Solid-State NMR Identification of Intermolecular Interactions in Amelogenin Bound to Hydroxyapatite

Article

Solid-State NMR Identification of Intermolecular Interactions in Amelogenin Bound to Hydroxyapatite Rajith Jayasinha Arachchige,1 Sarah D. Burton,1 Jun-Xia Lu,1 Bojana Ginovska,1 Larisa K. Harding,1 Megan E. Taylor,1 Jinhui Tao,1 Alice Dohnalkova,1 Barbara J. Tarasevich,1 Garry W. Buchko,1,2,* and Wendy J. Shaw1,* 1 Pacific Northwest National Laboratory, Richland, Washington and 2School of Molecular Biosciences, Washington State University, Pullman, Washington

ABSTRACT Biomineralization processes govern the formation of hierarchical hard tissues such as bone and teeth in living organisms, and mimicking these processes could lead to the design of new materials with specialized properties. However, such advances require structural characterization of the proteins guiding biomineral formation to understand and mimic their impact. In their ‘‘active’’ form, biomineralization proteins are bound to a solid surface, severely limiting our ability to use many conventional structure characterization techniques. Here, solid-state NMR spectroscopy was applied to study the intermolecular interactions of amelogenin, the most abundant protein present during the early stages of enamel formation, in self-assembled oligomers bound to hydroxyapatite. Intermolecular dipolar couplings were identified that support amelogenin dimer formation stabilized by residues toward the C-termini. These dipolar interactions were corroborated by molecular dynamics simulations. A b-sheet structure was identified in multiple regions of the protein, which is otherwise intrinsically disordered in the absence of hydroxyapatite. To our knowledge, this is the first intermolecular protein-protein interaction reported for a biomineralization protein, representing an advancement in understanding enamel development and a new general strategy toward investigating biomineralization proteins.

INTRODUCTION Biomineralization is the process used by living organisms to produce minerals with properties not always found in their equivalent inorganically produced counterparts (1,2). In vertebrates, mineralized tissues such as bone, cementum, dentin, and enamel all consist of calcium phosphate but have specialized functions as a result of the conditions during formation. Biomineralization proteins are an essential contributor to the resulting material properties, because they orchestrate the nucleation, growth, and organization of biominerals (1–3). Understanding the fundamental mechanisms responsible for this orchestration is important because this knowledge will guide the design of advanced

Submitted March 12, 2018, and accepted for publication August 22, 2018. *Correspondence: [email protected] or [email protected] Rajith Jayasinha Arachchige’s present address is Bridgestone Americas Center for Research and Technology, Akron, OH 44301. Jun-Xia Lu’s present address is School of Life Science and Technology, ShanghaiTech University, Shanghai, China. Editor: Elizabeth Komives. https://doi.org/10.1016/j.bpj.2018.08.027 Ó 2018 Biophysical Society.

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materials in general and new repair or regeneration strategies for biominerals (2,4). One of nature’s hardest materials is enamel, which is 95% (by weight) carbonated hydroxyapatite (HAP) (5). The HAP crystals are extremely long and narrow, packed into parallel arrays (enamel rods) interwoven into a distinctive lattice architecture (6). The exceptional functional properties of enamel arise from its intricate hierarchical structure and cannot be reproduced in vitro inorganically. In vivo, the formation of enamel depends on amelogenin, the predominant protein in the enamel matrix (5,7,8). Although amelogenin’s importance in enamel formation is widely accepted, the mechanism of growth and control at a molecular level is not established. In vitro studies suggest that amelogenin self-assembles in a stepwise process, forming dimers (9–11), followed by oligomers (10,11), then nanospheres (20–100 monomers) (11,12), and nanosphere chains (13). The nanosphere structure is the most common structure observed in solution (14–16); however, it is in an equilibrium between structures that can be interrupted by changes in pH, the presence of a surface, or the presence of calcium

