Engineering metal ion coordination to regulate amyloid fibril assembly and toxicity

Abstract

Protein and peptide assembly into amyloid has been implicated in functions that range from beneficial epigenetic controls to pathological etiologies. However, the exact structures of the assemblies that regulate biological activity remain poorly defined. We have previously used Zn²⁺ to modulate the assembly kinetics and morphology of congeners of the amyloid β peptide (Aβ) associated with Alzheimer's disease. We now reveal a correlation among Aβ-Cu²⁺ coordination, peptide self-assembly, and neuronal viability. By using the central segment of Aβ, HHQKLVFFA or Aβ(13–21), which contains residues H13 and H14 implicated in Aβ-metal ion binding, we show that Cu²⁺ forms complexes with Aβ(13–21) and its K16A mutant and that the complexes, which do not self-assemble into fibrils, have structures similar to those found for the human prion protein, PrP. N-terminal acetylation and H14A substitution, Ac-Aβ(13–21)H14A, alters metal coordination, allowing Cu²⁺ to accelerate assembly into neurotoxic fibrils. These results establish that the N-terminal region of Aβ can access different metal-ion-coordination environments and that different complexes can lead to profound changes in Aβ self-assembly kinetics, morphology, and toxicity. Related metal-ion coordination may be critical to the etiology of other neurodegenerative diseases.

Protein intermolecular assembly, especially formation of amyloid fibrillar structures, is correlated with a variety of human neurodegenerative diseases, including Alzheimer's, Parkinson's, Huntington's, and Creutzfeldt–Jakob diseases (1). More recently, amyloid has been tied to many nonpathological functional roles. For example, formation and self-perpetuation of amyloids in Saccharomyces cerevisiae regulate diverse yeast phenotypic expression as a positive response to environmental fluctuations (2), and amyloid may be involved in long-term memory and synapse maintenance in the marine snail, Aplysia (3, 4). Many proteins, including archetypical globular proteins such as myoglobin, can also form amyloid fibrils, suggesting that amyloidogenesis may be an intrinsic property of any α-amino acid polymer (5). Accordingly, these highly ordered paracrystalline protein self-assemblies have now been recognized as useful for nanostructure fabrication and biotechnology (6–8). Fully capturing these technological opportunities and understanding the biological roles of amyloid will depend on further definition of the organized structure and assembly pathway.

Increasing evidence now implicates transition metal ions, including Zn²⁺, Cu²⁺, and Fe³⁺, as contributors both to amyloid β (Aβ) assembly in vitro and to the neuropathology of Alzheimer's disease, AD (9). The obligatory region of metal ion (Zn²⁺/Cu²⁺) binding of Aβ has been mapped to the N terminus, amino acids 1–28 (10–16). In its soluble nonamyloid conformation, the peptide contains multiple intramolecular binding sites for Zn²⁺ and Cu²⁺ (9, 17), and intermolecular Zn²⁺ binding can promote Aβ aggregation (14, 18, 19). We have previously examined the role of these intermolecular Zn²⁺-binding sites in the truncated peptide HHQALVFFA-NH₂, Aβ(13–21)K16A. This peptide contains the His-13/His-14 dyad previously implicated in metal binding (14, 15, 19) and the core hydrophobic sequence, LVFFA (20, 21) that is crucial for Aβ assembly. This short peptide self-assembles in vitro into typical amyloid fibrils that are morphologically similar to the full-length Aβ peptide (22). Zn²⁺ ions accelerate the assembly by means of coordination with two imidazole side chains from different peptide molecules. These two peptide molecules could be within a single β-sheet (intrasheet coordination), or come from adjacent sheets (intersheet coordination). Different metal coordination structures result in distinct self-assembled morphologies, ranging from typical amyloid fibrils to twisted ribbons and homogeneous nanotubes (22).

