Membrane association: Working at the membrane–water interface

Membrane-associated GT-B enzymes represent a specialized group of GTs with the remarkable ability to recognize both the hydrophilic water-soluble nucleotide mono- or diphospho-sugar donor and acceptor substrates mostly in the form of hydrophobic lipids and membrane-associated proteins (Figure ​(Figure3;3; Forneris and Mattevi 2008). What are the structural requirements for a GT-B enzyme to be associated to the lipid bilayer? What is the impact of the membrane composition and structure on enzymatic activity? How is the active site of a membrane-associated GT-B GT made accessible to hydrophobic and hydrophilic substrates? Here, we focus on the two best-studied families of membrane-associated GT-B GTs: The monotopic and peripheral enzymes.

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Fig. 3.

Membrane-associated GT-B GTs. (A) A GT-B peripheral membrane GT as illustrated by the phosphatidyl-myo-inositol mannosyltransferase PimA (PM, GT4; Guerin et al. 2007; Guerin et al. 2009). (B) A GT-B monotopic GT as illustrated by the UDP-N-acetylglucosamine–N-acetylmuramyl-(pentapeptide) pyrophosphoryl-undecaprenol N-acetylglucosamine transferase MurG (MT, GT28; Hu et al. 2003) (C) A GT-B bitopic GT as illustrated by the multifunctional sialyltransferase PmST1 from Pasteurella multocida (Ni et al. 2006; Ni et al. 2007). (D) The GT-B dimeric monotopic α-2,3-sialyltransferase NST from Neisseria meningitidis (Gilbert et al. 1997; Lin et al. 2011).

Monotopic GT-B GTs

The 1,2-diacylglycerol 3-glucosyltransferase from Acholeplasma laidlawii (AlMGS) represents a good model reflecting our current knowledge of the molecular mechanism of membrane association for the monotopic GT-B enzymes. Acholeplasma laidlawii is a cell wall-less prokaryote that controls the surface charge density and curvature properties of its membrane through the action of two cytosolic side membrane-associated glucosyltransferases (Dahlqvist et al. 1995; Andersson et al. 1996; Lindblom et al. 1986; Lindblom et al. 1993; Andrés et al. 2012). Specifically, the AlMGS synthase and the diglucosyl-diacylglycerol synthase (AlDGS) synthesize, respectively, the two major nonbilayer-prone and bilayer-forming glucolipids, GlcDAG and GlcGlcDAG, by consecutively adding two glucose residues to a diacylglycerol acceptor backbone at the membrane interface, respectively (Karlsson et al. 1997; Vikstrom et al. 1999). The molar ratios of these glucolipids and acylated counterparts vary markedly in response to a variety of stimuli, including acyl chain types and ionic environment (Wieslander et al. 1986). AlMGS lack a hydrophobic trans-membrane segment, but detergents are needed for its solubilization (Li et al. 2003). Interestingly, the N-terminal domain of AlMGS displays a calculated pI value of 9.7 and hence is positively charged at physiological pH. In contrast, the C-terminal domain of the enzyme is dominated by the presence of acidic residues (Berg et al. 2001; Edman et al. 2003). A close inspection of its amino acid sequence revealed that the N-terminal domain of the enzyme contains a mixture of hydrophobic and basic residues, mainly in the form of arginine and lysine, in an amphiphatic α-helix (Lind et al. 2007). The amphipathic character of this α-helix was structurally verified by a combination of solution nuclear magnetic resonance (NMR) and circular dichroism, and proved to interact with both zwitterionic and anionic phospholipids by surface plasmon resonance (SPR; Lind et al. 2007). Very recently, a preferential binding of AlMGS to anionic lipids including phosphatidylglycerol (PG) and cardiolipin (CL) was observed in vivo (Ariöz et al. 2013).