Interactions of Amelogenin on HAP

and phosphate ions in solution (9,17–19), all conditions that vary during enamel formation. The dynamic self-assembly process of amelogenin is thought to be important in enamel formation with all the quaternary forms likely playing a role. Cryo-electron microscopy (cryo-EM) studies by Beniash and co-workers (10) identified an amelogenin dimer as a building block upon which the larger oligomer and nanosphere structures are built as enamel grows. The dimer subunit structure is proposed to be stabilized through the C-termini; in reconstituted cryo-EM images of amelogenin monomers, the C-terminus appears as a foot-like appendage extending from the globular core of the protein, appendages that overlap in the dimer subunit. Among supporting evidence for the C-terminal-stabilized dimer building block is the absence of the foot-like appendage and the lack of formation of dimer or oligomer subunits in cryo-EM experiments using an amelogenin construct with the 13 C-terminal residues removed (10). Although these cryo-EM studies offer insight into the tertiary and quaternary structure of amelogenin in the presence of HAP, the spatial resolution of cyro-EM and similar techniques do not provide an atomic-level characterization of the dimer interface, a view needed to fully understand and mimic the elegant process of enamel formation and to understand the role of biomineralization proteins in general. To acquire atomic-level insight, we turned to solid-state NMR (ssNMR) spectroscopy, a technique uniquely suited to study the structure and intermolecular interactions of surface-immobilized proteins at the atomic level (20–24). We recently demonstrated the power of ssNMR to reveal structural features in large biomineralization proteins on surfaces by evaluating the structure of select residues in full-length amelogenin mineralized with HAP (25). Here, we use ssNMR for the first time to identify intermolecular interactions between full-length amelogenin bound to its biologically relevant surface to provide a molecular level explanation for observed tertiary and quaternary interactions (10). We hypothesized that the dimer subunits observed by cryo-EM (10) were stabilized by intermolecular salt bridges between the 13 C-terminal residues aligned antiparallel as shown in Fig. 1. Intermolecular interactions near the salt bridges can be measured directly by introducing select amino acid residues with isotopic labels A

168 ...W D

B

169 170 171 172 173 174 175 176 P K R A T D K T

177 E

178 E

D

T

A

V

E

E

R

K

T

K

(13C and 15N) and evaluating the presence of dipolar couplings between those labeled residues. If two nuclei ˚ ), a crosspeak will are in close proximity (within
7 A be observed in tailored two-dimensional (2D) ssNMR spectra (26). If they are farther apart, no crosspeaks will be observed. MATERIALS AND METHODS Protein expression and purification Full-length murine amelogenin (M179) without an affinity purification tag was expressed using recombinant methods in Eschericia coli. The N-terminal methionine is removed in the recombinantly expressed protein by E. coli methionine aminopeptidase (27), and the side chain of S16 is not phosphorylated, counter to the in vivo observations in developing enamel, because of the absence of the proper phosphorylase in E. coli. Three samples were prepared with residue-specific 13C-, 15N-labeled amino acids incorporated into the primary amino acid sequence: threonine only (M179-T), arginine only (M179-R), and a 1:1 mixture of M179-T and M179-R (M179-T/R). This was accomplished biosynthetically using a modified ‘‘Redfield-medium’’ containing adenine (500 mg/L), guanosine (650 mg/L), thymine (200 mg/L), uracil (500 mg/L), cytosine (200 mg/L), anhydrous sodium acetate (1500 mg/L), succinic acid (1500 mg/L), ammonium chloride (750 mg/L), NaOH (850 mg/L), anhydrous K2HPO4 (10,500 mg/L), CaCl22H2O (2 mg/L), ZnSO47H2O (2 mg/L), MnSO4H2O (2 mg/L), thiamine (50 mg/L), niacin (50 mg/L), biotin (0.6 mg/L), ampicillin (150 mg/L), glucose (20,000 mg/L), MgSO4 (480 mg/L), and FeCl3 (1.6 mg/L) (28–30). The medium also contained 19 of the 20 common amino acids except asparagine, as follows: A (500 mg/L), R (400 mg/L), D (400 mg/L), C (50 mg/L), Q (400 mg/L), E (650 mg/L), G (550 mg/L), H (100 mg/L), I (230 mg/L), L (230 mg/L), K (420 mg/L), M (250 mg/L), F (130 mg/L), P (100 mg/L), S (2100 mg/L), T (230 mg/L), Y (170 mg/L), V (230 mg/L), and W (50 mg/L). To limit ‘‘scrambling’’ of the targeted amino acid(s) only onethird to one-quarter of the amount requested by the recipe, unlabeled, was added to the medium at the start of the culture growth. Ten minutes before induction with isopropyl-b-D-1-thiogalactopyranoside (0.26 mg/L) at an optical density OD600
1.0, the 13C- and 15N-labeled target amino acid(s) (approximately half of the amount requested by the recipe) was added to the media, and the temperature was lowered to 25 C. After further shaking for 30–60 min, the cells were harvested by centrifugation, and the cell pellet was frozen at 80 C until it was thawed for protein isolation. The frozen cell pellet of a 750 mL culture was resuspended in 6 M guanidinium hydrochloride (
35 mL), passed through a French press three times, sonicated, centrifuged, and the supernatant was dialyzed twice in 2% acetic acid (
1:140, v/ v) and was centrifuged and purified by preparative reverse-phase high-performance liquid chromatography (27). Protein purity was verified by sodium dodecyl sulfate polyacrylamide gel electrophoresis and analytical reversephase high-performance liquid chromatography. Residue-specific label incorporation and the amount of scrambling present was assessed by

179 180 V D P

W...

