Abstract

Introduction

The innate immune system represents the initial gateway to almost all mammalian inflammatory responses against invading microbes. Members of the transmembrane Toll-like receptor (TLR)⁴ family are specialized for recognizing pathogen-associated molecular patterns with diverse structural and physiochemical properties, ranging from microbial cell wall components to nucleic acids (1). A key pathogen-associated molecular pattern is lipopolysaccharide (LPS) from the outer membranes of Gram-negative bacteria. LPS is a powerful activator of the innate immune response, and in sepsis overstimulation can lead to endotoxicity and death (2). LPS recognition is carried out by TLR4 in concert with a series of accessory proteins that amplify the response (3). LPS-binding protein and CD14 help to transfer LPS from bacterial membranes or aggregates in serum to myeloid differentiation factor 2 (MD-2) (4). Unlike the rest of the TLR family, TLR4 does not recognize ligand in isolation, but when bound to MD-2. Upon formation of an active receptor complex at the cell surface, it is presumed that conformational changes lead to transmission of an activating signal to the intracellular Toll-interleukin 1 receptor domains, resulting in recruitment of adaptor proteins, and subsequent MAL-MyD88 and TRAM-TRIF-dependent inflammatory responses (3, 5).

LPS is a glycolipid, composed of an oligosaccharide core and a highly variable O antigen polysaccharide component, along with a hydrophobic lipid A segment containing multiple lipid acyl tails and a phosphorylated glucosamine disaccharide headgroup (6). The lipid A core of LPS is responsible for much of its bioactivity. The structure and composition of lipid A vary among bacterial species, and even subtle variations can profoundly impact the stimulatory activity of TLR4. For example, whereas the highly agonistic Escherichia coli lipid A (LPA) is hexaacylated, its biosynthetic intermediate lipid IVa (LPIVa) is tetraacylated and acts as an agonist in mice and horses (7) but as an antagonist in humans (8). The tetraacylated synthetic compound eritoran (Erit) appears to be a strong antagonist in all species (9, 10). Monophosphoryl lipid A (MPLA) resembles LPA but has a singly phosphorylated headgroup; it acts as a partial agonist by activating the TRIF pathway without stimulating the MyD88 response (11).

A variety of recent biochemical and crystallographic data have provided insight into the molecular mechanisms of LPS recognition and signaling by TLR4. Like all TLRs, TLR4 has a curved, solenoidal ectodomain containing multiple leucine-rich repeat (LRR) domains (12, 13). MD-2 has been shown to form a TLR4·MD-2 heterodimer by binding to the amino-terminal and central LRRs of TLR4, at the “primary interface” (12). Moreover, LPS-bound MD-2 facilitates formation of the active receptor complex (13), consisting of a “dimer of dimers,” i.e. [TLR4·MD-2]₂, stabilized by new interactions between MD-2/LPS and the carboxyl-terminal LRRs on the opposing TLR4 chain (referred to here as TLR4*), at the “secondary interface” (Fig. 1A).

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FIGURE 1.

TLR4 and MD-2 structures. A, active LPA-bound [TLR4·MD-2]₂ heterotetrameric complex (13), with protein shown in schematic format. Each TLR4 (red/blue) and MD-2 (green) chain is labeled, and bound LPA is shown in spacefill format (CPK colors). Key heterodimeric interfaces, TLR4 termini, and LRRs are highlighted. B, overlay of the closed conformation of MD-2 (MD-2c) bound to LPA (13) shown in red, and the open conformation of MD-2 (MD-2o) bound to LPIVa (14) shown in blue. Protein is shown in schematic format, with ligand (wireframe) and Phe-126 (spacefill) highlighted.

