Characterizing the Effect of Multivalent Conjugates Composed of AÎ²

Characterizing the Effect of Multivalent Conjugates Composed of Aβ-Specific Ligands and Metal Nanoparticles on Neurotoxic Fibrillar Aggregation Carmen Streich,†,△ Laura Akkari,‡,△ Christina Decker,§ Jenny Bormann,# Christoph Rehbock,† Andreas Müller-Schiffmann,⊥ Felix Carlsson Niemeyer,‡ Luitgard Nagel-Steger,§,¶ Dieter Willbold,§,¶ Barbara Sacca,∥ Carsten Korth,⊥ Thomas Schrader,*,‡ and Stephan Barcikowski*,† †

Technical Chemistry I and Center for Nanointegration Duisburg-Essen (CENIDE), University of Duisburg-Essen, 45141 Essen, Germany ‡ Institute for Organic Chemistry, University of Duisburg-Essen, 45117 Essen, Germany § Institut für Physikalische Biologie, Heinrich-Heine-University Düsseldorf, 40225 Düsseldorf, Germany # Chemical Biology, Zentrum für Medizinische Biotechnologie (ZMB), Fakultät für Biologie, Universtität Duisburg-Essen, 45117 Essen, Germany ∥ Bionanotechnology, Zentrum für Medizinische Biotechnologie (ZMB), Fakultät für Biologie, University of Duisburg-Essen, 45117 Essen, Germany ⊥ Department Neuropathology, University of Düsseldorf Medical School, 40225 Düsseldorf, Germany ¶ Institute of Complex Systems, Structural Biochemistry (ICS-6), Research Centre Jülich, 52425 Jülich, Germany S Supporting Information *

ABSTRACT: Therapeutically active small molecules represent promising nonimmunogenic alternatives to antibodies for specifically targeting diseaserelevant receptors. However, a potential drawback compared to antibody− antigen interactions may be the lower affinity of small molecules toward receptors. Here, we overcome this low-affinity problem by coating the surface of nanoparticles (NPs) with multiple ligands. Specifically, we explored the use of gold and platinum nanoparticles to increase the binding affinity of Aβspecific small molecules to inhibit Aβ peptide aggregation into fibrils in vitro. The interactions of bare NPs, free ligands, and NP-bound ligands with Aβ are comprehensively studied via physicochemical methods (spectroscopy, microscopy, immunologic tests) and cell assays. Reduction of thioflavin T fluorescence, as an indicator for β-sheet content, and inhibition of cellular Aβ excretion are even more effective with NP-bound ligands than with the free ligands. The results from this study may have implications in the development of therapeutics for treating Alzheimer’s disease. KEYWORDS: colloidal gold, cooperativity, multivalence, nanomedicine, misfolded proteins, protein fibrillization, neurodegenerative diseases

A

oligonucleotides, but they are prone to degradation in serum by nucleases if not further modified and require time- and laborconsuming synthesis. Small molecules represent promising nonimmunogenic alternatives for targeting at the expense of lower target-binding affinities.

ctive targeting represents an elegant way of drug delivery because it may greatly enhance the therapeutic efficacy of drugs by increasing drug concentrations at the site of the disease and reduce adverse effects by sparing healthy tissue exposure. Requirements for active targeting are high specificity and affinity of the ligands such as antibodies1 and aptamers.2 However, antibodies are immunogenic, and their large size may limit bioavailability or prevent access to biological compartments. Aptamers are small single-stranded © 2016 American Chemical Society

