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Pathogen Recognition Receptors Provide The First Line Of Detection For Microbial Antigen

As we learned in Chapter 1, the innate immune system employs an impressive battery of defense mechanisms that specifically detect the presence of invading microbes, to coordinate a series of rapid responses that deal directly with the invader, while at the same time sowing the seeds for a more specific and long‐lasting adaptive immune response. Over many millennia of co‐evolution, vertebrate immune systems have become impressively adept at accurately identifying the presence of potentially harmful microbes, through the detection of microbial structures that are essential for viability and, therefore, refractive to the pressures of natural selection. These conserved microbial antigens, called pathogen‐associated molecular patterns (PAMPs), are unique to individual classes of microbes, and as such, convey pathogen‐specific information to the innate immune system, to facilitate an appropriate response tailored to the particular threat at hand.

Detection of PAMPs is facilitated by a family of evolutionarily conserved germline‐encoded receptors called pathogen recognition receptors (PRRs), expressed on innate immune cells such as DCs, macrophages, and neutrophils. PAMP detection is often the first indication to the innate immune system of microbial presence and consequently, PAMP‐induced PRR activation rapidly promotes the production of a host of cytokines, chemokines, and type 1 interferons that mobilize innate immune cells to directly confront the invader. Additionally, PRR stimulation acts as a crucial line of communication between the innate and adaptive immune systems by instructing antigen‐presenting cells, such as DCs, to effectively
license a T‐cell‐mediated adaptive immune response against a particular antigen. As will be discussed in later chapters, the particular mode of T‐cell activation is further shaped by PRR‐ induced DC‐derived cytokines, which effectively tailor the T‐ cell‐mediated response to the particular type of microbe. As PRR signaling has also been shown to be important for instructing B‐cells to respond to particular types of microbial antigen, it should be clear that the recognition of microbial PAMP by PRRs plays a crucial role in coordinating both innate and adaptive immune responses to infection.
To date, several different classes of PRRs have been charac­ terized, including Toll‐like receptors (TLRs), NOD‐like receptors (NLRs), RIG‐1‐like receptors (RLRs), DNA receptors, and C‐type lectin‐like receptors, which together sense a wide range of conserved microbial antigen. TLRs are among the best‐characterized PRRs and we will next turn our attention to this important immune receptor family.
TLR family structure, ligand specificity, and signaling mechanism (a) Structures of TLRs bound to ligand and arranged into a phylogenetic tree.
Figure 4.30 TLR family structure, ligand specificity, and signaling mechanism (a) Structures of TLRs bound to ligand and arranged into a phylogenetic tree. The ligands are colored red, and TLRs are blue and green. (b) Overview of LPS recognition by TLR4/MD‐2. LPS binding induces dimerization of the TLR4/MD‐2 complex, which is proposed to enable dimerization of the intracellular TIR domains and recruitment of adapter molecules such as MyD88. Aggregation of the death domains (DD) of MyD88 brings four IRAK4 and four IRAK2 molecules together forming a large tower‐like structure called the “Myddosome.” (Source: Park B.S. et al. (2013) Experimental and Molecular Medicine 45(12), 1–9. Reproduced with permission of Nature Publishing Group.)

Toll‐like receptors detect a wide range of conserved microbial PAMP
Named after a Drosophila protein that was originally discovered as important for embryogenesis and later, as required for anti­fungal immunity, Toll‐like receptors (TLRs) are a key family of mammalian PRRs involved in the detection of a wide variety of PAMPs. To date, 10 TLRs have been described in humans, and 12 have been characterized in mice. TLR1, 2, 4, 5, and 6 are expressed on the cell surface and detect ligands from bacteria, fungi, protozoa, and certain self antigens, whereas expression of TLR3, 7, 8, and 9 are confined to intracellular endocytotic compartments, where they recognize nucleic acids signatures unique to bacteria and viruses (Figure 4.30a).
TLRs are type 1 integral membrane receptors composed of an extracellular ligand‐binding domain, a single transmembrane helix, and an intracellular Toll/IL‐1R (TIR) signaling domain, named because of its homology to the signaling domains of the interleukin‐1 receptor superfamily. Ligand binding induces dimerization of extracellular TLR domains, which in turn facilitates the localization and subsequent dimerization of intracellular TIR domains required for signaling. Dimerized TIR domains then recruit various adaptors, including myeloid differentiation primary response protein 88 (MyD88) (Figure 4.30b) and TIR‐domain‐containing adaptor inducing interferon‐β (TRIF), which ultimately promote activation of transcription factors such as nuclear factor kB (NFκB) and interferon regulatory factors (IRFs), responsible for inducing expression of cytokines, chemokines, and antimicrobial factors.