Membrane lipid surfaces are negatively charged essentially due to the presence of anionic lipids (McLaughlin 1989; McLaughlin and Murray 2005). The binding of AlMGS to lipid bilayers containing anionic lipids (e.g., phosphatidylglycerol) was measured by SPR and found to be essentially irreversible with an overall dissociation constant (K_D) of 10⁻¹⁰–10⁻¹² M (Vikstrom et al. 1999; Li et al. 2003). Moreover, the thickness of vesicular membrane fractions (lipid bilayer + interfacial water phase) containing AlMGS was determined to be ~4.30 nm by small angle X-ray diffraction (Eriksson et al. 2009; Ge 2011). Since the thickness of the inner membrane of Escherichia coli is about 3.75 nm and the size of AlMGS is roughly 4 × 5 nm, these observations are suggestive of the deep insertion of AlMGS in the membrane, as also indicated by molecular dynamic simulation of other monotopic membrane proteins (Mitra et al. 2004; Balali-Mood et al. 2009). Thus, experimental data clearly point to a permanent association to the bilayer surface, without any apparent shuttling of AlMGS between the cytoplasm and the membrane, typically observed for peripheral membrane-associated proteins (Li et al. 2003; Lind et al. 2007). Interestingly, anionic phospholipids seem to affect the binding and also the enzymatic activity of AlMGS. None of the uncharged GlcGlcDAG, the neutral-charged 1,2-dioleoyl-phosphatidylcholine and positively charged sphingosine lipids could replace 1,2-dioleoyl-phosphatidylglycerol (DOPG) as an activator for the purified AlMGS (Karlsson et al. 1997). In contrast, the anionic phospholipid 1,2-dioleoyl-phosphatidylserine could fully substitute for DOPG, indicating that the anionic-charged groups of the activator lipids hold the activating property (Karlsson et al. 1994; Dahlqvist et al. 1995).

In plants, photosynthetic membranes are mainly constituted by two galactolipids, the nonbilayer-prone GalDAG and the bilayer-prone GalGalDAG (Dormann and Benning 2002; Kelly and Dörmann 2004; Guskov et al. 2009; Kobayashi et al. 2013; Ge 2011). The relative amounts of these two glycolipids affect the membrane curvature as well as the lateral stress profiles of the membrane, thus likely influencing the functions of other membrane-associated proteins or protein complexes (Moellering and Benning 2011). Arabidopsis thaliana synthesizes GalDAG through the action of a monogalactosyldiacylglycerol synthase, in the form of three GT-B isoforms namely AtMGD1, AtMGD2 and AtMGD3 (Benning and Ohta 2005). The three isoforms transfer a galactose residue from UDP-Gal to the 3-position of sn-1,2-diacylglycerol (DAG) and differ in their substrate specificity and subcellular localization. Two GT-B enzymes were found to catalyze the biosynthesis of GalGalDAG, AtDGD1 and AtDGD2 (Ge 2011). AtDGD2, a monotopic enzyme displaying similar interface-binding structural elements as AlMGS, is apparently able to modulate the lipid environment by adjusting the biosynthetic activity of its product GalGalDAG (Ge et al. 2011). Nonbilayer-prone lipids such as GlcDAG, rac-1,2-dioleoylgly-cerol and dioleoyl-sn-glycero-3-phosphoethanolamine decreased the enzymatic activity of AtDGD2, suggesting that increasing curvature stress with these additives has a negative effect on GalGalDAG synthesis. In contrast, the anionic lipids PG, phosphatidic acid and phosphatidylserine displayed strong stimulatory effects on AtDGD2 activity; interestingly, CL did not even though it is structurally related to PG, suggesting the existence of PG-specific-binding site(s) in the protein. Three segments of AtDGD2 containing conserved tryptophan residues were demonstrated to be crucial not only for enzymatic activity but also for membrane interaction (Ge et al. 2011; Szpryngiel et al. 2011). It is worth noting that the enrichment of tryptophan residues in the lipid bilayer interface regions is well established for many membrane-associated proteins (Kozlov 2010). Interestingly, two of the tryptophan-containing segments (IV and V) of AtDGD2 are localized in the N-terminal domain, whereas another is in the C-terminal domain (VI), with all segments likely interacting with stimulatory anionic phospholipids and determining the orientation of AtDGD2 at the membrane interface. Altogether the experimental data allowed Wieslander et al. to propose a regulatory model for the anchoring of AtDGD2 at the membrane interface (Figure ​(Figure4).4). According to this model, the N-terminal segments IV and V may serve as permanent anchor points for the N-terminal domain at the lipid interface while segment V on the C-terminal domain interacts with the membrane interface in a more flexible way. The positively charged protein clusters located at the lipid interface are thought to play a determinant role in regulating the interaction of the whole enzyme with the membrane, leading to the different catalytic efficiencies (Ge 2011; Ge et al. 2011).

