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Structure

Article Crystal Structures of Human Saposins C and D: Implications for Lipid Recognition and Membrane Interactions Maxim Rossmann,1 Robert Schultz-Heienbrok,1 Joachim Behlke,2 Natascha Remmel,3 Claudia Alings,1 Konrad Sandhoff,3 Wolfram Saenger,1,* and Timm Maier1,4,* 1Institut

fu¨r Chemie und Biochemie, Kristallographie, Freie Universita¨t Berlin, Takustr. 6, 14195 Berlin, Germany Delbru¨ck Centrum fu¨r Molekulare Medizin, Robert-Ro¨ssle Str. 10, 13092 Berlin, Germany 3Kekule ´ -Institut fu¨r Organische Chemie und Biochemie der Universita¨t Bonn, Gerhard-Domagk Str. 1, 53121 Bonn, Germany 4Present address: Institut fu ¨ r Molekularbiologie und Biophysik, ETH Zurich, Schafmattstr.20, 8093 Zurich, Switzerland. *Correspondence: [email protected] (W.S.), [email protected] (T.M.) DOI 10.1016/j.str.2008.02.016 2Max

SUMMARY

Human saposins are essential proteins required for degradation of sphingolipids and lipid antigen presentation. Despite the conserved structural organization of saposins, their distinct modes of interaction with biological membranes are not fully understood. We describe two crystal structures of human saposin C in an ‘‘open’’ configuration with unusual domain swapped homodimers. This form of SapC dimer supports the ‘‘clip-on’’ model for SapC-induced vesicle fusion. In addition, we present the crystal structure of SapD in two crystal forms. They reveal the monomer-monomer interface for the SapD dimer, which was confirmed in solution by analytical ultracentrifugation. The crystal structure of SapD suggests that side chains of Lys10 and Arg17 are involved in initial association with the preferred anionic biological membranes by forming salt bridges with sulfate or phosphate lipid headgroups. INTRODUCTION Sphingolipid activator proteins (saposins or Sap) are highly homologous, small, nonenzymatic proteins. They belong to the large and divergent family of saposin-like proteins (SAPLIPs) and domains containing the ‘‘saposin fold’’ characterized by four or five adjacent amphipathic a helices forming bundles stabilized by conserved disulfide bridges or by cyclization (Gonzalez et al., 2000). The all-helical core structure of SAPLIPs is adapted to carry out a number of different functions at biological membranes. SAPLIP domains have been identified in proteins as diverse as cytolytic proteins from amoeba (Hecht et al., 2004), granulysin (Anderson et al., 2003), NK-lysin from pig (Liepinsh et al., 1997), jellyfish lens crystallin (Piatigorsky et al., 2001), plant aspartic protease (Kervinen et al., 1999), and neurotrophic factors (Bornhauser et al., 2003; O’Brien et al., 1995). The four human saposins SapA, B, C, and D are amphiphatic glycoproteins produced in late endosomes or lysosomes by sequential proteolysis of the precursor prosaposin. Within the

lysosomes, human saposins facilitate loading of hydrophobic lipid antigen from lysosomal membranes onto human CD1 molecules, which finally present them to T lymphocytes (Winau et al., 2004; Yuan et al., 2007; Zhou et al., 2004). Saposins are also essential cofactors in the degradation of glycosphingolipids (GSLs) in lysosomes and assist water-soluble-specific exohydrolases to access their membrane-bound GSLs and ceramide targets (Sandhoff and Kolter, 1996). Despite their similar structures, each saposin promotes the degradation of particular sphingolipids by a specific enzyme or partially overlapping sets of enzymes. SapD is involved in vivo in ceramide hydrolysis by acid ceramidase (Linke et al., 2001), whereas SapC is mainly responsible for potentiating the activity of acid b-glucosidase that degrades glucosylceramide (Qi et al., 1994), but additionally promotes degradation of ceramides (Linke et al., 2001), galactosylceramides (Harzer et al., 1997), and galactosylsphingosine (Harzer et al., 2001). In addition to their stimulatory effects, saposins have membrane binding and lipid transport properties. SapC and SapD participate in the destabilization of acidic phospholipid vesicles and modulate the membrane structure in a detergent-like manner (Ciaffoni et al., 2001; Vaccaro et al., 1994). Although the exact mode of hydrolase activation by these saposins is still unknown, it was proposed that SapC and lipid-degrading enzymes colocalize and directly interact at membrane sites remodeled by saposin binding (Wilkening et al., 1998). This model is supported by recent studies showing that SapC decreases the thickness of membranes (Alattia et al., 2006), thereby providing microenvironments optimized for sphingolipid hydolysis, and that SapC forms protein complexes with acid b-glucosidase at the bilayer (Alattia et al., 2007). In addition, SapC is capable of inducing the fusion of phospholipids-containing vesicles (Vaccaro et al., 1994; Wang et al., 2003), a feature not exhibited by the other saposins. Its membrane fusion activity is strictly dependent on pH and the presence of negatively charged phospholipids (Wang et al., 2003) because it relies on electrostatic interactions between negatively charged lipid head groups and positively charged amino acids in SapC (Qi and Chu, 2004) triggered by neutralization of glutamates in the acidic lysosomal environment (de Alba et al., 2003). Similarly, solubilization of membranes by SapD was shown to be effective only at low pH and in the presence of anionic lipids (Ciaffoni et al., 2001), suggesting that principles of initial membrane binding are common to all saposins.

Structure 16, 809–817, May 2008 ª2008 Elsevier Ltd All rights reserved 809

Structure Crystal Structures of SapC and SapD

Figure 1. Structure of SapC and SapD Monomers (A) Sequence alignment of saposins and SAPLIPS generated with ALSCRIPT (Barton, 1993). Positions of a helices a1 to a4 are shown on top. Conserved Cys and Pro are on orange background, other conserved amino acids are on blue background, and type-conserved hydrophobic residues are on white background and boxed. Conserved Tyr54/Phe54 is marked (*), and Lys conserved in SapD are marked (+). (B, C, and D) Ribbon representation of SapD and monomers of SapC, helices colored as in (A). Hinges between helices a1/a20 and a30 /a300 are indicated by arrows. Tyr54 with peptide torsion angles of 4, f at 123 , 15 for tetragonal SapC, 101 , 19 for orthorhombic SapC, 112 , 2 for SapD (Table S1) is at the hinge a30 /a300 .