TLRs belong to the leucine‐rich repeat (LRR) family of proteins, with extracellular domains characterized by tandem repeats of LRR modules of 20–30 amino acids in length, with the hydrophobic leucines spaced at defined intervals. The leucines face toward the interior of the protein, forming a hydrophobic core that acts to stabilize overall protein structure, with variable regions facing outward to form a β arrangement gives TLRs a classical solenoid‐like shape, with each LRR module organized into adjacent, coiled, circular structures, similar to the way nuclear DNA is wound around histones, while the β‐sheet of one LRR is arranged in parallel with the β‐sheet of an adjacent LRR. As the β‐sheets are more tightly packed than the rest of the LRR, the overall structure of the receptor is forced to bend into a horseshoe shape, with β‐ sheets arranged on the concave side (Figure 4.31a). Although the majority of LRR family proteins interact with protein ligands, TLRs are distinct in their interaction with nonprotein antigens, with ligands interacting at the concave or lateral sides of the receptor.
Although all TLRs share similar overall structure, they display considerable divergence in their ligand‐binding affinities, driven mainly by differences in the size and charge of ligand‐binding pockets, and their ability to engage in ligand‐induced homodimerization (TLR3, TLR7) and ligand‐driven heter- odimerization with other members of the TLR family (TLR2/1, TLR2/6), and with non‐TLR co‐receptors (TLR4/ MD‐2) (Figure 4.30a). Regardless of the ligand specificity of individual TLRs, ligand‐induced dimerization of adjacent receptors results in a characteristic “m‐shaped” conformation, with the TLRs interacting at their C‐termini to drive dimeriza­ tion of intracellular TIR domains. To look at the structure of TLRs more closely, we will next turn to possibly the best‐char­ acterized of these receptors, TLR4.

The TLR4/MD‐2 complex detects microbial lipopolysaccharide
Lipopolysaccharide (LPS) is an essential component of Gram‐negative bacterial cell walls, capable of inducing potent immune responses at extremely low concentrations, which, if left unchecked, can lead to septic shock and death. Such an acute response suggests that mammalian innate immune systems have evolved to detect this PAMP with exquisite sensitivity and this detection is carried out by TLR4, in conjunction with its co‐receptor MD‐2, both of which are abundantly expressed on the majority of innate immune cells, and on B‐ cells, and barrier tissues at the front line of infection. This double team forms a 1 : 1 heterodimer, with TLR4‐bound MD‐2 acting as the primary binding interface with LPS. Interaction between LPS and MD‐2 opens up MD‐2 residues that promote stable interaction with adjacent TLR4 molecules, promoting dimerization of adjacent TLR4/MD‐2 complexes, with the subsequent dimerization of intracellular TIR domains that triggers signaling.
Native LPS is buried in bacterial cell walls in a difficult‐to‐detect conformation, but is efficiently extracted by a serum factor called LPS‐binding protein (LBP) and facilitated by complement factors that punch holes in bacterial cell walls, dispersing bite‐sized chunks of LPS‐containing material into the bloodstream. LBP transfers LPS oligomers to CD14, which further splits them into monomers, for presentation to the TLR4/MD‐2 complex for efficient detection. Prior to LPS binding, TLR4 and MD‐2 are bound together as heterodimers, with the 21 LRR TLR4 ectodomain arranged in the typical horseshoe shape, and the smaller MD‐2 molecules bound to the lateral side, suspended downwards in a hanging, flower basket‐like arrangement (Figure 4.31a). MD‐2 is the main interactor with LPS and adopts a cup‐like structure, with two antiparallel β‐sheets forming a stable barrel‐shaped core that can accommodate lipid molecules of a defined size. LPS is a glycolipid with a hydrophobic lipid A region attached to a carbohydrate chain and the number of lipid chains in the lipid A segment appears to be a critical determinant of TLR4/MD‐2 complex activation, with six lipid chains forming the ideal number. Indeed, the lipid A region is responsible for the majority of inflammatory activity of LPS, with five lipid chains exhibiting 100‐fold lower activity and four lipid chains, such as eritoan, acting as inhibitors. The crystal structure of the TLR4 ectodomain/MD‐2/LPS complex illustrates the preference for six chains. Five lipid A chains of LPS are buried deeply in the hydrophobic β‐pocket of MD‐2, while the sixth lipid A residue is exposed, with negatively charged phosphate groups making critical contacts with positively charged residues on both MD‐2 and the TLR4 ectodomain. Importantly, these interactions re‐orien­tate MD‐2 such that its F126 and L87 loops become exposed and are now free to make contact with a separate, adjacent, TLR4 molecule, also bound to its own MD‐2, which, in turn, makes a reciprocal interaction. This site of interaction between adjacent LPS and MD‐2 molecules is called the dimerization interface and promotes dimerization of adjacent TLR4/MD‐2 molecules with the resulting heterotetrameric complex of TLR4–MD‐2–LPS, in a 2 : 2 : 2 ratio (Figure 4.31a). The net result of all these interactions results in stable interaction between the C‐termini of two TLR4 ectodomains, forming an m‐shaped structure that facilitates close interaction and subsequent dimerization of intracellular TIR domains (Figure 4.31a,b).