180’ 179’ 178’ 177’ 176’ 175’ 174’ 173’ 172’ 171’ 170’ 169’ 168’

MPLPPHPGSPGYINLpSYEVLTPLKWYQSMIRQPYPSYGYEPMGGWLHHQIIPVLSQQHPP SHTLQPHHHLPVVPAQQPVAPQQPMMPVPGHHSMTPTQHHQPNIPPSAQQPFQQPFQPQ AIPPQSHQPMQPQSPLHPMQPLAPQPPLPPLFSMQPLSPILPELPLEAWPATDKTKREEVD

FIGURE 1 (A) Hypothesized antiparallel alignment of the 13 C-terminal residues in amelogenin dimer subunits. This alignment highlights several possible salt bridges between arginine (R) or lysine (K) with aspartic acid (D) (shaded cyan) and the proximity of the T and R residues (shaded magenta) labeled in this study. (B) The primary amino acid sequence for full-length murine amelogenin (M179) with the labeled T and R residues colored blue and green, respectively. To see this figure in color, go online.

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Arachchige et al. resuspending samples in 2% acetic acid and collecting a 1H-15N heteronuclear single-quantum coherence spectrum. Typical protein yields were
15 mg/L.

Amelogenin-HAP complex preparation To obtain a homogeneous stock solution of monomeric M179, lyophilized protein was resuspended to a concentration of 1 mg/mL in 1 mM HCl, purged with nitrogen gas to remove dissolved CO2, and stored at 4 C overnight. To determine the optimal conditions to produce the M179-HAP complexes for the ssNMR experiments, the stock solution was diluted with 1 mM HCl to prepare five different solutions for study at the 2 mL scale: 20, 200, 400, 800, or 1000 mg/mL. To each 2 mL amelogenin solution, 38.8 mL of 130 mM CaCl2 stock solution was added to achieve a final Ca2þ concentration of 2.5 mM. After 5 min of stirring under an N2 purge, 37.5 mL of 80 mM KH2PO4 stock solution was added to the solution to attain a final concentration of 1.5 mM phosphate. During this process, the solution pH remained at
3.0. The solution pH was then increased to 7.8 using 0.1 M KOH while stirring with an N2 overlayer. The cap was placed on the centrifuge tube, and the mixture was then placed in a 37 C water bath and incubated for 24 h. The resultant precipitates were characterized by transmission electron microscopy (TEM) to verify the presence of elongated crystal structures indicative of amelogenin-HAP complexes (10,25,31). The 200 mg/mL amelogenin solution was chosen as the condition for scale-up to a volume of 40 mL for the ssNMR sample preparations because it maximized the amount of protein-HAP complex formed and maintained the crystal size while limiting coprecipitation of protein. The resulting ratio of protein/HAP was 1:18.5. The large-scale sample was spun down at 4500 rpm for 5 min, and the pellet was transferred to an ssNMR rotor using our previously established procedure (25).

SSNMR studies The one-dimensional (1D) and 2D NMR experiments at 25 C were conducted on either a Varian VNMRS spectrometer (Agilent Technologies, Santa Clara, CA) with a 19.96 T wide-bore magnet, operating at resonance frequencies of y0 (13C) ¼ 213 MHz and y0 (1H) ¼ 850 MHz, equipped with a quadruple-resonance, 3.2 mm HFXY probe (Agilent Technologies) with variable temperature capability or an 11.7 T narrow-bore magnet, operating at resonance frequencies of y0 (13C) ¼ 125 MHz and y0 (1H) ¼ 500 MHz, equipped with a triple-resonance, 4 mm HXY probe (Agilent Technologies) with variable temperature capability. Initial experimental parameters were optimized using adamantane, and the 13C-, 15N-labeled tripeptide, N-formyl-Met-Leu-Phe. Further power optimization of the 90 and decoupling pulse widths were calibrated using the labeled amelogenin-HAP samples. For cross-polarization (CP) experiments, the contact time ranged from 1 to 2 ms, the 1H 90 pulse ranged from 3.4 to 3.7 ms, the 13C 90 pulse ranged from 3.6 to 4.1 ms, and the 1H decoupling field strength ranged from 62 to 74 kHz. The DARR mixing time was 100 ms with a field strength equal to the spinning speed, which was 13 or 10 kHz. The time required for the CP, DP, and DARR experiments were 3, 3, and 72 h, respectively, for M179-T and M179-R, and 108 h for the CP-DARR experiment for M179-T/R.

TEM analyses TEM images for the mineral crystals were collected with a Tecnai F20 microscope (Field Electron and Ion Company, Hillsboro, OR) at an acceleration voltage of 200 kV. The preparation of the mineralized samples consisted of placing
20 mL droplets of the prepared amelogenin-HAP solutions on a Cu/C grid, washing them with a drop of deionized water, and blotting them with a kimwipe. The crystals were between 24 and 72 h old when the TEM images were recorded.

