or five transmembrane helices and five potential glycosylation sites in the N-terminal region. The hydrophilic C terminal (amino acids 282–570) of gp91phox contains the FAD- and NADPH-binding sites. The FAD is noncovalently bound (197). Mutations of this gene account for the most common forms of chronic granulomatous disease (CGD).
The gene for p22phox is located on chromosome 16q24. The protein has 194 amino acids, with hydrophobic helices in the N-terminal half of the molecule that span the membrane at least two (possibly four) times (198). A praline-rich domain (151,152,153,154,155,156,157,158,159,160) is important for binding the cytosolic components p47phox. A mutation of 156ProSGln, while giving normal amounts of protein, abolishes the translocation of p47phox and p67phox to the membrane and results in chronic granulomatous disease. Other sites, however, have also been implicated in interaction of p22phox with both p47phox and p67phox.
Cytosolic Components
There are three cytosolic components: p67phox, p47phox, and p40phox (Fig. 13.11, Table 13.3 and Table 13.4).
p67phox
The gene for p67phox is located on chromosome 1q25 and the protein has 526 amino acids with two SH3 domains, one central and one C-terminal, like those present in Src tyrosine kinases and other signaling molecules. They mediate protein–protein interactions by binding to proline-rich sequences (199), which are detected in all cytosolic components. The gp67phox also contains two praline-rich regions flanking the central SH3 domain, an N-terminal TPR repeat (tetratricopeptide repeat) and a PB1 domain C-terminal to the central SH3 domain, which interacts with octicosapeptide motifs (in p40phox) (Fig. 13.11B). The p67phox is absolutely required to induce electron transport through the flavocytochrome, and as a result it has been called the NOXA family (NOX A activator).
p47phox
The p47phox has two SH3 domains, at least one or up to three proline-rich sequences, and a PX domain. In the resting state, the C-terminal proline-rich sequence interacts with one of its own SH3 domains, and this interaction is considered a possible reason for the inability of resting p47phox to bind cytochrome b558. However, upon activation, the C-terminal proline-rich sequence interacts with an SH3domain of p67phox, which is important for their translocation to the membrane. The C-terminal region contains six to eight putative phosphorylation sites, and extensive phosphorylation upon activation initiates its translocation to the membrane, bringing along the p67phoxsubunit and serving as bridge between p67phox and p22phox for attachment to gp91. The p47phox stabilizes and organizes the oxidase (NOX2) and is known as NOXO2 (NOX2 organizer). Another NOXO protein, NOXO1, lacks the autoinhibitory domain and usually is coexpressed with NOX1 in the colon epithelium. p47phox binds through its N-SH3 domain to a Pro domain of p22 and stabilizes p67 attachment.
|
|
|
Figure 13.11. The cytosolic components of nicotinamide adenine dinucleotide phosphate (NADPH) oxidase. All three components, p40phox, p47phox, and p67phox, contain Src homology 3 (SH3; found in signaling molecules), and p47phox and p67phox, in addition, have proline-rich sequences (Pro). These domains mediate protein–protein interactions involved in the association of the cytosolic components, which then are translocated to the membrane for the assembly of the NADPH oxidase. p40 is associated with p67 in the serum tightly. p47 binds to p22phox and facilitates binding of p67phox to b558. |
p40phox
The p40phox subunit is a protein of 339 amino acids. It contains an N-terminal PX domain followed by an SH3 domain. The PX domain binds PI(3)P and targets localization of p40phox on endosomal membranes. Localization of p40phox is abolished as a result of inhibition of PI3K function due to mutation R57Q, which interferes with PI(3)P binding (200). Towards the C terminus, there is an octicosapeptide repeat (OPR), also known as PC domain. This domain seems to be involved in the binding of p40phox to p67phox.
|
Table 13.3 Properties of the Cytosolic Components of NADPH2 Oxidase |
||||||
|
The phox homology domain or PX of the cytosolic proteins is a 125– to 140–amino acid module, which is found in proteins involved in membrane targeting and trafficking, cytoskeletal organization, and protein sorting (201). It is the hallmark of the members of the protein-sorting nexin family (202) but is also found in phospholipases D1 and D2 and the protein kinase CISK, implicated in IL-3–dependent cell survival. Localization on vesicular compartments is strictly PX-dependent. The PX domain contains basic and hydrophobic residues and binds phosphoinositides. The p47phox pleckstrin homology domain binds strongly PI(3,4)P2 and weakly PI(3,5)P2 and PI(3,4,5)P3 (203). The binding site for phosphoinositides is contributed by β strands and a pair of α helices linked by a loop. Basic clusters and hydrophobic residues contribute to the binding, and two of the basic residues crucial for binding are conserved. Mutation of one of them, Arg42Gln, results in an autosomal recessive form of CGD. These findings provide a rational explanation for the impaired respiratory burst in mice deficient in the p110γ (class IB PI3K), which are unable to produce phosphoinositides (204).
Assembly and Activation of Nicotinamide Adenine Dinucleotide Phosphate Oxidase
Multiple phosphorylations of p47phox are thought to be critical for the assembly and translocation of the cytosolic subunits because they regulate a number of steps in the process. Phosphorylations are mediated by protein kinase A on C-terminal serines. Some of the serines, such as Ser379, are more important than others (205). Mutations Ser379Ala, Ser359Ala, and Ser370Ala result in loss of oxidase activity as well as block of phosphorylation of the remaining serines. Phosphorylation of Ser303/304 increases oxygen radical production, but the mechanism is not clear (206). In general, phosphorylation seems to increase direct binding of p67phox to b558 (207).
A very important consequence of p47phox phosphorylations is the opening of the phosphoinositide-binding pocket on the PX domain (208). The PX domain has been shown to mediate targeting of p47phox and p40phox to endosomal membranes (201,209,210,211). However, the “pocket” that binds the phospholipids is “closed” as a result of intramolecular interactions (208). Phosphorylations (Ser303/304/328/359/370/379) seem to change the molecular conformation and “open” the pocket. One may envision the sequence of events regulated by phosphorylations of p47phox as follows: conformational changes of p47phox, “opening” of the pocket for phosphoinositide binding, targeting of the subunits to endosomes, and facilitation of p67phox binding to b558. Others however found that phosphorylations do not modify SH3-binding affinity (212).
The assembly and translocation require interactions between all three cytosolic components. One interaction involves p47phox (C-terminal proline-rich sequence) and p67phox (C-terminal SH3 domain) (213) and a second interaction involves the SH3 domain of p47phox and proline sequences of p67phox (214,215,216) (Fig. 13.11A). Deletion of both SH3 domains of p67phox abolishes its membrane binding. The adaptor protein p40phox also participates in the assembly through direct binding to p67phox, which is linked to p47phox (217). p47phox is critical for the translocation of the cytosolic subunits, whereas the other subunits follow as a result of their interactions with p47phox. p47phox binds to p22phoxthrough one of the two SH3 domains, which are expressed in tandem in the middle of the protein. The N-terminal SH3 domain binds to a proline-rich sequence in the C terminus of p22phox (Fig. 13.11A). This interaction is essential for oxidase activity. However, the SH3 domain of p47phox is masked in the resting state by an interaction with an Arg/Lys domain in the C terminus. Phosphorylation of serines within the Arg/Lys domain unmasks the N-terminal SH3 domain of p47phox and allows binding to the C-terminal p22phox proline-rich sequence (218). The C-terminal praline-rich region of p22phoxmediates the interaction with the N-terminal SH3 domain of the p47phox. This interaction is enhanced by the C-terminal region of p22phox, which adopts an α-helix conformation and is involved in the full activation of the phagocytic oxidase by fortifying the interaction with the p47phox SH3 domains (219,220). It appears that the interactions of p47phox with the membrane component p22phox is not only critical for activation of NADPH oxidase but is quite complex, as some other work suggests that the N-SH3 domain of p47phox interacts with a helix in the cytoplasmic region of p22phox (221).
Another cytosolic component that regulates NADPH oxidase activity is Rac, which exists in two isoforms, Rac1 and Rac2, both being low-molecular-weight GTP-binding proteins. Upon activation of neutrophils, Rac2 becomes associated with the cell membrane. The translocation is independent of the other cytosolic subunits. There are two functional regions in Rac: the “effector” region and the “insert” region (222). The C terminus anchors Rac to the membrane, whereas the effector region binds to p67phox, and the insert region binds to b558 (223). Rac2 regulates oxidase activity (226) as a result of interaction directly with b558 and participation in the electron transfer (225). The NADPH oxidase has an “activation domain,” which is localized on p67phox (226) and is essential for the generation of superoxide (227).
In the assembled NADPH oxidase on the membrane, the important component is b558. Its p91phox subunit is the catalytic moiety that transfers the electrons from NADPH to O2, whereas the other components are regulatory. The pathway of electron flow consists of several steps, (Equation 9) (226).