Several crystal structures have confirmed that MD-2 adopts a nine-stranded immunoglobulin-like β-cup fold (with successive β-strands referred to as βA, βB, βC, etc.), composed of two curved β-sheets that house a deep hydrophobic pocket, specialized for binding the acyl tails of a range of lipid moieties (12–14). A variety of evidence suggests that one part of MD-2 plays a particularly important role in activation of TLR4, namely, the flexible βG-βH loop connecting strands βG and βH, which contains a key phenylalanine residue, Phe-126, located near the cavity mouth (Fig. 1B). An F126A mutant does not prevent ligand binding, but impairs formation of the active receptor complex and hence endotoxin-dependent signaling (15). Comparison of available structural data reveals that the βG-βH loop is conformationally dynamic, adopting an “open conformation” in the presence of nonactivating ligands, with Phe-126 oriented outside of the binding pocket, exposed to solvent (12, 14), and an alternative “closed conformation” in the presence of LPS, with Phe-126 pointing into the cavity, facilitating interaction with an exposed lipid tail (Fig. 1B) that may stabilize contacts between TLR4* and MD-2 in the [TLR4·MD-2]₂ complex (13). NMR analysis combined with metabolic labeling of a hexaacylated agonist recently provided support for the idea that Phe-126 acts as a “hydrophobic switch” (16). In both MD-2 and its F126A mutant, a single fatty acyl chain was shown to be more susceptible to paramagnetic attenuation, independent of TLR4 association, whereas the local environment of bound lipid tails was altered in the F126A mutant compared with wild type, suggesting that Phe-126 may be important in promoting formation of the active [TLR4·MD-2]₂ complex (16).

Despite the evident progress in characterizing TLR4 activation, there still remain outstanding questions. In particular, we lack information at atomic resolution about the nature of the agonist-bound MD-2 state prior to complex formation with TLR4, or conversely, an antagonist-bound receptor complex intermediate, both of which would help to clarify the nature of the “switch” in TLR4 molecular signaling. To gain further insights into the molecular signaling process, we now report an atomically detailed molecular simulation study of isolated MD-2 (iMD-2), as well as the entire active [TLR4·MD-2]₂ receptor complex, comprising almost half a million atoms (Fig. 2). The results obtained from this substantial computational effort, which constitutes ~3 μs of simulation time (Fig. 2), allow us to formulate detailed mechanistic hypotheses for the process of TLR4 (de)activation in the context of available experimental data.

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FIGURE 2.

Overview of each simulation system and corresponding naming convention, with domains represented schematically and colored as in Fig. 1. In isolated MD-2, the state of the gating loop containing Phe-126 is highlighted with a red dashed circle.

By first considering solvated human iMD-2 in the presence of a range of ligands (Fig. 3A and see Table 1), including agonistic LPA, partial agonist MPLA, antagonists LPIVa and Erit, and the inactive ligand myristate (Myr), we verify that lipid tail exposure precedes TLR4 association and is not exclusive to the agonist-bound state (16). Furthermore, we provide support for the notion that Phe-126 acts as a hydrophobic switch (16) within the βG-βH “gating loop.” We show that in both the closed (iMD-2c) and open (iMD-2o) gating loop states, the iMD-2 cavity is extraordinarily malleable, and can undergo “clamshell-like” motions, reversibly adjusting its volume to sensitively match the size and shape of the bound lipid moiety. Strikingly, MD-2 appears to be capable of transmitting the mechanical energy associated with cavity collapse and expansion to the gating loop, allosterically modulating its conformational state and determining potential ligand interactions with Phe-126. Furthermore, the apparent conformational plasticity of MD-2 is shown to be pertinent in the context of the complete [TLR4·MD-2]₂ heterotetramer. We directly observe spontaneous switching from the active to inactive signaling state in the absence of agonist (or to an intermediate state in the presence of partial agonist), coupled to destabilization of the receptor complex. In other words, a glimpse is provided of the Phe-126 molecular switch in action.

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FIGURE 3.