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MD studies Classical molecular dynamics (MD) studies evaluated the side chain interaction of the antiparallel structure proposed for amelogenin shown in Figs. 1, 4, and S9. Because of the limited structural knowledge of the protein, only the 13 C-terminal residues of amelogenin (W168–D180) shown to have b-sheet character (25) were used to model the tail. All simulations were performed with the Amber12 program (32,33) and the Amber03 force field. The environment around the structure was represented as a continuum solvent with a dielectric constant representative of a water environment (i.e., 78.5). The overall charge of the system was 2. To maintain a b-sheet structure with an antiparallel hydrogen bonding network, the distance between all backbone N and O atoms on opposing strands were constrained ˚ . All simulations used a temperature of 27 C, with a time step to 2.5 A of 1 ps. Statistics were collected from a total simulation time of 40 ns.

RESULTS AND DISCUSSION To simplify the spectral assignment and eliminate issues associated with collecting ssNMR data on uniformly 13 C-labeled protein samples (25), we used a biosynthetic, residue-specific labeling scheme based on the following three criteria: 1) sparsely represented residues in the primary amino acid sequence; 2) residues that do not ‘‘scramble,’’ i.e., the biosynthetic pathway does not result in labeled functional group transfer to other amino acids; and 3) residues that should be close enough in space in our proposed dimer subunit model to detect intermolecular interactions with our ssNMR experiments. Based on these criteria, arginine and threonine residue-specific labeling was chosen for our study, and the following three 13C-, 15N-residuespecific amelogenin samples were prepared: threonine only (M179-T), arginine only (M179-R), and a 1:1 mixture of M179-T/M179-R (M179-T/R). The 2D 1H-15N heteronuclear single-quantum coherence spectra for M179-R and M179-T collected in 2% acetic acid showed no evidence of significant label scrambling during biosynthesis (Fig. S1). The M179-R and M179-T samples were used to make spectral assignments and evaluate structure; the M179-T/R sample was used to identify intermolecular interactions. HAP was mineralized in the presence of one of each of the three labeled amelogenin samples, as previously described (10,25,31), to generate three amelogenin-HAP complexes. These complexes were evaluated by TEM to verify the presence of the expected elongated HAP crystals (Fig. S2) (10,25,31) and by 1D 13C ssNMR to verify the incorporation of the 13C-labeled amino acid. Integration of the 1D 13C ssNMR spectrum of the M179-T/R complex confirmed an
1:1 mixture of labeled M179-T and M179-R. The observation of similar signal-to-noise levels in the CP and direct-polarization (DP) experiments collected with a similar number of scans (Fig. S3) is indicative of restricted mobility (25), further verifying the generation of amelogenin-HAP complexes. Two-dimensional dipolar-assisted rotational resonance (DARR) experiments were conducted on the amelogeninHAP complexes to determine if there were dipolar

Interactions of Amelogenin on HAP

interactions between the T and R residues that would support the dimer configuration proposed to explain the cryoEM observations. The 2D-CP-DARR spectra of the three samples are compared in Figs. 2 and S4. To characterize intermolecular interactions, intramolecular intraresidue cross peaks first need to be identified and can be seen from the backbone carbonyls through many of the side chain carbons for each individual residue-specific labeled protein (M179-T and M179-R, colored blue and green, respectively, and indicated with dashed lines in Fig. 2).

ical shifts for each residue (
27 ppm), whereas only two arginine carbonyls (labeled R1 and R3) have dipolar couplings with their Cb carbons (
30.5 ppm). In no case was the C0 -to-Cd crosspeak observed. Chemical shifts for C0 and Ca are the most sensitive to secondary structure and relative to random coil values, move downfield in a-helices (left/down) and upfield for b-strands (right/up) (34,35). The C0 and Ca chemical shifts observed in the M179-R-HAP complex all shift upfield relative to the random coil values (R-C0 ¼ 176.6 ppm and Ca ¼ 56.7 ppm) (35); therefore, all four arginines are likely in a b-sheet environment when M179 is bound to HAP. This is in agreement with our ssNMR studies of lysine-labeled amelogenin bound to HAP, which indicated that the lysines were in a b-sheet conformation (25). To provide more definitive assignment of these residues, we prepared the single amino acid variants R31L and R31K (Figs. S6 and S7). The DARR spectra from these variants suggest that R1 and R3 are the resonances associated with R176. The structural characteristics of the six threonine residues, T21, T63, T95, T97, T171, and T174, are more complex in the CP-DARR spectrum of M179-T. Similar to the arginine residues, there are more threonine C0 resonances than there are residues, at least eight which can be easily identified as shown in Fig. 2 (solid blue lines) and Table S1. Of the observed C0 resonances, there are seven

