Institute for Immunology, Ruprecht-Karls-University, Heidelberg, Germany
Correspondence: Dr. Yvonne Samstag, Institute for Immunology, Ruprecht-Karls-University, Im Neuenheimer Feld 305, D-69120 Heidelberg, Germany. E-mail: o69{at}ix.urz.uni-heidelberg.de
 TOP ABSTRACT INTRODUCTIONDYNAMICS OF THE ACTIN...T CELL ACTIVATION AND...ACTIN REARRANGEMENT DURING...CONCLUSIONREFERENCES |
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Key Words: costimulation • cofilin • L-plastin • enhanced actin polymerization • immunological synapse
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TOPABSTRACTINTRODUCTIONDYNAMICS OF THE ACTIN...T CELL ACTIVATION AND...ACTIN REARRANGEMENT DURING...CONCLUSIONREFERENCES |
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Recognition of antigenic peptides bound to MHC molecules on APC through the T cell receptor/CD3 complex (TCR/CD3) elicits the "competence" signal for cellular activation. Full activation, however, requires additional signals (costimulation). Such "progression" signals result from the interaction of accessory receptors (e.g., CD2, refs. [9 , 10 ], or CD28, refs. [11 12 13 ], with their natural ligands, CD58, or CD80/CD86, respectively). Conversely, engagement of TCR/CD3 in the absence of costimulation leads to a state of unresponsiveness/anergy or even cell death through apoptosis [11 12 13 14 15 16 17 ], one physiological mechanism for tolerance induction.
Cumulative evidence suggests a regulatory role for the cytoskeletal matrix in early receptor-mediated, intracellular signaling events. Thus, T cell receptor stimulation results in actin polymerization [18 ]. Moreover, costimulation leads to rearrangements of the actin cytoskeleton, which promotes accumulation of receptors and raft membrane microdomains at the interface between T cells and APC [19 20 21 ], referred to as "immunological synapse" (IS). Depending on the experimental system, receptor clusters are called "caps" or "supramolecular activation clusters" (SMACs) [7 , 8 ] (for review, see ref. [22 ]). "Rafts" are lipid microdomains that are enriched in cholesterol and sphingolipids [23 , 24 ]. These microdomains exist as small clusters in the membrane of unstimulated T cells and flow together into bigger patches upon T cell activation. This process is also dependent on an intact actin cytoskeleton [25 , 26 ] (for review, see ref. [27 ]).
The cytoskeleton provides the matrix that enables recruitment and efficient formation of molecular signaling complexes [6 , 7 , 28 ]. In particular, the dynamic rearrangements of the actin cytoskeleton in response to T cell costimulation are indispensable for the formation of SMACs and for the stabilization of the immunological synapse [19 20 21 , 29 , 30 ]. Thus, actin dynamics are crucially involved in the decision between induction of lymphocyte effector functions versus lymphocyte anergy. This review will focus on actin-binding proteins, which link surface receptor engagement during T cell activation and migration to dynamic rearrangements in the actin cytoskeleton. Moreover, we propose a new model that may help to explain the molecular link between T cell costimulation and actin dynamics.
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TOPABSTRACTINTRODUCTIONDYNAMICS OF THE ACTIN...T CELL ACTIVATION AND...ACTIN REARRANGEMENT DURING...CONCLUSIONREFERENCES |
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Figure 1. (A) Circle of actin dynamics. Efficient rearrangements in the actin cytoskeleton require (a) the maintenance of a large pool of G-actin by the monomer-binding proteins thymosin-ß4 and profilin. (b) Actin nucleation (generation of new actin filaments) initiated by binding of Wasp to the Arp2/3 complex. Profilin-bound monomeric actin associates with the Wasp/Arp2/3 complex, resulting in actin polymerization. Capping proteins (capping protein or gelsolin) terminate the actin polymerization process by binding to the barbed ends of the actin filaments. (c) Enhanced actin polymerization requires uncapping or severing of the capped actin filaments. This leads to the creation of free barbed ends ready for filament elongation. One important mediator of the actin-severing activity is cofilin. (d) Depolymerization of actin filaments has to occur at sites where they are no longer required. This function is executed by cofilin as well. Released actin monomers are ready for new entry into the actin polymerization process. Thereby, ADP bound to monomeric actin is exchanged for ATP by profilin, resulting in new profilin-ATP complexes ready for actin nucleation. (B) Higher ordered F-actin structures. Actin filaments build up a dense network of cytoskeletal actin. (a) Binding Wasp/Arp2/3 to the side of pre-existing actin filaments leads to F-actin branches with an angle of 70°. (b) Actin-bundling proteins such as plastin, fascin, and  -actinin lead to parallel bundles of actin filaments. (c) -Actinin also cross-links actin filaments.
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Important for an efficient and structured reorganization within the actin cytoskeleton are, in addition, proteins with actin-depolymerizing function (Fig. 1A d)
. The most prominent candidates for actin severing and depolymerization are the cofilin/actin depolymerizing factor proteins (here referred to as cofilin). To become functional, cofilin needs to be activated through dephosphorylation of a serine residue in position 3. Monomeric actin (globular or G-actin), which attaches to the barbed ends of actin filaments (F-actin), is bound to ATP. 
-Phosphate dissociation from incorporated ATP-actin accelerates binding of cofilin to filamental adenosine 5'-diphosphate (ADP) actin. The results are severing and/or depolymerization of actin filaments. ADP-actin released from actin filaments is bound to cofilin. Closing the circle, profilin, which is an ATP-exchange factor for actin, then promotes the exchange of ADP for ATP, resulting in the release of G-actin from cofilin and the formation of new profilin-ATP complexes (Fig. 1A a)
[32
]. To maintain an available G-actin pool, thymosin-ß4, a monomer-sequestering protein, competes with profilin for binding to ATP-actin [33
].
How unstimulated cells maintain high amounts of unpolymerized actin
Flexible, multidirectional mobility of hematopoietic cells requires a high capability of actin polymerization within seconds in response to chemotactic stimuli or antigen presentation. To be prepared for these processes, cells maintain high concentrations of G-actin (e.g., 300 µM in unstimulated neutrophils, ref. [34
]; Fig. 1A a
). The vast majority of monomeric actin is bound to ATP [35
]. The minor fraction of monomeric actin is bound to ADP. In vitro, the critical concentration of ATP actin for polymerization [the concentration above which actin monomers polymerize (dissociation constant)] is 0.1 µM at the barbed end and 0.6 µM at the pointed end [31
]. The much higher concentrations of monomeric actin found in vivo must be protected from spontaneous polymerization. This protection is mediated by monomer-binding proteins that prevent incorporation of monomeric actin into actin filaments (Fig. 1A a)
and capping proteins that bind to the ends of actin filaments, thereby preventing the attachment of further actin monomers (Fig. 1A b)
.