The structure determination of saposins has provided some insights into their general functional mechanism (Ahn et al., 2003, 2006; de Alba et al., 2003; Hawkins et al., 2005). In all cases, ligand-free saposin adopted monomeric compact four-helix bundle-type structures. In contrast, the nuclear magnetic resonance (NMR) structure of SapC bound to sodium dodecyl sulfate (SDS) micelles depicted a very different boomerang-shaped configuration with an exposed hydrophobic surface (Hawkins et al., 2005). A similar, yet distinct fold was found for SapB, where two V-shaped monomers form a shell around encapsulated lipids bound in the hydrophobic cavity (Ahn et al., 2003). We present two crystal structures of SapC in a novel homodimeric extended open configuration with relevance for membrane interaction and fusion and the crystal structure of SapD, which crystallized as a homodimer in two different crystal forms, together with the structures of SapD-iodoTyr54 and the variant SapD-K9E. Together, these structures suggest a structural basis for the initial mode of membrane interaction and for the preferential binding to anionic membranes.

recombinant SapD remained mostly unglycosylated during expression in P. pastoris. MALDI-TOF spectra revealed a partial degradation of the His6 tag and partial presence of the N-terminal (Glu-Ala)2, indicating inefficient cleavage of the signal sequence by Ste13 protease. Structure Determination of SapD and SapD-K9E Because solution of the phase problem of SapD in space group P1 with four molecules per asymmetric unit failed when using molecular replacement, several approaches for heavy atom derivatization of SapD crystals were tested, among them vapor phase iodination that transformed space group P1 to C2221 with one molecule SapD-iodoTyr54 per asymmetric unit. Though trials failed to determine the structure of SapD-iodoTyr54 using the anomalous signal of iodine, molecular replacement searching with the partial structure of granulysin (Anderson et al., 2003) was successful. The iodination was confirmed by strong electron density for one iodine atom bound to Tyr54. The SapD-iodoTyr54 model served to determine the structures of SapD and SapD-K9E by molecular replacement.

RESULTS Expression and Purification of Human SapD and SapD-K9E We used our procedure (Schultz-Heienbrok et al., 2006) for SapC to prepare SapD and SapD-K9E at 5 mg purified protein per liter of P. pastoris culture medium. In contrast to the wild-type,

Tertiary Structures of SapD and SapD-K9E SapD forms a distorted four-helix bundle composed of a helices a1 (residues 4–18), a2 (residues 25–38), a3 (residues 43–62), and a4 (residues 68–76) (Figures 1B and 2A). Helix a2 is subdivided into a helix a20 (residues 25–32) and a 310 helix a200 (residues 33–38). A kink divides helix a3 in a30 (residues 43–54) and

810 Structure 16, 809–817, May 2008 ª2008 Elsevier Ltd All rights reserved


Figure 2. Crystal Structure of SapD (A) Proposed SapD dimer formation. SapD dimers are formed by contacts between hairpin turns connecting a200 , a30 and N-termini of helices a30 of molecules B and C, red ellipse indicates local C2 axis. In triclinic SapD, sulfate ions (marked SO4) are bound by Lys10 and Arg17 of molecules A, B, and C but not D. Tyr54 (iodinated in SapD-iodoTyr54), Phe4, and Phe50 (magenta) shield the hydrophobic inner cavity. (B) Sulfate binding sites formed inter- and intramolecularly by Lys10 and Arg17 of SapD molecules A and B. Electron density is contoured at 1.3 s level. The distance between the two sulfate sulfur atoms is 8.3 A˚.

a300 (residues 55–62), which, similar to the SapA structure, is caused by disruption of helix a3 between conserved Tyr54 and Pro56. Helix a1 is crosslinked with the C-terminal loop and helix a4 by disulfides Cys5–Cys78 and Cys8–Cys72, respectively, and disulfide Cys36–Cys47 connects the central helices a200 and a30 . All four a helices interact in a tight network of hydrophobic interactions with contributions of 39 of the 58 residues in the a and 310 helices. The kinks in helices a2 and a3 are located at residues Glu33 and Glu55 (Figure 1B), respectively, that are conserved in SapD (Figure 1A). Their carboxylates are hydrogen bonded with each other, suggesting that at least one of the glutamates is protonated at pH 5.9 used for crystallization. In the triclinic SapD structure, sulfate ions from the precipitant are coordinated by Lys10 and Arg17 on helix a1 (Figures 2A and 2B) for three of the four independent molecules. Because SapD interacts preferentially with membranes containing anionic phospholipids (Ciaffoni et al., 2003), the interaction with sulfate in the crystal likely mimics the physiological interaction of SapD with phosphate moieties. SapD contains a series of conserved positively charged residues (Figure 1A; see also Figure S1 in the Supplemental Data available with this article online), among them Lys9 and Lys10, and one might have assumed that the accumulation of positive charges in this region is required for anion binding. However, the inversion of charge at position 9 (variant SapDK9E) neither abolished binding of sulfate by Lys10 and Arg17 in the crystals nor affected the overall or local configuration of SapD. SapD Homodimer A molecular crystal packing analysis of SapD in the three different space groups (Table 1) revealed that the contact B/C (Figure 2A and Figure S2) prevails and buries 530 A˚2 of surface area per monomer (i.e., 11% of the total monomer surface). The B/C contact is formed by more than 30 mostly polar interactions between at least 11 residues on either monomer of the dimer. The central residues involved in the B/C contact (Phe4, Lys45, Gln46, and Asp48) are conserved in SapD (Figure 1A), and additional contacts are formed by conserved Gly2 and by residues in segments 41–49 and 52–53. Surprisingly, Gln49