Complex coordination environments for Cu²⁺ appear to exist in the longer Aβ(1–40) peptide. An intermolecular His residue-bridging binding site of Cu²⁺ in the amyloid fibril (14), similar to Zn²⁺-bridged His coordination (14, 18, 19, 22) has been indicated, whereas other results support an intramolecular Cu²⁺–Aβ complex existing in both soluble and fibrillar Aβ(1–40) (23). As both inhibitory and fibril-inducing activities have been reported for Cu²⁺ (12, 24, 25), it is very likely that the observed differences in Cu²⁺ coordination might be directly responsible for the differences in kinetics and morphologies and, further, that these different Cu²⁺–Aβ coordination structures can be accessed under slightly different experimental conditions. Indeed, Aβ(1–40) is capable of aggregating into a range of structures under slightly different assembly conditions (26). To simplify the study of Cu²⁺-induced Aβ self-assembly, we prepared a series of homogeneous Aβ(13–21) complexes with both Zn²⁺ (22) and Cu²⁺ incorporation. The structural models that emerged from visible, infrared, x-ray absorption and electron paramagnetic resonance (EPR) spectroscopies reveal a molecular basis for the self-assembly characteristics of different Aβ(13–21) peptides. These models and initial neurotoxicity results implicate a critical role of metal ions and common structural features of amyloid assemblies in a range of neurological diseases.

Results and Discussion

Zn²⁺/Cu²⁺ and the Assembly of Aβ(13–21)K16A.

We considered Aβ(13–21), HHQKLVFFA, a minimum sequence for investigating the role of metal-ion His binding in amyloid fibril formation because it contains the His-13/His-14 dyad previously implicated in metal binding (14, 15, 19) and a core hydrophobic sequence, LVFFA (20, 21), crucial for Aβ assembly. To isolate His-13/14 as the sole metal-binding site, Lys-16 was replaced with Ala, giving Aβ(13–21)K16A, which assembles into typical amyloid fibrils, following the characteristic conversion from random coil into β-strand (22). Zn²⁺ greatly accelerates the self-assembly rate and induces either typical amyloid fibrils or twisted ribbons and nanotubes depending on metal/peptide stoichiometry (22). These different morphologies arise from either intra- or intersheet His–Zn²⁺–His complexation for the fiber and ribbons/nanotubes, respectively (22). In contrast, Cu²⁺ induces no CD β-signature in this peptide over extended time periods [see supporting information (SI) Fig. 6A], even in the presence of excess Zn²⁺ (data not shown), and no aggregates were pelleted when centrifuged at 16,110 × g for 30 min (SI Fig. 6B), supporting the lack of observable self-assembly. More strikingly, Cu²⁺ rapidly disaggregates amyloid fibrils preformed by Aβ(13–21)K16A (SI Fig. 6C) but was not able to dissociate peptide assemblies preformed in the presence of equimolar Zn²⁺ (data not shown).

Aβ(13–21)K16A Cu²⁺ Coordination.

Inhibition of self-assembly in the presence of Cu²⁺ suggests that Cu²⁺ binds to the peptide. Isothermal titration calorimetry (ITC) measured the heat change upon Cu²⁺ binding and fit best to a one-site binding model, with each Cu²⁺ binding two Aβ(13–21)K16A molecules with an affinity constant (K_a) of 10⁸ M (SI Fig. 7). Displacement of water from the Cu²⁺(H₂O)₆ complex by a deprotonated amide nitrogen, amino group, imidazole nitrogen, carboxylate, and/or a water donor, causes a significant blue shift in λ_max (27) (see SI Materials and Methods). The Cu²⁺ transitions in the presence of one equivalent of Aβ(13–21)K16A in pH 5.6 buffer shifted to 600 nm (ε ≈ 40 M⁻¹ cm⁻¹), which is characteristic of three- or four-nitrogen coordination in a type (II) square-planar or distorted tetragonal complex (Fig. 1A). The four bands present in the CD spectrum (Fig. 1B), are assigned as a 264-nm transition for a strong NH₂ and/or π₂ imidazole-to-Cu²⁺ charge transfer, 300 nm as charge transfer from N⁻ to Cu²⁺, a weak 340-nm transition as charge transfer from π₁ imidazole to Cu²⁺, and a 596-nm Cotton effect for a Cu²⁺ d–d transition (28). Virtually identical Cu²⁺ ITC, UV, and CD data were obtained for wild-type Aβ(13–21) (data not shown), suggesting that Cu²⁺ adopts the same coordination with both peptides.