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Fig. 4.

A model for membrane association of monotopic GT-B GTs. Wieslander and colleagues proposed two modes of reorientation for AtDGD2 upon stimulation by anionic lipids: An up-down displacement (“dipping”) of the C-terminal domain (A) and a “rolling” transfer of the catalytic region into the interface (B). Both models contemplate the access of the soluble UDP-Gal donor substrate to the active site.

The relevance between electrostatic forces, both coulombic attraction/repulsion and dipolar interactions, hydrogen bond formation and hydrophobic interactions, as driving forces for membrane association of peptide/proteins has been well established (McLaughlin 1989; McLaughlin and Murray 2005). Peptide-membrane association seems to be mediated by different thermodynamic steps (Figure ​(Figure5;5; Jacobs and White 1989; White and Wimley 1999). In the first step, peptide binding is initiated by the electrostatic attraction of the positively charged residues to the anionic membrane. Depending on the peptide charge, and the strength of the membrane surface potential, the electrostatic attraction would increase significantly the protein concentration near the membrane surface, compared with the soluble fraction (Leventis and Silvius 1998; Seelig 2004). The next step implies the transition of the peptide likely into the plane of binding. The exact location of this layer is difficult to define and depends on the hydrophobic/hydrophilic balance of the molecular groups and forces involved. The third step in the binding process is a change of the conformation of the bound peptide (Figure ​(Figure5).5). In many cases, peptides are in a random coil conformation in solution but adopt an α-helical conformation when associated to the lipid membrane (e.g., illustrated with melittin and alpha-synuclein, the latest involved in Parkinson disease; Klocek et al. 2009; Lokappa and Ulmer 2011). These conformational transitions also include the random coil-to-β-structure transition (e.g., Alzheimer peptides; Meier and Seelig 2007) and secondary structure reshuffling as recently described for the pore-forming toxin pneumolysin (Tilley et al. 2005). The penetration of the peptide/protein depends on the chemical nature of the lipids, peptides and carbohydrates involved and also on the mechanistic nature of the processes investigated, in which both location and timing of membrane association can be tightly controlled.

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Fig. 5.

Molecular recognition of a peptide at the membrane surface. Schematic representation of different steps in the process of peptide binding to an anionic phospholipid bilayer. The positively charged peptide is attracted electrostatically to the membrane surface followed by a conformational change to an α-helix (White and Wimley 1999; Seelig 2004).