The b558 shows no activity unless the cytosolic subunits and Rac assemble in the membrane in a 1:1:1 ratio with b558. The p47phox is essential for translocation and provides binding sites for the other components as well as enhancing p67phox affinity (228). A protein complex constructed by genetically engineered fusion of p47phox and p67phox tested in vitro efficiently reconstituted the NADPH oxidase in terms of activity and stability (229). The fusion protein, tested in a cell-free system, had eightfold higher efficiency and produced higher activity than the individual proteins. Rac binds to both b558 and p67phox as an adaptor protein, but it is also required for electron transport. The activation domain of the NADPH oxidase is located within p67phox (i.e., sequence 199 to 210, which mediates the electron transfer from NADPH to flavin adenine dinucleotide) (226). A single mutation of residue 204 completely eliminates NADPH oxidase activity without affecting the interactions of the components of the oxidase. The activation domain does not influence NADPH binding.
|
Table 13.4 Critical Steps in NADPH2 Assembly |
|
|
In the second step, electrons are transferred to the hemes and then on to O2 for superoxide formation. Rac is required for both steps of the electron transfer.
Localization of Nicotinamide Adenine Dinucleotide Phosphate Oxidase
The original assembly and activation of the NADPH oxidase are carried out within intracellular vesicles and not on the plasma membrane. Human neutrophils contain a unique population of slender, rod-shaped vesicles that contain alkaline phosphatase (230). Neutrophil stimulation up-regulates these vesicles, which fuse to form tubular structures and eventually reach the cell membrane. Flavoprotein b558 is associated with these vesicles, which are known as secretory vesicles (231). Using cytochemical markers for alkaline phosphatase to identify these vesicles and cationized native ferritin as a marker for plasma membrane, it was clearly shown that superoxide (identified by a diamino benzidine cytochemical reaction) was first released within the secretory vesicles of the neutrophil (232) (Fig. 13.12). In human neutrophils undergoing phagocytosis, most of the oxygen radicals are detected in these vesicles, which then, by a process of exocytosis, reach the plasma membrane. The plasma membrane remains devoid of free-radical production. These findings provide an explanation for the lag period of the release of oxygen radicals after stimulation. Components of NADPH oxidase and Rac were detected by immunolabeling in punctate clusters of 0.03 to 0.10 μm2 on the cytoplasmic side of the membrane on neutrophils attached on glass surfaces (233). However, the process of adherence may have altered the distribution of the oxidase components, whereas in the previous study (232), the neutrophils were in suspension.
Defects of Oxidative Metabolism
Chronic Granulomatous Disease
CGD is a rare inherited disorder caused by defective NADPH oxidase that affects approximately 1 in 500,000 individuals. The main manifestations of the disease consist of infections of the lungs, gastrointestinal tract, and skin. In the majority of cases, the manifestations of the disease appear during the first year of life. Infections are caused by a variety of microorganisms such as bacteria and fungi. Some of the main pathogens contain catalase, the enzyme that converts the H2O2 to H2O, and, as a result, it cannot be used by the phagocytes for the formation of oxidants (234).
|
|
|
Figure 13.12. Localization of nicotinamide adenine dinucleotide phosphate oxidase. The NADPH oxidase is first assembled on the membrane of secretory vesicles of neutrophils, which then migrate and fuse with the cell membrane. A: The oxidant-positive vesicles (arrows) are separate from ferritin (small dark dots), identifying the cell surface (arrowheads). B: An oxidant-producing vesicle (dark arrows) opens to the cell surface (arrowhead) coated with cationic ferritin (dots, wide arrow). (Courtesy of Prof. Harumichi Seguchi, Department of Anatomy and Cell Biology, Kuchi Medical Center, Kohasu, Okoh-cho, Nankoku, Kochi, Japan; from Kobayashi T, Robinson JM, Seguchi H. Identification of intracellular sites of superoxide production in stimulated neutrophils. J Cell Sci 1998;111:81–91, with permission.) |
Microbicidal mechanisms other than the oxidants (lysosomal enzymes and peptides) are not as effective, and as a result, patients develop complications caused by chronic inflammatory stimulation, such as granulomatous obstruction of the esophagus and ileocolitis with diarrhea (235).
Early studies of the function of leukocytes in vitro from patients with CGD revealed that there is no respiratory burst on stimulation, suggesting that the defect is associated with the NADPH oxidase. The breakthrough in our understanding of the genetics of inheritance of CGD and the nature of NADPH oxidase came with the development of a cell-free system for activating the oxidase. It was shown in this system that both the membranes and the cytosol are needed for oxidase activity (236). With this system, it was found that some patients have a defect associated with the membranes, whereas others have a defect associated with the cytosol. Furthermore, it was established that patients with X-linked CGD were characterized by a membrane defect and absence of cytochrome b558. Autosomal inheritance, on the other hand, was associated with normal levels of b558 and a defect in the cytosol, which later was shown to have absence of p47phox or p67phox.
Analysis of a large number of patients and kindreds has shown that the majority of the patients (70%) have mutations affecting gp91phox or p22phox (236,237). Those with mutations affecting gp91phoxshow X-linked inheritance, and those with mutations affecting p22phox show autosomal inheritance. A great variety of mutations have been identified with gp91phox, including single-nucleotide insertions or deletions, creating frameshifts or premature stop codons, and deletions of part or all of an exon (238). The X-linked CGD is highly heterogeneous. Mutations of the CYBB gene usually lead to lack of gp91phox expression and absence of NADPH oxidase activity (known as X91°CGD). In others, the gp91phox protein is normally expressed but totally lacks oxidase activity (known as X91+CGD), and, finally, in another group of patients, gp91phox protein is decreased and oxidase activity is proportionally diminished (X91-CGD).
An interesting X91+ patient with an Arg54Ser mutation had a defect in the electron transport from flavin adenine dinucleotide to heme, but the translocation of p47phox to p67phox was normal (239).
In another patient, neutrophils, monocytes, and B cells had no oxidase activity, but eosinophils had normal activity (240). This patient probably had a mutation in a regulatory element of the CYBB gene directing the transcription to various cells. These experiments of nature have provided us with considerable information about the structure, assembly, and function of this very important enzyme, NADPH oxidase.
A patient has been reported with a mutation of Rac2, having deficiency in phagocytosis. The mutation (Asp57Asn) was negative-dominant and was associated with reduced production of superoxide and decreased cell movement (241). The mutant protein was unable to bind GTP. Only a small number of mutations of p47phox and p67phox have been reported. Deficiency of p47phox or p67phox is seen in 33% and 5% of all CGD cases, respectively. Infections in patients with CGD are treated aggressively with antibiotics and, more recently, with interferon-γ, which stimulates phagocyte activation (242).
Deficiency of Glutathione Reductase and Glutathione Synthetase
Deficiencies of GSH reductase and synthetase have been described. The patients have impaired neutrophil function but are not prone to bacterial infections. These enzymes are components of the GSH cycle, so that they convert H2O2 to H2O in the presence of reduced GSH, which is converted to GSSG. The GSSG regenerates reduced GSH in the presence of NADPH.
Defects in Granule Proteins
Myeloperoxidase
MPO is an important enzyme present in the azurophilic granules of neutrophils and in the lysosomes of monocytes. It generates oxidants from H2O2 and halides. The mature enzyme is a 150-kDa heterodimer composed of two heavy and two light chains. It is synthesized primarily during the promyelocytic stage of myeloid differentiation. Hereditary MPO deficiency is a benign abnormality that does not result in clinical illness despite the fact that MPO-deficient neutrophils exhibit delayed bacterial and fungal killing in vitro (243). MPO–messenger RNA transcripts are detected in neutrophils from patients, suggesting that deficiency of the enzyme is caused by some posttranslational defect. Because only neutrophils and monocytes have MPO, other cells (macrophages) may provide defense. It is also possible that the MPO-independent mechanisms are sufficient for protection (244).
Lactoferrin
Lactoferrin is present in the specific or secondary granules and is a member of the transferrin gene family. It is found in secretions such as tears, milk, saliva, and pancreatic secretions. It has antimicrobial activity through iron chelation and regulates neutrophil adhesiveness.
Some individuals have deficiency of the contents of specific granules. These patients are prone to recurrent bacterial and fungal infections of the skin and deep tissues. They are also deficient in defensins, a component of the primary granules.
Phagocytosis of Apoptotic Cells
Macrophages discriminate between viable and apoptotic cells. Uptake of apoptotic cells is noninflammatory, although certain receptors that recognize apoptotic cells are critical for innate immunity. The same receptors engulfing microbial organisms, however, trigger an inflammatory response. Receptors contributing to uptake of apoptotic cells are scavenger receptors class A and class B, receptors for oxidized low-density lipoprotein, integrins, and so forth (245,246). Scavenger receptors class A and B are multiligand and multifunctional receptors. Class A scavenger receptors have been implicated for recognition of apoptotic thymocytes and activated platelets. The cells undergoing apoptosis lose phospholipid asymmetry and expose phosphatidyl serine (PS). This change is required for recognition and engulfment (247). Normally, phospholipid asymmetry is maintained by the activity of an amino phospholipid translocase, a Mg2+-dependent ATPase. However, viable macrophages were shown to express PS constitutively on their surface. Annexin V, a PS-specific binding protein, inhibited phagocytosis of apoptotic cells but not of other particles (248). The anti-inflammatory effect in the clearance of apoptotic cells is an active process that is due to the release of transforming growth factor-β, IL-10, and prostaglandin E2 (245,249).