As noted above, TIR domain dimerization is required for the recruitment of the TIR domain‐containing adaptor MyD88, which recruits IRAK4 and IRAK2 in a defined structure that has been dubbed the Myddosome, which relays the inflammatory signal into the cell. We will look more closely at how the structure of the Myddosome is organized to perform this task but first let us take a look at a TLR with binding properties that are different from those of the TLR4/ MD‐2 complex, TLR2.
Overall structure of the TLR4–MD‐2–LPS complex
Figure 4.31 Overall structure of the TLR4–MD‐2–LPS complex. (a) Top view of the symmetrical dimer of the TLR4–MD‐2–LPS complex. The primary interface between TLR4 and MD‐2 is formed before binding LPS, and the dimerization interface is induced by binding LPS. (b) Side view of the complex. The lipid A component of LPS is colored red, and the inner core carbohydrates of LPS are colored pink. The module numbers of the LRRs in TLR4 and the names of the β‐strands in MD‐2 are written in black. TLR4 is divided into N‐, central, and C‐ terminal domains. The LRRNT and LRRCT modules cover the amino and carboxy termini of the LRR modules. (c) Structure of the primary and dimerization interfaces of the TLR4–MD‐2–LPS complex. The lipid chains of LPS are labeled. MD‐2 is colored gray. The lipid chains and phosphate groups of LPS are shown in red. The glucosamine backbone is pink. (Source: Park B.S. and Lee J.O. (2009) Nature 458, 1191–1195. Reproduced with permission of Nature Publishing Group.)

TLR2 plays a crucial role in the recognition of microbial lipo- peptides, and mice deficient in this receptor are at increased risk of infection with a variety of bacteria, including S. pneumoniae and M. tuberculosis. Bacterial lipoproteins are composed of a glycerol backbone with either two or three attached acyl (fatty acid) chains. Gram‐negative bacteria possess tri- acylated lipoproteins with two fatty acid chains, attached by ester bonds to an N‐terminal cysteine, with the third lipid chain connected to the cysteine by an amide bond, whereas lipoproteins from Gram‐positive bacteria and mycoplasma are diacylated as they lack the amide‐bound lipid chain and thus have just two fatty acid chains. Early gene knockout studies showed that macrophages from TLR2‐deficient mice lost the ability to respond to both di‐and triacylated lipoproteins from a variety of bacteria. Interestingly, TLR1‐deficient macrophages lost the ability to respond to triacylated lipoproteins only, whereas macrophages deficient in TLR6 failed to respond to the diacylated form. These results strongly suggeste that TLR2 worked in conjunction with TLR1 to detect triacylated lipoproteins from Gram‐positive bacteria, while it paired up with TLR6 for detection of Gram‐positive bacteria bearing diacylated lipoproteins. Indeed, subsequent crystal structures confirmed this data, showing that triacylated lipoproteins simultaneously bound both TLR1 and TLR2, effectively acting as a bridge to draw the two receptors close enough together for dimerization to occur, while diacylated lipoproteins formed a complex with both TLR2 and TLR6.
Although TLR2 can directly bind both di‐ and triglycerides without the need for intervention from TLR1 or TLR6, this binding does not promote an optimal interaction between individual lipoprotein‐bound TLR2 receptors and thus, the dimerization of adjacent TLR2 ectodomains required for intra­ cellular signaling fails to occur. This is due to the fact that TLR2 efficiently binds the first two lipid chains on a lipoprotein, leaving the rest of the molecule free to undergo specific interactions with TLR1, in the case of the triacylated form, or TLR6 for diaceylated lipoproteins. Indeed it is the specificity of TLR1 for triacylated lipoproteins and TLR6 for diacylated lipoproteins that confers specificity on the TLR2/1 and TLR2/6 complexes.