Actin monomer-binding proteins in leukocytes are thymosin-ß4 and profilin (Fig. 1A a) , which attach to overlapping sites on actin [36 37 38 39 40 ]. Both proteins have a higher affinity for ATP-actin than for ADP-actin. Monomeric actin, when bound to thymosin-ß4, cannot polymerize, but it is sequestered in an inactive state to maintain a large, intracellular pool of monomeric actin. In contrast, ATP-actin bound to profilin can be added onto the barbed ends of actin filaments [33 , 41 , 42 ], however, does not elongate the slowly growing pointed ends of actin filaments. Note that profilin/ATP-actin complexes by themselves do not spontaneously nucleate new filaments [43 , 44 ]. Rather, for actin-profilin complexes to proceed toward actin nucleation, additional proteins and stimuli are required (e.g., Arp2/3 complex and Wasp family proteins; see below). Given that profilin is able to add monomeric actin to free barbed ends of actin filaments (filament elongation) in the absence of stimuli that trigger actin polymerization, the barbed ends must be protected from spontaneous actin polymerization. This function is fulfilled by capping proteins (e.g., capping protein-ß2 or gelsolin) [34 , 45 ] (see below).
Actin polymerization in vivo involves actin nucleation and elongation of pre-existing actin filaments
Actin polymerization requires free barbed ends [46
]. They can arise by nucleation of new actin filaments, by uncapping barbed ends in existing actin filaments through release of capping proteins, or by severing of noncovalent bonds between actin monomers within filamentous actin, resulting in shorter filaments with free barbed ends.
Actin nucleation: de novo generation of actin filaments by assembly from actin monomers
Actin dimers and trimers are not stable. To proceed toward the generation of new actin filaments, they must associate with nucleation-promoting factors. The Arp2/3 complex has been identified as an important barbed end-nucleating factor. It consists of seven subunits: Arp2 and Arp3, p40 (ARPC1), p35 (ARPC2), p19 (ARPC3), p18 (ARPC4), and p14 (ARPC5). The Arp2/3 complex is highly conserved amongst species [47
48
49
50
]. The nucleation activity of the Arp2/3 complex is stimulated by binding the conserved C-terminal verprolin-homology-cofilin-homology-acidic (VCA) domain of Wasp/Scar family proteins to the ARPC3 subunit of the Arp2/3 complex [51
, 52
] (for review, see ref. [53
]). The Wasp/Scar family of proteins includes Wasp, which is expressed in leukocytes and platelets. It was identified as a result of its malfunction in patients with Wiskott Aldrich syndrome, a primary immunodeficiency in humans (for review, see refs. [54
, 55
]). N-Wasp, which is closely related to Wasp, is expressed in the brain as well as in other tissues [56
]. Additional family members that were originally discovered in Dictyostelium discoideum are the Wave proteins, vertebrate homologues of Scar [57
, 58
]. The Wasp/Scar family proteins, besides binding to the Arp2/3 complex, interact with actin monomers as well as a multitude of signaling molecules. The structures and functions of these proteins, i.e., with regard to their activation of Arp2/3, have been extensively reviewed elsewhere [53
, 59
]. The topic of Arp2/3 activation through Wasp in lymphocytes will be discussed below. The nucleation activity of the Arp2/3 complex also results in actin filament branches with an angle of 70° [60
, 61
].
Elongation of actin filaments: "enhanced" actin polymerization
Generation of free barbed ends by uncapping actin filaments.
Elongation of actin filaments requires free barbed ends. Experiments with neutrophils showed that less than half of the barbed ends created in vivo result from Arp2/3 stimulation (actin nucleation). This implies that additional mechanisms are required to enable actin polymerization, i.e., uncapping and severing [62
] (for review, see ref. [63
]). The two most abundant barbed end-capping proteins are gelsolin and capping protein, which prevent association as well as dissociation of actin monomers [34
, 45
]. Removal of capping proteins from actin filaments can be mediated by polyphosphoinositides (e.g., PIP2 or PIP3 ) [64
65
66
]. In neutrophils where capping protein-ß2 seems to play the major role for barbed end capping [34
], 
50% of the free barbed ends generated following formyl-Met-Leu-Phe (fMLP) stimulation result from uncapping filaments [62
]. Generation of free barbed ends by uncapping of gelsolin or capping protein was shown to contribute to thrombin-induced actin polymerization in platelets [65
, 67
]. The major mechanism inducing actin polymerization in platelet activation is, however, severing actin filaments [67
].
Generation of free barbed ends by severing actin filaments.
Gelsolin and cofilin are actin-binding proteins with severing activity. A rise in intracellular free calcium, which occurs upon receptor triggering in many cell types, activates gelsolin to sever ("cleave") actin filaments in platelets. At the same time, however, gelsolin caps the barbed ends of the filament fragments, which is contraproductive for actin polymerization. Therefore, elongation of actin filaments can only occur if gelsolin is simultaneously uncapped, e.g., by binding PIP2 [65
]. However, experiments performed with gelsolin-knockout mice (which develop normally) as well as many other experimental findings suggest that gelsolin itself is not essential for the generation of free barbed ends in vivo [63
, 68
].
Recently, it was recognized that the actin-severing function of cofilin is critical for efficient actin polymerization [69 70 71 ] (for review, see ref. [63 ]). Cofilin is ubiquitously expressed in all eukaryotic cell types, and genetic studies have shown that cofilin is essential for cell viability [72 73 74 75 ]. In vitro, cofilin binds to G- and F-actin and induces actin severing and depolymerization [76 77 78 79 ]. Moreover, binding cofilin to F-actin leads to alterations in the F-actin filament twist, which promotes F-actin severing and influences the interaction of other molecules with F-actin [80 ]. The short filament fragments generated by cofilin severing are likely stabilized and cross-linked by the Arp2/3 complex [81 ].
The actin-binding capacity of cofilin is negatively regulated by phosphorylation at serine-3 [82 ]. LIM-kinase 1 and 2 [83 , 84 ] as well as the serine kinases testis-specific protein kinases 1 and 2 [85 , 86 ] inactivate cofilin through phosphorylation. Conversely, the okadaic acid-sensitive, cyclosporin A-resistant serine/threonine phosphatases of type 1 (PP1) and type 2A (PP2A) associate with and dephosphorylate cofilin in T lymphocytes, thereby mediating cofilin activation [87 ]. An additional cofilin phosphatase (slingshot), which was recently identified in Drosophila, is okadaic acid-resistant [88 ]. In unstimulated leukocytes, cofilin is mainly phosphorylated. Cofilin dephosphorylation is induced upon costimulation of untransformed human T lymphocytes [89 , 90 ] as well as following stimulation of neutrophil-like, differentiated HL-60 cells [91 92 93 ] and polymorphonuclear leukocytes (PMNs) derived from peripheral blood [94 ].
Severing by cofilin produces free barbed ends, which can favor actin polymerization. Thus, cofilin, which localizes to the leading edge membrane (i.e., lamellipodia) in carcinoma cells and fibroblasts, promotes barbed end generation and lamellipod protrusion, e.g., upon epidermal growth factor (EGF) stimulation [50 , 70 ]. Accordingly, phosphorylation/inactivation of cofilin as a consequence of overexpression of the kinase domain of LIM-kinase in carcinoma cells abolishes EGF-induced appearance of free barbed ends and actin polymerization at the leading edge and thus, subsequent lamellipod extension [95 ]. Taken together, these findings clearly demonstrate that the severing activity of cofilin is important for the polymerization of actin filaments in the process of pseudopod extension in vivo.
Actin depolymerization
Upon cell migration and/or the formation of cell-cell contacts (e.g., surface scanning of APC by lymphocytes), cells rapidly change their direction of movement. This requires that pseudopod extension at one site of the cell has to stop within seconds, and at the same time, new pseudopod formation at a different site of the cell has to occur. Flexible cell polarization as well as directed movement therefore require not only actin polymerization but equally important, actin depolymerization.