aligns with Cys48 in the related saposin-like surfactant protein B (SP-B) from human lung that forms a homodimer stabilized by a Cys48–Cys480 disulfide between the monomers. In the theoretical model of homodimeric SP-B, residues that are engaged in dimerization (Zaltash et al., 2000) are equivalent in sequence to those in the SapD dimer, strongly suggesting that this SapD contact region is physiologically relevant. To analyze the oligomeric state of SapD in solution, sedimentation equilibrium experiments were carried out at different concentrations of SapD under conditions resembling the lysosomal (10 mM sodium acetate, 150 mM NaCl, [pH 4]) and the cytosolic milieu (10 mM Tris-HCl, 150 mM NaCl [pH 7.5]). The obtained molecular mass values at acidic and neutral pH are concentration dependent and typical for a monomer-dimer equilibrium with an average Kd of 118.1 ± 19.6 mM and Kd of 103.6 ± 24.4 mM, respectively, indicating that at protein concentrations around or above 1 mg/ml, a significant proportion of SapD molecules occurs as dimers. As shown in Figure S3, the surface of the proposed SapD dimer is polar. The ‘‘top’’ of the dimer is enriched in hydrophobic and uncharged residues, whereas lysines and one arginine are located at the ‘‘bottom’’ of the dimer where sulfate ions are located, suggesting that this is the positively charged initial binding site for phospholipids. Structure Determination and Tertiary Structure of Human SapC In both space groups, SapC forms a homodimer with domainswapped boomerang-shaped monomers in an extended, open configuration. The SapC monomers form a distorted four-helix bundle defined as for SapD except for helix a1 that is two residues longer (residues 2–18), helices a2, a3, and a4 are broken into the same segments, and disulfides are formed as described for SapD (Figures 1C and 1D). The four helices interact in a tight interface of hydrophobic interactions (Figure 3A) with contributions from 39 of 60 helical residues. In both crystal forms, the monomers in the domain-swapped SapC dimers adopt a nearly identical configuration, with RMS deviations for the Ca atoms of residues 2–78 at 0.9 A˚ and 1.3 A˚



Table 1. Data Collection and Refinement Statistics Orthorhombic SapC

Tetragonal SapC

SapD

SapD-iodoTyr54

SapD-K9E

Wavelength (A˚)

1.54179

0.93300

1.54179

1.12714

0.95373

Space group Unit cell (A˚); ( )

C2221

P41212

P1

C2221

P21

57.0, 88.9, 93.5;

49.0, 49.0, 155.6;

40.2, 41.1, 65.9; 89.9, 88,8, 71.5

40.5, 74.9, 66.8;

40.4, 66.0, 42.5; 114.8 6695

Data collection

Unique reflections

8953

7011

21627

3659

Completeness (%)a

98.9 (99.8)

98.4 (96.3)

94.5 (91.7)

98.7 (97.7)

94.6 (71.6)

I/s(I)

16.9 (2.1)

19.0 (6.5)

12.7 (3.3)

13.5 (3.1)

14.3 (7.3)

Rsym

5.5 (46.5)

9.3 (54.5)

5.0 (18.3)

10.4 (49.1)

7.5 (12.0)

Refinement

a

Resolution (A˚)

40.0–2.45

25.0–2.5

22.0–2.10

19.0–2.5

38–2.5

R/Rfree

22.9/28.0

23.9/27.1

21.1/27.4

24.8/27.8

21.0/28.7

Rmsd bond length (A˚)

0.012

0.011

0.009

0.011

0.009

Rmsd bond angles ( )

1.41

1.33

1.25

1.39

1.20

Values in parentheses refer to outer shell.

for the tetragonal and orthorhombic crystal forms, respectively. However, between the two crystal forms the monomers differ in their opening angle measured between helix pairs a1/a20 and a30 /a300 (Figures 3B and 3C). In orthorhombic SapC, the opening angle at the hinge (arrows in Figures 1C and 1D) between a1/a20 and a30 /a300 is wider (116 ) than in the tetragonal form (112 ), whereas in hexagonal SapC and in the complex between SapC and SDS, the opening angles are 97 and 34 , respectively (Figures 1C and 1D, and 3B and 3C). The configuration of helix pair a1/a4 is comparable in both SapC crystal structures, whereas helix a3 in orthorhombic SapC is less bent than in tetragonal SapC where it is broken. The dimerization interfaces of orthorhombic and tetragonal SapC bury 1721 A˚2 and 1965 A˚2 surface area per monomer, respectively. Most hydrophobic residues in the helical parts are involved in the formation of a tight dimer interface, resembling the packing of the common closed saposin four-helix bundle. In tetragonal SapC, 35 residues from either monomer form the dimer through more than 100 intermolecular contacts, of which 51% are hydrophobic. The hinge of molecule A in orthorhombic SapC is less strongly involved in crystal contacts than in the tetragonal space group and shows higher flexibility as indicated by elevated B-factors. This dimer is formed by only 56 intermonomer contacts, although the amount of hydrophobic contacts remains about the same as in the tetragonal space group. The ‘‘top’’ of the SapC dimer is enriched in polar residues, and positively charged lysines are located mostly on the ‘‘top’’ and ‘‘sides’’ of the dimer (Figure 3A and Figure S3). The ‘‘bottom’’ surface of the dimer is hydrophobic and contains Tyr4 and Tyr54 (Figure 3A), which act as gatekeepers of the hydrophobic inner cavity. DISCUSSION Configurational Flexibility and Oligomerization of Saposins Two configurations have been described previously for saposins: the substrate-free canonical closed four-helix bundle and