The ectodomains of all three TLRs display the characteristic TLR horseshoe shape, with 20 LRR modules each containing 24 residues, and can be divided into three distinct subdomains: N‐terminal, central, and C‐terminal (Figure 4.32a). Although the N‐terminal domain shares homology with other LRRs, the central and C‐terminal domains of TLR1 and TLR2 deviate from the norm, with the border between these two domains molded into ligand‐binding pockets, lined with hydrophobic residues. The ligand‐binding pocket on TLR2 is large enough to accommodate the first two fatty acid chains of a triacylated lipoprotein, while the third acyl chain fits into a similar but smaller pocket on TLR1. The bound triacylated ligand now effectively acts as a bridge to pull both TLRs close together, allowing hydrophobic residues that surround the binding pockets on both TLRs to form hydrogen bonds that further stabilize the interaction, pulling both TLRs closer together (Figure 4.32). These ligand–TLR and TLR–TLR interactions result in dimerization of TLR1 and TLR2 at their C‐termini, forming the distinctive “m shape” that facilitates localization of intracellular TIR domains.
Although TLR1/2 complexes efficiently bind triacylated lipoproteins, why are TLR2/6 complexes specific for diacylated ligands? The answer lies in a number of important structural differences between TLR1 and TLR6 in their ligand‐binding and dimerization surfaces. Whereas TLR1 can accommodate an acyl chain in its C‐terminal ligand‐binding pocket, this pocket in TLR6 is partially blocked by the bulky side chains of two phenylalanine residues, reducing the pocket size by half and restricting ligand entry. Indeed, mutation of this region of TLR6 to mimic that found in TLR1 allows TLR6 to efficiently bind triacylated ligands, underlying the importance of these C‐terminal phenylalanine residues in conferring specificity for diacylated lipoproteins. Although TLR6 lacks a ligand‐binding pocket that could accommodate an acyl chain, it makes up for it in a superior ability to bind the peptide part of diacylated lipoproteins. As in the TLR1/2 complex, the two acyl chains of the lipopeptide are buried in the C‐terminal pocket of TLR2, while the exposed peptide region of the ligand forms a number of strong hydrogen bonds with both TLR2 and TLR6 (Figure 4.32). In addition, an extensive region on TLR6 also makes direct contact with TLR2, forming stable hydrogen bonds that account for an increase in protein–protein inter-action of at least 80% when compared with TLR1/2. These interactions combine to drive TLR2 and TLR6 close enough together for dimerization and intracellular signaling to occur.
Although we have focused on the extracellular TLR domain interactions that are brought about by ligand binding, the associated re‐orientation of intracellular domains required to drive signaling is equally as important and it is to this that we will next turn our attention.

Dynamic structural rearrangements propagate intracellular TLR signaling
Regardless of the nature of ligand‐induced dimerization of individual TLR ectodomains, dimerization at the C‐termini re‐orientates the receptors such that the intracellular TIR domains colocalize and undergo the dimerization required to recruit TIR domain‐containing adaptors. Interestingly, extensive artificial truncation of TLR ectodomains triggers receptor auto‐activation, which suggests that in their unbound forms, the ectodomains may act to inhibit an intrinsic tendency for the transmembrane and intracellular domains to dimerize.
There are five TIR domain‐containing adapters that transmit TLR signals into the cell, with MyD88 required at a proximal level for the signaling of all TLRs except TLR3, which uses TRIF exclusively. In the case of TLR4, ligand binding promotes ectodomain dimerization, allowing the TIR domains to dimerize and recruit six molecules of MyD88, in conjunction with the bridging molecule MyD88 adaptor‐like protein (MAL). Close contact between the death domains of MyD88 is then thought to facilitate recruitment of four molecules of the death domain‐containing adaptor IRAK4, which in turn, recruits four molecules of IRAK2, forming a higher order, column‐like structure that has been dubbed the Myddosome, which is responsible for activating NFκB.
TIR domain structure can be subdivided into a central β‐ sheet, organized into four or five parallel β‐strands (the βA–βE strands), with five α‐helices (αA–αE helices), connected to the edges of the sheet by a series of loops. Some of these loops play a critical role in signal transduction, such as the BB loop that joins the βB strand of the β‐sheet with the αB α‐helix. A polymorphism in this region in TLR4 in the CHC3H/HeJ strain of laboratory mice completely kills signaling from the receptor and renders these mice incapable of responding to LPS. Although dimerized TIR domains have proved difficult to crystallize, mutational and inhibitor studies have shed light on the method of TIR domain dimerization, with the BB loop of adjacent TIRs predicted to form an extensive interface. In addition, regions within the BB loop also make direct contact with the TIR domain of MAL, which acts as a bridging molecule to stabilize TLR4–MyD88 interaction.
