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. 2009 May 6;28(9):1351-61.
doi: 10.1038/emboj.2009.63. Epub 2009 Mar 12.

The Structure of the Integrin alphaIIbbeta3 Transmembrane Complex Explains Integrin Transmembrane Signalling

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The Structure of the Integrin alphaIIbbeta3 Transmembrane Complex Explains Integrin Transmembrane Signalling

Tong-Lay Lau et al. EMBO J. .
Free PMC article

Abstract

Heterodimeric integrin adhesion receptors regulate cell migration, survival and differentiation in metazoa by communicating signals bi-directionally across the plasma membrane. Protein engineering and mutagenesis studies have suggested that the dissociation of a complex formed by the single-pass transmembrane (TM) segments of the alpha and beta subunits is central to these signalling events. Here, we report the structure of the integrin alphaIIbbeta3 TM complex, structure-based site-directed mutagenesis and lipid embedding estimates to reveal the structural event that underlies the transition from associated to dissociated states, that is, TM signalling. The complex is stabilized by glycine-packing mediated TM helix crossing within the extracellular membrane leaflet, and by unique hydrophobic and electrostatic bridges in the intracellular leaflet that mediate an unusual, asymmetric association of the 24- and 29-residue alphaIIb and beta3 TM helices. The structurally unique, highly conserved integrin alphaIIbbeta3 TM complex rationalizes bi-directional signalling and represents the first structure of a heterodimeric TM receptor complex.