the V-shaped, open, and ligand-bound configurations in the presence of lipids or SDS as observed for SapB and SapC, respectively. Though the hydrophobic insides form the substrate-binding cavity in the open configurations of SapB and SapC, in SapC structure they are involved in dimer formation in the domain-swapped monomers of the SapC dimer. The hinge angle between helix pairs (legs) is 34 in monoclinic SapC and 97 in the SapC-SDS complex and widened to 112 and 116 in the orthorhombic and tetragonal crystal forms, respectively, of unliganded SapC (Figures 3B and 3C, and Figures S4A and S4B) reported in this study. These differences in hinge bending demonstrate an exceptional level of configurational flexibility that appears to be restrained only by the three disulfides and possibly required at various stages of membrane interaction with SapC. Previously, in the absence of detergents, a dimerization of SapC has not been observed. The crystal structure of SapC (Ahn et al., 2006), as well as its solution structure (de Alba et al., 2003), depicted a compact monomeric fold common to other members of the family, whereas the NMR solution structure in presence of anionic detergent SDS revealed an opening of helix pairs without formation of oligomers (Hawkins et al., 2005). By contrast, analytical ultracentifugation at low pH revealed oligomerization of SapC in the presence of the detergent C8E5 supposedly driven by significant structural changes of the tertiary structure of individual chains (Ahn et al., 2006). In our study, no detergents were added during the purification or crystallization of SapC. However, SapC has been expressed here in the yeast P. pastoris and not in E. coli as in previous studies. It is possible that the interaction of SapC with endogenous lipids, as oberserved in case of the SapB crystal structure (Ahn et al., 2003), influences the folding and structure of SapC. For crystallization, SapC samples were temporarily incubated at their physiological temperature of 37 C. This procedure may have favored a particular configuration of SapC, similar to that observed by NMR spectroscopy that indicated a shift toward a more open configuration and dimerization of SapC in aqueous detergentfree solution at elevated temperatures (John et al., 2006).



Figure 3. Crystal Structure of SapC (A) SapC dimer formation. In the SapC dimer, domains a1/a20 /a200 , and a30 /a300 /a4 are swapped, monomers intertwine to form a dimer, and the red ellipse indicates a local C2 axis. (B) Superimposition of the four SapC molecules. Atoms of residues 2–19 were used for superimposition (SapC+SDS: PDB ID: 1SN6; hexagonal SapC: PDB ID: 2GTG; tetragonal and orthorhombic SapC: this study). SapC undergoes remarkable bending at hinges (arrows) that transform it from a compact closed configuration with hydrophilic exterior (hexagonal SapC) to an open configuration exposing hydrophobic residues for lipid and membrane interaction. (C) Schematic presentation of the hinge-bending motion of SapC. The hinge is located between a1 and a20 at Asn22. The angle is measured between Ca atoms of Val3, Asn22, and Ser37.

Oligomerization seems to be a common feature of saposins. SapA and B form dimers in solution (Ahn et al., 2006). Analytical ultracentrifugations showed that SapD, which crystallized in the compact closed form, is in a monomer-dimer equilibrium in solution at pH 4.0 and pH 7.5. As indicated by a crystal-packing analysis and by analogy to SP-B, the homodimer interface is formed by the N termini of helices a1 and a30 . This interface involves regions of the structure that are at the tips of both legs and separated farthest in the V-shaped or extended open configurations of other saposins. Thus, this interface must be disturbed upon ligand-induced opening of SapD, suggesting that the dimer represents a storage form of SapD that is stabilized against unspecific substrate binding by homophilic interactions. Initial Interactions of Saposins with Lipid Headgroups Among the various SAPLIPs, the biological functions and details of interactions with membrane components are diverse. The most detailed model for initial interaction of SAPLIPs with membranes has been proposed on the basis of sulfate binding sites in crystalline granulysin and biophysical studies on NK-Lysin (Gutsmann et al., 2003). A general importance of positively charged clusters on the surface of saposins that interact with negatively charged lipid head groups has been recognized for saposinmembrane interactions (Ciaffoni et al., 2001; de Alba et al., 2003; Liu et al., 2005). However, the crystal structure of SapD shows that the granulysin mechanism cannot apply for all saposins because the patch of positively charged amino acids for initial interaction with phospho- or sulfolipids is similar in granulysin and NK-Lysin, but not in SapD and SapC (Figure S1). In turn, both Lys10 and Lys17 that interact with sulfate anions in SapD are not conserved in granulysin (only Lys17) or in NK-Lysin (only Lys10) (Figure 1A). The anion-binding patch of positively charged residues in granulysin and NK-Lysin is located on helix a30 , oriented toward a2 and the kink in helix a3 at conserved Tyr54/Phe54. By contrast, sulfatebinding Lys10 and Lys17 on helix a1 in SapD point toward the

loop between helices a200 and a30 at the tip (hairpin) of one of the legs of the open configuration of saposins. Notably, both sulfate/phosphate binding patches are orientated toward regions with particular importance for the hinge-bending motion, either the hinge around Tyr54 in granulysin or the tip of one of the two legs, where helices separate farthest during opening. Only three polar residues that are not cysteines are conserved between the four saposins: Asn22, Thr24, and Tyr54/Phe54 (Figure 1A), and the conservation of the first two of these residues is only due to the presence of glycosylation sites N-X-T in prosaposin. Because aromatic Tyr and Phe side chains were found to be involved in glucolipid binding in diverse proteins (Mahfoud et al., 2002; Malinina et al., 2004; Wright et al., 2005), they consequently are also candidates for glycolipid recognition centers in saposins. The solvent accessible, adjoined and conserved Tyr54, Phe50, and Phe4 seal the buried hydrophobic inside of SapC and SapD (Figures 2A and 3A), and configurational flexibility at the opening to the hydrophobic cavity has been suspected to be important for the recognition and extraction of lipids by saposins and other lipid-transport proteins (Ahn et al., 2003; Niere et al., 2001; Wright et al., 2005). In addition, the kinking of helix a3 at conserved Tyr54 was proposed to be a key for the plasticity of saposins (Ahn et al., 2006). The conservation of this kink in all ligand-free saposins and also in the novel extended open configuration of SapC (Table S1) emphasizes its importance. Membrane Interactions by SapD and SapC Biophysical studies documented that the interaction of SapD with membranes is a complex process critically dependent on pH and the presence of anionic lipids (Ciaffoni et al., 2001, 2003). Acting as a ‘‘solubilizer,’’ SapD can break down phospholipids-containing membranes at critical phospholipid concentrations (Ciaffoni et al., 2001). Of special importance is bis(monoacyl)glycerophosphate (BMP), the marker lipid of intralysosomal vesicles (Mo¨bius et al., 2003) where the physiological degradation of glycosphingolipid takes place (Fu¨rst and Sandhoff,