Fig. 1.

Characterization of Cu²⁺–Aβ(13–21) (HHQKLVFFA) congener complexes. (A and B) UV-Vis absorbance (A) and CD spectra (B) of 0.4 mM Cu²⁺ in the presence of 0.4 mM Aβ peptide. Black, Aβ(13–21)K16A; green, Aβ(Q15A); red, Aβ(H14A); and blue, Aβ(Ac-N). (C) CW-EPR spectra of the Cu²⁺–Aβ(13–21)K16A complex with different Cu²⁺-to-peptide ratios. The concentration of Aβ(13–21)K16A peptide is 0.5 mM. (D) Fourier transform (solid line) and simulation (dashed line) of three-pulse ESEEM for the soluble Cu²⁺–Aβ(13–21)K16A complex prepared from 0.4 mM Cu²⁺ in the presence of 0.5 mM Aβ(13–21)K16A.

The assignments were further tested with three fiber-forming single-point substitutions in Aβ(13–21)K16A: H14A, Q15A, and acetylation of the N-terminal amino group, Ac-N. Cu²⁺ bound and inhibited self-assembly of all three peptides (data not shown). Q15A retained the UV-Vis λ_max absorption at 600 nm (ε ≈ 40 M⁻¹ cm⁻¹) (Fig. 1A) and a CD spectrum very similar to Aβ(13–21)K16A (Fig. 1B). These results argue against involvement of Gln-15 in Cu²⁺ binding. In contrast, both H14A and Ac-N exhibited red-shifted UV maxima and decreased absorption intensity (Fig. 1A) as well as dramatic changes in the CD spectra (Fig. 1B). These results (consistent with Eq. 5 in SI Materials and Methods and SI Table 1) implicate both His-14 and the N-terminal amine as coordinating ligands.

Cu²⁺ was interrogated directly with continuous wave—EPR (CW-EPR) after titration of Aβ(13–21)K16A with CuCl₂ (Fig. 1C). Identical line shapes were observed for samples with 0.2 and 0.4 molar equivalent (eq) of Cu²⁺. At 0.8 eq, weak features characteristic of the aqueous copper complex, Cu²⁺(H₂O)₆, emerged that broadened and distorted the line shape. At 1.0 eq Cu²⁺, the m_I = −3/2 copper hyperfine feature of Cu²⁺(H₂O)₆ at 2,605 G and the narrow derivative feature of Cu²⁺(H₂O)₆ (peak, 3,200 G; trough, 3,250 G) appeared. The amplitude of the Cu²⁺(H₂O)₆ spectrum becomes dominant over the Cu²⁺–Aβ(13–21)K16A spectrum with 2 and 3 eq of Cu²⁺. These results are consistent with ITC confirming a stoichiometry of 0.5 eq of Cu²⁺ per peptide. The EPR spectrum with 0.4 eq of Cu²⁺ (Fig. 1C) best represents the line shape of the Cu²⁺–peptide complex and is characterized by g_‖ = 2.236, and a copper hyperfine splitting at a_‖ of 196 G, consistent with coordination by 2N2O or 3N1O or 4N equatorial ligands (29) (summarized in SI Table 1).

The three-pulse electron spin echo envelope modulation (ESEEM) spectrum of the Aβ(13–21)K16A complex with Cu²⁺ (Fig. 1D) is characteristic of superhyperfine coupling (shfc) between the electron on Cu²⁺ and a ¹⁴N nucleus, where the shfc and nuclear Zeeman contributions approximately cancel for one electron spin manifold (A/2 ≈ ν_N, where A is the shfc constant and ν_N is the ¹⁴N free nuclear frequency) (30, 31). This “exact cancellation” condition gives rise to the sharp lines at 0.6, 0.8, and 1.4 MHz in the ESEEM spectrum, which are assigned to the ν₀, ν₋ and ν₊ ¹⁴N nuclear quadrupole transitions, respectively (32). The broad feature at 4 MHz arises from the Δm_I = ±2 splitting in the electron spin manifold where the shfc and nuclear Zeeman contributions are additive. The pattern in the ESEEM spectrum (Fig. 1D) and the simulation values (SI Table 2) are characteristic of imidazole coordination to Cu²⁺, and arise from the uncoordinated, remote nitrogen of imidazole (30, 31, 33). These results establish that one histidine imidazole coordinates Cu²⁺ in an equatorial position.