MD-2 ligand binding. A, chemical structures of ligands employed in this study. B, MD-2c structure (transparent schematic format) with bound ligand shown in wireframe format. For each system, the initial location of bound ligand (red) is shown, with the final conformation in iMD-2 (blue) and [TLR4·MD-2]₂ (green) simulations overlaid. The initial conformation of Phe-126 is shown in gray wireframe format, and for the LPA-bound system, Phe-126 and amino (NT) and carboxyl (CT) termini are labeled.

COMPUTATIONAL PROCEDURES

RESULTS

Protrusion of a Single Lipid Tail Is Insufficient for Signaling to TLR4

NMR-based relaxation enhancement was recently used to demonstrate the increased solvent accessibility of the mouth-proximal lipid tail in a hexaacylated agonist (16). To explore these observations further, we carried out molecular simulations for both active (iMD-2c) and inactive (iMD-2o) conformations of the gating loop and for the complete active [TLR4·MD-2]₂ complex and measured ligand tail solvent exposure (Fig. 4A). For LPA and MPLA, the positions occupied by the lipid tails were relatively stable throughout the simulation (Fig. 3B), with the R2 chain remaining solvent-exposed. The oppositely orientated lipid A state, with the glucosamine disaccharide headgroup rotated by 180° (LPAo), was also simulated; in this case, following relaxation of the initial tail conformations (Fig. 3B), the R3″ chain remained relatively exposed. Thus, for the hexaacylated ligands, the lipid tail nearest to the cavity mouth was consistently solvent-exposed, typically by approximately ~2–3 times that of the remaining fatty acyl chains. Importantly, this was the case for both the iMD-2c and iMD-2o states, in support of the NMR relaxation data for both wild type and F126A (16). In contrast with the relatively stable tails, the glucosamine disaccharide headgroups were able to shift across the cavity by up to ~0.5 nm, to satisfy direct or counterion-mediated salt bridges with the phosphate groups. In the absence of stabilizing electrostatic interactions with TLR4, the phosphate most distant from the cavity entrance became complexed with residues including Arg-96, Asp-99, Asp-100, and Asp-101 on a highly charged patch of MD-2, whereas the other phosphate tended to interact with Glu-92/Arg-90, resulting in an orientation more similar to the crystal structure of LPIVa-bound iMD-2.

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FIGURE 4.

Lipid tail exposure. Solvent exposure is defined as the mean number of waters within 0.6 nm of each lipid tail. A and B, histograms are shown for each iMD-2 system (A) and for the two symmetry-related MD-2 sites in each [TLR4·MD-2]₂ complex system (B). The lipid tails on the histogram are listed in order of their proximity to the cavity mouth, from closest (left) to furthest (right). C, the relations between lipid tail exposure and the conformation of Phe-126 (spacefill format) of MD-2 (schematic format) are highlighted for iMD-2^LPA (left) and iMD-2^Erit (right), at 0 ns (gray) and 100 ns (red).

Intriguingly, a similar pattern of lipid tail exposure was observed for the antagonistic and inactive ligands (LPIVa, Erit, and Myr) in iMD-2 as for the agonist-bound state (Fig. 4A). It is noteworthy that in all systems, ligand exposure was consistently greater in the closed gating loop conformation, suggesting that Phe-126 provides a platform to support fatty acyl protrusion. Moreover, although absolute exposure was reduced upon interaction with TLR4, the same overall pattern was again reproduced within the [TLR4·MD-2]₂ complex, irrespective of the nature of the bound agonist or antagonist (Fig. 4B). In summary, increased solvent exposure, relative to the remainder of the ligand, of the first lipid tail (or single fatty acid chain in the case of Myr) is not dependent upon the conformation of the gating loop or the nature of bound ligand, and precedes receptor association. It is therefore unlikely to be sufficient for TLR4 (in)activation, and hence, other molecular signals must be involved.