Secondary structure The arginines in M179 appear structurally distinct. Amelogenin contains two arginine residues present at opposite ends of the primary amino acid sequence, R31 and R176. The CP-DARR spectrum for M179-R (green) contains four distinct C0 resonances at 173.7, 174.7, 175.2, and 175.8 ppm associated with different Ca chemical shifts (between
53 and 56 ppm), indicating that all arginines are in different chemical environments (Table S1). The presence of more than two unique arginine carbonyl spin systems suggests that one or both of the arginine residues exist in two unique structures, as previously reported for lysine residues (25). All four of the arginine carbonyls have dipolar couplings that extend to their Cg carbons with similar chemT1 T2 T3 T4 T5 T6 T7 T8

R1/T R3/T

R1 R2 R3 R4

T2/R T3/R

1 2 34 1 2 345678

T-Cγ

20

Cβ

Cβ Cγ

RC’-TCγ

Cα

Cα

RC’-TCγ

Cγ

OH

Cδ

R-Cγ

N Cε

30

C (ppm)

R-Cβ

N

TC’-RCγ

N

TC’-RCβ TC’-RCδ

13

R-Cδ 50

TC’-RCα R-Cα

RC’-TCα T-Cα

T-Cβ

60

RC’-TCα

70 176

174 172

176

174

172

176

174

FIGURE 2 CP-13C-13C-DARR coherence spectra for left: M179-T (blue); and middle: M179-R (green). M179-T and M179-R were collected at a 500 MHz 1H resonance frequency at 25 C with a 100 ms mixing time. The blue or green lines coincide with C0 resonances and highlight the crosspeaks observed with these resonances. The bold vertical and horizontal dashed lines indicate the random coil values of the associated C0 and Ca resonances, respectively. The chemical shifts upfield of the random coil values indicate that the majority of the Tand R residues are in a b-sheet environment. Right: M179-T/R (red) overlaid with M179-T and M179-R is shown. M179-T/R was collected at an 850 MHz 1H resonance frequency at 25 C with a 100-ms mixing time. Cross peaks are observed between two backbone T-C0 and arginine side chain carbons R-Ca, R-Cb, R-Cd and R-Cg, and the complementary R-C0 backbone with threonine side chain carbons T-Ca and T-Cg, establishing that there is an intermolecular interaction between the C-termini of amelogenin molecules in the protein-HAP complex. To see this figure in color, go online.

172

13

C (ppm)

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unique threonine C0 -to-Cg (
22 ppm) cross peaks, eight cross peaks for C0 to Ca (
58–63 ppm), and six crosspeaks for C0 to Cb (
69 ppm). Five of the C0 resonances have cross peaks to Ca, Cb, and Cg, consistent with these five residues existing in motionally restricted environments or being closer in space (T1, T2, T4, T5, T8), whereas the other three residues (T3, T6, T7) are likely in more mobile environments or farther away, based on the weaker/missing cross peaks. This interpretation would agree with the observations for the arginine spectra, suggesting that at least three of the threonine residues exist in multiple conformations, and is consistent with previous studies with labeled lysine residues (25). If this interpretation is correct, the observation that both arginines and threonines exist in multiple conformations suggests that multiple structures are in both the Nand C-termini and are not isolated to one part of the protein. A second interpretation consistent with the data is that some of the additional resonances in M179-T are due to interresidue interactions, rather than multiple structures. It should be noted that neither of these possibilities exclude the other, and a combination of the two is also conceivable. All of the identified C0 chemical shifts are upfield (right) of 174.8 ppm (Table S1), the random coil chemical shift value for threonine C0 . Six of the threonine Ca chemical shifts, including the strongest cross peaks, are upfield of their respective random coil value (62.0 ppm; dashed lines; Fig. 2). Again, because C0 and Ca chemical shifts are most sensitive to secondary structure, the majority of the threonines are also all likely in b-sheet environments. Note that no new arginine or threonine C0 spin systems are observed in the DP-DARR spectrum (Fig. S5), consistent with the interpretation that R31 and R176, and five of the threonines are located in rigid regions of the protein. To more definitively assign these residues, we prepared a variant with the four N-terminal threonine residues (T21, T63, T95, T97) mutated to serine, leaving only T171 and T174. The results are consistent with assignments of T171 and T174 to wildtype T2, T3, T4, and possibly T5 (Fig. S8). The comparison to the structure of amelogenin in solution is stark (Fig. 3; Table S2) (9,29). The solution data were previously collected in 2% acetic acid (pH
2.8) in which M179 is a monomer (29). Under this condition, there is little evidence of traditional secondary structure (9,36,37). As discussed, C0 and Ca are good indicators of secondary structure, and the larger the chemical shifts are from random coil values, the more canonical structure present. The solution state C0 and Ca chemical shifts for both R and T had minimal dispersion from random coil chemical shift values (<1.0 ppm). On the other hand, in the solid state, the C0 and Ca chemical shift dispersion from random coil values increased significantly for both R and T (>1 ppm for most resonances and >2 ppm for over half the resonances). Although amelogenin undergoes changes in oligomerization state with increasing pH (monomers to dimers to oligomers to nanospheres), circular dichroism spectroscopy indicates