Therefore, ITC established that Cu²⁺ binds two Aβ(13–21)K16A peptides. The CD blue shifted d–d transition and ellipticity at 596 nm strongly support Cu²⁺ coordination to a deprotonated amide nitrogen (11, 34–36). Cu²⁺ is known to chelate backbone amides, particularly if available histidine residues or an α amino group provide an anchor point for the metal ion (37). In these cases, Cu²⁺ can promote peptide nitrogen ionization at pH values as low as 5–6, even though the pK_a is 12–13 in solution. The value of Δε/n for the charge transfer between N⁻ and Cu²⁺ at 300 nm is fairly constant, in the range of 0.2–0.3, where n is the number of negatively charged peptide nitrogens involved in coordination (28). Therefore, Δε (300 nm) ≈ 0.23 (Fig. 1B) indicates Cu²⁺ coordination with one deprotonated backbone amide. Coordination to the N-terminal amine, the amide backbone, and one equatorial histidine residue requires His-13 and His-14 to form a tridentate metal complex, making structure 1 (SI Fig. 8) the most energetically accessible structure. The asymmetric α-carbon of His-14, held in the chelating ring between a main-chain amide nitrogen and a histidine imidazole ring, provides a vicinal effect (38, 39), consistent with the intense CD signal at 596 nm for the Cu²⁺ d–d transition. Finally, the conclusion from ITC that Cu²⁺ binds with two Aβ(13–21)K16A peptides indicates that the open coordination site is occupied by a ligand from the second peptide.

Design of Ac-Aβ(13–21)H14A for Cu²⁺-Induced Self-Assembly.

In contrast to Zn²⁺ (22), Cu²⁺ inhibits assembly of Aβ(13–21)K16A by deprotonating a backbone amide nitrogen and rearranging the peptide backbone to create a chelated metal complex. Even when the amino group is acetylated, as in Aβ(Ac-N), which results in loss of the amide nitrogen as a ligand, as suggested by the disappearance of the CD absorption at 600 nm (Fig. 1B and SI Table 1), amyloid fibril formation remains inhibited. In this case, Cu²⁺ appears to bind with both His-13 and His-14 intramolecularly, preventing extended β-strand formation and self-assembly (SI Table 1). Therefore, we designed a peptide Ac-Aβ(13–21)H14A, CH₃CO-HAQKLVFFA-NH₂, where removal of both His-14 and the free amino group would limit the ligands accessible to Cu²⁺ in each peptide molecule to a single His-13.

During incubation at pH 7.0, Ac-Aβ(1–40). Although the absent or present metal-ion fibrils displayed similar structure, their morphology differed. In the absence of metal ions, typical amyloid fibrils and tightly twisted fibers were apparent by atomic force microscopy (SI Fig. 9C) and transmission electron microscopy (TEM, data not shown) with diameters of 8 nm. The fibrils formed in the presence of Zn²⁺ or Cu²⁺ were both nontwisted and smooth, with diameters of 8–9 nm (SI Fig. 9 D and E).

Structural Characterization of Metal Ion-Induced Amyloid Fibrils.

Isotope-edited FT-IR (Fig. 2) was used to probe the local structures of the metal-free and metal-induced amyloid fibrils (41–45). In Ac-Aβ(13–21)H14A, ¹³CO labels were placed in the middle of the sequence, [1-¹³C]L17, or close to the C terminus, [1-¹³C]F20. In metal-free amyloid fibrils, the [1-¹³C]F20 amide stretch has an extremely weak shoulder at 1,605 cm⁻¹. The [1-¹³C]L17 peptide also has a band at 1,605 cm⁻¹, but with increased intensity, and an absorption maximum shift from 1,630 to 1,637 cm⁻¹ for ¹²CO. This shift results from disturbance of the ¹²C hydrogen-bonding network by ¹³C (46) and is not observed for [1-¹³C]F20-labeled fibrils. The stronger intensity for the [1-¹³C]L17 carbonyl carbon is consistent with Leu-17 being buried in the fibril hydrophobic core with more ordered hydrogen bonding. The isotope-edited IR spectra of fibrils formed in the absence and presence of Zn²⁺ or Cu²⁺ ions are identical in both labeling schemes. This result further confirms a very similar, if not identical, peptide conformation and packing order within the fibril β-sheets.