To maintain actin polymerization, Arp2/3 complexes have to be activated. In the absence of Arp2/3-activating signals, the balance shifts toward filament disassembly by cofilin [69 , 96 ]. ATP hydrolysis within the actin filaments marks older filaments for severing and depolymerization by cofilin. Mechanisms by which cofilin increases depolymerization are the formation of unpolymerizable heterodimers (for review, see ref. [97 ]) as well as the enhancement of the off-rate of actin monomers from the pointed end or both ends of actin filaments [77 , 98 ]. Moreover, changes in the F-actin twist induced by cofilin binding also favor release of actin monomers [80 ].
Given that cofilin regulates actin polymerization as well as actin depolymerization, this essential protein may represent a master switch for actin dynamics. The exact mechanisms that determine, besides the availability of monomeric actin, whether cofilin favors actin polymerization or depolymerization remain to be determined. They may differ in various cell types and in addition, may be influenced by the subcellular localization of cofilin. Thus, in some cell types (e.g., fibroblasts), cofilin localizes to the leading edge membrane where nucleation of actin polymerization is predominant [50 , 70 ]. In contrast, in keratinocytes, cofilin exists at the cytoplasmic side of the actin filament network of lamellipodia [50 ], which points toward the involvement of cofilin in the disassembly of actin filaments upon forward movement. As lamellipodia move very fast (up to 1 µm/s), hundreds of actin subunits per s have to be removed from actin filaments [99 ]. Filament turnover is likely enhanced by additional proteins that interact with cofilin. One interesting candidate is actin-interacting protein 1, which has been found in lamellipodia of D. discoideum [100 ], yeast [101 ], and Xenopus [102 ]. It forms ternary complexes with actin and cofilin. Additional mechanisms that may influence the function of cofilin rely on its interaction with PIP2 [103 , 104 ] or changes in the intracellular pH [105 ].
Bundling actin filaments
Radiating bundles of actin filaments with diameters of 0.1–0.2 µm and a length of several micrometers lead to membrane protrusions called filopodia. Similar actin bundles, which, however, do not protrude the plasma membrane, are named microspikes (for review, see ref. [106
]). It is interesting that proteins that localize to the tips of microspikes and filopodia differ from those localizing to the tips of lamellipodia: Whereas Vav is recruited to filopodia and microspikes [107
], Arp2/3 seems to be excluded [50
]. Here, elongation of preexisting filaments, instead of actin nucleation, seems to dominate [108
]. Note that mechanisms regulating the assembly of microspikes and filapodia in vivo are largely unknown.
The actin-bundling proteins fascin and plastin (fimbrin) are thought to play a major role for microspike and filopodia formation [109 110 111 ]. The actin-bundling activity of fascin is antagonized by phosphorylation on serine-39 mediated through protein kinase C (PKC) [112 113 114 ] or by binding of caldesmon coupled with tropomyosin [115 ]. Plastin belongs to the family of fimbrins, members of the calponin-homology superfamily of actin-binding proteins. So far, three different plastin isoforms have been identified: L-plastin is exclusively expressed in leukocytes; I-plastin is produced in intestinal and renal brush borders; and T-plastin localizes to solid tissues [116 ]. In addition to two repetitive actin-binding sequences, L-plastin contains an N-terminal domain containing two calcium-binding sites (EF hand motifs) and a potential calmodulin-binding domain [117 , 118 ]. At least in vitro, actin bundling through L-plastin is inhibited if the concentration of calcium is increased [119 ]. In several cell types, differential phosphorylation patterns of L-plastin were observed. The serine in position 5 and/or in position 7 can be phosphorylated [120 ]. The relationship between phosphorylation of L-plastin and its actin-bundling activity remains to be clarified.
Aside from filopodia formation, bundling (Fig. 1B b)
and, in addition, cross-linking actin filaments (Fig. 1B c)
, e.g., by -actinin (for review see [121
, 122
]), are required for the stabilization of cellular actin filament networks.
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TOPABSTRACTINTRODUCTIONDYNAMICS OF THE ACTIN...T CELL ACTIVATION AND...ACTIN REARRANGEMENT DURING...CONCLUSIONREFERENCES |
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, and the peripheral SMAC, where lymphocyte function-associated antigen-1 (LFA-1) and talin are enriched [7
, 124
, 125
] (for review, see ref. [126
]). Such a composition may lead to segregation of phosphatases from kinases to allow sustained signaling [6
, 30
, 127
].
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Figure 2. The actin cytoskeleton in T cell activation. (A) Schematic view of a resting T cell (left) and a T cell contacting an APC (right) with regard to F-actin distribution. The drawings indicate the morphology of the T cell: round-shaped when resting (left) and polarized toward the APC with formation of an immunological synapse at the cell-cell interface (right). The F-actin distribution within the T cells is indicated in red. (B) Actin cytoskeletal remodeling proteins involved in TCR/CD3 stimulation alone (left) or in costimulation (right), respectively. Actin polymerization driven by TCR/CD3 engagement alone requires Wasp, profilin, and Arp2/3. Moreover, the linker proteins ezrin, moesin, and paxillin as well as talin, vinculin, and -actinin [123] play a role. In addition, costimulation leads to activation of the actin-binding proteins cofilin and L-plastin.
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Clustering receptors and signaling molecules is crucial for T cell activation and function. Costimulation through accessory receptors is indispensable for the formation of SMACs. This view is supported by a recent finding demonstrating that if the interaction of CD2 with its ligand CD58 is blocked in T cells interacting with APC, the exclusion of the phosphatase CD45 from the immunological synapse is strongly reduced [136 ]. Moreover, costimulation induces cytoskeletal rearrangements that are responsible for receptor and lipid raft redistribution toward the TCR/MHC contact site [21 , 137 ]. Actin cytoskeleton-disrupting agents (i.e., cytochalasin) abrogate SMAC formation and prevent T cell activation [125 , 130 , 138 ]. These findings emphasize the importance of costimulation-dependent actin cytoskeleton dynamics for T cell activation [7 , 21 , 125 , 130 , 137 , 139 140 141 ].
There is evidence that F-actin formation can be induced by signaling events that result from TCR/CD3 triggering alone. An initial actin polymerization, most likely driven by Wasp/Arp2/3-dependent actin nucleation, can be observed following TCR engagement [18 , 142 , 143 ]. True actin dynamics, however, require costimulation through accessory receptors. In this regard, distinct actin-binding proteins were identified that are only activated through costimulation via accessory receptors such as CD2 or CD28, namely cofilin and L-plastin [89 , 90 , 144 ]. These proteins have the ability to sever and/or depolymerize F-actin, in the case of cofilin, or to bundle F-actin, as demonstrated for L-plastin. F-actin severing generates free barbed ends required for an enhancement of actin polymerization processes. This ability of cofilin together with the actin-bundling activity of L-plastin could stabilize SMACs. Moreover, the depolymerization of actin filaments by cofilin at sites where they are no longer required allows flexibility in the contact zone and provides additional actin monomers for new actin polymerization processes.