WASP is recruited to the phagocytic cup and plays an important and nonredundant role in clearance of apoptotic cells (250). Cdc42, which activates WASP, also contributes in this process (251). Defective clearance of apoptotic cells may explain the susceptibility of WAS patients to autoimmune disease, because aggregation of lipids on apoptotic cell membranes and their inefficient removal have been linked to development of autoantibody production (252) and autoimmune disorders.
A flow cytometric method for measuring phagocytosis of apoptotic cells has been developed (253). It has the advantages of assessment of large numbers of cells and high sample throughput and can distinguish bound from internalized apoptotic cells. Apoptotic cells are labeled with 5-chloromethylfluorescein diacetate.
Phagocytosis of Senescent Erythrocytes
Several processes have been implicated as causes of senescence of red blood cells (RBCs), such as loss of carbohydrates by desialylation, modification of natural components such as band 3, and glycophorin and phospholipid asymmetry with the appearance on the surface of PS (254). There are several reports regarding the nature of the antigen that triggers removal of the RBC as it approaches the end of its lifespan. The nature of a senescent epitope is not yet clear, but all major classes of organic matter have been implicated for the primary signal (i.e., proteins, carbohydrates, and lipids). Senescence is considered by some to be the unmasking of amino acid sequences by proteolysis or rearrangement due to clustering or oligomerization of band-3 protein (255). Others, however, have shown that the senescent antigen is not present on the polypeptide of band 3 but on a glycan (256). Strong evidence supports that β-galactosyl residues stimulate IgG autoantibodies and phagocytosis of aging RBCs (256). Presence of PS on the cell surface correlates directly with the propensity of RBCs to bind to monocytes and be rapidly cleared in the spleen (257).
A flow cytometric method has been developed for an accurate measurement of erythrophagocytosis (258).
Assessment of Phagocytic Function
Nitroblue Tetrazolium
The standard method of laboratory testing for phagocytosis is the reduction of a colorless substance known as nitroblue tetrazolium to a blue-black formazan-insoluble deposit within the neutrophils. This indicates the presence of an activated respiratory burst oxidase. In normal persons, <10% of blood neutrophils are nitroblue tetrazolium–positive, whereas in patients with bacterial infections, >10% of neutrophils are nitroblue tetrazolium–positive.
Flow Cytometry
Phagocytic function can be assessed by flow cytometry, which has the advantage over other methods because of the smaller number of cells and fewer preparative procedures required. One useful method uses fluorescein heat-killed Candida albicans. After phagocytosis, ethidium bromide is added. Because the ethidium bromide (red color) does not penetrate viable cells, only the external organisms stain red, whereas those phagocytosed stain green (fluorescein) (259).
Flow cytometry can also be used to measure oxidative mechanisms. The most useful method is the measurement of intracellular H2O2 by the dichlorofluorescein (DCFH) diacetate probe (260). The DCFH diacetate dye is incorporated into the lipid regions of the cell, where the acetate side chains are cleaved by hydrolytic enzymes to the nonfluorescence molecule, 2′,7′-DCFH, which becomes trapped within the cell. On cell activation, the RBO generates superoxide anion, which is further reduced to H2O2. In the presence of H2O2, peroxidases oxidize the DCFH to 2′,7′-dichlorofluorescein, which fluoresces at 530 nm (the same emission as fluorescein thioisocyanate). The green fluorescence produced is proportional to the amount of H2O2 generated.
Chemiluminescence
Some of the reactive oxygen intermediates generated during phagocytosis exist for a short period of time in a higher energy state but release this energy in the form of light known as CL, which can be quantified (261).
CL can be amplified by the use of bystander molecules called chemiluminescent probes, which react with the O2 radicals to produce excited products in high yield. The intensity of CL is directly proportional to the functional capacity of the phagocytes. The two most commonly used probes are luminol (5-amino-2,3-dihydro-1,4-phthalazinedione) and lucigenin (10,10-dimethyl-9,9-biacridinium dinitrate), and the most commonly used stimulus is zymosan (yeast particles).
CL is proportional to phagocytic activity as long as the CL probes and the stimulus are not rate-limiting. Contamination by erythrocytes reduces CL because the extinction coefficient of hemoglobin is in the blue region of the visible region and light is absorbed.
References
1. Chernyak L, Tauber AI. The birth of immunology: Metchnikoff, the embryologist. Cell Immunol 1988;117:218–233.
2. Robinson MS, Watts C, Zerial M. Membrane dynamics in endocytosis. Cell 1996;84:13–21.
3. Mukherjee S, Ghosh RN, Maxfield FR. Endocytosis. Physiol Rev 1997;77: 759–803.
4. Scales SJ, Gomez M, Kreis TE. Coat proteins regulating membrane traffic. Int Rev Cytol 2000;195:67–144.
5. Stuart LM, Ezekowitz RAB. Phagocytosis: elegant complexity. Immunity 2005; 22:539–550.
6. Platt N, Gordon S. Scavenger receptors: diverse activities and promiscuous binding of polyanionic ligands. Chem Biol 1998;5:R193–R203.
7. Indik ZK, Park JG, Hunter S, Schreiber AD. The molecular dissection of Fcγ receptor mediated phagocytosis. Blood 1995;86:4389–4399.
8. Ravetch JV, Kinet JP. Fc receptors. Annu Rev Immunol 1991;9:457–492.
9. Cambier JC. New nomenclature for the Reth motif (or ARH1/TAM/ARAM/ YXXL). Immunol Today 1995;16:110.
10. Indik ZK, Hunter S, Huang MM, et al. The high affinity Fcγ receptor (CD64) induces phagocytosis in the absence of its cytoplasmic domain: the γ subunit of FcγRIIIA imparts phagocytic function to FcγRI. Exp Hematol 1994;22: 599–606.
11. Ernst LK, Duchemin AM, Anderson CL. Association of the high-affinity receptor for IgG (FcγRI) with the γ subunit of the IgE receptor. Proc Natl Acad Sci U S A 1993;90:6023–6027.
12. Brooks DG, Qiu WQ, Luster AD, Ravetch JV. Structure and expression of human IgG FcRII (CD32). Functional heterogeneity is encoded by the alternatively spliced products of multiple genes. J Exp Med 1989;170:1369–1385.
13. Cassel DL, Keller MA, Surrey S, et al. Differential expression of FcγRIIA, FcγRIIB and FcγRIIC in hematopoietic cells: analysis of transcripts. Mol Immunol 1993;30:451–460.
14. Indik ZK, Pan XQ, Huang MM, et al. Insertion of cytoplasmic tyrosine sequences into the nonphagocytic receptor FcγRIIB establishes phagocytic function. Blood 1994;83:2072–2080.
15. Letourneur O, Kennedy IC, Brini AT, et al. Characterization of the family of dimers associated with Fc receptors (FcγRI and FcγRIII). J Immunol 1991; 147:2652–2656.
16. Wirthmueller U, Kurosaki T, Murakami MS, Ravetch JV. Signal transduction by FcγRIII (CD16) is mediated through the γ chain. J Exp Med 1992;175: 1381–1390.
17. Ravetch JV, Perussia B. Alternative membrane forms of FcγRIII (CD16) on human natural killer cells and neutrophils. Cell type-specific expression of two genes that differ in single nucleotide substitutions. J Exp Med 1989;170: 481–497.
18. Huizinga TW, Kerst M, Nuyens JH, et al. Binding characteristics of dimeric IgG subclass complexes to human neutrophils. J Immunol 1989;142:2359–2364.
19. Huizinga TW, de Haas M, Kleijer M, et al. Soluble Fc gamma receptor III in human plasma originates from release by neutrophils. J Clin Invest 1990;86:416–423.
20. Daeron M. Fc receptor biology. Annu Rev Immunol 1997;15:203–234.
21. Sampson LL, Heuser J, Brown EJ. Cytokine regulation of complement receptor-mediated ingestion by mouse peritoneal macrophages. M-CSF and IL-4 activate phagocytosis by a common mechanism requiring autostimulation by IFN-β. J Immunol 1991;146:1005–1013.
22. Griffin FM Jr, Mullinax PJ. Augmentation of macrophage complement receptor function in vitro. III. C3b receptors that promote phagocytosis migrate within the plane of the macrophage plasma membrane. J Exp Med 1981;154: 291–305.
23. Griffin FM Jr. Effects of soluble immune complexes on Fc receptor- and C3b receptor-mediated phagocytosis by macrophages. J Exp Med 1980;152: 905–919.
24. Nepomuceno RR, Henschen-Edman AH, Burgess WH, Tenner AJ. cDNA cloning and primary structure analysis of C1qR(P), the human C1q/MBL/SPA receptor that mediates enhanced phagocytosis in vitro. Immunity 1997;6:119–129.
25. Malhotra R, Willis AC, Jensenius JC, et al. Structure and homology of human C1q receptor (collectin receptor). Immunology 1993;78:341–348.
26. Malhotra R, Thiel S, Reid KB, Sim RB. Human leukocyte C1q receptor binds other soluble proteins with collagen domains. J Exp Med 1990;172:955–959.
27. Tenner AJ, Robinson SL, Ezekowitz RA. Mannose binding protein (MBP) enhances mononuclear phagocyte function via a receptor that contains the 126,000 M(r) component of the C1q receptor. Immunity 1995;3:485–493.