The crystal structures of TagF (GT2; CDP-Gly, Weidenmaier and Peschel 2008; Lovering et al. 2010), WaaG (GT4; Raetz and Whitfield 2002; Martinez-Fleites et al. 2006), WsaF (GT4; Steiner et al. 2010), WaaC (GT9; Grizot et al. 2006), WaaF (GT-9; pdb code 1PSW), Vpar_0760 (pdb code 3TOV), MurG (GT28; Hu et al. 2003), WaaA (GT30; Schmidt et al. 2012) and GumK (GT70; Barreras et al. 2008), all of which have been proposed to behave similar to the monotopic membrane-associated family of GT-B GTs, clearly display common structural features with AlMGS and AtDGD2. The N-terminal domains are enriched in positively charged residues, mostly in the form of amphiphilic helices, yielding high calculated pI values (Table ​(TableII;II; Figure ​Figure6).6). Surface-exposed aromatic hydrophobic residues as tryptophan and phenylalanine are also observed on the N-terminal domain of the crystal structures of TagF, WaaG, WsaF, WbaZ, WaaC, WaaF, MurG and GumK. Interestingly, in TagF, preceding the GT-B region, there are two α-helices that are strong candidates for membrane association and protein oligomerization. The first α-helix (α1; residues 316–331) is highly hydrophobic while the second α-helix (α2; residues 335–344) forms an extremely positively charged environment with neighboring residues (Lovering et al. 2010). In contrast, the Helicobacter pylori α1,3 fucosyltransferase (FucT) is composed of an N-terminal catalytic domain, 2–10 heptad repeats likely involved in protein dimerization and a C-terminal tail which is rich in both basic and hydrophobic residues and would mediate protein–membrane interactions (Sun et al. 2007). These structural features are believed to be involved in anchoring the enzymes to specific lipid environments and to also influence their activities. Moreover, multivariate and bioinformatic sequence analyses identified the occurrence of equivalent structural elements in the N-terminal domain of many structurally uncharacterized membrane-associated GT-B enzymes suggestive of conserved molecular mechanisms of membrane association (Table ​(TableII;II; Lind et al. 2007). Major contributions to membrane interaction free energies arise from the desolvation of protein and membrane as they associate and from the electrostatic attraction between protein's basic residues and membranous acid phospholipids (Figure ​(Figure7;7; Wimley and White 1996; White and Wimley 1999).

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Fig. 6.

Electrostatic surface potential representation of membrane-associated GT-B GTs, as visualized in GumK, MurG, WaaA, WaaC and PimA.

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Fig. 7.

Calculation of the desolvation energy values derived from octanol/water experiments. The hotspots indicate areas with the most favorable energy change upon binding, likely to be buried in the membrane. The largest hotspots correspond to energy values lower than −15.0 kcal mol⁻¹, medium-sized spots represent values between −15 and −10 kcal mol⁻¹ and the smallest points represent values higher than −10 kcal mol⁻¹.

Peripheral GT-B GTs

The membrane association and dissociation rate constants of many GT-B GTs remain largely unknown, making their unambiguous classification as monotopic or peripheral membrane-associated enzymes difficult. Consequently, the determination of the protein sub-cellular localization became, in most of the cases, the sole criteria to discriminate between both groups of enzymes. The phosphatidylinositol mannosyltransferase A, which has been shown to co-localize both in the membrane and in cytosolic fractions, represents a clear example of a peripheral membrane-associated GT-B GT (PimA; Korduláková et al. 2002; Guerin et al. 2009). PimA is an essential enzyme for mycobacterial growth that initiates the biosynthetic pathway of key structural elements and virulence factors of Mycobacterium tuberculosis, the phosphatidyl-myo-inositol mannosides (PIM), lipomannan and lipoarabinomannan (Kaur et al. 2009; Guerin et al. 2010; Morita et al. 2011). The enzyme catalyzes the transfer of a mannose residue from GDP-Man to the 2-position of inositol phosphate ring in phosphatidyl-myo-inositol (PI) to form phosphatidyl-myo-inositol monomannoside (PIM₁) on the cytoplasmic side of the plasma membrane (Guerin et al. 2009). The crystal structure of PimA in complex with its donor sugar substrate pointed to a region located in the N-terminal domain and adjacent to the two-domain cleft, as potentially mediating protein–membrane interactions. Specifically, the interfacial-binding surface contains a solvent-exposed hydrophobic patch close to a cluster of basic residues in an amphiphatic α-helix and to the connecting loop β3-α2, which is disordered in the lipid-free structure (Guerin et al. 2007). Supporting this notion, a PimA mutant in which residues Arg77, Lys78, Lys80 and Lys81 on α-helix 2 were substituted by serine completely inactivated the enzyme and drastically impaired the ability of the protein to bind PI (Guerin et al. 2007; Guerin et al. 2009). It is worth noting that the positively charged cluster is far from the catalytic center and exposed to solvent, and therefore, it is not expected to interfere with the catalytic machinery. Moreover, the involvement of this cluster in protein–membrane interactions is consistent with the observed activity enhancement of PimA in the presence of nonsubstrate anionic phospholipids including 1,2-dipalmitoyl-sn-glycero-3-phosphate or cardiolipin (Guerin et al. 2007). It may also explain previous results showing that a salt wash of mycobacterial plasma membranes significantly reduced the biosynthesis of PIM₁ (Morita et al. 2005).