Figure 4. Models of Lipid Activation by Saposins (A) Schematic model for SapD-stimulated lipid activation. Step 1: water-soluble SapD monomers and dimers bind to negatively charged membrane surface. Step 2: SapD rotates such that the hydrophobic ‘‘top’’ of the dimer (Figure 2A and Figure S3) faces the membrane surface. Step 3: SapD changes configuration to a boomerang shape found for SapC (Figure 3A), and amphipathic a helices stretch parallel to the lipid bilayer, exposing polar residues to the solvent. The hydrophobic surface dips into the membrane and perturbs its structure. Step 4: SapD changes configuration to the closed form, lifts a lipid out of the membrane, and may leave the membrane with bound lipid. (B) Clip-on model for SapC-induced vesicle fusion proposed by Wang et al. (2003). SapC molecules anchored to phospholipid bilayers of vesicles clip to each other through domain swapping, bringing the vesicles close enough for fusion. The size of the vesicle and saposins are not on the same scale.

1992). Based on the results presented, and other structures, we suggest the following model for SapD functioning: prior to interaction with membranes, SapD rests in a monomer-dimer equilibrium in its closed compact configuration (Figure 4A). The low pH in lysosomes neutralizes negatively charged glutamates, inhibits repulsion of SapD by negatively charged membrane surface, and makes SapD more hydrophobic. The interaction of the positively charged patch of amino acids at the ‘‘bottom’’ of SapD aligns the molecule on the surface of intralysosomal membranes, which are rich in negatively charged lipids, characteristically BMP (Mo¨bius et al., 2003; Figure 4A, step 1). The ‘‘top’’ of the proposed SapD dimer is rich in apolar residues (Figure S3) favoring a hydrophobic surrounding, which may promote rolling of the dimer by 180 around its long axis on the membrane surface, thereby burying hydrophobic residues in the membrane bilayer and exposing positively charged residues to the solvent (Figure 4A, step 2). During the initial interaction with the membrane, monomer-monomer interactions in the SapD dimer are possibly weakened by structural rearrangements in SapD. The interaction of carbohydrate and alkyl moieties with

the gatekeeper amino acids in SapD (Phe50, Phe4, and Tyr54), which seal the hydrophobic interior, causes a hinge-bending opening of SapD, and the hydrophobic surfaces of the a helices dip into the membrane, thereby perturbing the membrane structure (Figure 4A, step 3). Supported by thermal motion, the orientation process could be accompanied by opening and closing cycles of the SapD monomer at hinges between helices a1/a20 and a300 /a4 and locking of lipid molecules within the hydrophobic cavity. The breakdown of the lipid bilayer in the solubilizer mode (Ahn et al., 2003) could be sufficient for lipid activation, but a simple perturbation of the membrane structure or the ‘‘liftase’’ mode (Sandhoff and Kolter, 1996) could also be involved (Figure 4A, step 4). In contrast to SapD, SapC does not act as a lipid solubilizer, but rather participates in fusion and destabilization of acidic vesicles (Vaccaro et al., 1994; Wang et al., 2003). The initial binding of SapC to negatively charged membranes may occur similarly as for SapD by positively charged lysines located mostly on helices a1 and a20 (Figures 1A and 3A, and Figure S1), followed by gate opening, membrane insertion, and transition to an open



configuration of SapC. In the only reported crystal structure of a SAPLIP in open configuration, two monomers of SapB form a dimeric V-shaped shell containing a population of E. coli lipids. A similar structure may also exist for SapC (John et al., 2006; Wang et al., 2003); however, the crystal structure determined in the present study demonstrates the possibility for an alternative, extended open configuration that has been postulated to occur in the hypothetical clip-on model for SapC-mediated vesicle fusion (Qi and Chu, 2004; Wang et al., 2003). In this model, helix pairs a1/a4 at both ends of SapC dimers clip to opposing liposomal vesicles, thereby bringing the vesicles close enough for fusion (Figure 4B). The clip-on model was supported by experiments (Abu-Baker et al., 2005; Wang et al., 2003), that showed helices a1 and a4 to be embedded in negatively charged membrane surfaces, whereas helices a2 and a3 were found to be solvent exposed. Such a configuration for a SapC dimer is now supported by our crystallographic data. Based on the conserved disulfide bridges and amphipathic helices, the question arises whether other saposins could form similar dimers or oligomers through domain swapping, and whether heterodimers formed between different saposins could exist as well in vitro and in vivo. EXPERIMENTAL PROCEDURES Cloning and Mutagenesis of SapD Coding sequences for human SapD were polymerase chain reaction (PCR) amplified from vector pBHE0 (Henseler et al., 1996), introducing C-terminal His6-tag codons and restriction sites. The PCR product was subcloned into the yeast expression vector pPIC9 (Invitrogen) using the XhoI and EcoRI restriction sites that added Tyr and Val at the N terminus of SapD and Arg at the C terminus between the mature sequence and the His6 tag. This plasmid was digested with SacI and SalI, and the fragment containing the SapD gene was purified and religated with plasmid pPIC9K (Invitrogen) to generate vector pPIC9K-D. The Lys9 to Glu9 (K9E) mutation was introduced in the SapD gene using QuikChange mutagenesis kit (Stratagene) with vector pPIC9K-D as template. Expression and Purification of Human SapD and of SapD-K9E Expression and purification SapD and SapD-K9E followed the protocol developed for SapC (Schultz-Heienbrok et al., 2006). His6-tagged SapD and its variant were expressed in P. pastoris and purified by anion exchange, metal chelate, and gel filtration chromatography using 10 mM sodium acetate (pH 4.0), and 150 mM NaCl buffer and then concentrated to 8 mg protein/ml.