28. Ezekowitz RA, Sastry K, Bailly P, Warner A. Molecular characterization of the human macrophage mannose receptor: demonstration of multiple carbohydrate recognition-like domains and phagocytosis of yeasts in Cos-1 cells. J Exp Med 1990;172:1785–1794.
29. Schlesinger LS, Hull SR, Kaufman TM. Binding of the terminal mannosyl units of lipoarabinomannan from a virulent strain of Mycobacterium tuberculosis to human macrophages. J Immunol 1994;152:4070–4079.
30. Cross CE, Bancroft GJ. Ingestion of acapsular Cryptococcus neoformans occurs via mannose and β-glucan receptors, resulting in cytokine production and increased phagocytosis of the encapsulated form. Infect Immun 1995;63:2604–2611.
31. Prigozy TI, Sieling PA, Clemens D, et al. The mannose receptor delivers lipoglycan antigens to endosomes for presentation to T cells by CD1b molecules. Immunity 1997;6:187–197.
32. Wright SD, Ramos RA, Tobias PS, et al. CD14, a receptor for complexes of lipopolysaccharide (LPS) and LPS binding protein. Science 1990;249:1431–1433.
33. Wright SD, Tobias PS, Ulevitch RJ, Ramos RA. Lipopolysaccharide (LPS) binding protein opsonizes LPS-bearing particles for recognition by a novel receptor on macrophages. J Exp Med 1989;170:1231–1241.
34. Griffin FM Jr, Silverstein SC. Segmental response of the macrophage plasma membrane to a phagocytic stimulus. J Exp Med 1974;139:323–336.
35. Griffin FM Jr, Griffin JA, Leider JE, Silverstein SC. Studies on the mechanism of phagocytosis. I. Requirements for circumferential attachment of particle-bound ligands to specific receptors on the macrophage plasma membrane. J Exp Med 1975;142:1263–1282.
36. Griffin FM Jr, Griffin JA, Silverstein SC. Studies on the mechanism of phagocytosis. II. The interaction of macrophages with anti-immunoglobulin IgG-coated bone marrow-derived lymphocytes. J Exp Med 1976;144:788–809.
37. Garcia-Garcia E, Rosales C. Signal transduction during Fc receptor-mediated phagocytosis. J Leukoc Biol. 2002;72:1092–1108.
38. Korade-Mirnics Z, Corey SJ. Src kinase-mediated signaling in leukocytes. J Leukoc Biol 2000;68:603–613.
39. Garcia-Garcia E, Rosales C. Signal transduction during Fc receptor-mediated phagocytosis. J Leukoc Biol 2002;72:1092–1108.
40. Sobota A, Strzelecka-Kiliszek A, Glodkowska E, et al. Binding of IgG-opsonized particles to FcγR is an active stage of phagocytosis that involves receptor clustering and phosphorylation. J Immunol 2005;2175:4450–4457.
41. Crowley MT, Costello PS, Fitzer-Atlas CJ, et al. A critical role for Syk in signal transduction and phagocytosis mediated by Fcg receptors on macrophages. J Exp Med 1997;186:1027–1039.
42. Levecrier Y, Okkenhaug K, Sawyer C, et al. Class I phosphoimositide 3-kinase p110β is required for apoptotic cell and Fcγ receptor-mediated phagocytosis by macrophages. J Biol Chem 2003;278:38437–38442.
43. Wymann MP, Sozzani S, Altruda F, et al. Lipids on the move: phosphoinositide 3-kinases in leukocyte function. Immunol Today 2000;21:260–264.
44. Cox D, Tseng CC, Bjekic C, Greenberg S. A requirement for phosphatidylinositol 3-kinase on pseudopod extension. J Biol Chem 1999;274:1240–1247.
45. Cox D, Berg JS, Cammer M, et al. Myosin X is a downstream effector of PI(3)K during phagocytosis. Nat Cell Biol 2002;4:469–477.
46. Lundqvist-Gustafsson H, Gustafsson M, Dahlgren C. Dynamic Ca2+ changes in neutrophil phagosomes. A source for intracellular Ca2+ during phagolysosome formation. Cell Calcium 2000;27:353–362.
47. Boelho RJ, Teruel M, Diekman R, et al. Localized biphasic changes in phosphatidyl-4,5-biphosphate at sites of phagocytosis. J Cell Biol 2000;151: 1353–1368.
48. Cox D, Chang P, Zhang D, et al. Requirement for both Rac1 and Cdc42 in membrane ruffling and phagocytosis in leukocytes. J Exp Med 1997;186: 1487–1489.
49. Caron E, Hall A. Identification of two distinct mechanisms of phagocytosis controlled by different Rho GTPases. Science 1998;282:1717–1721.
50. Bokoch GM. Regulation of innate immunity by Rho GTPases. Trends Cell Biol 2005;15:163–171.
51. Chimini G, Chavrier P. Function of Rho family and protein actin dynamics during phagocytosis and engulfment. Nat Cell Biol 2000;2:E191–E196.
52. Serokina EM, Chernoff J. Rho-GTPases: new members, new pathways. J Cell Biochem 2005;94:225–231.
53. Hoppe AD, Swanson JA. Cdc42, and Rac2 display distinct patterns of activation during phagocytosis. Mol Biol Cell 2004;15:3509–3519.
54. Bokoch GM, Diebold BA. Current molecular models for NADPH oxidase regulation by RacGTPase. Blood 2002;100:2692–2696.
55. Gasque P. Complement: a unique innate immune sensor for danger signals. Mol Immunol 2004;41:1089–1098.
56. Wiedemann A, Patel JC, Lim J, et al. Two distinct cytoplasmic regions of the β2 integrin chain regulate RhoA function during phagocytosis. J Cell Biol 2006;172:1069–1079.
57. Bristelo XR. Regulatory and signaling properties of the Vav family. Mol Cell Biol 2000;20;1461–1477.
57a. Hall AB, Gakidis MAM, Gloganer M, et al. Requirements for Vav guanine nucleotide exchange factors and Rho GTPases in FcγR- and complement-mediated phagocytosis. Immunity 2006;24:305–316.
58. Lorenzi R, Brickell PM, Katz DR, et al. Wiskott-Aldrich syndrome protein is necessary for efficient IgG-mediated phagocytosis. Blood 2000;95:293–294.
59. Leverrier Y, Lorenzi R, Blundell MP, et al. Cutting edge: the Wiskott-Aldrich syndrome protein is required for efficient phagocytosis of apoptotic cells. J Immunol 2001;166:4831–4834.
60. Long EO. Regulation of immune responses through inhibitory receptors. Annu Rev Immunol 1999;17:875–904.
61. Bruhns P, Vely F, Malbec O, et al. Molecular basis of the recruitment of the SH2 domain-containing inositol 5-phosphatases SHIP-1 and SHIP-2 by FcγIIB. J Biol Chem 2000;275:37357–37364.
62. Lesourne R, Bruhns P, Fridman WH, Daeron M. Insufficient phosphorylation prevents FcγRIIB from recruiting the SH2 domain-containing protein tyrosine phosphatase SHP-1. J Biol Chem 2001;276:6327–6336.
63. Cox D, Dale BM, Kashiwada M, et al. A regulatory role for Src homology 2 domain-containing inositol 5′-phosphatase (SHIP) in phagocytosis mediated by Fcγ receptors and complement receptor 3 (αMβ2; CD11b/CD18). J Exp Med 2001;193:61–72.
64. Trindandapani S, Siefker K, Teilland JL, et al. Regulated expression and inhibitory function of FcγRIIb in human monocytic cells. J Biol Chem 2002;277:5082–5089.
65. Holevinsky KO, Nelson DJ. Membrane capacitance changes associated with particle uptake during phagocytosis in macrophages. Biophys J 1998;75: 2577–2586.
66. Beningo KA, Wang Y-L. Fc receptor-mediated phagocytosis is regulated by mechanical properties of the target. J Cell Sci 2002;115:849–856.
67. Greenberg S. Modular components of phagocytosis. J Leukoc Biol 1999;66: 712–715.
68. Lennartz MR, Yuen AF, Masi SM, et al. Phospholipase A2 inhibition results in sequestration of plasma membrane into electron lucent vesicles during IgG-mediated phagocytosis. J Cell Sci 1997;110:2041–2052.
69. Niedergang F, Chevrier P. Signaling and membrane dynamics during phagocytosis: many roads lead to the phagos (R) ome. Curr Opin Cell Biol 2004;16: 422–428.
70. Booth JW, Trimble WS, Grinstein S. Membrane dynamics in phagocytosis. Semin Immunol 2001;13:357–364.
71. Araki N, Johnson MT, Swanson JA. A role of phosphoinositide-3 kinase in the completion of macropinocytosis and phagocytosis by macrophages. J Cell Biol 1996;135:1249–1260.
72. Cox D, Lee DJ, Dale BM, et al. A Rab 11-containing rapidly recycling compartment in macrophages that promotes phagocytosis. Proc Natl Acad Sci U S A 2000;97:680–685.
73. McNew JA, Parlati F, Fukuda R, et al. Compartmental specificity of cellular membrane fusion encoded in SNARE proteins. Nature 2000;407:153–159.