However, peripheral GT-B GTs seem to have acquired different strategies to associate to the membrane, according to the specific functions they perform inside the cell (Figure ​(Figure8;8; Lemmon 2008; Moravcevic et al. 2012). In this regard, it is interesting to mention the case of Alg13/Alg14, an heterodimeric enzyme involved in the biosynthesis of N-glycans in Saccharomyces cerevisiae (Gao et al. 2005). Alg13/Alg14 mediates the second step of the pathway by transferring a GlcNAc residue from UDP-GlcNAc donor to the Dol-PP-GlcNAc acceptor. Alg14 is a bitopic membrane-associated protein, with a cytosolic-predicted Rossmann-like fold domain. According to the NMR solution structure, Alg13 adopts a nonconventional Rossmann-fold. Titration of Alg13 with UDP-GlcNAc demonstrated that Alg13 interacts with the sugar donor through residues located on the C-terminal half of the protein. Alg14 recruits Alg13 to the cytoplasmic face of the endoplasmic reticulum to form the bipartite enzyme, where the C-terminal region seems to modulate Alg13/Alg14 protein–protein interactions (Gao et al. 2007). Another good example is the plant sucrose synthase (SUS). SUS participates in the modulation of sucrose flux by rapidly altering its cellular location from the cytosol to other sites including with various organelle membranes (Subbaiah et al. 2006). However, the molecular mechanisms by which SUS binds to its cellular targets, the structural aspects controlling SUS partitioning and the structural impact of partitioning on its catalytic function are currently unknown. In Zea mays, the phosphorylation of the SUS1 isoform at the Ser-15 residue increases membrane association and catalytic activity at acidic pH in Zea mays (Hardin et al. 2004). Interestingly, phosphorylation of SUS1 modifies the oligomerization state of SUS1, from dimers to tetramers. The recently solved crystal structure of SUS1 from A. thaliana suggests a model that explains how SUS catalysis and the interaction with its cellular targets might be regulated (Zheng et al. 2011). A conserved sequence surrounding the equivalent Ser-13 is positively charged (RXHSX(R/K)ER), whereas the neighboring helix α8 has both anionic and cationic patches (LKRAEEYL) most probably altering the electrostatic environment in this region.

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Fig. 8.

Peripheral GT-B GTs adopt different strategies for membrane association. (A) The mycobacterial mannosyltransferase PimA interacts with the phospholipid bilayer by a combination of positively charged and hydrophobic residues exposed on the N-terminal domain of the protein. An important conformational change is expected to occur during/after protein–membrane interaction (Guerin et al. 2007; Guerin et al. 2009; Giganti et al. 2013). (B) A model for the Alg7/13/14 complex formation. This functional multienzyme complex catalyzes the first two steps of lipid-linked oligosaccharide on the cytosolic face of the ER membrane. Alg7, a politopic protein, which transfers a GlcNAc-phosphate to dolichol phosphate, interacts with the transmembrane α-helix of the bitopic GT-GT Alg14 (Lu et al. 2012). Biochemical and structural data support a model in which Alg13-Ag14 interaction is mediated by residues located on the C-terminal α-helix of Alg13 and C-terminal amino acids of Alg14 to form a dimeric Alg13/14 (Wang et al. 2008).