Fig. 2.

Isotope edited FT-IR. Absorption of amide I bands of Ac-Aβ(13–21)H14A (Ac-HAQKLVFFA) fibrils formed with free peptide (P) and in the presence of Zn²⁺ or Cu²⁺. UL, unlabeled peptide; F20, ¹³CO was introduced into the peptide at Phe-20; L17, ¹³CO was introduced into the peptide at Leu-17.

Extent of Metalation of the Ac-Aβ(13–21)H14A Fibrils.

ITC cannot deconvolute metal binding from peptide–peptide association during Ac-Aβ(13–21)H14A amyloid assembly. Therefore, amino acid analysis (AAA) and inductively coupled plasma mass spectrometry (ICP-MS) were performed to determine the metal-to-peptide ratio in the metalated Ac-Aβ(13–21)H14A fibrils (SI Table 3). Initial peptide concentration was 2 mM, with differing initial metal concentrations ranging from 0.5 to 2 mM. After maturation, fibers were pelleted, washed, and subjected to AAA and ICP-MS measurements. The Zn²⁺-to-peptide ratio varied from 0.48 to 0.64, and the Cu²⁺-to-peptide ratio varied from 0.55 to 0.73. The imprecision in incorporation ratios probably arises from nonspecifically bound metal ions. The results establish a peptide-to-metal ratio close to 2:1 in both the Zn²⁺ and Cu²⁺ assemblies.

Coordination Environment of Metal Ions in Ac-Aβ(13–21)H14A Fibrils.

To specifically characterize the Cu²⁺ coordination responsible for induction of amyloid fibrils, Cu²⁺–fibrillar complexes were pelleted, rinsed, and resuspended in fresh buffer. The three-pulse ESEEM spectrum of the Cu²⁺–fibrillar complex (Fig. 3A) shows ν₀, ν₋, and ν₊ nuclear quadrupole features at the same frequency positions, 0.6, 0.8, and 1.4 MHz, as observed for the Cu²⁺–Aβ(13–21)K16A complex (Fig. 1D). In addition, weaker peaks are present that correspond to combinations of the fundamental frequencies, including bands centered at 2.1 MHz ([ν₀+ν₊] and [ν₋ + ν₊]) and at 2.8 MHz ([ν₊ + ν₊]). Combination lines in ESEEM spectra indicate the presence of multiple nuclear couplings, and the relative amplitudes of the fundamental and combination features (47) are consistent with two coupled ¹⁴N nuclei, with e²qQ/h = 1.55 MHz and η = 0.751 (SI Table 2). The simulation parameters and spectral pattern, including the enhanced amplitude of the Δm_I = ±2 feature at 4 MHz, demonstrate coupling of Cu²⁺ to two remote ¹⁴N atoms of distinct histidine imidazoles, establishing that two histidine imidazoles are coordinated equatorially to Cu²⁺ in the fibrillar complex.

Fig. 3.

Copper ion analysis in the Ac-Aβ(13–21)H14A (Ac-HAQKLVFFA) fibrillar complex. (A) Fourier transform (solid line) and simulation (dashed line) of three-pulse ESEEM for Cu²⁺-Ac-Aβ(13–21)H14A fibrils. Simulation values are summarized in SI Table 2. (B) The Fourier transform of extended x-ray absorption fine-structure (EXAFS) (Inset) of Cu²⁺-Ac-Aβ(13–21)H14A fibrils. Solid line, experiment; dashed line, fit 8 (values listed in SI Table 4).