The MD-2 Cavity Allosterically Signals to the Gating Loop

Yu et al. showed via chemical shift perturbation that the local environment of labeled acyl tails in bound MD-2 agonist is somehow altered by Phe-126 (16). To characterize the behavior of the Phe-126 loop, we aligned the β-strands of simulated MD-2 onto the x-ray structure and then calculated the pairwise root mean square deviation (r.m.s.d.) of the loop Cα atoms; this provides an indication of the relative motion of the loop with respect to the rest of the protein. For iMD-2c^LPA, this r.m.s.d. was ~0.3 nm (Fig. 5A), and the side chain of Phe-126 retained its location for stable interaction with the proximal acyl tails (Fig. 5C). This was similar to iMD-2c^LPAo, consistent with minor NMR cross-peaks observed for the reverse orientation of endotoxin (16). However, all other ligand-bound states exhibited significant destabilization of the closed gating loop conformation, relative to the β-cup fold, with an r.m.s.d. of 0.45–0.5 nm (Fig. 5A), including MPLA, whose lack of a 1′-PO₄ “anchor” loosened glucosamine attachment and hence tail stability (Fig. 3) at the cavity mouth. Consequently, partial agonist, antagonist, and inactive ligands induced loss of the Phe-126 conformation common to the agonist-bound state (Fig. 5C) prior to receptor association, leading to a loss of interaction with the respective protruding lipid tail (Fig. 4C). This was concomitant with a loss of buried surface area (BSA) between the Phe-126 side chain and lipid tails by the end of each simulation, ranging from 1.47 ± 0.15 nm² and 1.05 ± 0.30 nm² for iMD-2c^LPA and iMD-2c^LPAo, respectively, to just 0.58 ± 0.22 nm² for iMD-2c^LPIVa. Moreover, there was effectively zero BSA observed for iMD-2c^MPLA (0.01 ± 0.03 nm²), iMD-2c^Erit (0.01 ± 0.04 nm²), and iMD-2c^Myr (0 ± 0.01 nm²). Thus, the conformational stability of the active gating loop conformation is reduced in the absence of agonist (see Table 1), supporting the proposal of Yu et al. that local rearrangements of this loop may determine the likelihood of receptor complex formation (16).

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FIGURE 5.

Conformational changes in MD-2. A, mean Cα r.m.s.d. over final 20 ns for all β-strand residues, or for the gating loop, in iMD-2c simulations, relative to the x-ray structure of MD-2 in the active receptor complex. B, mean Cα r.m.s.d. for all β-strand residues or gating loop of each MD-2 protein, and for structural regions of each TLR4 chain, over final 20 ns of [TLR4·MD-2]₂ simulations, relative to the x-ray structure of MD-2 in the active receptor complex. C, final conformation of Phe-126 in iMD-2c simulations; frame chosen according to proximity to the average buried area between Phe-126 and lipid over the final 20 ns. MD-2 is shown in schematic representation, with Phe-126 (green) and ligand (CPK colors) shown in wireframe format. The gating loop is highlighted in red-dashed format.

Although no large scale unfolding was evident (Fig. 5A), the strands surrounding the exit to the cavity exhibited local conformational heterogeneity in the absence of agonist, adapting their conformation according to the ligand present (Fig. 6A). Careful inspection of the MD-2 cavity over the course of each trajectory revealed that flexing and unfolding of βC-βD and βG-βH (i.e. leading into the gating loop), combined with adjustments in the separation between opposing β-sheets, led to significant changes in the internal cavity shape for some systems (Fig. 6A). Strikingly, a near perfect correlation was observed between the number of aliphatic carbons present in the fatty acyl tail portion of each ligand and the resultant cavity volume (Fig. 6B), for both the closed and open states of MD-2. The internal volume remained close to that of the LPS-bound x-ray structure in the presence of hexaacylated species (~2 nm³ for LPA, LPAo, and MPLA), shrunk to ~1.5 nm³ (Erit) and ~1.25 nm³ (LPIVa) for the tetraacylated ligands, and collapsed to half its original size (~1 nm³) when housing three myristate fatty acids (see Table 1). Thus, while avoiding major conformational rearrangements, MD-2 is able to sensitively adapt to each respective ligand via clamshell-like motions, adjusting the separation between opposing β-sheets; this is coupled to (de)stabilization of β-hairpins at the cavity mouth and consequently, the gating loop, thereby providing a possible mechanism for its role as a “molecular switch” in receptor (de)activation.