was rebuilt with ARP/wARP (Perrakis et al., 2001). Solvent molecules were added manually, Tyr54 was modeled as Iodo-Tyr, and the structure was refined using REFMAC (Murshudov et al., 1997) and CNS (Brunger et al., 1998). SapD-iodoTyr54 was employed in a MOLREP (Vagin, 1997) search against the SapD data, and this structure was refined using REFMAC. The structure of SapD-K9E was determined using the structure of SapD as a search model in MOLREP and refined with REFMAC and CNS. For refinement statistics, see Table 1. Analytical Ultracentrifugation The molecular masses of SapD diluted in acidic (10 mM sodium acetate, 150 mM NaCl [pH 4.0]) or in basic buffer (10 mM Tris-HCl, 150 mM NaCl [pH 7.5]) were determined with an XL-A Analytical Ultracentrifuge (Beckman, Palo Alto, CA) equipped with UV absorbance optics, externally loaded sixchannel cells with 12 mm optical path length and the capacity to handle three solvent-solution pairs of about 70 ml liquid each. The sedimentation equilibrium was reached after 2 hr of overspeed at 30,000 rpm followed by equilibrium speed of 26,000 rpm for 30 hr at 10 C. Depending on the loading concentration, the radial absorbance in each compartment was recorded at three different wavelengths between 270 and 285 nm. Molecular masses were determined using the program POLYMOLE (Behlke et al., 1997). Cloning, Expression, Purification, Crystallization, Data Collection, and Processing of SapC SapC was purified as described previously (Schultz-Heienbrok et al., 2006). His6-tagged SapC was expressed in P. pastoris and purified by metal chelate and anion exchange chromatography followed by gel filtration. Crystals belonging to space groups P41212 and C2221 were obtained in 50 mM sodium acetate (pH 4.0), 50 mM MgSO4, 42% (v/v) pentaerythritol ethoxylate 15/4 or 50 mM sodium acetate (pH 4.0), 50 mM (NH4)2SO4, 30% (v/v) pentaerythritol ethoxylate 15/4, respectively. To improve mixing with the viscous pentaerythritol ethoxylate 15/4 reservoir solution, samples were prewarmed to 37 C for crystallization setup and then incubated at 19 C. X-ray data of the orthorhombic crystal form were collected at room temperature using an in-house EnrafNonius FR571 rotating-anode generator. Diffraction data from the tetragonal crystal form were collected at beamline ID14-2 at ESRF (Grenoble, France). Data processing and reduction were done with the HKL suite. Structure Determination and Refinement of SapC The structure of SapC was determined with MOLREP using data collected from tetragonal crystals and searching with a poly-Ser model of SapC monomer in closed configuration (PDB ID: 2GTG). The obtained SapC dimer was improved with ARP/wARP and manual rebuilding with COOT (Emsley and Cowtan, 2004) followed by restrained refinement with REFMAC and CNS. Solvent molecules were added using ARP/wARP, and the final model was validated using composite omit maps calculated with CNS. SapC dimer was used as search model with MOLREP for SapC in the orthorhombic space group, the structure being refined as described for tetragonal SapC (see Table 1).

Crystallization, Data Collection, and Processing of SapD and SapD-K9E SapD and SapD-K9E were crystallized employing hanging drop vapor diffusion by mixing 2.5 ml of precipitant solution (2.3 M (NH4)2SO4, 0.1 M sodium cacodylate [pH 6.0] or 0.1 M BisTris pH 5.9) and 2.0 ml of protein solution and equilibrating the droplets with 750 ml precipitant solution. Mother liquor containing 22.5% (v/v) glycerol was used as a cryoprotectant. X-ray diffraction data were collected at 100K in-house on an Enraf-Nonius FR571 rotatinganode generator (Enraf-Nonius B.V., Rotterdam, The Netherlands) and Marresearch (Norderstedt, Germany) image plate and using the PSF beamline ID14-1 of Freie Universita¨t Berlin at synchrotron BESSY, Berlin. Data were reduced with the HKL suite (Otwinowski and Minor, 1997). For heavy atom derivatization, crystals were treated with triiodide as described (Evans and Bricogne, 2002) to yield SapD-iodoTyr54.

ACKNOWLEDGMENTS

Structure Determination and Refinement of SapD and SapD-K9E The phase problem was solved for SapD-iodoTyr54 by molecular replacement with AMoRe (Navaza, 1994) using a poly-Ser model of the N- and C-terminal a helices of granulysin. After rigid-body fitting of individual helices, the model

We thank Peter Franke (Berlin) for performing MALDI-TOF-MS and Wilhelm Weihofen and the staff of the PSF beamline of Freie Universita¨t Berlin at BESSY (Berlin) and of ESRF (Grenoble, France) for technical assistance. This study was supported by the Deutsche Forschungsgemeinschaft Grant Sa196/41.

ACCESSION NUMBERS Atomic coordinates and structure factors have been deposited in the Protein Data Bank with the entry codes 2Z9A for tetragonal SapC, 2QYP for orthorhombic SapC, 2RB3 for triclinic SapD, 2R1Q for SapD-iodoTyr54, and 2R0R for SapD-K9E variant. SUPPLEMENTAL DATA Supplemental Data include four figures and one table and can be found with this article online at http://www.structure.org/cgi/content/full/16/5/809/DC1/.



Received: November 7, 2007 Revised: January 30, 2008 Accepted: February 6, 2008 Published: May 6, 2008

Harzer, K., Paton, B.C., Christomanou, H., Chatelut, M., Levade, T., Hiraiwa, M., and O’Brien, J.S. (1997). Saposins (sap) A and C activate the degradation of galactosylceramide in living cells. FEBS Lett. 417, 270–274.

REFERENCES

Hawkins, C.A., de Alba, E., and Tjandra, N. (2005). Solution structure of human saposin C in a detergent environment. J. Mol. Biol. 346, 1381–1392.