74. Zerial M, McBride H. Rab proteins as membrane organizers. Nat Rev Mol Cell Biol 2001;2:107–117.
75. Gold ES, Underhill DM, Morrisette NS, et al. Dynamin 2 is required for phagocytosis in macrophages. J Exp Med 1999;190:1849–1856.
76. Gold ES, Morrissette NS, Underhill DM, et al. Amphiphysin IIm, a novel amphiphysin II isoform, is required for macrophage phagocytosis. Immunity 2001;12:285–292.
77. Cougoule C, Hoshino S, Dart A, et al. Dissociation of recruitment and activation of the small G-protein Rac during Fcγ receptor-mediated phagocytosis. J Biol Chem 2006;281:8756–8764.
78. Berg JS, Powell BC, Cheney RE. A millennial myosin census. Mol Biol Cell 2001;12:780–784.
79. Pitt A, Mayorga LS, Stahl PD, Schwartz AL. Alterations in the protein composition of maturing phagosomes. J Clin Invest 1992;90:1978–1983.
80. Ward DM, Perou CM, Lloyd M, Kaplan J. “Synchronized” endocytosis and extracellular sorting in alveolar macrophages: the early sorting endosome is a transient organelle. J Cell Biol 1995;129:1229–1240.
81. Gotthardt D, Warnatz HJ, Henschel O, et al. High resolution dissection of phagosome maturation reveals distinct membrane trafficking phases. Mol Biol Cell 2002;13:3508–3520.
82. Desjardins M, Huber LA, Parton R, Griffiths G. Biogenesis of phagosomes proceeds through a sequential series of interactions with the endocytic apparatus. J Cell Biol 1994;124:677–688.
83. East L, Isacke CM. The mannose receptor family. Biochim Biophys Acta 2002;1572;364–386.
84. LeCabec V, Emorine LJ, Toesca I, et al. The human macrophage mannose receptor is not a professional phagocytic receptor. J Leukoc Biol 2005;77:934–943.
85. Pollard TD, Blanchoin L, Mullins RD. Molecular mechanisms controlling actin filament dynamics in non-muscle cells. Annu Rev Biophys Biomol Struct 2000;29:545–576.
86. Welch MD, Mullins RD. Cellular control of actin nucleation. Annu Rev Cell Dev Biol 2002;18:247–288.
87. Higgs HN, Pollard TD. Regulation of actin filament network formation through Arp2/3 complex: activation by a diverse array of proteins. Annu Rev Biochem 2001;70:649–675.
88. Robinson RC, Turbedsky K, Kaiser DA, et al. Crystal structure of Arp2/3 complex. Science 2001;294:1679–1684.
89. Shao D, Forge A, Munro PMG, Bailly M. Arp2/3 complex-mediated actin polymerization occurs on specific pre-existing networks in cells and requires spatial restriction to sustain functional lamellipod extension. Cell Motility Cytoskeleton 2006;63:395–414.
90. Martin AC, Welch MD, Drubin DG. Arp2/3 ATP hydrolysis-catalysed branch dissociation is critical for endocytic force generation. Nat Cell Biol 2006 8:826–832.
91. Mullins RD, Heuser JA, Pollard TD. The interaction of Arp2/3 complex with actin: nucleation, high-affinity pointed end capping, and formation of branching network filaments. Proc Natl Acad Sci U S A 1998;95:6181–6186.
92. Takenawa T, Miki H. WASP and WAVE family proteins: key molecules for rapid rearrangement of cortical actin filaments and cell movement. J Cell Sci 2001;114:1801–1809.
93. Stradal TEB, Scita G. Protein complexes regulating Arp 2/3-mediated actin assembly. Curr Opin Cell Biol 2005;18:4–10.
94. Higgs HN, Pollard TD. Activation by Cdc42 and PIP2 of Wiskott-Aldrich syndrome protein (WASP) stimulates actin nucleation of Arp2/3 complex. J Cell Biol 2000;150:1311–1320.
95. Rohatgi R, Ho HY, Kirschner MW. Mechanism of N-WASP activation by Cdc42 and phosphatidylinositol-4,5-biphosphate. J Cell Biol 2000;150:1299–1310.
96. Anton IM, Jones GE. WIP: a multifunctional protein involved in actin cytoskeleton regulation. Eur J Cell Biol 2006;85:295–304.
97. Soderling SH, Scott JD. WAVE signaling: from biochemistry to biology. Biochem Soc Trans 2006;34:73–76.
98. Marchand JB, Kaiser DA, Pollard TD, Higgs HN. Interaction of WASP/Scar proteins with actin and vertebrate Arp2/3 complex. Nat Cell Biol 2001;3:76–82.
99. Hüfer K, Higgs HN, Pollard TD, et al. The verpolin-like central (VC) regions of Wiskott-Aldrich syndrome protein induces Arp2/3 complex-dependent actin nucleation. J Biol Chem 2001;276:35761–35767.
100. Weaver AM, Young ME, Lee W-L, Cooper JA. Integration of signals to the Arp2/3 complex. Curr Opin Cell Biol 2003;15:23–30.
101. Thrasher AJ. WASp in immune-system organization and function. Nat Rev Immunol 2002;2:635–646.
102. Prehoda KE, Scott JA, Mullins RD, Lim WA. Integration of multiple signals through cooperative regulation of the N-WASP-Arp2/3 complex. Science 2000;290:801–806.
103. Rohatgi R, Nollau P, Ho HY, et al. Nck and phosphatidylinositol-4,5-biphosphate synergistically activate actin polymerization through the N-WASP-Arp2/3 pathway. J Biol Chem 2001;276:26448–26452.
104. Svitkina TM, Borisy GG. Arp2/3 complex and actin depolymerizing factor/cofilin in dendritic organization and treadmilling of actin filament array in lamellipodia. J Cell Biol 1999;145:1009–1026.
105. Machesky LM, Mullens RD, Higgs HN, et al. Scar, a WASP-related protein, activates nucleation of actin filaments by the Arp2/3 complex. Proc Natl Acad Sci U S A 1999;96:3739–3744.
106. Pollard TD, Blanchoin L, Mullins RD. Molecular mechanisms controlling actin filament dynamics in non-muscle cells. Annu Rev Biophys Biomol Struct 2000;29:545–576.
107. Pantaloni D, Boujemaa R, Didry D, et al. The Arp2/3 complex branches filament barbed ends: functional antagonism with capping proteins. Nat Cell Biol 2000;2:385–391.
108. Blanchoin L, Amann KJ, Higgs HN, et al. Direct observation of dendritic actin filament networks nucleated by Arp2/3 complex and WASP/Scar proteins. Nature 2000;404:1007–1011.
109. Amann KJ, Pollard TD. Direct real-time observation of actin filament branching mediated by Arp2/3 complex using total internal reflection fluorescence microscopy. Proc Natl Acad Sci U S A 2001;98:15009–15013.
110. Ichetovkin I, Grant W, Condeelis J. Cofilin produces newly polymerized actin filaments that are preferred for dendritic nucleation by the Arp2/3 complex. Curr Biol 2002;12:79–84.
111. Amann KJ, Pollard TD. The Arp2/3 complex nucleates actin filament branches from the sides of pre-existing filaments. Nat Cell Biol 2001;3: 306–310.
112. Uruno T, Liu J, Zhang P, et al. Activation of Arp2/3 complex-mediated actin polymerization by cortactin. Nat Cell Biol 2001;3:259–266.
113. Weaver AM, Karginov AV, Kinley AW, et al. Cortactin promotes and stabilizes Arp2/3-induced actin filament network formation. Curr Biol 2001;11: 370–374.
114. Weaver AM, Heuser JE, Karginov AV, et al. Interaction of cortactin and N-WASp with Arp2/3 complex. Curr Biol 2002;12:1270–1278.
115. Higgs HN. Actin nucleation: cortactin caught in the act. Curr Biol 2002;12: R593–R595.
116. Kim AS, Kakalis LT, Abdul-Manan N, et al. Autoinhibition and activation mechanisms of the Wiskott-Aldrich syndrome protein. Nature 2000;404:151–158.
117. May RC, Machesky LM. Phagocytosis and the actin cytoskeleton. J Cell Sci 2001;114:1061–1077.
118. Castellano F, Chavrier P, Caron E. Actin dynamics during phagocytosis. Semin Immunol 2001;13:347–355.
119. Snapper SB, Rosen FS. The Wiskott-Aldrich syndrome protein (WASP). Roles in signaling and cytoskeletal organization. Annu Rev Immunol 1999;17: 905–929.
120. Machesky LM, Insall RH. Scar1 and the related Wiskott-Aldrich syndrome protein, WASP, regulate the actin cytoskeleton through the Arp2/3 complex. Curr Biol 1998;8:1347–1356.
121. Castellano F, Le Clainche C, Patin D, et al. A WASp-VASP complex regulates actin polymerization at the plasma membrane. EMBO J 2001;20:5603–5614.
122. Lorenzi R, Brichell PM, Katz DR, et al. Wiskott-Aldrich syndrome protein is necessary for efficient IgG mediated phagocytosis. Blood 2000;95:2943–2946.
123. Natarangelo LD. Missense mutations of the WASP gene cause intermittent X-linked thrombocytopenia. Blood 2002;99:2268–2269.