Nucleotide sugar donor-binding site

The nucleotide mono- or diphospho-sugar-binding sites essentially comprise a hydrophobic pocket and a conserved α/β/α motif located on the more conserved C-terminal domain of the enzymes, and a region constituted by the β₁-α₁ loop and the top of the α₁ on the N-terminal domain, with all regions facing the interdomain cleft (Hu and Walker 2002; Figure ​Figure9A9A and B, Table ​TableI).I). The nucleobase heterocycle accommodates into the hydrophobic pocket where it makes electrostatic interactions and hydrogen bonds with the GT. These noncovalent interactions account for the nucleobase selectivity of this family of enzymes (Figure ​(Figure9C).9C). The first α-helix (α_F) of the α/β/α motif (α_F/β/α_S) mainly interacts with the O2 and O3 hydroxyl groups of the ribose ring, while the second α-helix (α_S) and the β-α₂ connecting loop makes key contacts with the sugar ring. It is worth noting that α-helices possess a permanent dipole moment with a positively charged N-terminus and a negatively charged C-terminus. As a consequence, it has been observed that α-helices favor the interaction with ionic species in which the first and second turn seem to be critical for the electrostatic interaction to occur (Aqvist et al. 1991). In that sense, it is believed that in GT-B GTs the negatively charged pyrophosphate group is stabilized by α₁ (N-terminal domain) and α_S (C-terminal domain) through helix dipole effect (Figure ​(Figure9C;9C; Hol et al. 1978; Hu and Walker 2002). In many cases, the α- and β-phosphates are further stabilized by positively charged side chain residues (Arg and/or Lys; Figure ​Figure9C;9C; Lairson et al. 2008) and/or the β₁-α₁ loop located in the N-terminal domain. This loop is known to undergo significant conformational changes upon sugar-nucleotide binding in many GT-B GTs (Buschiazzo et al. 2004; Vetting et al. 2008; Sheng et al. 2009).

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Fig. 9.

The nucleotide sugar donor-binding site. (A) Overall structure of the phosphatidylinositol mannosyltransferase PimA (PM, GT4) from M. smegmatis showing the β₁-α₁ motif (residues Met1 to Ala31) in orange and the α/β/α motif (α_F/β/α_S; residues Asp252 to Gly287) in yellow (Guerin et al. 2007). (B) Surface representation of the active site of PimA showing the hydrophobic binding pocket and Asp253 that account for nucleoside heterocycle specificity (Guerin et al. 2007). (C) The negatively charged pyrophosphate group is stabilized by α1 (Gly15 to Ala31 at the N-terminal domain) and αS (Gly277 to Gly287 at the C-terminal domain) through helix dipole effect (Aqvist et al. 1991; Hol et al. 1978; Hu and Walker 2002). The α- and β-phosphates are further stabilized by two positively charged residues, Arg196 and Lys202, and the α₁-β₁ loop.

Sugar acceptor-binding site

To date, no direct structural information is available for any integral membrane-associated GT-B GT, including monotopic, bitopic and polytopic proteins, in the presence of their acceptor substrates. However, the acceptor-binding site is expected to be located in the N-terminal domain as occur with the soluble enzymes (Lairson et al. 2008). Supporting this notion, the crystal structure of WaaG revealed a deep and open cavity in which the sugar moiety of the donor is found to cover the deepest point of its floor. This pocket encloses an area of ~940 Å, able to accommodate the first two sugar moieties of the acceptor LPS and eventually the glycosyl ramification on the first of these residues. The walls of the cavity are lined by four tyrosine residues and with various lysine and arginine residues pointing directly to the cavity, in line with the generic signature of carbohydrate-binding sites (Martinez-Fleites et al. 2006). Similarly, the N-terminal domain of WaaA exhibits a depression that extends from the putative membrane-association site to the central groove between the two domains. A PEG molecule has been detected in the |Fo| − |Fc| difference density map, strongly suggesting that this particular region may serve as the acceptor-substrate-binding site for the lipid A precursor (Schmidt et al. 2012). A close inspection of the monotopic AviGT4, WaaA, WaaG, WsaF, WbaZ, WaaC, WaaF and MurG, and the bitopic Fut8 crystal structures also revealed the presence of a cavity, of variable shape and volume and located nearby the sugar donor-binding site, suitable for binding the acceptor molecules. However, to improve the efficient glycosylation of membrane-bound acceptors such as lipids/glycolipids, a close proximity between the lipid bilayer and the acceptor-binding sites is likely to be required. Structural comparison of the N-terminal domains of the monotopic GT-B structures currently available showed that the amphipathic α-helices display very different orientations, with all of them being exposed to the solvent. Thus, important conformational changes are expected to occur in the proteins after membrane/acceptor substrates association.