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FIGURE 6.

Malleability of the MD-2 cavity. A, mean surface of the internal hydrophobic cavities over the final 20 ns for iMD-2c systems. Cavity surfaces with >90% probability are shown in blue, with the equivalent >50% probability overlaid in transparent gray. Protein is shown in schematic representation, with Phe-126 in spacefill format. B, correlation between MD-2 cavity volume size and total number of aliphatic carbons within the tail of each bound ligand. R² = 0.95 for all data points; R² ≥0.95 for iMD-2c, iMDo, or [TLR4·MD-2]₂ systems only.

MD-2 Plasticity Is Reversible and Reproducible

In the absence of high resolution experimental data it is important to establish the reproducibility and extent of the apparent malleability of the MD-2 β-cup. Thus, 600 ns of additional simulation sampling was generated, split into six independent trajectories, consisting of MD-2 in the closed (iMD-2c^apo) or open state (iMD-2o^apo), in the absence of any lipid. Three further 100-ns iMD-2c^apo simulations were generated to test force field dependence, using the all-atom AMBER99SB-ILDN (21) and OPLS-AA/L (22) parameters, and the united-atom GROMOS53A6 (23) parameter set. Finally, additional 100-ns control simulations using CHARMM protein parameters with the TIP4P and TIP5P water models were also performed, for which comparable results were obtained (data not shown). In all cases, complete loss of the internal cavity was observed, with the volume initially reducing from ~1.5 nm³ to ~0.5 nm³ in the first ~50 ns (Fig. 7A), and to ≤0.25 nm³ toward the end of each simulation, i.e. the cavity was effectively absent (Fig. 6A). The final conformations of each apo system across force fields closely resembled one another (Fig. 7B), exhibiting significant narrowing of the cavity entrance and destabilization of the β-hairpins at the mouth.

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FIGURE 7.

Reproducibility and reversibility of MD-2 conformational plasticity. A, collapse of the binding cavity of ligand-free iMD-2c and iMD-2o with the CHARMM force field over 100 ns compared with that with AMBER, GROMOS, and OPLS. B, final MD-2 conformation (shown in schematic format) colored according to A. C, starting and final structures of iMD-2c^apo, with nonpolar hydrophobic side chains within the cavity shown in spacefill format, highlighting the process of hydrophobic collapse. D and E, change in volume over 100 ns for the experimental myristate-bound iMD-2^Myr, compared with collapsed iMD-2c^apo upon addition and subsequent binding of myristate (iMD-2^+Myr), with initial and final structures highlighted in E.

Closure in the absence of ligand resulted from hydrophobic collapse of the internal cavity (Fig. 7C), with concomitant loss of ~30–40 resident water molecules over the same time scale as cavity closure (Fig. 7A). To test the reversibility of this process, the final snapshot from an iMD-2c^apo system was extracted, and six myristate fatty acids were placed a minimum of 1 nm above the cavity opening (Fig. 7E). A subsequent simulation of this system (iMD-2c^+Myr) led to adsorption of four myristate chains onto the surface of the cavity opening within ~20 ns, a gradual reopening of the protein between 30 and 60 ns as three fatty acyl tails spontaneously tunneled into the hydrophobic core of the cavity, and a final relaxation phase with a mean volume of ~1.1 nm³ (Fig. 7D). It is helpful to compare this with the 100-ns iMD-2c^Myr simulation, based on a crystal structure in which unidentified ligand density was modeled as three myristic acid molecules (14), where a final cavity volume of ~1.2 nm³ was observed (Fig. 7D). Moreover, the backbone r.m.s.d. of the β-cup fold compared with the x-ray structure decreased from ~0.3 nm observed for iMD-2c^apo to ~0.15 nm in the iMD-2c^+Myr simulation over 100 ns, comparable with that observed for iMD-2c^Myr. Thus, the extreme malleability of the MD-2 β-cup fold is absolutely reproducible across a range of protein force fields and water models, and it may collapse or expand, depending upon the local presence of hydrophobic molecules.