Abu-Baker, S., Qi, X., Newstadt, J., and Lorigan, G.A. (2005). Structural changes in a binary mixed phospholipid bilayer of DOPG and DOPS upon saposin C interaction at acidic pH utilizing 31P and 2H solid-state NMR spectroscopy. Biochim. Biophys. Acta 1717, 58–66.

Hecht, O., Van Nuland, N.A., Schleinkofer, K., Dingley, A.J., Bruhn, H., Leippe, M., and Grotzinger, J. (2004). Solution structure of the pore-forming protein of Entamoeba histolytica. J. Biol. Chem. 279, 17834–17841.

Ahn, V.E., Faull, K.F., Whitelegge, J.P., Fluharty, A.L., and Prive, G.G. (2003). Crystal structure of saposin B reveals a dimeric shell for lipid binding. Proc. Natl. Acad. Sci. USA 100, 38–43. Ahn, V.E., Leyko, P., Alattia, J.R., Chen, L., and Prive, G.G. (2006). Crystal structures of saposins A and C. Protein Sci. 15, 1849–1857. Alattia, J.R., Shaw, J.E., Yip, C.M., and Prive, G.G. (2006). Direct visualization of saposin remodelling of lipid bilayers. J. Mol. Biol. 362, 943–953. Alattia, J.R., Shaw, J.E., Yip, C.M., and Prive, G.G. (2007). Molecular imaging of membrane interfaces reveals mode of b-glucosidase activation by saposin C. Proc. Natl. Acad. Sci. USA 104, 17394–17399. Anderson, D.H., Sawaya, M.R., Cascio, D., Ernst, W., Modlin, R., Krensky, A., and Eisenberg, D. (2003). Granulysin crystal structure and a structure-derived lytic mechanism. J. Mol. Biol. 325, 355–365. Barton, G.J. (1993). ALSCRIPT: a tool to format multiple sequence alignments. Protein Eng. 6, 37–40. Behlke, J., Ristau, O., and Schonfeld, H.J. (1997). Nucleotide-dependent complex formation between the Escherichia coli chaperonins GroEL and GroES studied under equilibrium conditions. Biochemistry 36, 5149–5156. Bornhauser, B.C., Olsson, P.A., and Lindholm, D. (2003). MSAP is a novel MIR-interacting protein that enhances neurite outgrowth and increases myosin regulatory light chain. J. Biol. Chem. 278, 35412–35420. Brunger, A.T., Adams, P.D., Clore, G.M., DeLano, W.L., Gros, P., GrosseKunstleve, R.W., Jiang, J.S., Kuszewski, J., Nilges, M., Pannu, N.S., et al. (1998). Crystallography and NMR system: a new software suite for macromolecular structure determination. Acta Crystallogr. D Biol. Crystallogr. 54, 905–921.

Harzer, K., Hiraiwa, M., and Paton, B.C. (2001). Saposins (sap) A and C activate the degradation of galactosylsphingosine. FEBS Lett. 508, 107–110.

Henseler, M., Klein, A., Glombitza, G.J., Suziki, K., and Sandhoff, K. (1996). Expression of the three alternative forms of the sphingolipid activator protein precursor in baby hamster kidney cells and functional assays in a cell culture system. J. Biol. Chem. 271, 8416–8423. John, M., Wendeler, M., Heller, M., Sandhoff, K., and Kessler, H. (2006). Characterization of human saposins by NMR spectroscopy. Biochemistry 45, 5206–5216. Kervinen, J., Tobin, G.J., Costa, J., Waugh, D.S., Wlodawer, A., and Zdanov, A. (1999). Crystal structure of plant aspartic proteinase prophytepsin: inactivation and vacuolar targeting. EMBO J. 18, 3947–3955. Liepinsh, E., Andersson, M., Ruysschaert, J.M., and Otting, G. (1997). Saposin fold revealed by the NMR structure of NK-lysin. Nat. Struct. Biol. 4, 793–795. Linke, T., Wilkening, G., Sadeghlar, F., Mozcall, H., Bernardo, K., Schuchman, E., and Sandhoff, K. (2001). Interfacial regulation of acid ceramidase activity. Stimulation of ceramide degradation by lysosomal lipids and sphingolipid activator proteins. J. Biol. Chem. 276, 5760–5768. Liu, A., Wenzel, N., and Qi, X. (2005). Role of lysine residues in membrane anchoring of saposin C. Arch. Biochem. Biophys. 443, 101–112. Mahfoud, R., Garmy, N., Maresca, M., Yahi, N., Puigserver, A., and Fantini, J. (2002). Identification of a common sphingolipid-binding domain in Alzheimer, prion, and HIV-1 proteins. J. Biol. Chem. 277, 11292–11296. Malinina, L., Malakhova, M.L., Teplov, A., Brown, R.E., and Patel, D.J. (2004). Structural basis for glycosphingolipid transfer specificity. Nature 430, 1048– 1053. Mo¨bius, W., van Donselaar, E., Ohno-Iwashita, Y., Shimada, Y., Heijnen, H.F., Slot, J.W., and Geuze, H.J. (2003). Recycling compartments and the internal vesicles of multivesicular bodies harbor most of the cholesterol found in the endocytic pathway. Traffic 4, 222–231.

Ciaffoni, F., Salvioli, R., Tatti, M., Arancia, G., Crateri, P., and Vaccaro, A.M. (2001). Saposin D solubilizes anionic phospholipid-containing membranes. J. Biol. Chem. 276, 31583–31589.

Murshudov, G.N., Vagin, A.A., and Dodson, E.J. (1997). Refinement of macromolecular structures by the maximum- likelihood method. Acta Crystallogr. D Biol. Crystallogr. 53, 240–255.

Ciaffoni, F., Tatti, M., Salvioli, R., and Vaccaro, A.M. (2003). Interaction of saposin D with membranes: effect of anionic phospholipids and sphingolipids. Biochem. J. 373, 785–792.