The crystal structure of SUS1, a peripheral membrane-associated GTB GT, has been recently solved in the presence of UDP and its sugar acceptor fructose at 2.85 Å resolution (Zheng et al. 2011). The enzyme transfers a glucose residue to the 2-position of fructose to form sucrose in plants. The fructose is firmly bound in the β-furanose form within a pocket formed exclusively by residues from the N-terminal domain of the protein. Along the same line, the molecular architecture of the lipid acceptor-binding site of PimA has been proposed to occur on the N-terminal domain by using in silico molecular docking approaches (Guerin et al. 2009). The inositol phosphate group accommodates into a well-defined pocket with its O₂ atom favorably positioned to receive the mannosyl residue from GDP-Man. A similar scenario has been proposed for PimB’, the second enzyme in the PIM biosynthetic pathway (Batt et al. 2010). However, the interplay between the membrane and acceptor-binding site in peripheral membrane-associated GT-B GTs seems to be diverse at the molecular level.

Conformational changes

To achieve the enzyme-transition state complex, a spatial rearrangement of the active site is very often required, highlighting the importance of protein dynamics and conformational changes in substrate recognition (Hammes-Schiffer and Benkovic 2006; Schramm 2011). Specifically, protein conformational changes not only involve local reorganization of flexible loops and side-chain residues, but also, in many cases, domain motions and protein oligomerization events. Moreover, as a consequence of protein–protein interactions, post-translational modifications and noncovalent associations with small-molecule inhibitors or activators, conformational dynamics critically modulates enzyme catalysis.

The “open”-to-“closed” motion

Structural, biochemical and biophysical studies have demonstrated that members of the GT-B superfamily, including soluble and membrane-associated enzymes, can adopt both open and closed forms (Buschiazzo et al. 2004; Guerin et al. 2009; Giganti et al. 2013). It has been proposed that nucleotide sugar donor binding triggers a closure movement that involves the rotation of the N- and C-terminal domains, which brings together critical residues making up a functionally competent catalytic center. This conformational flexibility can be dramatic, as is illustrated by the structures of the ligand-free and UDP-GlcNAc bound complex of MshA from Corynebacterium glutamicum in which the relative orientation of the N- and C-terminal domains involves a rotational movement of 97° (Vetting et al. 2008). Other very good examples are given by the glycogen synthase (Buschiazzo et al. 2004; Sheng et al. 2009) and the monotopic MurG (Hu et al. 2003; Figure ​Figure10A).10A).

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Fig. 10.

Conformational changes on GT-B GTs. (A) Structural comparison between the “open” (molecular surface representation) and “closed” (schematic cartoon) states of glycogen synthase (Sheng et al. 2009) and (C) MshA from C. glutamicum (Vetting et al. 2008). (B, D) Average low-resolution structure of PimA-GDP complex with the high-resolution crystal structure of PimA-GDP complex fitted by rigid body docking (Giganti et al. 2013).