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FIGURE 3 Comparison of the C0 and Ca chemical shifts to random coil values (35) for amelogenin in 2% acetic acid solution (29) and in the amelogenin-HAP complex studied here, where Ds13C ¼ srandom coil  sobserved. The significantly larger changes in chemical shifts for the amelogenin-HAP complex indicate that it becomes more structured when bound to HAP with the typically positive values indicative of a b-sheet structure. To see this figure in color, go online.

that amelogenin undergoes little change in secondary structure between a pH 3.5 and 8 (12), suggesting the protein, by itself, remains unstructured over this pH range. Our ssNMR chemical shift observations for the ameloginen-HAP complex imply that this intrinsically disordered protein becomes structured when interacting with HAP under physiological conditions that mimick early stages of enamel development. We are applying more advanced techniques in ongoing studies to unambiguously assign each residue in the amelogenin-HAP complex to obtain a more detailed description of the amelogenin-HAP interface. Intermolecular interactions The M179-T/R sample offers the ability to confirm our hypothesis of intermolecular interactions stabilizing amelogenin dimers when bound to HAP. Importantly, as highlighted in the right panel in Fig. 2, there are additional cross peaks between the T-C0 and the arginine side chain carbons R-Ca, R-Cb, and R-Cg, as well as between the R-C0 and the

Interactions of Amelogenin on HAP

threonine side chain carbons T-Ca and T-Cb. The M179-T/R T-C0 cross peaks at 173.9 (T2) and 173.5 (T3) ppm to arginine carbons are tentatively assigned to T171 or T174 as discussed above. Reciprocal M179-R/T R-C0 crosspeaks at 175.7 (R1) and 174.9 (R3) ppm to threonine carbons are consistent with the assignment of these resonances as R176 in two different orientations. These cross peaks clearly demonstrate that a T and an R residue from different mole˚ of each other. This observation supports cules are within 7 A the proposed antiparallel, C-terminal salt bridge stabilized dimer alignment illustrated in Fig. 1. Multiple different alignments of two amelogenin C-termini could generate the C-terminal tails observed in the cryo-EM studies. To provide insight into the C-termini alignment consistent with our ssNMR data, MD studies were performed on all possible combinations, and the arginine to threonine distances were calculated. A snapshot of the fully aligned structure is shown in Fig. 4 with the details provided in the Supporting Materials and Methods. If the structure is antiparallel matched with no amino acid overhangs as shown in Figs. 1, 4, and S9, we would expect R176 to interact strongly with T1710 and weakly with T1740 (0 denotes the second chain in the dimer). If the alignment of the two strands is shifted by one residue, with one residue, W168, under alignment (Fig. S9 B), we would expect R176 to interact strongly with T1740 and weakly with T1710 , an alignment that would generate similar NMR data as the fully matched alignment but because of reversed interactions. Both of these alignments agree with the ssNMR data described above. All other possible C-terminal alignments are predicted to generate, at most, a single strong set of arginine crosspeaks between only one, but not two, threonine residues (Figs. S9, C and D), eliminating other possible alignments. The MD simulations show that the fully aligned and the W168 residue under alignment are consistent with our experimental data, but the degeneracy of the predicted interactions prevents us from unambiguously assigning the T1710 and T1740 cross peaks. CONCLUSIONS Using ssNMR, we have identified C-terminal atomic level intermolecular interactions consistent with a C-termini

antiparallel dimer alignment proposed from cryo-EM observations of amelogenin-HAP complexes. The R and T residues predominantly adopt b-strand structure in the amelogenin-HAP complex. Such an increase in canonical structure is markedly distinct from the unstructured disorder that is reported for solubilized amelogenin at all pH values. Similarities in the DP and CP ssNMR data show that the majority of the R and T residues are in motionally restricted regions consistent with the formation of a stable structured complex with HAP. These atomic level details contribute to our larger effort to fully elucidate the mechanism describing the amelogenin-HAP interface under conditions that more closely mimic the biological environment. Not only is this an important advancement toward understanding the mechanism used by amelogenin to control the growth of enamel at the molecular level, but it also offers, to our knowledge, a new general strategy toward investigating the formation of hierarchical hard tissues by biomineralization proteins.