TLR4 can also signal through the TIR domain‐containing adaptor TRIF, in conjunction with the bridging molecule TRIF‐related adaptor molecule (TRAM), to drive activation of IRF3 and expression of interferon genes. TRAM is recruited to TLR4 only after receptor endocytosis, suggesting that a possible conformational change in the receptor, driven by the acidic environment of the endosome, may be required for TRAM binding and subsequent TRIF recruitment. Interestingly, the TIR domain of TLR3, which signals exclusively through TRIF, contains an alanine in the BB loop, rather than a proline like all the other TLRs, and mutation of this residue in TLR3 to proline changes specificity of TLR3 from TRIF to MAL/ MyD88, with associated NFκB signaling as opposed to IRF‐ dependent events.
Overall structure of the human TLR1–TLR2–Pam3CSK4 complex and the mouse TLR2–TLR6–Pam2CSK4 complex.
Figure 4.32 Overall structure of the human TLR1–TLR2–Pam3CSK4 complex and the mouse TLR2–TLR6–Pam2CSK4 complex. To facilitate crystallization and structure determination the LRR C‐terminal and the last one or two LRRs of TLRs 1, 2, and 6 were replaced by corresponding regions of a hagfish VLR. The TLR1, TLR2, TLR6, and VLR fragments in the TLR–VLR hybrids are shown schematically in green (TLR1 and TLR6), blue (TLR2), and gray (VLR). Pam3CSK4 and Pam2CSK4 are shown in red. Some LRR modules are numbered and the N‐terminal, central, and C‐terminal subdomains are labeled. (a) Side view, (b) top view. (Source: Jin M.S. et al. (2007) Cell 130, 1071–1082 and Kang J.Y. et al. (2009) Immunity 31, 873–884. Reproduced with permission of Nature Publishing Group.)
MyD88 and TRIF form higher order complexes
In addition to a TIR domain, MyD88 also contains a death domain (DD), common in proteins associated with apoptosis as well as immunity. The MyD88 DD provides a platform for recruitment of the DD‐containing IRAK4, which in turn recruits IRAK2 via DD interactions. Death domains bestow on these proteins the ability to form hetero‐oligomers and the crystal structure of the MyD88–IRAK4–IRAK2 complex has illuminated the impressively ordered nature of this signaling platform (Figure 4.33). Six–eight molecules of MyD88 recruit four molecules of IRAK4, which in turn recruit four molecules of IRAK2 in a helical, three‐layered complex called the Myddosome, driven by DD–DD interactions. The importance of this com­ plex for TLR signaling is illustrated by a naturally occurring polymorphism in the DD of MyD88 that renders these complexes defective for both signaling and Myddosome formation.
In contrast to MyD88, the larger TRIF molecule lacks a DD, instead containing an α‐helical N‐terminal domain (TRIF‐NTD) that is thought to autoinhibit activation of the resting TRIF protein by obscuring the binding sites of down­stream adaptors. Binding of TRIF to TLR3 or TLR4/TRAM displaces the TRIF‐NTD and frees up a proline‐rich region in the protein, which facilities recruitment of tumor necrosis fac­ tor receptor‐associated factor 2 (TRAF3) and TANK‐binding kinase 1 (TBK1) for activation of IRFs. In addition, the TRIF receptor‐interacting protein (RIP) homotypic interaction motif (RHIM) is also liberated to recruit RIP kinase 1, resulting in both FADD‐dependent apoptosis and NFκB activation. Crystal structures of the TRIF complexes have not yet been resolved to answer the question of whether or not they form higher order complexes like the Myddosome, but the current thinking is that a similar TRIF‐containing complex may be formed.
Myddosome structure.
Figure 4.33 Myddosome structure. (a) Ribbon diagram of Myddosome structure, with the six MyD88 molecules in cold colors, the four IRAK4 molecules in earth‐tone colors, and the four IRAK2 molecules in warm colors. (b) Surface diagram of the complex with each subunit labeled using the same color coding as in (a). M, MyD88; I4, IRAK4; I2, IRAK2. (Source: Lin S.C. et al. (2010) Nature 465, 885–890. Reproduced with permission of Nature Publishing Group.)