However, perhaps the most well-documented studies about the conformational changes a GT-B enzyme undergoes upon substrate binding corresponds to the peripheral membrane-associated PimA from mycobacteria. The binding of the sugar nucleotide donor GDP-Man induces an interdomain rearrangement from an “open” to a “closed” state that stabilizes the enzyme. The interaction of PimA with the β-phosphate of GDP-Man was essential for this conformational change to occur (Guerin et al. 2009). Several lines of experimental evidence strongly support this notion. First, the crystal structures of the PimA-GDP and PimA-GDP-Man complexes revealed that GDP-Man is buried at the N- and C-terminal domains interface, with residues Gly16, Arg196 and Lys202 stacking the β-phosphate (Guerin et al. 2007). It is worth mentioning that the α-PO4 of the donor substrate does not interact with any particular residue from the enzyme. Secondly, PimA-limited proteolysis studies confirmed that the enzyme was rapidly degraded after incubation with elastase. N-terminal sequencing of the two predominant species of 23 and 15 kDa revealed two exposed sites located in α9, involved in GDP-Man recognition, and the connecting loop β7-β8 at the junction between N- and C-terminal domains. GDP or GDP-Man, but not guanosine where the α-PO4 and β-PO4 are missing, protected PimA from the action of the protease even after 90 min of incubation. The “open-to-closed” motion was further monitored in the absence and presence of GDP and guanosine by sedimentation velocity analytical ultracentrifugation studies of pure PimA (Guerin et al. 2009). PimA sedimented as a single homogeneous species with an average sedimentation coefficient of 3.22 s, which is consistent with an asymmetric monomeric protein. Upon addition of equimolar GDP, the sedimentation coefficient increased to 3.53 s, indicating the formation of a more symmetrical and compact structure. As expected, the addition of guanosine did not significantly affect the sedimentation coefficient value of the PimA. Moreover, ITC measurements revealed that guanosine binds to PimA with a binding constant ~10³-fold smaller than that of GDP. The GDP binding also produces a stabilizing effect on PimA characterized by an increment of ~3.5°C in the melting temperature value as observed by differential scanning calorimetry and circular dicroism (Guerin et al. 2009). More recently, the ab initio low-resolution envelopes obtained from small-angle X-ray scattering of the unliganded PimA and the PimA-GDP complexed forms clearly demonstrate that the “open” and “closed” conformations of the GT-B enzyme are largely present in solution (Giganti et al. 2013; Figure ​Figure10B).10B). Specifically, the radii of gyration (R_g) obtained for PimA apo and the PimA-GDP complex revealed a clear reduction in R_g (ΔR_g) of −1.0 (1) Å. In summary, the sugar nucleotide donor induces the closure movement of PimA, with a clear increment of the sedimentation coefficient value which is correlated with a reduction of the radii of gyration, and accompanied by a markedly stabilization of the enzyme.

In contrast, the binding of the acceptor substrate PI to the enzyme had an effect opposite to that observed for the sugar nucleotide donor. When incubated with PI, PimA became highly sensitive to elastase, even in the presence of GDP, indicating that PI triggers a yet significant conformational change that modifies the closed GDP-Man-induced conformation (Guerin et al. 2009). Furthermore, analytical ultracentrifugation experiments demonstrated that the addition of PI to the enzyme resulted in a significant change in the sedimentation coefficient values of both unliganded PimA and PimA-GDP complex consistent with the formation of less compact structures, which correlates with a reduction of the melting temperature by 1.5 and 0.4°C, respectively, in agreement with the limited proteolysis experiments. Interestingly, single-molecule force spectroscopy indicated that PimA unfolds following heterogeneous multiple-step mechanical unfolding pathways at low force, akin to molten globule states. Small-angle X-ray scattering data revealed that PimA experiences remarkable flexibility that undoubtedly corresponds to the N-terminal domain, which has been proposed to participate in lipid acceptor and protein–membrane interactions (Guerin et al. 2009; Giganti et al. 2013). Altogether, the experimental data support a model wherein the flexibility and conformational transitions confer the adaptability of PimA to the donor and acceptor substrates, and to the membrane, which seems to be of importance during catalysis.

We gratefully acknowledge the valuable scientific discussions with Prof. Åke Wieslander (Department of Biochemistry and Biophysics, Stockholm University) and Prof. Antoni Planas (IQS, Barcelona, Spain) and all members of the Structural Glycobiology Group (Spain). We thank Michael Nilges (Institut Pasteur, Paris, France), who provided the ICM software for the calculation of the desolvation energy values. D.A.J. and D.G. acknowledge the support from the Fundación Biofísica Bizkaia (Spain).