TCR-inducible actin rearrangements: actin nucleation via the Wasp/Arp2/3 pathway indicates the site of contact formation
Currently, it is thought that TCR/CD3 triggering alone, in the absence of costimulation, is not efficient to induce the formation of SMACs and to stabilize the IS [30
, 145
]. Nevertheless, a certain degree of actin polymerization has been observed following TCR/CD3 triggering alone, likely as a result of actin nucleation via the Wasp/Arp2/3 signaling pathway [18
, 146
]. One could speculate that initial actin polymerization at the T cell/APC contact zone determines the area to which receptors and signaling/effector molecules need to be moved during the activation process. Transport of the latter molecules is, however, dependent on costimulation and, at least in part, achieved via myosin motor molecules, i.e., myosin II [137
]. TCR/CD3 triggering leads to activation of multiple cytoplasmic proteins and their recruitment to the plasma membrane in the vicinity of the engaged TCR. The TCR-related signaling cascades lead to protein tyrosine phosphorylation and the release of second messengers (for review, see elsewhere, refs. [147
, 148
]). Briefly, the tyrosine kinase 
-associated protein 70 (ZAP-70) [132
] is activated and recruited to the plasma membrane immediately after TCR/CD3 triggering [149
, 150
]. Activated ZAP-70 phosphorylates linker for activation of T cells (LAT) [151
], a member of the transmembrane-adaptor protein family, and likewise, SLP-76 [152
] is phosphorylated. SH2 domain-containing leukocyte protein of 76 kDa (SLP-76) is a cytosolic adaptor protein that bridges TCR engagement to the actin cytoskeleton via binding to Vav1 and thereby to the Wasp/Arp2/3 complex (Fig. 3
) [127
]. The whole signaling complex is, in addition, fixed to the plasma membrane via anchor proteins, for example talin and vinculin (see below).
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Figure 3. T cell signaling events that result in activation of actin-reorganizing proteins. On the left side, signal transduction cascades are shown that can be initiated by TCR/CD3 triggering alone. These signaling events lead to activation of Vav and, likely via Cdc42 activation, of Wasp. Wasp activates the actin nucleation complex Arp2/3, thereby initiating actin polymerization. Costimulation, which is signal 1 plus signal 2, leads to activation of additonal actin-remodeling molecules, namely cofilin and L-plastin. Cofilin is activated via dephosphorylation by the serine phosphatases PP1/PP2A. This event is sensitive to inhibitors of phosphatidylinositol 3-kinase (PI3-kinase). Once activated, cofilin contributes to enhanced polymerization as well as depolymerization of F-actin fibers. Another protein that is activated only via costimulation is the actin-bundling protein L-plastin. The actin cytoskeleton rearrangements after T cell costimulation are crucial for receptor clustering and stabilization of the IS, which is a prerequisite for T cell activation. Fyb/Slap, Fyn-binding protein/SLP-76-associated protein; Ena/Vasp, Ena/vasodilator-stimulated phosphoprotein; Wip, Wasp-interacting protein.
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Vav is activated by phosphorylation. The enzymatic activity of phosphorylated Vav in vitro is increased by about twofold in the presence of PI3-kinase products (i.e., PtdIns-3,4-P2 and PIP3). There are several membrane receptors described that are involved in Vav activation. In T lymphocytes, signals initiated by TCR/CD3 triggering alone lead to Vav phosphorylation [156 ]. Engagement of the coreceptor CD28 alone is also sufficient to induce Vav phosphorylation. Accordingly, costimulation via CD3 plus CD28 leads to a higher proportion of phosphorylated Vav molecules and prolongs phosphorylation [153 , 162 163 164 165 166 167 ].
In vitro studies have identified Lck, Fyn, and ZAP-70 as Vav kinases. In vivo, Lck seems not essential, as Vav is phosphorylated in Lck-deficient Jurkat T lymphoma cells [143
, 161
]. Therefore, some investigators consider ZAP-70 to be the major Vav kinase [168
169
170
]. Recently, a scenario was suggested in which two waves of Vav phosphorylation take place after TCR/CD3 triggering [143
]: Fyn is the major kinase for an early phosphorylation of a small amount of Vav, and ZAP-70 is responsible afterward for the phosphorylation of the bulk of Vav. A minor proportion of Vav is constitutively associated with CD3, independent of Vav phosphorylation. The phosphorylation of the latter Vav molecules after TCR engagement is independent of ZAP-70 and most likely mediated by Fyn. This first wave of Vav phosphorylation lasts for 
5 min and does not require lipid raft clustering. It is thought that this initial Vav phosphorylation helps to promote raft clustering by T cell costimulation, which is consistent with previous findings demonstrating that lipid raft polarization to the IS depends on an intracellular pathway that involves Vav and Rac1. Note that lipid raft clustering in vav(-/-) T cells is impaired [25
].
The second wave of Vav phosphorylation depends on ZAP-70 and Fyn and is sensitive to the raft-disrupting agent methyl-ß-cyclodextrin. Together with SLP-76, ZAP-70 recruits larger amounts of Vav to the plasma membrane [143 ]. Thus, Vav participates in establishing SMACs. In this regard, Vav has been shown to associate with the membrane anchors talin and vinculin [156 ]. The latter proteins are involved in the attachment of F-actin to the cell membrane. It is thought that through this link between activated Vav, which is connected to SLP-76, and the cytoskeleton, the signaling complex is, at least in part, attached to the focal site of T cell activation [161 , 171 ].
Experiments with Vav-deficient mice verified that vav(-/-) T cells are defective in cytoskeletal reorganization. As a consequence, receptor cap formation upon CD3
cross-linking is severely impaired in vav(-/-) T cells [155
, 156
]. This phenotype is a consequence of the inability to couple the TCR/CD3 signaling complex via talin and vinculin to the plasma membrane. In addition, the Vav substrate Rac1 is not activated. Rac1 is involved in cytoskeletal rearrangements that result in lipid-raft coalescence, a prerequisite for cap formation [25
]. Moreover, as Vav is one candidate that positively regulates Wasp via Cdc42 activation, cap inhibition in vav(-/-) T cells could in part be a result of Wasp inactivity. Conversely, overexpression of Vav leads to an increase of actin polymerization. As described above, Wasp family proteins stimulate the Arp2/3 complex to nucleate actin polymerization.
Wasp family proteins connect T cell receptor signaling to the actin nucleation complex Arp2/3
The Arp2/3 complex plays a pivotal role in T cell activation, as it nucleates actin polymerization through interaction with Wasp family proteins [146
, 172
173
174
175
]. Wasp proteins belong to a family of proteins containing a polyproline-binding domain of the Ena/Vasp homology 1 (EVH1) type (recently reviewed in ref. [175
]). This EVH1 domain is located near the N-terminus and is often referred to as Wasp homology domain 1 (WH1 domain). An additional proline-rich region is capable of recruiting profilin bound to monomeric ATP-actin. This ternary complex enhances the ability of Wasp proteins to nucleate actin polymerization [176
]. Upon direct interaction with the small GTPase Cdc42, Wasp stimulates actin polymerization by activating the Arp2/3 complex.
T cells from Wasp-deficient mice are defective in actin polymerization [141 ], demonstrating the important role of Wasp in this process. In resting T cells, Wasp keeps itself in an inactive state by autoinhibition through cis- or trans-binding of the C-terminal VCA region to the N-terminal GTPase-binding domain/Cdc42 and Rac interactive-binding (GBD/CRIB) region and the WH1 domain [177 , 178 ]. This junction between the N- and C-terminus of the protein masks the binding site for Arp2/3, and as a consequence, the Arp2/3 complex remains in a quiescent state. The inactive conformation of Wasp is released by Cdc42 binding to the GBD/CRIB domain. Activated Wasp then binds to the Arp2/3 complex via the VCA region initiating actin nucleation.