Figures

Figure 1
Figure 1
Sequence alignment of the transmembrane segments of all human integrin subunits. (A) The 18 α subunits and (B) 8 β subunits are depicted. Proposed minimal lipid tail-to-headgroup borders for monomeric and heterodimeric α and β subunits are depicted (c.f. Figure 7) (Armulik et al, 1999; Stefansson et al, 2004; Lau et al, 2008a, 2008b). Conserved amino acids are coloured by the Jalview multiple alignment editor (Clamp et al, 2004) using the ClustalX colour scheme.
Figure 2
Figure 2
Illustration of αIIb–β3 heterodimerization at the protein backbone level. (A) H–N correlation spectra for each labelled subunit in the presence of its unlabelled partnering subunit. For each transmembrane resonance, a second one is obtained, as indicated by connecting lines for resonances in well-resolved spectral regions. For comparison, spectra for monomeric αIIb and β3 peptides are shown in Supplementary Figure 1. Spectra were recorded at 23°C and a 1H frequency of 700 MHz using bicelles composed of 385 mM 1,2-dihexanoyl-sn-glycero-3-phosphocholine, 83 mM 1-palmitoyl-2-oleoyl-sn-glycero-3-phosphocholine, 41 mM 1-palmitoyl-2-oleoyl-sn-glycero-3-[phospho-L-serine]. (B) Average H–N chemical shift difference for a given residue between monomeric and heterodimeric resonances. The shift difference for a residue i is calculated as {[Δδi(1HN)2 + (Δδi(15N)/5)2]/2}1/2 (Grzesiek et al, 1997). (C) Verification of specific αIIb–β3 heterodimerization. The signal intensities of the monomeric and heterodimeric peaks, for example, V695 and I704 of the 2H/13C/15N-labelled β3 subunit, correlate with peptide concentrations, but their positions are independent of peptide concentrations, showing slow exchange on the NMR timescale. Moreover, a mutant αIIb(R995A) peptide does not induce any significant second set of resonances, verifying the specificity of the αIIbβ3 heterodimeric association in bicelles and corroborating strong αIIb(R995)–β3(D723) electrostatic interaction (c.f. main text).
Figure 3
Figure 3
Illustration of αIIb–β3 heterodimerization at the protein side chain level. H–C correlation spectra of the β3 isoleucine methyl groups are shown for β3 alone, and for β3 in the presence of increasing concentrations of its partnering αIIb subunit. For I693 and I719, a second heterodimeric resonance is obtained, as indicated by the connecting lines (slow exchange limit). For the remaining four isoleucines, small or no shifts of only one resonance are detected (fast exchange limit). The behaviour of a resonance to exhibit fast or slow exchange depends, among other factors, on the chemical shift difference between monomeric and heterodimeric states.
Figure 4
Figure 4
Comparison of 13Cα secondary chemical shifts between monomeric and heterodimeric integrin αIIb–β3 transmembrane states. (A, B) Secondary 13Cα chemical shifts, defined as the difference between the observed and tabulated random-coil 13Cα shift of a residue, correlate with the underlying backbone conformation but are also sensitive to local backbone dynamics (Spera and Bax, 1991; Ulmer et al, 2005). The minor differences between monomeric and heterodimeric shifts indicate the absence of significant backbone rearrangements on heterodimerization and the stabilization of secondary structure at the intracellular side in the presence of αIIb(R995)–β3(D723) electrostatic interactions (Figure 2C). Chemical shifts were measured in bicelles composed of 385 mM 1,2-dihexanoyl-sn-glycero-3-phosphocholine, 83 mM 1-palmitoyl-2-oleoyl-sn-glycero-3-phosphocholine, 41 mM 1-palmitoyl-2-oleoyl-sn-glycero-3-[phospho-L-serine].
Figure 5
Figure 5
Structure of the integrin αIIbβ3 transmembrane complex. (A) Superposition of the ensemble of 20 calculated simulated annealing structures. αIIb(I966-R995) and β3(I693-D723) adopt well-structured conformations. (B, C, E) Selected views of the energy minimized, average structure. (D) The outer membrane clasp: illustration of glycine packing. αIIb(G972), αIIb(G976) and β3(G708) are shown in green spheres, with their β3 and αIIb packing residues shown in blue and red spheres, respectively. (F) The inner membrane clasp: stabilization of the α–β TM helix arrangement by αIIb(F992-F993)-mediated interhelical packing and αIIb(R995)–β3(D723) electrostatic interaction. β3(W715) and β3(D723) are shown in blue, αIIb(R995) in red, and αIIb(F992-F993) in green spheres.
Figure 6
Figure 6
Site-directed mutagenesis of the αIIbβ3 dimer interface. (A) Selected mutations were introduced to an αIIb construct, αIIb™-TAP, consisting of the αIIb TM region plus cytosolic tail (Q954-E1008) with an N-terminal flag-tag, a C-terminal TAP (tandem affinity purification; calmodulin-binding domain and IgG-binding domain) tag, and a β3 construct, Tac-β3™, encompassing the β3 TM region plus cytosolic tail (V681-T762), with the extracellular domain of interleukin-2 receptor α (Tac) fused to the N-terminus (B) After co-transfection of αIIb™-TAP and Tac-β3™ constructs into CHO cells, their association was analysed by capturing αIIb™-TAP using calmodulin beads, and subsequently detecting bound Tac-β3™ through western blotting using an anti-Tac antibody (upper panels). Expression of Tac-β3™ (middle panels) and captured αIIb™-TAP (bottom panels) was verified by western blots using anti-Tac and anti-flag antibodies, respectively. The interaction was quantified by calculating the amount of bound Tac-β3™ divided by the amount of expressed Tac-β3™ and captured αIIb™-TAP. The mean and standard error from three independent experiments are depicted. (C) To verify the analogy of the assay with the NMR studies, the strongly TM-dissociating nature of the salt bridge mutations, αIIb™-TAP(R995A) and Tac-β3™(D723A), was confirmed.