124. Snapper SB, Rosen FS, Mizoguchi E, et al. Wiskott-Aldrich syndrome protein-deficient mice reveal a role for WASP in T but not B cell activation. Immunity 1998;9:81–90.
125. Cianferoni A, Massaad M, Feske S, et al. Defective nuclear translocation of nuclear factor of activated T cells and extracellular signal-regulated kinase underlies deficient IL-2 gene expression in Wiskott-Aldrich syndrome. J Allergy Clin Immunol 2005;116:1364–1371.
126. May RC, Caron E, Hall A, Machesky LM. Involvement of Arp2/3 complex in phagocytosis mediated by Fc gamma R or CR3. Nat Cell Biol 2000;2:246–248.
127. Taunton J. Actin filament nucleation by endosomes, lysosomes, and secretory vesicles. Curr Opin Cell Biol 2001;13:85–91.
128. Taunton J, Rowning BA, Coughlin ML, et al. Actin-dependent propulsion of endosomes and lysosomes by recruitment of N-WASP. J Cell Biol 2000; 148:519–530.
129. Merrifield CJ, Moss SE, Ballestrem C, et al. Endocytic vesicles more at the tips of actin tails in cultured mast cells. Nat Cell Biol 1999;1:72–74.
130. Dramsi S, Cossart P. Intracellular pathogens and the actin cytoskeleton. Annu Rev Cell Dev Biol 1998;14:137–166.
131. Cameron LA, Svitkina TM, Vignjevic D, et al. Dendritic organization of actin comet tails. Curr Biol 2001;11:130–135.
132. Borisy GG, Svitkina TM. Actin machinery: pushing the envelope. Curr Opin Cell Biol 2000;12:104–112.
133. Nachmias VT. Small actin-binding proteins: the β-thymosin family. Curr Biol 1993;5:56–62.
134. Pantaloni D, Carlier MF. How profilin promotes actin filaments assembly in the presence of thymosin-β4. Cell 1993;75:1007–1014.
135. Bamburg JR. Proteins of the ADF/cofilin family: essential regulators of actin dynamics. Annu Rev Cell Dev Biol 1999;15:185–230.
136. Carlier M-F, Ressad F, Pantaloni D. Control of actin dynamics in cell motility. Role of ADF/cofilin. J Biol Chem 1999;274:33827–33830.
137. Lappalainen P, Fedorov EV, Fedorov AA, et al. Essential functions and actin-binding surfaces of yeast cofilin by systematic mutagenesis. EMBO J 1997;16:5520–5530.
138. Puius YA, Mahoney NM, Almo SC. The modular structure of actin-regulatory proteins. Curr Opin Cell Biol 1998;10:23–24.
139. Kusano K, Abe H, Obinata T. Detection of a sequence involved in actin-binding and phosphoinositide-binding in the N-terminal side of cofilin. Mol Cell Biochem 1999;190:133–141.
140. McGough A, Pope B, Chiu W, Weeds AG. Cofilin changes the twist of F-actin: implications for actin filament dynamics and cellular function. J Cell Biol 1997;138:771–781.
141. Teubner A, Wagner A. Kinetic evidence for a readily exchangeable nucleotide at the terminal subunit of the barbed ends of actin filaments. Biochemistry 1998;37:7532–7538.
142. Didry D, Carlier M-F, Pantaloni D. Synergy between actin depolymerizing factor/cofilin and profilin in increasing actin filament turnover. J Biol Chem 1998;273:25602–25611.
143. Sun HQ, Yamamoto M, Mejillano M, Yin HL. Gelsolin, a multifunctional actin regulatory protein. J Biol Chem 1999;274:33179–33182.
144. Selden LA, Kinosian HJ, Newman J, et al. Severing of F-actin by the amino-terminal half of gelsolin suggests internal cooperativity in gelsolin. Biophys J 1998;75:3092–3100.
145. Southwick FS. Gelsolin and ADF/cofilin enhance the actin dynamics of motile cells. Proc Natl Acad Sci U S A 2000;97:6936–6938.
146. Kwiatkowski DJ. Functions of gelsolin: motility, signaling, apoptosis, cancer. Curr Opin Cell Biol 1999;11:103–108.
147. Choe H, Burtnick LD, Mejillano M, et al. The calcium activation of gelsolin: insights from the 3Å structure of the G4-G6/actin complex. J Mol Biol 2002;324:691–702.
148. Dawson JF, Sablin EP, Spudich JA, Fletterick RJ. Structure of an F-actin trimer disrupted by gelsolin and implications for the mechanism of severing. J Biol Chem 2003;278:1229–1238.
149. Jammey PA, Iida K, Yin HL, Stossel TP. Polyphosphoinositide micelles and polyphosphoinositide-containing vesicles dissociate endogenous gelsolin-actin complexes and promote actin assembly from the fast-growing end of actin filaments blocked by gelsolin. J Biol Chem 1987;262:12228–12236.
150. Hartwig JH, Bokoch GM, Carpenter CL, et al. Thrombin receptor ligation and activated Rac uncap actin filament barbed ends through phosphoinositide synthesis in permeabilized human platelets. Cell 1995;82:643–653.
151. Barkalow K, Witke W, Kwiatkowski DJ, Hartwig JH. Coordinated regulation of platelet actin filament barbed ends by gelsolin and capping protein. J Cell Biol 1996;134:389–399.
152. Li Z, Kim ES, Bearer EL. Arp2/3 complex is required for actin polymerization during platelet shape change. Blood 2002;99:4466–4474.
153. Young JD, Ko SS, Cohn ZA. The increase of intracellular free calcium associated with IgG gamma 2b/gamma 1 Fc receptor-ligand interactions: role in phagocytosis. Proc Natl Acad Sci U S A 1984;81:530–534.
154. Zimmerli S, Majeed M, Gustavsson M, et al. Phagosome-lysosome fusion is a calcium-independent event in macrophages. J Cell Biol 1996;132:49–61.
155. Brumell JH, Howard JC, Craig K, et al. Expression of the protein kinase C substrate pleckstrin in macrophages: association with phagosomal membranes. J Immunol 1999;163:3388–3395.
156. Carballo E, Pitterle DM, Stumpo DJ, et al. Phagocytic and macropinocytic activity in MARCKS-deficient macrophages and fibroblasts. Am J Physiol 1999;277:C163–C173.
157. Czech MP. PIP2 and PIP3: complex roles at the cell surface. Cell 2000;100:603–606.
158. Vanhaesebroeck B, Leevers SJ, Ahmadi K, et al. Synthesis and function of 3-phosphatidylinositol lipids. Annu Rev Biochem 2001;70:535–602.
159. Gillooly DJ, Simonsen A, Stenmark H. Phosphoinositides and phagocytosis. J Cell Biol 2001;155:15–18.
160. Martin TF. Phosphoinositide lipids as signaling molecules: common themes for signal transduction, cytoskeletal regulation, and membrane trafficking. Annu Rev Cell Dev Biol 1998;14:231–264.
161. Strzelecka A, Pyrzynska B, Kwiatkowska K, Sobota A. Syk kinase, tyrosine phosphorylated proteins and actin filaments accumulate at forming phagosomes during Fc gamma-receptor-mediated phagocytosis. Cell Motil Cytoskeleton 1997;38:287–296.
162. Kwiatkowska K, Sobata A. Signaling pathways in phagocytosis. Bioassays 1999;21:422–431.
163. Cox T, Tseng CC, Bjekic G, Greenberg S. A requirement for phosphatidylino-sitol 3-kinases in pseudopod extension. J Biol Chem 1999;274:1240–1247.
164. Vieira OV, Botelho RJ, Rameh J, et al. Distinct roles of class I and III phosphatidylinositol 3-kinases in phagosome formation and maturation. J Cell Biol 2001;155:19–26.
165. Fratti RA, Backer JM, Reenberg J, et al. Role of phosphatidylinositol 3-kinase and Rab5 effectors in phagosome biogenesis and mycobacterial phagosome maturation arrest. J Cell Biol 2001;154:631–644.
166. Hall A. Rho GTPases and the actin cytoskeleton. Science 1998;279:509–514.
167. Tapon N, Hall A. Rho, Rac and Cdc42 GTPases regulate the organization of the actin cytoskeleton. Curr Opin Cell Biol 1997;9:86–92.
168. Cox D, Chang P, Zhang Q, et al. Requirements for both rac1 and Cdc42 in membrane ruffling and phagocytosis in leukocytes. J Exp Med 1997;186: 1487–1494.
169. Massol P, Montcourrier P, Guillemot JC, Chavrier P. Fc receptor-mediated phagocytosis required CDC42 and Rac1. EMBO J 1998;17:6219–6229.
170. Caron E, Hall A. Identification of two distinct mechanisms of phagocytosis controlled by different Rho GTPases. Science 1995;282:1717–1721.
171. Machesky LM, Insall RH. Scar1 and the related Wiskott-Aldrich syndrome protein, WASP, regulate the actin cytoskeleton through the Arp2/3 complex. Curr Biol 1998;8:1347–1356.
172. Higgs HN, Pollard TD. Activation by Cdc42 and PIP(2) of Wiskott-Aldrich syndrome protein (WASp) stimulates actin nucleation by Arp2/3 complex. J Cell Biol 2000;150:1311–1320.