Wasp localizes to the T cell/APC contact zone [171 , 179 ]. Its subcellular localization, however, seems to be independent of the activation by Cdc42. It is mediated by the cytosolic adaptor protein Nck, which itself is recruited by SLP-76 [179 , 180 ]. The actin nucleation machinery is thereby directed to the site where signaling is initiated, which together with Vav, contributes to formation of the immunological synapse. This is one reason why Wasp knockout T cells are unable to produce TCR caps [141 ].
Proteins of the enabled/vasodilator-stimulated phosphoprotein (Ena/Vasp) family were also implicated in TCR signaling to the actin cytoskeleton. Like Wasp, these proteins contain an N-terminal EVH1 domain that binds to the cytosolic adaptor protein Fyb/SLAP [171 ] and a central proline-rich region, which can bind to profilin [181 ]. In vitro data showed that Ena/Vasp proteins directly interact with F-actin via their C-terminal EVH2 domain [182 ]. In CD3-activated T cells, Ena/Vasp proteins are recruited to the site of receptor stimulation [171 , 183 ]. Here, the adaptor proteins Fyb/SLAP, SLP-76, and Nck form a large complex together with Vasp, Wasp, and Arp2/3. Inhibition of the Fyb/Vasp interaction disturbs the TCR-mediated F-actin accumulation at the contact site, demonstrating the importance of Ena/Vasp proteins in actin cytoskeletal reorganization [171 ].
Recently, it was shown that similar to wasp(-/-) lymphocytes, wip knockout T cells fail to increase their F-actin content [184 ]. This finding supports earlier reports that demonstrated that Wip stimulates actin polymerization if introduced into permeabilized cells [185 ]. In addition, there is evidence that wip(-/-) T cells are defective in the formation of a proper T cell/APC contact [184 ]. Like Wasp and Vasp, Wip binds to the G-actin-binding protein profilin. However, the underlying molecular mechanisms of how Wip may orchestrate T cell activation are so far largely unknown. Likewise, the relevance of Wave, another Wasp-related protein family, for T cell activation is as yet unresolved.
Actin cytoskeletal anchor proteins are activated by TCR/CD3 and are involved in IS stabilization
For the fixation of the actin meshwork to the APC contact zone, various F-actin anchor proteins are discussed. For instance, vinculin and talin connect Vav with the actin cytoskeleton and the plasma membrane. Additional anchor proteins are known that participate in connecting TCR/CD3 signaling to the actin cytoskeleton: ezrin-radixin-moesin (ERM) family proteins and paxillin [186
, 187
]. ERM proteins contain an N-terminal Four.1 ERM domain, which binds in unphosphorylated status to the C-terminus. Thereby, ERM protein binding to F-actin and membrane proteins is blocked. Phosphorylation releases this conformation and allows ERM-dependent linking of F-actin to membrane proteins [188
]. Ezrin and moesin but not radixin concentrate at the periphery of the T cell/APC contact zone but are clearly excluded from the IS [186
, 189
]. For ezrin, it was demonstrated that TCR-mediated signals are sufficient to induce its relocalization to the plasma membrane [186
, 188
]. Moreover, overexpression of the N-terminus (membrane-binding domain) of ezrin disturbs TCR clustering. Therefore, Roumier et al. [186] suggest that ezrin participates in the coalescence of small TCR clusters into larger clusters. Another mechanism by which ERM proteins could function in T cell activation is the exclusion of the sialoglycoprotein CD43 (which is involved in T cell adhesion) from the IS [186
, 189
, 190
]. Paxillin is phosphorylated/activated after TCR/CD3 triggering, but it is not phosphorylated by CD28 ligation. It appears that paxillin might act as a kind of adaptor protein, as it binds to tyrosine kinases and cytoskeletal proteins. The relevance of paxillin for T cell activation remains to be determined [187
, 191
].
Costimulation conducts receptor and raft segregation and provides signals for the stabilization of the immunological synapse that cannot be achieved by TCR engagement alone
The former chapter described molecular events driven by TCR/CD3 engagement that result in remodeling of the actin cytoskeleton, i.e., by directed actin polymerization. However, TCR engagement alone is definitely not sufficient to induce the whole functional repertoire of untransformed T lymphocytes such as cell proliferation and interleukin-2 production. To achieve full activation, second/costimulatory signals are, in addition to TCR/CD3-mediated first signals, indispensable (two-signal model). Without second signals, which are delivered through accessory receptors such as CD2 or CD28, T cells proceed to a state of anergy (unresponsiveness) or even programmed cell death (apoptosis) [11
12
13
14
15
16
17
]. Costimulation is necessary to enhance the TCR signal most likely by receptor clustering and SMAC formation plus stabilization of the IS [30
, 127
]. The view that TCR engagement is itself not the decisive factor for developing an immunological synapse is, in addition, supported by the fact that an immunological synapse at the T cell/APC contact zone can be developed in the absence of nominal antigen [145
]. Here, TCR-independent activation of Vav via CD28 [166
] resulting in Wasp/Arp2/3-mediated actin nucleation could substitute for TCR signaling. Important parameters for stable, organized receptor clustering, however, are the availability of costimulatory signals and the integrity of the actin cytoskeleton as well as myosin motor proteins [137
]. It is interesting that actin remodeling proteins have been described that are, in marked contrast to Vav, Wasp/Arp2/3 and ERM family proteins, exclusively activated after T cell costimulation (TCR/CD3 plus CD28 or CD2, respectively). Thus, only costimulation leads to dephosphorylation/activation of the 19-kDa actin-severing and -depolymerizing protein cofilin as well as to phosphorylation of the 67-kDa actin-bundling protein L-plastin [10
, 144
]. This could explain costimulation-dependent actin rearrangements.
Cofilin, the missing link between T cell costimulation and rearrangement of the actin cytoskeleton
In untransformed, resting human peripheral blood T lymphocytes (PBL-T), the bulk of cofilin is constitutively phosphorylated on serine-3 and is thus inactive. Following costimulation via accessory receptors (e.g., CD2 or CD28), but not following T cell receptor triggering alone, cofilin undergoes activation through dephosphorylation [10
, 89
, 90
, 192
]. Recently, the serine/threonine phosphatases PP1 and PP2A were identified to mediate cofilin activation in PBL-T cells [87
]. These phosphatases function through a Cyclosporin A/FK506-resistant costimulatory signaling pathway that is common for the accessory receptors CD2 and CD28. Likewise, this costimulatory signaling pathway is not affected by a series of additional, immunosuppressive agents (i.e., rapamycin, dexamethason, leflunomide, or mycophenolic acid) [87
].
Note that in preactivated cells (e.g., the malignant T lymphoma line Jurkat), activation of cofilin through dephosphorylation occurs spontaneously, i.e., independent of external stimuli [72 , 87 , 89 , 90 ]. This enables actin cytoskeletal dynamics following TCR/CD3 stimulation even in the absence of costimulation. Moreover, in activated cells, dephosphorylation of cofilin can be accompanied by its nuclear translocation [72 , 89 , 193 ]. The role of cofilin within the nucleus is so far unknown. Given that cofilin contains a nuclear localization signal and binds to G-actin, it may serve as a transport system for actin into the nucleus. There, actin might act as a scaffolding component of the nuclear matrix. In addition, a role of the nuclear actin cytoskeleton for chromatin remodeling and splicing processes seems likely (for review, see ref. [194 ]).