Figure 7
Figure 7
Model of integrin αIIbβ3 membrane embedding. (A, B) Signal broadening due to paramagnetic relaxation enhancement arising from the presence of net neutral Mn2+EDDA2− in the aqueous phase. The normalized ratio of H–N TROSY signal intensities, in the presence and absence of 1 mM Mn2+EDDA2−, I/I0, is used to quantify signal broadening for each monomeric and heterodimeric residue in slow exchange. Datasets were recorded at two temperatures (28 and 33°C) to verify the obtained broadening pattern. αIIb(I966-F993) and β3(I693-I721), marked by dashed lines, are considered membrane embedded (Lau et al, 2008a, 2008b). The αIIb TM helix border, αIIb(K989-V990), is also marked. Mn2+EDDA2− protection was evaluated in bicelles composed of 385 mM 1,2-dihexanoyl-sn-glycero-3-phosphocholine, 83 mM 1-palmitoyl-2-oleoyl-sn-glycero-3-phosphocholine, 41 mM 1-palmitoyl-2-oleoyl-sn-glycero-3-[phospho-L-serine]. (C) Predicted orientations of the αIIb and β3 TM segments, shown in red and blue, in their monomeric and associated states, respectively, relative to the indicated functional groups of a lipid bilayer composed of 1,2-dioleoyl-sn-glycero-3-phosphocholine (Wiener and White, 1992), which is closely related to the long-chain lipids used herein. In addition, the activating β cytosolic tail–talin F3 complex (Wegener et al, 2007) (PDB ID 2h7e) has been fused to the monomeric β3 TM segment (PDB ID 2rmz). The depicted Lys322 and/or Lys324 side chains of talin are positioned to interfere with the IMC, particularly αIIb(R995)–β3(D723) electrostatic interactions.
Figure 8
Figure 8
Stabilization of the αIIbβ3 transmembrane complex by the resting integrin ectodomain. The αIIb(G972S), αIIb(G976A), β3(L712A) and β3(W715Y) substitutions, which were TM dissociating in the absence of the integrin ectodomain (Figure 6B), were evaluated to determine whether they were capable of activating full-length integrin αIIbβ3 receptors. The point mutation-bearing αIIb and β3 subunits were co-transfected into CHO cells. Subsequent to 24 h of transfection, cells were stained with PAC1 (an activation-specific αIIb–β3 antibody) to measure activation, and with D57 (an αIIb–β3 complex-specific antibody) to measure surface expression. D57-positive cells were analysed to calculate the Activation Index, defined as (F0Fr)/(FMnFr), where F0 is the mean fluorescence intensity (MFI) of PAC1 binding, Fr is the MFI of PAC1 binding in the presence of competitive inhibitor (integrilin), and FMn is the MFI of PAC1 binding in the presence of 2 mM Mn2+. In contrast to activating control mutations, LXXXL denoting αIIb(G972 L)/αIIb(G976 L) and αIIb(R995D) (Hughes et al, 1996; Luo et al, 2005), the substitutions were unable to trigger significant integrin activation, showing the stabilization of the αIIbβ3 TM complex by the resting integrin ectodomain.
Figure 9
Figure 9
Illustration of proposed integrin receptor functional states. (A) Inside-out activated integrin composed of an activated extracellular headpiece (Xiao et al, 2004) (PDB ID 2vdn), extended I-EGF1-2 domains (Shi et al, 2007) (PDB ID 2p28) and modelled tailpiece, fused to the monomeric αIIb and β3 TM structures (PDB ID 2k1a and 2mrz), and connected to the activating β cytosolic tail–talin F3 complex (Wegener et al, 2007) (PDB ID 2h7e). Talin F3 domain binding stabilizes α-helical structure subsequent to the β TM helix. (B) Resting integrin composed of the bent integrin αVβ3/αIIbβ3 structure (Xiong et al, 2001; Adair et al, 2005; Zhu et al, 2008) (PDB ID 1jv2), the αIIbβ3 TM complex (PDB ID 2k9j), and dynamically unstructured cytosolic tails (Ulmer et al, 2001; Li et al, 2002). (C) Outside-in activated integrin composed of an activated extracellular headpiece (Xiao et al, 2004) (PDB ID 2vdn), extended I-EGF1-2 domains (Shi et al, 2007) (PDB ID 2p28) and modelled tailpiece, fused to the monomeric αIIb and β3 TM structures (PDB ID 2k1a and 2mrz), and connected to dynamically unstructured cytosolic tails (Ulmer et al, 2001; Li et al, 2002). As no high-resolution structures of an entire, activated ectodomain exists, the depicted domain–domain orientations only serve to illustrate a presumed extended geometry (Adair and Yeager, 2002; Takagi et al, 2003; Zhu et al, 2008). In each functional transition, the TM helices have been rotated in addition to their dissociation. During both inside-out and outside-in signalling, additional ectodomain intermediates are likely to exist (Takagi et al, 2003; Xiao et al, 2004; Zhu et al, 2008) and some debate regarding the structural rearrangement of the ectodomain on activation remains (Arnaout et al, 2005; Rocco et al, 2008; Ye et al, 2008; Zhu et al, 2008).

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