C‐type lectin‐like receptors detect fungal antigen
C‐type lectin‐like receptors (CLRs) form a large and varied family of receptors that share in common a C‐type lectin‐like domain (CTLD) and function in a variety of scenarios, from cell–cell adhesion to immune signaling and apoptosis. Although the CTLD bears structural homology to the carbohydrate‐ binding domains found in carbohydrate‐binding proteins, CTLDs are more varied and are not necessarily restricted to carbohydrate ligands. This family of receptors can be loosely subdivided by their requirement for calcium for functional ligand binding and on the type of intracellular signaling domain that can possess activating ITAMs or inactivating ITIMs. Ligand recognition and signal transduction by activating CLRs is broadly similar to the TLR scenario; ligand binding promotes receptor ectodomain dimerization, which then dimerizes and activates the intracellular ITAM motifs to recruit
ITAM‐containing adapter molecules such as Syk kinase, to promote activation of proinflammatory transcription factors such as NFκB.
Although many members of the CTLD family bind a variety of carbohydrates from a number of different microorganisms (e.g., dectin‐1 binds β‐glucan and dectin‐2 recognizes mannose) other members, such as the lipid‐binding mincle, can also bind non carbohydrate ligands. The fungal β‐glucan‐ binding receptor dectin‐1 is the best‐characterized CTLD receptor and we will now look more closely at its mode of action.

Dectin‐1 recognizes fungal β‐glucan
Immune responses to fungal infections are mediated mainly by CTLD receptors, with the detection of β‐glucans by dectin‐1 playing a particularly important role in antifungal immunity. Mice deficient in this receptor display marked defects in immune cell infiltration during fungal challenge and are highly susceptible to infection with Candida albicans, while dectin‐1 also detects β‐glucans from a range of other fungi, including Saccharomyces, Penicillium, and Aspergillus. As highly conserved and essential components of the cell wall of certain fungi and baker’s yeast, β‐glucans certainly fit the bill as classical PAMPs. Dectin‐1 can recognize β‐1,3 and β‐1,6‐linked glucans from fungi, plants, and bacteria, with the best‐characterized ligand, zymosan from yeast cell walls, binding with high affinity. The expression of dectin‐1 on dendritic cells, monocytes, macrophages, and neutrophils places it on the front line of antifungal immunity, where receptor activation can trigger pathogen phagocytosis or the generation of antifungal cytokines and chemokines.
With a single extracellular CTLD, a transmembrane region and a cytoplasmic ITAM, ligand binding is thought to promote dimerization of the dectin‐1 ectodomain, required to activate intracellular ITAMs. Unlike other members of the CTLD receptor family, ligand binding occurs in the absence of calcium. Crystal structure of the extracellular portion of dectin‐1 illustrates that it adopts a similar conformation to other CTLD‐containing receptors, with two antiparallel β‐sheets and two α‐helices, with the N‐ and C‐termini in close proximity (Figure 4.34). Sequence analysis has highlighted a number of surface hydrophobic residues that could play a role in ligand binding, and mutational studies have identified two residues, Trp221 and His223 in the third ticularly important for ligand recognition. Mutation of these residues to an alanine blocked the interaction of β‐glucan with the receptor, while a dectin‐1 antibody that efficiently inhibited β‐glucan binding failed to bind to the W221A mutant, suggesting the region plays a key role in ligand interaction. This region adopts a shallow hydrophobic groove in the crystal structure of dectin‐1, but no ligands were observed binding in this pocket, possibly due to technical constraints in achieving crystallization of β‐glucan ligands of sufficient size. Indeed, cell‐based studies have suggested that the minimum size of β‐glucan sufficient to bind the receptor is no smaller than 10‐mer, which could certainly be accommodated in this groove. Although the current crystal structure is inconclusive, it remains likely that β‐glucan binding acts to bridge adjacent dectin‐1 molecules to facilitate ITAM dimerization and recruitment of Syk kinase and, potentially, Raf, which can both drive immune signaling through NFκB activation. Activated Syk also drives calcium‐dependent outcomes such as NFAT activation, with associated cytokine secretion.
Two dectin‐1 monomers form a dimer into which a short β‐glucan binds.
Figure 4.34  Two dectin‐1 monomers form a dimer into which a short β‐glucan binds. A cartoon diagram of the dectin‐1 dimer, with each monomer colored from blue at the N‐terminus to red at the C‐terminus. (Source: Brown J. et al. (2007) Protein Science 16, 2. Reproduced with permission of Wiley.)

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