Cofilin/F-actin interactions are not detectable in resting T cells. Activation-induced dephosphorylation of cofilin is accompanied by its transient translocation to the actin cytoskeleton. In the actin cytoskeletal fraction of activated T cells, cofilin is exclusively present in its dephosphorylated form [90 ]. This is likely a result of direct binding of cofilin to F-actin, as cofilin-derived peptides representing the actin binding sites of cofilin are able to block the association of cofilin with F-actin (own unpublished).
PI3-kinase and its products are involved in the regulation of cytoskeletal structures [195 196 197 ]. It is interesting that specific PI3-kinase inhibitors block cofilin dephosphorylation and its association with the actin cytoskeleton in PBL-T. Thus, cofilin might represent one of the mediators of PI3-kinase function, which couples signaling cascades to the cytoskeleton, thereby affecting cytoskeletal restructuring during T cell activation. In Jurkat T lymphoma cells, cofilin is spontaneously dephosphorylated and permanently associated with the actin cytoskeleton. In the latter cells, the PI3-kinase inhibitor wortmannin does not block the cofilin/F-actin association, suggesting that in this tumor cell line an autonomous process distal of PI3-kinase activity is ongoing [90 , 198 ].
The in vivo relevance of cofilin is difficult to investigate, as cofilin is an essential protein for cell survival [72 73 74 75 ]. This fact rules out the possibility to construct knockout mice. Recent experiments, however, have strongly supported the importance of cofilin in T cell activation: Cofilin-derived cell-permeable peptides that block the interaction between cofilin and actin prevent receptor cap formation and T cell activation (own unpublished). Thus, these studies indicate that cofilin likely represents an important mediator of costimulation-induced actin cytoskeletal rearrangements. As delineated above, such rearrangements are required for receptor clustering and stabilization of the IS.
Based on the information that is available to date, the following model explaining the molecular link between T cell costimulation and actin dynamics is proposed (Fig. 4 ): TCR engagement initiates Arp2/3-dependent actin nucleation. This actin polymerization is obviously not sufficient for SMAC formation. Costimulation activates cofilin, which has two major functions: It enhances actin polymerization by generation of free barbed ends through its actin-severing activity and depolymerizes actin filaments at sites where they are no longer required. The latter activity enables the T cell to flexibly reorientate actin polymerization, as breaking down F-actin fibers that are no longer required, in addition, creates more actin monomers for enhanced polymerization. Thus, through its dual function, cofilin could trigger actin dynamics within the IS.
 View larger version (40K): [in a new window] |
Figure 4. Model of molecular mechanisms linking costimulation-induced actin dynamics to the formation and stabilization of the immunological synapse. (a) TCR/CD3 stimulation alone induces a certain degree of actin polymerization resulting from nucleation of actin filaments by binding Wasp to the Arp2/3 complex. This process is usually rapidly terminated by capping proteins (gelsolin or capping protein), which bind to the barbed ends, thereby preventing the elongation of actin filaments (not shown). (b) Enhanced polymerization in response to T cell costimulation can be explained by the severing activity of cofilin. This provides new barbed ends for filament elongation, thereby enabling the formation of a dense network of actin filaments, which may contribute to the stabilization of the immunological synapse. In addition, efficient formation of cell-cell contacts requires the depolymerization of filaments at sites where they are no longer required. This, in turn, leads to the generation of newly available actin monomers, supporting the nucleation and elongation of actin filaments.
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Most experiments concerning L-plastin function have been performed in PMNs. Jones et al. [200 ] analyzed the function of L-plastin by introducing L-plastin-derived peptides containing the N-terminus into PMNs. These experiments demonstrated that L-plastin is somehow involved in integrin-mediated adhesion to immune complex. Therefore, a critical role for L-plastin in the avidity regulation of ß2 integrins was suggested [199 200 201 ]. Only little is known about the functional consequences of L-plastin phosphorylation after T-lymphocyte activation. However, as L-plastin undergoes phosphorylation specifically after costimulation, a role in adherence of T cells to APC or a function in SMAC formation and stabilization, respectively, can be predicted. Such a function could be mediated by bundling actin filaments in the area of the IS.
|
TOPABSTRACTINTRODUCTIONDYNAMICS OF THE ACTIN...T CELL ACTIVATION AND...ACTIN REARRANGEMENT DURING...CONCLUSIONREFERENCES |
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Polarization of migrating T lymphocytes
T lymphocytes are highly mobile cells, reaching speeds as high as 7–10 µm/min, which is approximately 10 times faster than migrating fibroblasts (0.3–1.3 µm/min) [202
]. Moving T cells have morphological and functional characteristics reminiscent of amoeba. Their shape has been described as "hand mirrors", as they display a highly polarized morphology with three distinct compartments (Fig. 5
): a leading edge at the front, followed by the main cell body with the nucleus and the trailing edge, consisting of a narrow, cytoplasmatic, backward projection, also referred to as the "uropod" [204
205
206
207
]. The compartmentalization of polarized T cells has been discussed extensively elsewhere [140
, 207
, 208
] and is not subject of this review, except for a brief summary of the most important features.
 View larger version (52K): [in a new window] |
Figure 5. Schematic view of a migrating T cell showing the morphology and functional compartments (upper part) as well as actin-binding proteins with their location and function during migration (lower part). The drawings illustrate the "hand-mirror" shape of a migrating T cell in side-view and top-view, displaying leading edge and uropod (side view) and the functions of the different compartments (top view), respectively. F-actin distribution (red) and the direction of migration (arrow) are indicated. The diagram below gives a summary of the distribution and major functions of the actin-binding proteins mentioned in the text. At the leading edge, Wasp, profilin, Arp2/3, and cofilin drive membrane protrusion through actin polymerization/depolymerization. Ena/Vasp proteins may also play a role here, as they were shown to localize to the leading edge membrane of various cell lines [203
]. Actin-binding proteins of the leading edge linking adhesion molecules to the actin cytoskeleton are talin, vinculin, and -actinin. Ena/Vasp and profilin-mediated actin polymerization are thought to contribute to linking adhesive contacts to membrane protrusion. An important function of the uropod is forward movement of the whole cell body containing the nucleus (traction). Actin myosin II-based contraction is responsible for the force generation. The membrane/F-actin linker of the ERM family, radixin colocalizes with myosin II in the neck of the uropod [213
]. Plectin binds to F-actin and vimentin intermediate filaments, creating a link between the nucleus and the actin cytoskeleton. Plectin is thought to enhance deformability of the cell body, facilitating, e.g., diapedesis [215
]. In the tip of the uropod, the ERM proteins moesin and ezrin colocalize. They link adhesion molecules to the actin cytoskeleton and may therefore be crucial for the recruitment of other cells.