Following previously developed procedures for probing Zn²⁺ coordination environments (22), Ac-Aβ(13–21)H14A fibrils were characterized by extended x-ray absorption fine structure (EXAFS) spectroscopy. Curve fitting indicates 3N/1O or 2N/2O or 1N/3O atoms in the first shell (SI Table 4). The coincident appearance and intensities of the 3-Å and 4-Å peaks in the Fourier transform EXAFS spectra [Fig. 3B for Cu²⁺ and ref. 22 for Zn²⁺ Ac-Aβ(13–21)H14A fibrillar complexes] are diagnostic of two imidazole ligands. The best fit (dashed line in Fig. 3B) was generated by using two His imidazoles and two additional first-shell light atoms (N or O) (SI Table 4, Fit 8). Therefore, both Zn²⁺ and Cu²⁺ induce Ac-Aβ(13–21)H14A fibril formation by bridging two His-13 residues between parallel hydrogen-bonded β-strands in a His–metal²⁺–His chelated structure.

Toxicity of Cu²⁺–Peptide Complexes.

Initial neurotoxicity assays in cultured neurons suggested that the two Cu²⁺–peptide coordination environments [Cu²⁺–Aβ(13–21)K16A complex and Cu²⁺–Ac-Aβ(13–21)H14A fibrils] were biologically distinct (data not shown). To more specifically characterize the activity, a myocyte enhancer factor 2 (MEF2) assay was developed in SN4741, a dopaminergic neuron cell line derived from the mouse midbrain. The MEF2 family of transcription factors play critical roles in diverse cellular processes including neuronal survival (48). MEF2s are an endpoint for several neurotoxic signaling pathways that control the molecular machinery of cellular apoptosis (49, 50). As shown in Fig. 4Left, Aβ(1–42) regulates MEF2. In addition, inhibition of MEF2 activity by mature peptide assemblies is dose-dependent and peptide-specific. The Cu²⁺–Ac-Aβ(13–21)H14A fibrils are equally as inhibitory as Aβ(1–42), whereas the Cu²⁺-free Ac-Aβ(13–21)H14A fibrils and Cu²⁺ ions alone are ineffective. The soluble Cu²⁺–Aβ(13–21)K16A complex is also nontoxic, suggesting that different Cu²⁺ chelation environments, as mediated by different Aβ conformations, compromise neuron viability.

Fig. 4.

MEF2 luciferase reporter gene assay. (Left) Aβ(1–42) dose-dependent response. (Right) Cu²⁺–Ac-Aβ(13–21)H14A fibrils are as toxic as Aβ(1–42) and Aβ(10–35) fibrils and Cu²⁺–Aβ(13–21)K16A complex is nontoxic. Peptide concentration is 10 μM.

Summary.

Metal ion association with Aβ has frequently been correlated with Alzheimer's disease (11, 35, 51, 52), and the link among metal-coordination structure, the impact on assembly kinetics, and overall aggregation remains critical to understanding disease etiology. Our attempts to identify individual metal sites within Aβ segments now reveal that Cu²⁺ coordination can radically alter assembly kinetics and morphology. In the short Aβ fragment Aβ(13–21)K16A, backbone deprotonation occurs even at low pH to generate a high affinity (K_a = 10⁸ M) Cu²⁺-binding site. The five- {NH₂, N_amide⁻} and six-membered {N_amide⁻, N_Im} metal chelate in structure 1 (SI Fig. 8) coordinates with a second peptide, probably through a histidine residue. A virtually identical {NH₂, N_amide⁻, N_Im} complex was previously observed for several short peptides that contain His as the second residue from the peptide N terminus, such as X-His (X = Ala, Gly, His) or X-His-Y (X = Ala, Gly, and Y = Ala, Leu, Lys) (53–56). In Aβ(13–21)K16A, this X-His (X = His) motif retards β-strand and β-sheet assembly, but this inhibition may be unique to Cu²⁺ because Zn²⁺ readily promotes amyloid self-assembly (22). Even with acetylation of the N terminus as in Ac-Aβ(13–21)K16A, Cu²⁺ coordinates with both His-13 and His-14 intramolecularly to retard peptide assembly. This inhibition of assembly was previously observed for Ac-Aβ(11-X) (X = 16, 20, 28) (57, 58), probably through a similar complex.