SUPPORTING MATERIAL Nine figures and two tables are available at http://www.biophysj.org/ biophysj/supplemental/S0006-3495(18)30981-0.

AUTHOR CONTRIBUTIONS R.J.A., S.D.B., J.-X.L., and M.E.T. prepared the mineralized amelogeninHAP complexes and performed all ssNMR spectroscopy. B.G. performed the MD simulations. L.K.H and G.W.B. prepared the proteins. J.T., A.D., and B.J.T performed and interpreted TEM images. R.J.A., G.W.B., and W.J.S. wrote the manuscript. All authors contributed to the experimental design, manuscript revision, and discussion of the results.

ACKNOWLEDGMENTS This research was supported by National Institutes of Health/National Institute of Dental and Craniofacial Research grant DE-015347. The research was conducted at Pacific Northwest National Laboratory, a facility operated by Battelle for the US Department of Energy, with a portion of it performed at the W. R. Wiley Environmental Molecular Sciences Laboratory, a national scientific user facility sponsored by the US Department of Energy Biological and Environmental Research program. Pacific Northwest National Laboratory institutional computing was used for the computational portion of this work.

FIGURE 4 Representative MD structure of two amelogenin C-termini in a fully aligned, antiparallel b-sheet alignment that is consistent with the current and previous structural characterizations. The arginine and threonine residues are shown as elementcolored sticks (carbon ¼ cyan, nitrogen ¼ blue, oxygen ¼ red). The backbone residues are shown as blue (T) and green (R) ribbon. One set of the intermolecular interactions observed by ssNMR is highlighted with a pink-shaded ellipse. To see this figure in color, go online.

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REFERENCES 1. Addadi, L., and S. Weiner. 1985. Interactions between acidic proteins and crystals: stereochemical requirements in biomineralization. Proc. Natl. Acad. Sci. USA. 82:4110–4114. 2. Lowenstam, H. A., and S. Weiner. 1989. On Biomineralization. Oxford University Press, New York. 3. Hunter, G. K. 1996. Interfacial aspects of biomineralization. Curr. Opin. Solid State Mater. Sci. 1:430–435. 4. Palmer, L. C., C. J. Newcomb, ., S. I. Stupp. 2008. Biomimetic systems for hydroxyapatite mineralization inspired by bone and enamel. Chem. Rev. 108:4754–4783. 5. Simmer, J. P., and A. G. Fincham. 1995. Molecular mechanisms of dental enamel formation. Crit. Rev. Oral Biol. Med. 6:84–108. 6. Fincham, A. G., J. Moradian-Oldak, and J. P. Simmer. 1999. The structural biology of the developing dental enamel matrix. J. Struct. Biol. 126:270–299. 7. Fincham, A. G., and J. Moradian-Oldak. 1995. Recent advances in amelogenin biochemistry. Connect. Tissue Res. 32:119–124. 8. Margolis, H. C., E. Beniash, and C. E. Fowler. 2006. Role of macromolecular assembly of enamel matrix proteins in enamel formation. J. Dent. Res. 85:775–793. 9. Buchko, G. W., B. J. Tarasevich, ., W. J. Shaw. 2008. A solution NMR investigation into the early events of amelogenin nanosphere self-assembly initiated with sodium chloride or calcium chloride. Biochemistry. 47:13215–13222.

20. Drobny, G. P., J. R. Long, ., P. S. Stayton. 2003. Structural studies of biomaterials using double-quantum solid-state NMR spectroscopy. Annu. Rev. Phys. Chem. 54:531–571. 21. Stayton, P. S., G. P. Drobny, ., M. Gilbert. 2003. Molecular recognition at the protein-hydroxyapatite interface. Crit. Rev. Oral Biol. Med. 14:370–376. 22. Shaw, W. J. 2015. Solid-state NMR studies of proteins immobilized on inorganic surfaces. Solid State Nucl. Magn. Reson. 70:1–14. 23. Goobes, G. 2014. Past and future solid-state NMR spectroscopy studies at the convergence point between biology and materials research. Isr. J. Chem. 54:113–124. 24. Tsai, T. W. T., and J. C. C. Chan. 2011. Recent progress in the solidstate NMR studies of biomineralization. Annu. Rep. NMR Spectrosc. 73:1–61. 25. Lu, J. X., Y. S. Xu, ., W. J. Shaw. 2013. Mineral association changes the secondary structure and dynamics of murine amelogenin. J. Dent. Res. 92:1000–1004. 26. Castellani, F., B. van Rossum, ., H. Oschkinat. 2002. Structure of a protein determined by solid-state magic-angle-spinning NMR spectroscopy. Nature. 420:98–102. 27. Buchko, G. W., and W. J. Shaw. 2015. Improved protocol to purify untagged amelogenin - application to murine amelogenin containing the equivalent P70/T point mutation observed in human amelogenesis imperfecta. Protein Expr. Purif. 105:14–22.