|
-actinin, and adhesion receptors such as LFA-1 (Lß2 integrin) are concentrated [209
210
211
]. The main cell body is round-shaped and contains the nucleus. "Behind" the cell body, a narrowing transition zone termed the "polarizing compartment", which contains the microtubule-organizing centre (MTOC), precedes the uropod [208
, 212
]. The latter is located at the trailing edge of the cell and protrudes from the contact area with the substratum. There, the motor protein myosin II, the Golgi apparatus, and a meshwork of microtubules directed from the MTOC are located [202
, 212
, 213
]. The intercellular adhesion molecules (ICAM)-1, -2, and -3, CD43, and CD44, the cytoskeletal linker protein plectin, the actin-binding proteins of the ERM family, as well as ß1 integrins are also concentrated in the uropod [210
, 213
214
215
]. Other molecules, e.g., TCR/CD3 and CD2, as well as the ß3 integrins, show no apparent enrichment in either compartment [210
, 216
]. The chemokine receptors at the leading edge direct the migration of the cells, whereas adhesion molecules in the uropod mediate recruitment of other cells [209
, 217
]. This polarized organization of migrating T lymphocytes seems to be generated, at least in part, by a coordinated actin cycle: Polymerization of actin at the leading edge pushes the front of the cell forward, and filamentous actin close to the membrane moves backward [218
]. Radixin, ezrin, and moesin (ERM family) serve as physical linkers between the plasma membrane and F-actin. In the inactive state, intramolecular interactions of N- and C-termini within ERM proteins prevent membrane- and F-actin-binding. Activation of ERM proteins depends on Rho signaling and requires phosphorylation of a C-terminal threonine as well as PIP2 binding to the N-terminus [219
]. ERM proteins link adhesion molecules to F-actin and redistribute to the uropod by backward movement of actin filaments. Supporting this view, in polarized T cells, moesin and ezrin coimmunoprecipitate with molecules such as CD43, CD44, and ICAM-3, which are enriched in the uropod [213
, 220
]. Only recently, lipid rafts were also implicated in the polarization of migrating T cells [221 , 222 ]. The raft-associated lipids GM3 and GM1 were shown to segregate into distinct leading edge rafts and uropod rafts, respectively. Raft integrity is required not only for the polarization of the cells but also for protein redistribution and chemotaxis. A functional actin cytoskeleton seems to be necessary, as the F-actin-disrupting agent latrunculin-B abolishes raft segregation [221 ].
Amoeboid locomotion of T lymphocytes
Amoeboid locomotion of animal cells over a substratum is understood as a multistep process comprising the following steps: formation of a membrane protrusion at the cell front (lamellipodium, "leading edge"), which is mainly driven by actin polymerization; attachment of the new extension to the substratum and formation of adhesion sites, in which actin filaments and actin-binding proteins such as vinculin, -actinin, and paxillin, adhesion receptors (integrins), and kinases such as focal-adhesion kinase are assembled; force generation by myosin-based motor proteins along actin filaments resulting in traction of the cell body; and detachment from the substratum at the end of the cell and retraction of the trailing edge [208
, 223
]. Two types of amoeboid locomotion seem to exist: First, a slow movement of fibroblast-like cells involving the establishment of strong focal adhesion contacts to the substratum and second, a fast movement of white blood cells, which have more diffuse contacts with the substratum and apparently lack stress fibers and true focal adhesions [208
, 224
]. For monocytes/macrophages and immature dendritic cells, the formation of another type of adhesion structures, the so called "podosomes", has been reported [225
226
227
]. Migrating T cells seem to use both types of amoeboid locomotion, depending on the functional status of the cell and the respective environment encountered [208
, 228
]: The fibroblast-like, integrin-mediated movement is used for migration over "2D ligand-coated surfaces", a model system for the crawling of T cells along vessel walls [229
, 230
], whereas the fast movement is used during "3D migration" of T cells within solid tissues. T cell migration in a 3D environment involves multiple, simultaneous cell matrix and/or cell-cell interactions and is, in contrast to 2D migration, largely integrin-independent [208
, 210
].
The molecular machinery underlying polarization and migration
As little is known about the molecular mechanisms underlying integrin-independent 3D movement of T cells, we discuss here the integrin-mediated 2D migration of leukocytes. Moreover, given that T cell specific data are not yet available for all steps of migration, we will extend our description on leukocytes in general.
Induction of cell polarization
Chemokines, small proteins with chemoattractant properties for leukocytes, are bound by seven-transmembrane domain chemokine receptors on their target cells. These receptors are coupled to heterotrimeric Gi proteins. Upon receptor engagement, they initiate a multitude of signaling cascades (reviewed in refs. [4
, 207
]). Thereby, a shallow chemokine gradient in the environment of the cell is transformed into a strong asymmetry within the cell, resulting in the polarized phenotype described above. Chemokine receptors, e.g., CCR2, CCR5, and CXCR4, were shown to enrich at the leading edge of migrating T cells [209
, 221
]. Other chemoattractant receptors remain evenly distributed, as demonstrated for the complement receptor C5a during chemotaxis of a neutrophil-like cell line [231
]. In either case, asymmetrical triggering of surface receptors produces an intracellular asymmetry of signaling components. For instance, a fusion protein consisting of the PH domain of the PI3-kinase effector protein kinase B linked to green fluorescent protein accumulates at the leading edge of migrating neutrophils [232]
. Through positive feedback during signaling, the asymmetry is further increased [233
, 234
]. Eventually components of the actin polymerization machinery such as Arp2/3 are directed to the anterior pole of the cell and drive membrane protuberance [235
].
Generation of membrane protrusions at the leading edge
The forward thrust of membrane protrusions (lamellipodia) is supported by actin polymerization at the leading edge followed by stabilization of the new membrane extensions by adhesive contacts [236
, 237
]. As described above, actin polymerization can occur via two distinct pathways: first, actin nucleation through activation of the Arp2/3 complex resulting in de novo polymerization of actin filaments. Direct evidence for the importance of wasp-mediated Arp2/3 activation is presented by the fact that wasp(-/-) T lymphocytes display impaired chemotaxis [238
]. Second is the elongation of pre-existing actin filaments ("enhanced actin polymerization") requiring uncapping and severing of actin filaments. Thereby, generated free barbed ends allow addition of new actin subunits. For a remodeling of newly formed membrane protrusions, in addition, actin filaments have to be disassembled at sites where they are no longer needed. Cofilin plays a central role in these processes, through its ability to sever and depolymerize F-actin (see above) [63
]. Accordingly, the induction of barbed ends of actin filaments and lamellipodia protrusions [70
, 95
] as well as an increased cell motility in response to different chemoattractants [97
] correlate, at least in nonhematopoietic cells, with dephosphorylation/activation of cofilin.
A very early event observed following activation of chemokine receptors is a rapid and transient increase in the total F-actin content of the cell. Transient inactivation of cofilin by phosphorylation at serine-3 by LIM-kinase 1 could, at least in part, account for this phenomenon. That binding of stromal cell-derived factor-1 to the chemokine receptor CXCR4 leads to activation of LIM-kinase 1 supports this notion [239
].
Formation of cell-substratum contact sites
Membrane protrusions are stabilized by the formation of adhesion sites for ECM in the leading edge, involving adhesion molecules of the integrin family. Triggering of surface receptors such as TCR or CD2 or ligand binding to integrins leads to integrin clustering and subsequent recruitment of filamentous actin to cytoplasmic integrin domains. Signaling to and by integrins is a complex process and has been the subject of various recent reviews [201
, 237
, 240]
; therefore, here, we shall focus on those proteins that mediate cross-linking of actin filaments to integrins.