173. Rohatgi R, Ho H-Y, Kirschner MW. Mechanisms of N-WASP activation by CDC42 and phosphatidylinositol 4,5-biphosphate. J Cell Biol 2000;150:1299–1310.
174. Kim AS, Kahalis LT, Abdul-Manan N, et al. Autoinhibition and activation mechanisms of the Wiskott-Aldrich syndrome protein. Nature 2000;404:151–158.
175. Arcaro A. The small GTP-binding protein Rac promotes dissociation of gelsolin from actin filaments in neutrophils. J Biol Chem 1998;273:805–813.
176. Tsukita S, Yonemura S. ERM (ezrin/radixin/moesin) family: from cytoskeleton to signal transduction. Curr Opin Cell Biol 1997;9:70–75.
177. Arpin M, Algrain M, Louvard D. Membrane-actin microfilament connections: an increasing diversity of players related to band 4.1. Curr Opin Cell Biol 1994;6:136–141.
178. Gary R, Bretscher A. Ezrin self-association involves binding of an N-terminal domain to a normally masked C-terminal domain that includes the F-actin binding site. Mol Biol Cell 1995;6:1061–1075.
179. Gupton SL, Waterman-Storer CM. Spatiotemporal feedback between actomyosin and focal-adhesion systems optimizes rapid cell migration. Cell 2006;125:1361–1374.
180. Greenberg S, Burridge K, Silverstein SC. Co-localization of F-actin and talin during Fc receptor-mediated phagocytosis in mouse macrophages. J Exp Med 1990;172:1853–1856.
181. Lehrer RI. Primate defensins. Nat Rev Microbiol 2004;2:727–738.
181a. Klotman ME, Chang TL. Defensins in innate antiviral immunity. Nat Rev Immunol 2006;6:447–456.
182. Babior BM, Kipnes RS, Curnutte JT. Biological defense mechanisms. The production by leukocytes of superoxide, a potential bactericidal agent. J Clin Invest 1973;52:741–744.
183. Baldridge CW, Gerard RW. The extra respiration of phagocytosis. Am J Physiol 1933;103:235–236.
184. Klebanoff SJ. Oxygen metabolism and the toxic properties of phagocytes. Ann Intern Med 1980;93:480–489.
185. Klebanoff SJ. Antimicrobial mechanisms in neutrophilic polymorphonuclear leukocytes. Semin Hematol 1975;12:117–142.
186. Babior BM. Oxidants from phagocytes: agents of defense and destruction. Blood 1984;64:959–966.
187. Fantone JC, Ward PA. Role of oxygen-derived free radicals and metabolites in leukocyte-dependent inflammatory reactions. Am J Pathol 1982;107:397–418.
188. Geiszt M. NADPH oxidases: new kids on the block. Cardiovascular Res 2006;71:289–299.
188a. Donko A, Peterfi Z, Sum A, et al. Dual oxidases. Phil Trans R Soc B 2005;360: 2301–2308.
189. Cross AR, Segal AW. The NADPH oxidase of professional phagocytes-prototype of the Nox electron transport chain systems. Biochim Biophys Acta 2004; 1657:1–22.
190. Yu L, Quinn MT, Cross AR, Dinauer MC. Gp91-phox is the heme binding subunit of the superoxide-generating NADPH oxidase. Proc Natl Acad Sci U S A 1998;95:7993–7998.
191. Hurst JK, Loehr TM, Curnutte JT, Rosen H. Resonance raman and electron paramagnetic resonance structural investigations of neutrophil cytochrome b558. J Biol Chem 1991;266:1627–1634.
192. Segal AW, West I, Wientjes F, et al. Cytochrome b-245 is a flavocytochrome containing FAD and the NADPH-binding site of the microbicidal oxidase of phagocytes. Biochem J 1992;284:781–788.
193. Rotrosen D, Yeung CL, Leto TL, et al. Cytochrome b558. The flavin-binding component of the phagocyte NADPH oxidase. Science 1992;256:1459–1462.
194. Parkos CA, Dinauer MC, Jesaitis A, et al. Absence of both the 91 KD and 22 KD subunits of human neutrophil cytochrome b in two genetic forms of chronic granulomatous disease. Blood 1989;73:1416–1420.
195. Porter C, Parkar M, Verhoeven A, et al. p22-phox-deficient chronic granulomatous disease: reconstitution by retrovirus-mediated expression and identification of a biosynthetic intermediate of gp91-phox. Blood 1994;84: 2767–2775.
196. Yu L, Zhen L, Dinauer MC. Biosynthesis of the phagocyte NADPH oxidase cytochrome b558. Role of heme incorporation and heterodimer formation in maturation and stability of gp91phox and p22phox subunits. J Biol Chem 1997;272:27288–27294.
197. Yu L, Cross AR, Zhen L, Dinauer MC. Functional analysis of NADPH oxidase in granulocytic cells expressing a Δ488-497 gp91phox deletion mutant. Blood 1999;94:2497–2504.
198. Susimoto H, Hata K, Mizuki K, et al. Assembly and activation of the phagocytic NADPH oxidase. J Biol Chem 1996;271:22152.
199. Leto TL, Adams AG, de Mendez I. Assembly of the phagocyte NADPH oxidase: binding of Src homology 3 (SH3) domains to proline-rich targets. Proc Natl Acad Sci U S A 1994;91:10650–10654.
200. Ellson CD, Gobert-Gosse S, Anderson KE, et al. PtdIns(3)P regulates the neutrophil oxidase complex by binding to the P domain of p40phox. Nat Cell Biol 2001;3:679–682.
201. Penting CP. Novel domains in NADPH oxidase subunits, sorting nexins and PtdIns 3-kinases: binding partners of SH3 domains? Protein Sci 1996;5: 2353–2357.
202. Cheever ML, Sato TK, de Beer T, et al. Phox domain interaction with PtdIns(3)P targets the Vam7 t-SNARE to vacuole membranes. Nat Cell Biol 2001;3:613–618.
203. Kanai F, Liu H, Field SJ, et al. The PX domains of p47phox and p40phox phagosome formation and maturation bind to lipid products of PI(3)K. Nat Cell Biol 2001;3:675–678.
204. Hirsch E, Katanaev VL, Garlanda C, et al. Central role for G protein-coupled phosphoinositide 3-kinase gamma in inflammation. Science 2000;287: 1049–1053.
205. Foust LP, El Benna J, Babior BM, et al. The phosphorylation targets of p47phox, a subunit on the respiratory burst oxidase. Functions of the individual target serines as evaluated by site-directed mutagenesis. J Clin Invest 1995;96: 1499–1505.
206. Inanami O, Johnson JL, McAdara JK, et al. Activation of the leukocyte NADPH oxidase by phorbol ester requires the phosphorylation of p47phox on serine 303 and 304. J Biol Chem 1998;273:9539–9543.
207. Dang PM-C, Cross AR, Quinn MT, Babior BM. Assembly of the neutrophil respiratory burst oxidase: a direct interaction between p67phox and cytochrome b558 II. Proc Natl Acad Sci U S A 2002;99:4262–4265.
208. Karathanassis D, Stahelin RV, Bravo J, et al. Binding of the PX domain of p47phox to phosphatidylinositol 3,4-bisphosphate and phosphatidic acid is masked by an intramolecular interaction. EMBO J 2002;21:5057–5068.
209. Zhan Y, Virbasius JV, Song X, et al. The p40phox and p47phox PX domains of NADPH oxidase target cell membranes via direct and indirect recruitment by phosphoinositides. J Biol Chem 2002;277:4512–4518.
210. Ellson CD, Andrews S, Stephens LR, Hawkins PT. The PX domains: a new phosphoinositide-binding module. J Cell Sci 2002;115:1099–1105.
211. Ago T, Takeya R, Hiroaki H, et al. The PX domain as a novel phosphoinositide-binding molecule. Biochem Biophys Res Commun 2001;287:733–738.
212. Massenet C, Chenaras S, Cohen-Adad C, et al. Effects of p47phox terminus phosphorylations on binding interactions with p40phox and p67phox. J Biol Chem 2005;280:13752–13761.
213. Mizreki K, Takeya R, Kwibayashi F, et al. A region C-terminal to the proline-rich core of p47phox regulates activation of the phagocyte NADPH oxidase by interacting with the C-terminal SH3domain of p67phox. Arch Biochem Biophys 2005;444:185–194.
214. McPhail LC. SH3-dependent assembly of the phagocyte NADPH oxidase. J Exp Med 1994;180:2011–2015.
215. Finan P, Shimizu Y, Gout I, et al. An SH3 domain and proline-rich sequence mediate an interaction between two components of the phagocyte NADPH oxidase complex. J Biol Chem 1994;269:13752–13755.
216. de Mendez I, Garrett MC, Adams AG, Leto TL. Role of p67phox SH3 domain in assembly of the NADPH oxidase system. J Biol Chem 1994;269:16326–16332.
217. Kuribayashi F, Nunoi H, Wakamatsu K, et al. The adaptor protein p40phox as a positive regulator of the superoxide-producing phagocyte oxidase. EMBO J 2002;21:6312–6320.
218. Huang J, Kleinberg ME. Activation of the phagocyte NADPH oxidase protein p47phox. Phosphorylation controls SH3 domain dependent binding to p22phox. J Biol Chem 1999;274:19731–19737.