The MD-2 Allosteric Switch Determines Receptor Complex Stability

To explore the role of the proposed MD-2 allosteric switch within the active receptor complex (13), the stability of LPA-bound [TLR4·MD-2]₂ was compared with that in the presence of MPLA, LP4A, Erit, or the lipid-free apo state, in the context of key interaction interfaces at the primary (TLR4·MD-2) and secondary (TLR4*·MD-2) sites (Fig. 1A). In all ligand-bound systems, the primary interface was well maintained (Fig. 8C), with a constant BSA ≥8 nm². This was supported in part by the presence of a salt bridge consistently formed between phosphate and Arg-264 (and in LPA only, Asp-294, via a bridging Na⁺ ion that spontaneously bound to this site in the absence of crystallographic Mg²⁺) (Fig. 8A). Even in the ligand-free, apo state, this contact interface was relatively stable, with a BSA ≥7 nm², as a result of a series of stable interactions formed between conserved ionizable residues on TLR4 and MD-2 (Fig. 9). Thus, formation of the TLR4·MD-2 heterodimer is not conditional upon the presence of stimulatory endotoxin, consistent with experiment (12).

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FIGURE 8.

Molecular signaling within the receptor complex. A, the x-ray structure of LPS-bound [TLR4·MD-2]₂ at one of the MD-2 bound sites is compared with the final simulation structure of the complex when bound to a range of ligands, or in the ligand-free state. Protein chains are shown in schematic representation and colored as in Fig. 1, with bound ligand and adsorbed sodium ions in wireframe or spacefill format, respectively. Key interacting residues are labeled for the x-ray structure. The orientation of Phe-126 is highlighted with a red, dashed ellipse. B, loss of docked, active conformation of Phe-126 (wireframe format) and concomitant disruption of TLR4*·MD-2 hydrophobic cluster (spacefill format) is highlighted for starting (left) and final (right) structures of [TLR4·MD-2]₂^apo. Protein chains are shown in schematic representation, colored according to A. C, the mean contact surface area buried between MD-2 and the receptor at the primary and secondary interfaces was calculated over the final 20 ns, each shown for two equivalent sites within the receptor complex.

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FIGURE 9.

Schematic representation of the common interactions observed within each [TLR4·MD-2]₂ system over the final 20 ns of simulation.

However, the secondary interface was only consistently stable in the LPA complex, which reproduced key MD-2·TLR4* interactions from the crystallographic LPS-bound structure (13) (Fig. 8A) and maintained a BSA of ≥5 nm² at both sites (Fig. 8C). Destabilization of this interface in the agonist-free systems was somewhat asymmetric with respect to the two TLR4*·MD-2 contact surfaces, with one receptor site apparently compensating for weakening at the other, supported in the context of the relative internal rigidity of the solenoidal TLR4 framework, as observed both in the complex (Fig. 5B) and in the isolated TLR4 (iTLR4) state, for which an r.m.s.d. of ≤0.2 nm was observed over ten simulation replicas (data not shown). For example, one of the sites in the MPLA and Erit systems exhibited a reduction in BSA of ~1 and ~3 nm², respectively (Fig. 8C). Moreover, the LPIVa and apo systems each exhibited a reduction of ~2 nm² at one site, and as much as ~4 nm² at the other (Fig. 8C). This partly resulted from a loss of “tethering” of the glucosamine associated with the more exposed acyl tails, either due to the absence of a 1-PO₄ group in MPLA or contraction of the MD-2 cavity in response to tetraacylated Erit and LPIVa (Fig. 6). Thus, in contrast with LPA where its 1-PO₄ phosphate group interacted electrostatically with Lys-388 on TLR4*, only the more “buried” phosphate group was consistently coordinated in the agonist-free systems, by Lys-362 and/or Lys-341 of the TLR4 chain (Figs. 8A and ​and9);9); intriguingly, these residues are part of a hypervariable region important for species-dependent specificity (31).