Although little is known about the mechanisms by which adhesive structures assemble, evidence points to a sequential recruitment of individual adhesion components [237
, 241
] rather than to the recruitment of large, preassembled complexes. Actin-binding proteins that are assembled upon integrin clustering include talin, vinculin, -actinin, and filamin. These proteins do not only link integrins to F-actin but also provide a scaffold on which molecules involved in integrin signaling are sequestered, as for example, the focal adhesion kinase [242
]. The best-characterized integrin F-actin cross-linking proteins are probably talin and vinculin.
Talin is a dimeric actin-cross-linking protein that binds to the cytoplasmic domains of many integrins. Talin contains an F-actin binding site close to its carboxy-terminal region, a binding site for FAK in its globular head domain, and three nonoverlapping binding sites for vinculin [243 ]. Talin interacts with the cytoplasmic tail of dimerized integrins via its head domain and its rod domain [244 ]. In PMNs, activation of ß2 integrins such as LFA-1 by PMA or fMLP has been shown to be associated with cleavage of talin by the calcium-activated protease calpain to produce the head and rod domain fragments of talin [245 ]. It was, therefore, speculated that intact talin keeps integrins in an inactive state [244 ].
Vinculin is composed of a large, globular head domain and an extended tail. The head domain contains binding sites for talin and the actin-binding protein -actinin; the tail domain contains binding sites for F-actin and paxillin. Head and tail domains are separated by a proline-rich region providing a binding site for Vasp [243
]. The vinculin head domain can participate in an intramolecular interaction with the extended tail domain, masking the binding sites for talin, -actinin, F-actin, and Vasp but not for paxillin. This intramolecular interaction is relieved by acidic phospholipids, which bind to the tail domain, exposing the binding sites for talin, -actinin, F-actin, and Vasp [246
247
248
]. In chicken embryo fibroblasts, a model was suggested in which Rho-dependent synthesis of PIP2 promotes recruitment of vinculin to the membrane via interaction of the vinculin tail domain with the negatively charged headgroups of the phospholipids [246
]. As a result of its numerous interaction domains, vinculin may contribute to the assembly of focal adhesions in different ways: Vinculin may stabilize the interactions between talin and actin as well as between talin and the plasma membrane [243
]. Moreover, binding of Vasp to vinculin may recruit profilin/ATP-G-actin complexes, thereby increasing the rates of actin polymerization at the barbed ends of actin filaments [249
]. Indeed, Vasp was shown to colocalize with vinculin to adhesion sites [248
]. This indicates that in rapidly migrating cells, the formation of cell-substratum contact sites at the leading edge and the formation of new membrane protrusions via actin polymerization might be directly linked [250
]. Although most of the research on vinculin has been performed with other cell types, a role in T cells is likely. In fact, it could be demonstrated that Vav is constitutively associated with vinculin and also with talin [156
]. Vav regulates members of the Rho family of small G-proteins and may therefore couple adhesive interactions with changes of the actin cytoskeleton [251
].
Forward movement of the cell body and retraction of the tail (uropod)
Besides extension of new membrane protrusions at the leading edge by actin polymerization and cross-linking to integrins, a contractile force is needed to move the cell body forward. This is, at least in part, generated by myosin-based motors. Myosin II is a double-headed, rod-like molecule, capable of self-associating into bipolar filaments. Myosins can bind to actin filaments and produce ATP-dependent motion, pulling two actin filaments against each other [252
]. There is evidence that the contractile force generated by myosin II acts strongly at the uropod [253
]. This is consistent with the spatial distribution of myosin II in migrating T cells: It is concentrated in the neck of the uropod [211
, 213
]. A major function of myosin II-based contraction might be to promote the breakdown of adhesive interactions in the uropod by direct application of physical stress [253
]. This tension force disrupts intracellular interactions of integrins with the actin cytoskeleton and/or interactions of integrins with their extracellular ligands. In leukocytes, the latter is more frequently observed than the former [254
].
One important mechanism in tail retraction is stimulation of actin-myosin II-based contractility via calcium signaling [255 , 256 ]. Calcium activates myosin II through myosin light chain (MLC) kinase-mediated MLC phosphorylation [257 ]. In migrating leukocytes, calcium concentrations are lowest at the leading edge and highest in posterior regions [258 ] so that myosin II-mediated contraction would be greatest in the tail region, where release of cell-substratum contacts must occur. Accordingly, inhibition of myosin II by prevention of MLC phosphorylation has been shown to block retraction of the neutrophil uropod [256 ]. In addition, there is evidence that RhoA and one of its effectors, p160ROCK, are required for leukocyte tail retraction during transendothelial migration, possibly also through myosin II activation [259 , 260 ]. Thus, inhibition of RhoA results in trapping monocytes between endothelial cells, the former being unable to complete transmigration as a result of a failure to retract their tail. Furthermore, inhibition of RhoA results in the accumulation of ß2 integrins at the rear end of migrating monocytes. Moreover, the RhoA effector p160ROCK has been shown to negatively regulate integrin-dependent adhesion in leukocytes. This suggests that RhoA/p160ROCK are involved in the turnover of integrins at the tail of leukocytes [259 ]. Note that these findings are in contrast to the situation in fibroblasts. Here, RhoA is known to promote adhesion [254 ]. This emphasizes that the same signaling molecules can have opposite effects in different cell types.
Migration during T cell activation: the "serial encounter model"
Initially, migration and antigen-specific T cell activation seemed to be separated events. Specific TCR/MHC interactions were shown to deliver migratory stop signals for the T cell [5
]. The T cell/APC contact time required for full activation of naive T cells was reported to be at least 2 h [128
, 131
, 261
, 262
]. Recently, a new model of antigen recognition was suggested: the "serial encounter model" (for review, see ref. [263
]). This model was established in an experimental system, in which naive T cells move through a three-dimensional collagen matrix in search for antigen-presenting dendritic cells. Here, short-lived T cell/dendritic cell contacts (average 6–12 min) alternate with phases of vigorous T cell migration. Surprisingly, T cells can be fully activated, although they only transiently reduce their migratory velocity [264
]. Thus, depending on the respective environment and the T cell/APC system used, single T cell/APC contacts may last several hours or only a few minutes. These findings indicate that the initial phase of T cell activation, namely antigen recognition on APC, may comprise both components: stop signals leading to the formation of an immunological synapse plus T cell migration resulting in a serial encounter of APC.
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TOPABSTRACTINTRODUCTIONDYNAMICS OF THE ACTIN...T CELL ACTIVATION AND...ACTIN REARRANGEMENT DURING...CONCLUSIONREFERENCES |
|---|
Received June 4, 2002; accepted July 25, 2002.
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TOPABSTRACTINTRODUCTIONDYNAMICS OF THE ACTIN...T CELL ACTIVATION AND...ACTIN REARRANGEMENT DURING...CONCLUSIONREFERENCES |
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A. Zanin-Zhorov, R. Hershkoviz, I. Hecht, L. Cahalon, and O. Lider Fibronectin-Associated Fas Ligand Rapidly Induces Opposing and Time-Dependent Effects on the Activation and Apoptosis of T Cells J. Immunol., December 1, 2003; 171(11): 5882 - 5889. [Abstract] [Full Text] [PDF]
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