Received: April 19, 2016 Accepted: July 12, 2016 Published: July 12, 2016 7582

DOI: 10.1021/acsnano.6b02627 ACS Nano 2016, 10, 7582−7597

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Here we investigate the interactions of bare NPs, free ligands, and NP-bound ligands on fibrillary Aβ aggregation. From the two major species Aβ1−40 and Aβ1−42, Aβ1−42 was employed as model peptide since it is the predominant species in neuritic plaques and shows a higher aggregation propensity.23 Pulsed laser ablation in liquid24,30 was employed to generate colloidal NPs from bulk metal targets. Since this nanoparticle fabrication procedure excludes the use of any artificial stabilizers, colloids are initially ligand-free and feature a high purity. Especially, small AuNPs were chosen as a nanoparticle platform due to their unique properties, including high biocompatibility, colloidal stability, good control over particle size, and surface plasmon resonance.30 Importantly, AuNPs can be easily surface-modified via covalent thiol bonds with functional molecules. As ligand, the Aβ-specific D3 dodecapeptide was immobilized on AuNPs. D3 is a protease-resistant, Denantiomeric peptide that was identified as an Aβ-binder through mirror-image phage display. Affinities of 1.1 and 0.12 μM (Kd) toward two different binding sites within Aβ25 have been reported, and D3 was shown to inhibit formation of fibrils and reduce Aβ-induced cytotoxicity in vitro. Furthermore, when injected into brains or orally administered to mice, D3 reduced plaque load as well as inflammation and improved cognitive function,26,27 indicating that the ligand can cross the blood− brain barrier. For this project, D3 was equipped with a C-terminal cysteine for Au immobilization and an increasing number of cationic residues for varied net charge. The interaction of the loaded nanocarrier with the Aβ peptide was analyzed as a function of ligand modification (ligand net charge) and ligand load per nanoparticle. To differentiate between effects of pure nanoparticles and pure ligands, reference samples of bare nanoparticles and free ligands were employed. The binding event of Aβ on D3-coated nanoparticles was characterized with physicochemical methods, and the ability of nanoconjugates to interfere with the aggregation of synthetic Aβ was assessed by biophysical assays. Finally, ligand-coated nanoparticles were incubated with cells naturally secreting neurotoxic Aβ species, and the biological effect was monitored.28,29

In nature, the lack of affinity and selectivity of free ligands is overcome by multivalency, i.e., simultaneous binding of multiple ligands to multiple receptors, creating interactions much stronger than corresponding monovalent systems due to a “collective” affinity (avidity).3 Examples of multivalent systems include the attachment of viruses to host cells or the binding of circulating neutrophil cells to vascular endothelium cells upon inflammation.4 Interestingly, nanoparticles have a high surface area to volume ratio, and because of this, the surface can be coated with multiple ligands to generate multivalency. For example, Shaw and colleagues characterized a library of iron oxide nanoparticles conjugated with small molecules that had a 4 orders of magnitude enhancement of binding affinity to the target.5 Hong et al. performed a study on the avidity of folate-coated dendrimers targeting the folate receptor overexpressed in epithelial cancer cells. They showed that affinity could be drastically increased up to 170 000-fold as compared to the free compound, by increasing the number of folate molecules bound per dendrimer. Moreover, they attributed the improvement of in vivo efficacy by this nanocarrier over free folate to the multivalent interaction.6 Intracellular gene regulation with oligonucleotide-modified gold nanoparticles (AuNPs) was studied by Rosi et al. They found elevated affinities toward complementary nucleic acids for oligonucleotide-modified nanoparticles compared to free oligonucleotides. Interestingly, particles with higher affinities also reduced target gene expression more effectively.7 Bowman et al. showed that the immobilization of a weakly binding, biologically inactive small molecule on AuNP results in a multivalent conjugate that effectively inhibits HIV-1 fusion to human T cells.8 Here we explore the concept of nanoparticle-based multivalency for use in the field of neurodegenerative diseases, which are characterized by protein misfolding and aggregation. As the most prominent example, the amyloid β-peptide (Aβ) can play a central role in the development and progression of Alzheimer’s disease (AD).9 Senile plaques composed of aggregated Aβ species are histopathological hallmarks found in the brains of AD patients. More recently, small soluble Aβ oligomers were suggested as major neurotoxic species.10 To date only symptom-relieving drugs are clinically approved for AD treatment. However, agents targeting Aβ and thus interfering with the disease mechanism would be desirable. Research approaches include the reduction of Aβ production a priori, the increase of Aβ removal via anti-Aβ immunotherapy, and direct interference with Aβ aggregation.11 The latter can be achieved with small molecules, peptides, and/or inorganic nanoparticles. With regard to the effect of nanoparticles themselves, physicochemical properties such as particle size, charge, shape, and surface modification including stabilizing ligands such as dihydrolipoic acid12 and the protein corona13 determine their inhibitory or acceleratory effect on Aβ fibrillization. Wu and colleagues report a promoting effect of TiO2 nanoparticles on Aβ fibril formation.14 In contrast, negatively charged AuNPs have been shown to inhibit fibrillization and to induce fibril dissociation.15 Tailored surface modifications of nanoparticles to achieve a specific Aβ targeting have also been described. Thus, nanoparticles have been functionalized with phosphatidic acid,16 curcumin,17 Aβ-oligomer-specific antibody NU4,18 and Aβ-selective LPFFD,19,20 LVFFARK,21 or D1 peptide,22 respectively.