Consistent with the stable cavity volume (Fig. 6B) and low structural heterogeneity of the gating loop in [TLR4·MD-2]₂^LPA (Fig. 5B), the conformation of Phe-126 remained close to that of the x-ray structure, forming a hydrophobic cluster of interactions with the exposed LPA lipid tail along with residues including Ile-80, Val-82, and Leu-87 in MD-2, and Phe-440, Leu-444, and Phe-463 in TLR4* (Figs. 8A and ​and9),9), supported by a salt bridge formed between nearby Lys-125 of MD-2 and Glu-422 of TLR4*. This is in striking contrast to the agonist-free interfaces; following the reduction in MD-2 cavity volume (Fig. 6) and gating loop stability (Fig. 5B), spontaneous reorientation of Phe-126 from the closed to open state was observed to occur, in the presence of LPIVa, Erit, and in the ligand-free apo state (Fig. 8A and Table 1). This led to loss of interaction with lipid and destruction of the TLR4*·MD-2 hydrophobic cluster evidently critical for maintaining the receptor heterodimerization interface. In particular, the open orientation of Phe-126 disrupted the arrangement of nearby side chains of Leu-87, Val-82, and Met-85 in MD-2, leading to loss of a pocket that supported the phenyl ring of Phe-463 from TLR4* in the closed state (Fig. 8B). In the case of the MPLA system, partial agonist induced a reoriented Phe-126 state that was intermediate between the agonist and antagonist systems (Fig. 8A).

DISCUSSION

Our findings help to clarify the molecular signaling mechanism in TLR4, the details of which are consistent with and extend recent observations from biochemical and biophysical experiments. In particular, we have built on the work of Yu et al. (16), confirming that lipid tail exposure alone is insufficient for determining formation of the active receptor complex, and showing that tuned (de)stabilization of the gating loop in response to bound ligand influences stability of Phe-126, prior to receptor association. Furthermore, for the first time we observe the spontaneous reorientation of Phe-126 from the closed to open conformation in various agonist-free states of the active receptor complex, with concomitant destabilization of the key TLR4*·MD-2 heterodimeric interface, confirming the role of Phe-126 as a molecular switch. We speculate that this loss of stability at the heterodimeric interface in the absence of agonist would lead to changes in the relative arrangement of TM domains, which play a key functional role in oligomerization (32–34), and loss of organization of the activated cytoplasmic TIR domain scaffold necessary for recruitment of adaptor proteins and subsequent signaling. Indeed, in comparison with [TLR4·MD-2]₂^LPA, which maintained the separation found within the x-ray structure (13) between the pair of membrane-proximal carboxyl-terminal regions on each TLR4 chain, all agonist-free systems demonstrated significant divergence from this, by 0.2–0.3 nm. Thus, the conformational plasticity that we observe for each system at the TLR4*·MD-2 interface correlates with the relative arrangement of the carboxyl-terminal regions, in agreement with recent structural data obtained for TLR8 which revealed changes in dimerization interface and carboxyl-terminal separation upon ligand binding (35). However, it should be borne in mind that the time scales associated with downstream signal propagation are likely to be significantly longer than we have sampled in our simulations. To our knowledge, the kinetics of TLR4 assembly and extracellular-to-intracellular signaling is unknown, but almost certainly spans microsecond to millisecond time scales or beyond.

