Action and Traction: Cytoskeletal Control of Receptor Triggering at the Immunological Synapse

Abstract

It is well known that F-actin dynamics drive the micron-scale cell shape changes required for migration and immunological synapse (IS) formation. In addition, recent evidence points to a more intimate role for the actin cytoskeleton in promoting T cell activation. Mechanotransduction, the conversion of mechanical input into intracellular biochemical changes, is thought to play a critical role in several aspects of immunoreceptor triggering and downstream signal transduction. Multiple molecules associated with signaling events at the IS have been shown to respond to physical force, including the TCR, costimulatory molecules, adhesion molecules, and several downstream adapters. In at least some cases, it is clear that the relevant forces are exerted by dynamics of the T cell actomyosin cytoskeleton. Interestingly, there is evidence that the cytoskeleton of the antigen-presenting cell also plays an active role in T cell activation, by countering the molecular forces exerted by the T cell at the IS. Since actin polymerization is itself driven by TCR and costimulatory signaling pathways, a complex relationship exists between actin dynamics and receptor activation. This review will focus on recent advances in our understanding of the mechanosensitive aspects of T cell activation, paying specific attention to how F-actin-directed forces applied from both sides of the IS fit into current models of receptor triggering and activation.

Keywords: T cell receptor; actin; adhesion; costimulation; cytoskeleton; immunological synapse; integrin; mechanotransduction.

Figure 1

Organization and actin dynamics within the IS. Diagram showing the architecture of a radially symmetric “bulls-eye” IS such as that formed between a B cell and an antigen-specific mature T cell. Based on molecular segregation, the IS can be divided into three regions: (1) a peripheral actin-rich region termed the distal supramolecular activation cluster (D-SMAC), (2) a deeper region rich in LFA-1 and actomyosin arcs termed the peripheral supramolecular activation cluster (pSMAC), and (3) a central region rich in PKCθ and other signaling molecules termed the central supramolecular activation cluster (cSMAC). Signaling microclusters containing TCR and other signaling molecules (gold balls) form and begin to signal in the IS periphery and are transported by the cytoskeleton toward the cSMAC region, where signal extinction takes place. Microcluster movement is coupled to centripetal flow of the actin network (blue arrows). Actin flow is driven primarily by addition of actin monomers to the barbed ends of branched actin filaments, which lie just under the plasma membrane. This generates a pushing force that drives the network inward. In addition, myosin-driven sliding of actin filaments causes contraction of the network. This provides a pulling force that stabilizes the network and maintains radial symmetry.

Figure 2

Regulation of the F-actin network at the IS. Ligation of multiple receptors, including the TCR, the costimulatory molecule CD28, and the adhesion molecule LFA-1, results in the induction of robust actin polymerization at the IS. The pathways that mediate F-actin polymerization are highly interdependent. For example, the TCR-dependent activation of Lck is involved in initiating CD28-mediated signaling. Moreover, both TCR-induced activation and CD28-induced recruitment of PKCθ contribute to LFA-1 activation and downstream signaling. Vav1, a GEF for the critical actin regulators Rac1 and CDC42 and their respective effectors WAVE2 and WASp, is triggered in a co-operative fashion downstream of each of these key surface receptors. In addition, signaling events downstream of CD28 lead to inhibition of capping protein and activation of cofilin, events that allow growth and remodeling of the branched actin network. Together with HS1, WAVE2 and WASp activate ARP2/3 complex-dependent growth of branched actin filaments, forming lamellipodial protrusions and invadopodium-like protrusions, respectively. Collectively, these events lead to cell spreading, retrograde actin flow, and formation of the mature IS, along with assembly and centripetal flow of TCR-associated signaling microclusters. In addition to triggering actin nucleation, TCR, CD28, and LFA-1 all associate with the F-actin network (proteins known to interact with F-actin are denoted by yellow stars). In many cases, these interactions with the F-actin network serve to drive additional signaling events via actin-dependent feedback loops.