Removing one histidine, as in Ac-Aβ(13–21)H14A, generates a peptide in which both Zn²⁺ and Cu²⁺ accelerate amyloid fibril formation by capturing intermolecular His–metal–His coordination (Fig. 5C). The isotope-edited FT-IR results predict a parallel, in-register β-sheet organization, an assignment further supported by preliminary solid-state NMR results (data not shown). The same conformation was previously determined for the well characterized Aβ(10–35) peptide assembled in the absence of metal ions (21, 59). Here, both Zn²⁺ and Cu²⁺ associate with two His-13 residues from adjacent peptides arranged along the growing β-sheet surface (22), increasing the growth rate and stability of the growing fibril. Stoichiometry analyses are consistent with the organized linear array of metal ions along the fibril surface (Fig. 5C). Therefore, the ability of Cu²⁺ to access the intermolecular coordination environment depends on subtle changes in the intramolecular site for Cu²⁺.

Fig. 5.

Structural models for the Cu²⁺–peptide complexes. (A) Proposed model for Cu²⁺ coordination with Aβ(13–21)K16A showing only the first four residues (HHQA). (B) Crystal structure for Cu²⁺–HGGGW complex, the N-terminal repeat sequence of human prion protein (63). Only equatorial ligands are illustrated in both models. Purple, Cu ion; green, carbon; red, oxygen; blue, nitrogen; gray, unidentified ligand. (C) Structural models for Cu²⁺ arrangement in Ac-Aβ(13–21)H14A fibrils. (Left) View of three stacked β-sheets. (Right) View down peptide backbone. H-bonds are parallel to fibril long axis, and fibrils consist of parallel in-register β-sheets in both the absence and presence of metal ions. Sheets stack perpendicular to the H-bond direction. Both Zn²⁺ and Cu²⁺ coordinate two histidines along the H-bonding dimension within the same β-sheet. Red, His-13; blue, remaining Ac-Aβ(13–21)H14A residues; magenta, Zn²⁺, Cu²⁺; and green, H-bonds between backbone carbonyl and amide. Amino acid side chains, except His-13, are removed for clarity.

In this regard, Aβ(1–40) are not included in the core β-sheet of the fibrils but, rather, are structurally disordered and exposed along the fibril surface (60). Intramolecular metal complexation close to this structurally flexible region may slow assembly (24), but, as observed in these short peptides, does not block assembly.

“Switching” to achieve different Cu²⁺-binding sites and peptide conformations has also been considered for the mammalian prion protein (PrP). Cu²⁺ binds primarily to the N terminus of PrP, a region that is structurally flexible and outside of the core assembly region (61). Therefore, metal-binding regions in both Aβ and PrP are in the structurally flexible N-terminal domain and lie outside the amyloid-determining core (21, 62). The N terminus of PrP contains four copies of the Cu²⁺-binding octarepeat sequence, PHGGGWGQ. In the presence of two or more molar equivalents of Cu²⁺, the metal ions bind in an intrarepeat manner, with ligands supplied by one histidine imidazole, two deprotonated amides from the next two glycines, and the amide carbonyl of the second glycine (63) (Fig. 5B). At low Cu²⁺ concentrations or under acidic pH conditions, multiple His imidazoles from different octarepeats coordinate with Cu²⁺, forming interrepeat binding sites (64, 65). These interrepeat sites could be intramolecular, intermolecular, or a combination (65).

Moreover, several proteins associated with neurodegenerative diseases also bind metal ions, and metal binding modulates the aggregation behavior of these proteins. α-Synuclein (α-Syn), a protein associated with Parkinson's disease (PD), is intrinsically unfolded and Cu²⁺ binding results in rapid aggregation (66). Superoxide dismutase 1 (SOD1), a CuZn metalloprotein that catalyzes the dismutation of the superoxide anion, is associated with the progressive neurodegenerative disease amyotrophic lateral sclerosis (ALS). Cu²⁺ and Zn²⁺ bind and organize a loop region of SOD1, and mutations around the binding site that result in alterations of the coordination environment of Cu²⁺ and Zn²⁺ lead to misfolding and aggregation of SOD1 (67, 68). The oxidative modification and aggregation of SOD1 has been correlated with both AD and PD (69). Thus, it seems reasonable to speculate that AD, PD, and ALS share common mechanisms of metal ion regulation and/or metal-ion induction of protein aggregation, which might lead to a common, or at least overlapping, pathogenic mechanism(s).