We have also provided evidence that MPLA results in an intermediate level of Phe-126 switching, which may help to explain why it is only able to weakly activate the MAL-MyD88 signaling pathway, due to diminished recruitment of MyD88 to TLR4. Indeed, the resultant, partially destabilized TLR4*·MD-2 interface is consistent with recent data from Casella and Mitchell (36). An antibody assay revealed that MPLA species drove low potency heterotetramerization in comparison with lipid A. However, the signaling capacity of MPLA was reduced by the F126A MD-2 mutant (which allows lipid binding but interferes with active TLR4·MD-2 complex formation), suggesting that an intermediate level of TLR4·MD-2 heterotetramerization is the likely explanation for the weak, biased agonism of MPLA, in agreement with our atomic level observations.

Key to the proposed signaling mechanism is the apparent malleability of the MD-2 scaffold. Its β-cup fold exhibits reversible, clamshell-like motions that enable it to allosterically transmit its ligand-bound state to the Phe-126 molecular switch. Our conclusions have been made on the basis of several protein conformations, a wide variety of ligand-bound states, and multiple simulation replicas of the ligand-free state, using four force fields and three water models. The reproducibility of the conformational plasticity of MD-2 is in partial agreement with previous theoretical studies revealing flexibility around the cavity mouth (37) and collapse of apo MD-2 (38), though no discernible trends with regard to ligands or the gating loop were reported, likely due to the significantly shorter time scales used. Only limited evidence has so far been found for the plasticity of MD-2 from high resolution structural approaches. Comparison of various ligand-bound structures and associated B-factors (12–14) at least hints at the potential for flexibility around the mouth of the β-cup, whereas it is possible that the presence of disordered/unidentified hydrophobic ligands within the cavity (14) and extensive crystal lattice contacts may mask the conformational plasticity of MD-2. Significantly, the MD-2-related lipid-recognition family also includes mite allergen proteins such as Der f 2 and Der p 2, for which both collapsed/closed and open experimental structures do exist, separated by clamshell-like motions (39). These conserved dynamics also provide evidence that mite allergenicity may result from TLR4 coercion.

In terms of time scale, the experimental characterization of hydrophobic collapse has typically been hampered by difficulties in isolating the process from other events such as folding. However, Sadqui et al. were able to directly follow the dynamics of hydrophobic collapse in a simple protein, in the absence of competing processes, via fluorescence resonance energy transfer (FRET) measurement between two terminal fluorophores upon a laser-induced temperature jump trigger (40). The relaxation time for hydrophobic collapse at room temperature was on the order of tens of nanoseconds. Moreover, comparable time scales as described here for hydrophobic collapse were recently reported for atomistic simulations of another lipid-binding protein family, CD1, suggesting that this may be a common mechanism in hydrophobic ligand binding and release (41).

From the biomedical viewpoint, our results advocate the importance of considering conformational plasticity when attempting to design novel therapeutic molecules for manipulation of TLR4 signaling pathways. Thus, molecular simulation represents a genuinely useful strategy for predicting the stimulatory outcome of novel compounds, particularly because endotoxic ligands typically possess “non-drug-like” properties. We have demonstrated a striking correlation between the size of the hydrophobic portion of the ligand and cavity volume of MD-2, which should be useful for rapidly estimating the degree of endotoxicity; and we have shown that this directly, but asymmetrically, can determine receptor complex stability and hence activation. Although the time scales sampled here are insufficient to delineate the complete regulatory pathway, if the observed asymmetry of receptor de(activation) is indeed a general feature of TLR4 signaling, this has potential consequences for ligand design, as well as for refinement strategies of crystallographic receptor structures. It is therefore hoped that the knowledge gained from this study will contribute both to understanding receptor signaling mechanisms and to the development of new molecules for pharmacological manipulation of TLRs, as their association with many infectious, allergic, inflammatory and malignant disease continues to intensify (2).

Acknowledgments

REFERENCES