Navaza, J. (1994). AMoRe: an automated package for molecular replacement. Acta Crystallogr. D Biol. Crystallogr. A50, 157–163.

de Alba, E., Weiler, S., and Tjandra, N. (2003). Solution structure of human saposin C: pH-dependent interaction with phospholipid vesicles. Biochemistry 42, 14729–14740. Emsley, P., and Cowtan, K. (2004). Coot: model-building tools for molecular graphics. Acta Crystallogr. D Biol. Crystallogr. 60, 2126–2132. Evans, G., and Bricogne, G. (2002). Triiodide derivatization and combinatorial counter-ion replacement: two methods for enhancing phasing signal using laboratory Cu Ka X-ray equipment. Acta Crystallogr. D Biol. Crystallogr. 58, 976–991. Fu¨rst, W., and Sandhoff, K. (1992). Activator proteins and topology of lysosomal sphingolipid catabolism. Biochim. Biophys. Acta 1126, 1–16. Gonzalez, C., Langdon, G.M., Bruix, M., Galvez, A., Valdivia, E., Maqueda, M., and Rico, M. (2000). Bacteriocin AS-48, a microbial cyclic polypeptide structurally and functionally related to mammalian NK-lysin. Proc. Natl. Acad. Sci. USA 97, 11221–11226. Gutsmann, T., Riekens, B., Bruhn, H., Wiese, A., Seydel, U., and Leippe, M. (2003). Interaction of amoebapores and NK-lysin with symmetric phospholipid and asymmetric lipopolysaccharide/phospholipid bilayers. Biochemistry 42, 9804–9812.

Niere, M., Dettloff, M., Maier, T., Ziegler, M., and Wiesner, A. (2001). Insect immune activation by apolipophorin III is correlated with the lipid-binding properties of this protein. Biochemistry 40, 11502–11508. O’Brien, J.S., Carson, G.S., Seo, H.C., Hiraiwa, M., Weiler, S., Tomich, J.M., Barranger, J.A., Kahn, M., Azuma, N., and Kishimoto, Y. (1995). Identification of the neurotrophic factor sequence of prosaposin. FASEB J. 9, 681–685. Otwinowski, Z., and Minor, W. (1997). Processing of X-ray diffraction data collected in oscillation mode. In Methods in Enzymology, Volume 276, C.W. Carter, Jr. and R.M. Sweet, eds. (New York: Academic Press), pp. 307–326. Perrakis, A., Harkiolaki, M., Wilson, K.S., and Lamzin, V.S. (2001). ARP/wARP and molecular replacement. Acta Crystallogr. D Biol. Crystallogr. 57, 1445– 1450. Piatigorsky, J., Norman, B., Dishaw, L.J., Kos, L., Horwitz, J., Steinbach, P.J., and Kozmik, Z. (2001). J3-crystallin of the jellyfish lens: similarity to saposins. Proc. Natl. Acad. Sci. USA 98, 12362–12367. Qi, X., and Chu, Z. (2004). Fusogenic domain and lysines in saposin C. Arch. Biochem. Biophys. 424, 210–218. Qi, X., Leonova, T., and Grabowski, G.A. (1994). Functional human saposins expressed in Escherichia coli. Evidence for binding and activation properties of saposins C with acid b-glucosidase. J. Biol. Chem. 269, 16746–16753.



Sandhoff, K., and Kolter, T. (1996). Topology of glycosphingolipid degradation. Trends Cell Biol. 6, 98–103. Schultz-Heienbrok, R., Remmel, N., Klingenstein, R., Rossocha, M., Sandhoff, K., Saenger, W., and Maier, T. (2006). Crystallization and preliminary characterization of three different crystal forms of human saposin C heterologously expressed in Pichia pastoris. Acta Crystallogr. Sect. F Struct. Biol. Cryst. Commun. 62, 117–120. Vaccaro, A.M., Tatti, M., Ciaffoni, F., Salvioli, R., Serafino, A., and Barca, A. (1994). Saposin C induces pH-dependent destabilization and fusion of phosphatidylserine-containing vesicles. FEBS Lett. 349, 181–186. Vagin, A.T. (1997). MOLREP: an automated program for molecular replacement. Journal of Applied Crystallography 30, 1022–1025. Wang, Y., Grabowski, G.A., and Qi, X. (2003). Phospholipid vesicle fusion induced by saposin C. Arch. Biochem. Biophys. 415, 43–53. Wilkening, G., Linke, T., and Sandhoff, K. (1998). Lysosomal degradation on vesicular membrane surfaces. Enhanced glucosylceramide degradation by lysosomal anionic lipids and activators. J. Biol. Chem. 273, 30271–30278.

Winau, F., Schwierzeck, V., Hurwitz, R., Remmel, N., Sieling, P.A., Modlin, R.L., Porcelli, S.A., Brinkmann, V., Sugita, M., Sandhoff, K., et al. (2004). Saposin C is required for lipid presentation by human CD1b. Nat. Immunol. 5, 169–174. Wright, C.S., Mi, L.Z., Lee, S., and Rastinejad, F. (2005). Crystal structure analysis of phosphatidylcholine-GM2-activator product complexes: evidence for hydrolase activity. Biochemistry 44, 13510–13521. Yuan, W., Qi, X., Tsang, P., Kang, S.J., Illarionov, P.A., Besra, G.S., Gumperz, J., and Cresswell, P. (2007). Saposin B is the dominant saposin that facilitates lipid binding to human CD1d molecules. Proc. Natl. Acad. Sci. USA 104, 5551– 5556. Zaltash, S., Palmblad, M., Curstedt, T., Johansson, J., and Persson, B. (2000). Pulmonary surfactant protein B: a structural model and a functional analogue. Biochim. Biophys. Acta 1466, 179–186. Zhou, D., Cantu, C., 3rd, Sagiv, Y., Schrantz, N., Kulkarni, A.B., Qi, X., Mahuran, D.J., Morales, C.R., Grabowski, G.A., Benlagha, K., et al. (2004). Editing of CD1d-bound lipid antigens by endosomal lipid transfer proteins. Science 303, 523–527.