Our studies now establish that the N-terminal region of Aβ can access a range of metal-coordination structures. These results suggest that the effect of metal ions in neurodegenerative diseases is not simply a result of metal ion on/off binding, but is due to switching between different metal-binding modes. Clearly, the short peptides allow these structures to be constructed and evaluated directly, whereas in the larger Aβ peptides, the accessible conformations and subtle environmental conditions can alter ligand accessibility and modulate both amyloid assembly and cellular toxicity. The striking quasilinear array of Cu²⁺ ions along the surface of the biologically active Ac-Aβ(13–21)H14A fibril suggests that assembly within a complex cellular milieu will certainly produce novel protein architectures, architectures that can now be constructed and evaluated directly for functional neuron toxicity and prion function.

Materials and Methods

Fibril Formation.

Each Aβ(13–21) congener peptide was dissolved completely in distilled deionized H₂O, sonicated for 10 min, and centrifuged at 16,110 × g for 10 min. The supernatant was used as stock solution for each peptide. Samples with desired peptide and metal concentration in 25 mM Mes buffer (pH 5.6) or Hepes buffer (pH 7.2) and 10 mM NaCl were prepared from stock solutions.

Spectroscopy.

UV-Vis absorption spectra were obtained on a V-530 UV/VIS spectrometer (Jasco, Easton, MD) with a 1-cm path-length quartz cuvette. CD spectra were recorded on a Jasco 810 CD spectropolarimeter. Ellipticity (θ, m degrees) was converted to mean residue molar ellipticity (MRME) [θ] = θ/(10 × n × C × l) or to molar ellipticity Δε = θ/(32980 × C × l), where n is number of amide bonds per peptide, C is molar concentration (mol/liter), and l is path length (cm). FT-IR spectra were collected on a Magna 560 IR spectrometer (Nicolet, Madison, WI). Mature fibrils were spun down, lyophilized, mixed, and ground with KBr and pressed into transparent disks. Typically, 100 scans were averaged with 4-cm⁻¹ resolution. X-ray absorption spectra were collected at the Stanford Synchrotron Radiation Laboratory on beamline 9-3. Mature fibrils were spun down and washed with fresh buffer solution three times. The hydrated pellet was transferred to a 25-well sample holder, frozen, and stored in liquid nitrogen until data collection. See SI Materials and Methods for instrumental setup and data analysis.

EPR.

Samples for both EPR and ESEEM were mixed with an equal volume of ethylene glycol, transferred to 4 mm o.d. quartz EPR tubes, frozen in liquid nitrogen-chilled 2-methylbutane (≈150 K), and stored in liquid nitrogen. Continuous-wave EPR spectra were collected on a ER200D EPR spectrometer (Bruker, Billerica, MA). ESEEM spectra were collected on a home-built pulsed-EPR spectrometer by using the three-pulse stimulated echo pulse sequence. Details of EPR and ESEEM experiments and simulations (70, 71) are described in SI Materials and Methods.

Microscopy.

AFM samples were placed on silicon chips and imaged under dry conditions in tapping mode on a JSPM-4210 AFM (JEOL, Tokyo, Japan) by using ultrasharp noncontact silicon cantilevers with typical frequencies from 240 to 350 kHz.

MEF2 Assay.

SN4741 cells were transiently transfected with a DNA construct containing a luciferase gene under control of MEF2 enhancer by using the Lipofectamine 2000 transfection system (Invitrogen, Carlsbad, CA). Total transfected DNA was kept constant. After 24 h of transfection, cells were treated with Aβ peptides that had been assembled as described above. Twenty-four hours after Aβ treatment, cell lysates were analyzed for luciferase activity by using Luciferase reporter gene assay kit (Roche, Mannheim, Germany).

Abbreviations

Aβ(13–21)K16A

HHQALVFFA-NH₂

Ac-Aβ(13–21)H14A

CH₃CO-HAQKLVFFA-NH₂

ESEEM

electron spin echo envelope modulation

shfc

superhyperfine coupling.