10. Fang, P. A., J. F. Conway, ., E. Beniash. 2011. Hierarchical self-assembly of amelogenin and the regulation of biomineralization at the nanoscale. Proc. Natl. Acad. Sci. USA. 108:14097–14102.

28. Buchko, G. W., G. Lin, ., W. J. Shaw. 2013. A solution NMR investigation into the impaired self-assembly properties of two murine amelogenins containing the point mutations T21/I or P41/T. Arch. Biochem. Biophys. 537:217–224.

11. Du, C., G. Falini, ., J. Moradian-Oldak. 2005. Supramolecular assembly of amelogenin nanospheres into birefringent microribbons. Science. 307:1450–1454.

29. Buchko, G. W., J. Bekhazi, ., W. J. Shaw. 2008. 1H, 13C, and 15N resonance assignments of murine amelogenin, an enamel biomineralization protein. Biomol. NMR Assign. 2:89–91.

12. Bromley, K. M., A. S. Kiss, ., J. Moradian-Oldak. 2011. Dissecting amelogenin protein nanospheres: characterization of metastable oligomers. J. Biol. Chem. 286:34643–34653.

30. Griffey, R. H., A. G. Redfield, ., F. W. Dahlquist. 1985. Nuclear magnetic resonance observation and dynamics of specific amide protons in T4 lysozyme. Biochemistry. 24:817–822.

13. Wiedemann-Bidlack, F. B., E. Beniash, ., H. C. Margolis. 2007. pH triggered self-assembly of native and recombinant amelogenins under physiological pH and temperature in vitro. J. Struct. Biol. 160:57–69. 14. Moradian-Oldak, J., M. L. Paine, ., M. L. Snead. 2000. Self-assembly properties of recombinant engineered amelogenin proteins analyzed by dynamic light scattering and atomic force microscopy. J. Struct. Biol. 131:27–37. 15. Moradian-Oldak, J., and M. Goldberg. 2005. Amelogenin supra-molecular assembly in vitro compared with the architecture of the forming enamel matrix. Cells Tissues Organs (Print). 181:202–218.

31. Beniash, E., J. P. Simmer, and H. C. Margolis. 2005. The effect of recombinant mouse amelogenins on the formation and organization of hydroxyapatite crystals in vitro. J. Struct. Biol. 149:182–190.

16. Moradian-Oldak, J. 2001. Amelogenins: assembly, processing and control of crystal morphology. Matrix Biol. 20:293–305. 17. Tarasevich, B. J., S. Lea, ., W. J. Shaw. 2009. Adsorption of amelogenin onto self-assembled and fluoroapatite surfaces. J. Phys. Chem. B. 113:1833–1842. 18. Tarasevich, B. J., S. Lea, ., W. J. Shaw. 2009. Changes in the quaternary structure of amelogenin when adsorbed onto surfaces. Biopolymers. 91:103–107. 19. Chen, C. L., K. M. Bromley, ., J. J. DeYoreo. 2011. In situ AFM study of amelogenin assembly and disassembly dynamics on charged surfaces provides insights on matrix protein self-assembly. J. Am. Chem. Soc. 133:17406–17413.

1672 Biophysical Journal 115, 1666–1672, November 6, 2018

32. Salomon-Ferrer, R., A. W. Go¨tz, ., R. C. Walker. 2013. Routine microsecond molecular dynamics simulations with AMBER on GPUs. 2. Explicit solvent particle mesh ewald. J. Chem. Theory Comput. 9:3878–3888. 33. Case, D. A., T. A. Darden, ., P. A. Kollman. 2012. AMBER 2012. University of California, San Francisco, CA. 34. Wishart, D. S., and B. D. Sykes. 1994. The 13C chemical-shift index: a simple method for the identification of protein secondary structure using 13C chemical-shift data. J. Biomol. NMR. 4:171–180. 35. Szilagyi, L. 1995. Chemical shifts in proteins come of age. Prog. Nucl. Magn. Reson. Spectr. 27:325–443. 36. Delak, K., C. Harcup, ., J. S. Evans. 2009. The tooth enamel protein, porcine amelogenin, is an intrinsically disordered protein with an extended molecular configuration in the monomeric form. Biochemistry. 48:2272–2281. 37. Zhang, X., B. E. Ramirez, ., T. G. Diekwisch. 2011. Amelogenin supramolecular assembly in nanospheres defined by a complex helixcoil-PPII helix 3D-structure. PLoS One. 6:e24952.