Figure 3

Mechanisms through which actin-dependent forces can contribute to proposed models controlling TCR triggering. (A) The kinetic-segregation model of TCR signaling is dependent on the separation of molecules with small extracellular regions, such as the TCR, from those with large extracellular regions, such as the phosphatase CD45. Actin-dependent protrusions would serve to bring the T cell and APC plasma membranes into close proximity, thereby driving molecular segregation. This should occur even in the presence of low numbers of cognate pMHC on the APC surface. (B) The kinetic proofreading model proposes that TCR triggering is based on longer bond lifetimes for strong agonists than weak agonists. The force-dependent catch-bond behavior of the TCR with strong, but not weak, agonist pMHC complexes can enhance bond lifetime for strong agonists, while serving to rupture the slip-bonds formed by TCRs engaging weak or non-agonist pMHC. (C) The serial triggering/serial engagement molecule could benefit from the presence of multiple F-actin interactions with the TCR. Though it may not be force dependent, the association of the TCR with the F-actin network could lead to clustering of the TCR on the plasma membrane, allowing for rapid successive unbinding and rebinding, and serial triggering of multiple TCRs by a single pMHC complex. (D) Several conformational changes that have been described for the TCR may be induced or enhanced by the application of force by the T cell actin cytoskeleton. The first posits a mechanical lever-type action of the TCR under the application of a tangential force. According to this model, bending of the stiff CD3 chains propagates to the intracellular domain and results in signal initiation. The second model suggests that actin associations with the CD3 complex help to pull the CD3 chains away from the inner leaflet of the plasma membrane, thus exposing the ITAMs for phosphorylation and binding of essential regulators such as the kinase ZAP70. The third model is based on catch-bond molecular interactions between TCR and cognate pMHC complexes. According to this variant of the kinetic proofreading model, cytoskeletal force causes a conformational change in the TCR that results in stronger pMHC binding and prolonged or enhanced signaling.

Figure 4

Integrin regulation by cytoskeletal forces. (A) Known α and β integrin chains and pairings. Integrin pairs expressed in leukocytes are depicted in red. (B) Domain structure of the integrin LFA-1. The α chain consists of an intracellular tail (cyan), a transmembrane domain (orange), two calf and one thigh domains (gray), and a β-propeller domain (purple) with an inserted ligand-binding I domain (pink). The β chain consists of an intracellular tail (cyan), a transmembrane region (orange), a β tail domain (brown), four EGF repeats (yellow), a hybrid domain (blue) with an inserted βI domain (green), and a PSI domain (red). (C) Retrograde actin flow drives LFA-1 into the IS from the cell periphery. This increases local concentrations of LFA-1, thereby increasing the valency of the interaction and strengthening cell–cell adhesion. (D) LFA-1 can exist in roughly three conformations: a bent, low affinity conformation; an extended intermediate affinity conformation; and an extended conformation, where the hybrid domain on the β chain is swung outward, allowing for downward movement of the α7 helix in the βI domain. This downward movement allows the βI domain to bind an internal ligand in the αI domain, causing downward movement of the αI domain α7 helix and opening of the ligand-binding site. These changes generate a high affinity, extended-open conformation. Maintenance of this conformation at the IS is dependent on ongoing actin flow, presumably because connection of the β chain intracellular domain to the dynamic F-actin network is enough to drive swing-out of the hybrid domain. The resulting force-dependent increase in affinity would promote and augment changes induced by ligand binding. (E) In addition to regulating LFA-1 affinity for ligand, applied force can also strengthen the connection of LFA-1 to the underlying actin cytoskeleton. Talin, a key protein that links integrins to the actin network, can stretch upon the application of force. This stretching reveals up to 11 cryptic vinculin-binding sites. Vinculin, itself an actin-binding protein, then binds to the exposed sites and reinforces linkage to the F-actin network.

Figure 5

Regulation of CD28 signaling by applied forces. (A) CD28 clustering at the IS may occur in a signaling-independent manner through the kinetic segregation of bound and unbound molecules. Actin polymerization could contribute to this process by generating protrusions that bring the T cell plasma membrane into close proximity with that of the APC. In the case of a receptor and a surface bound ligand, the free receptor would occupy less extracellular space than the bound receptor–ligand pair, such that bound receptors would be forced to cluster within areas of low membrane proximity. (B) Filamin A (FLNa) is a scaffolding protein and an essential CD28 effector that can undergo force-dependent conformational change. Force causes the extension of the second rod domain of filamin A, eliminating binding sites that exist in the relaxed protein, and exposing a different set of binding sites that only exist in the extended protein. Thus, under tension, FLNa can release low force-binding partners and recruit new proteins to the CD28 signaling complex.