219. Nobuhisa I, Takeya R, Ogura K, et al. Activation of the superoxide-producing phagocyte NADPH oxidase requires cooperation between the tandem SH3 domains of p47phox in recognition of a polyproline type II helix and an adjacent α-helix of p22phox. Biochem J 2006;396:183–192.
220. Ogura K, Nobuhisa I, Yuzawa S, et al. NMR solution structure of the tandem Src homology 3 domains of p47phox complexed with a p22phox-derived proline-rich peptide. J Biol Chem 2006;281:3660–3668.
221. Watanabe Y, Tsuboi H, Koyama M, et al. Molecular dynamics study on the ligand recognition by tandem SH3 domains of p47phox regulating NADPH oxidase activity. Comp Biol Chem 2006;30:303–312.
222. Freeman JL, Abo A, Lambeth JD. Rac “insert region” is a novel effector region that is implicated in the activation of NADPH oxidase, but not PAK65. J Biol Chem 1996;271:19794–19801.
223. Nisimoto Y, Freeman JLR, Motalebi SA, et al. Rac binding to p67phox. Structural basis for interactions of the Rac1 effector region and insert region with components of the respiratory burst oxidase. J Biol Chem 1997;272: 18834–18841.
224. Kim C, Dinauer MC. Rac2, an essential regulator of neutrophil nicotinamide adenine dinucleotide phosphate oxidase activation in response to specific signaling pathways. J Immunol 2001;166:1223–1232.
225. Diebold BA, Bokoch GM. Molecular basis for Rac2 regulation of phagocyte NADPH oxidase. Nat Immun 2002;2:211–215.
226. Nisimoto Y, Motalebi S, Han C-H, Lambeth J. The p67phox activation domain regulates electron transport flow from NADPH to flavin in flavocytochrome b558. J Biol Chem 1999;274:22999–23055.
227. Han C-H, Freeman JLR, Lee T, et al. Regulation of the neutrophil respiratory burst oxidase. Identification of an activation domain in p67phox. J Biol Chem 1998;273:16663–16668.
228. De Leo FR, Ulman KV, Davis AR, et al. Assembly of the human neutrophil NADPH oxidase involves binding of p67phox and flavocytochrome b to a common functional domain in p47phox. J Biol Chem 1996;271:17013–17020.
229. Ebisu K, Nagasawa T, Watanabe K, et al. Fused p47phox and p67phox truncations efficiently reconstitute NADPH oxidase and higher activity and stability than the individual components. J Biol Chem 2001;276:24498–24505.
230. Kobayashi T, Robinson JM. A novel intracellular compartment with unusual secretory properties in human neutrophil. J Cell Biol 1991;113:743–756.
231. Calafat J, Kuijpers TW, Janssen H, et al. Evidence of small intracellular vesicles in human blood phagocytes containing cytochrome b558 and the adhesion molecules CD11b/CD18. Blood 1993;81:3122–3129.
232. Kobayashi T, Robinson JM, Seguchi H. Identification of intracellular sites of superoxide production in stimulated neutrophils. J Cell Sci 1998;111: 81–91.
233. Wientjes FB, Segal AW, Harting JH. Immunoelectron microscopy shows a clustered distribution of NADPH oxidase components in the human neutrophil plasma membrane. J Leukoc Biol 1997;61:303–312.
234. Mandell GL, Hook EW. Leukocyte bactericidal activity in chronic granulomatous disease: correlation of bacterial hydrogen peroxide production and susceptibility to intracellular killing. J Bacteriol 1969;100:531–532.
235. Odell EW, Segal AW. Killing of pathogens associated with chronic granulomatous disease by the non-oxidative microbicidal mechanisms of human neutrophils. J Med Microbiol 1991;34:129–135.
236. Casimir C, Chetty M, Bohler MC, et al. Identification of the defective NADPH-oxidase component in chronic granulomatous disease: a study of 57 European families. Eur J Clin Invest 1992;22:403–406.
237. Curnutte JT. Chronic granulomatous disease: the solving of a clinical riddle at the molecular level. Clin Immunol Immunopathol 1993;67:S2–S15.
238. Roos D, de Boer M, Kuribayashi F, et al. Mutations in the X-linked and autosomal recessive forms of chronic granulomatous disease. Blood 1996;87:1663–1681.
239. Cross AR, Heyworth PG, Rae J, Curnette JT. A variant X-linked chronic granulomatous disease patient (X91+) with partially functional cytochrome b. J Biol Chem 1995;270:81–94.
240. Kuribayashi F, Kumatori A, Suzuki S, et al. Human peripheral eosinophils have a specific mechanism to express gp91phox, the large subunit of cytochrome b558-1. Biochem Biophys Res Commun 1995;209:146–152.
241. Williams DA, Tao W, Yang F, et al. Dominant negative mutation of the hematopoietic-specific Rho GTPase, Rac2, is associated with a human phagocyte immunodeficiency. Blood 2000;96:1646–1654.
242. The International Chronic Granulomatous Disease Cooperative Study Group. A controlled trial of interferon gamma to prevent infection in chronic granulomatous disease. N Engl J Med 1991;324:509–516.
243. Klebanoff SJ. Myeloperoxidase: friend and foe. J Leukoc Biol 2005;77: 598–625.
244. Hampton MB, Kettle AJ, Winterbourn CC. Inside the neutrophil phagosome: oxidants, myeloperoxidase and bacterial killing. Blood 1998;92:3007–3017.
245. Fadok VA, Chimini G. The phagocytosis of apoptotic cells. Semin Immunol 2001;13:365–372.
246. Fadok VA, Bratton DL, Henson PM. Phagocyte receptors for apoptotic cells: recognition, uptake, and consequences. J Clin Invest 2001;108:957–962.
247. Fadok VA, de Cathelineau A, Daleke DL, et al. Loss of phospholipid asymmetry and surface exposure of phosphatidylserine is required for phagocytosis of apoptotic cells by macrophages and fibroblasts. J Biol Chem 2001;276: 1071–1077.
248. Callahan MK, Williamson P, Schlegel RA. Surface expression of phosphatidylserine on macrophages is required for phagocytosis of apoptotic thymocytes. Cell Death Differ 2000;7:645–653.
249. Henson PM, Bratton DL, Fadok VA. The phosphatidylserine receptor: a crucial molecular switch? Nat Rev Mol Cell Biol 2001;2:627–633.
250. Leverrier Y, Lorenzi R, Blundell MP, et al. Cutting edge: the Wiskott-Aldrich syndrome protein is required for efficient phagocytosis of apoptotic cells. J Immunol 2001;166:4831–4834.
251. Leverrier Y, Ridley AJ. Requirement for Rho GTPases and PI-3 kinases during apoptotic cell phagocytosis by macrophages. Curr Biol 2001;11:195–199.
252. Casciola-Rosen L, Rosen A, Petri M, Schlissel M. Surface blebs on apoptotic cells are sites of enhanced procoagulant activity: implications for coagulation events and antigenic spread in systemic lupus erythematosus. Proc Natl Acad Sci U S A 1996;93:1624–1629.
253. Jersmann HPA, Ross KA, Vivers S, et al. Phagocytosis of apoptotic cells by human macrophages: analysis by multiparameter flow cytometry. Cytometry 2003;51A:7–15.
254. Bratosin D, Mazurier J, Tissier JP, et al. Cellular and molecular mechanisms of senescent erythrocyte phagocytosis by macrophages. A review. Biochimie 1998;80:173–195.
255. Turrini F, Arese P, Yuan J, Low PS. Clustering of integral membrane proteins of the human erythrocyte membrane stimulates autologous IgG binding complement deposition and phagocytosis. J Biol Chem 1991;266:23611–23617.
256. Ando K, Kikugawa K, Beppu M. Binding of an anti-Band 3 autoantibody to sialylated poly-N-acetyllactosaminyl sugar chains of band 3 glycoprotein on polyvinylidene difluoride membrane and sepharose gel: further evidence for anti-band 3 autoantibody binding to the sugar chains of oxidized and senescent erythrocytes. J Biochem 1996;119:639–647.
257. Allen TM, Williamson P, Schlegel RA. Phosphatidylserine as a determinant of reticuloendothelial recognition of liposome models of the erythrocyte surface. Proc Natl Acad Sci U S A 1988;85:8067–8071.
258. Bratosin D, Mazurier J, Slomianny C, et al. Molecular mechanisms of erythrophagocytosis: flow cytometric quantitation of in vitro erythrocyte phagocytosis by macrophages. Cytometry 1997;30:269–274.
259. Robinson JP, Carter WO. Flow cytometric analysis of granulocytes. In: Bauer KDA, Duque RE, Shankey TV, eds. Clinical flow cytometry. Principles and application. Baltimore: Williams & Wilkins, 1993:405–433.
260. Lehman AK, Sörnes S, Haltensen A. Phagocytosis: measurement by flow cytometry. J Immunol Methods 2000;243:229–242.
261. Allen RC, Stjernholm RL, Steele RH. Evidence for the generation of an electronic excitation state(s) in human polymorphonuclear leukocytes and its participation in bactericidal activity. Biochem Biophys Res Commun 1972;47:679–684.