RESULTS AND DISCUSSION Characterization of Laser-Generated, Ligand-Free Nanoparticles. Colloidal nanoparticles were generated via pulsed laser ablation in aqueous solution from a bulk metal target without the use of any artificial stabilizers.30,33 The fabrication method was chosen since laser-generated, ligandfree particles feature high purities, high biocompatibilities, and a readily accessible surface for functionalization.30 After laser synthesis nanoparticles feature average diameters of 11 ± 4 nm. Through centrifugation the width of the size distribution could be reduced, resulting in overall monodisperse particles of 7 ± 1 nm in size (Figures S1 and S3). Typical working concentrations were 50 μg colloid/mL, resulting in number concentrations of 1.44 × 1013 NP/mL colloid and surface concentrations of 22.2 cm2/mL colloid. Due to partial surface oxidation and anion adsorption,31 the laser-generated colloid features a net charge of −42 mV (Figure S9). Long-term measurements revealed that the particle size and concentration remained constant for at least 50 d, underlining the high colloidal stability (Figures S1c, S4). Moreover, the colloid was highly stable over a wide range of pH (pHIEP = 2), with highest stability at neutral to alkaline pH,32 and stable at low-mM sodium chloride concentrations (max. tolerable ionic strength was 50 mM, corresponding to 2.1 7583


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Figure 1. Conjugation of AuNP with different variants of the D3 ligand. The ligand net charge varies from +5 to +10. Colloidal stability upon ligand addition was assessed as a function of ligand to nanoparticle ratio via zeta potential measurements and UV/vis extinction spectroscopy (a). Stable conjugates can be fabricated by dosing ligands by avoiding the regime of zero net charge (red boxes) to create electrosterically stabilized, overall anionic or cationic conjugates. Conjugation of ligand-free nanoparticles (TEM image, top of (b)) with a tryptophan-labeled modification of Trp-D3-Cys (5+) allowed the quantification of conjugation efficiency (black) and ligand surface density expressed as number of bound ligands per nanoparticle (green, bottom of (b)). The dashed green line indicates maximum surface density. Note that conjugates for functionality tests were typically conjugated with 418 ligands per NP, of which approximately 125 are bound to the surface.

× 106 NaCl/NP, Figure S1d and e). When conjugated with the stabilizing ligand mPEG-SH, the colloid could even be frozen and thawed several times or freeze-dried and resuspended without impairing colloidal stability (Figures S5 and S6). Synthesis and Characterization of AuNP−Peptide Conjugates. Gold colloids were covalently conjugated with the Aβ-targeting D3 ligand, by including a terminal cysteine residue in the ligand structure. Three D3 derivatives with increasing positive net charge were prepared in order to explore their influence on nanoparticle stability. To this end, three or five lysine residues were inserted between the N-terminal dodecapeptide and the C-terminal cysteine. Peptides were synthesized via Fmoc strategy on cysteine-preloaded Wang resin, by automated SPPS with microwave technology. The free peptides were all characterized by 1H NMR spectroscopy, HPLC, and MS (Figures S11−S14).

conjugating the cationic peptides to the anionic AuNP, conjugate net charge was reversed from negative to positive charge with increasing ligand to nanoparticle ratio.33 As observed previously for conjugates of cell-penetrating peptides and AuNP, each conjugate has a specific ligand to nanoparticle ratio at which the isoelectric point is reached and the surface plasmon resonance (SPR) peak is maximally red-shifted.34 At this point, particles precipitate due to charge compensation and subsequent aggregation. However, it should be noted that there are two concentration regimes of high colloidal stability in which either the charge of the nanoparticle or the charge of the ligand molecules dominates and prevents nanoparticles from precipitating (Figure 1a). Furthermore, by increasing the ligand charge through the addition of cationic amino acids, the isoelectric point is shifted toward lower ligand concentrations from D3_5+ to D3_8+ and D3_10+. For further analysis, D3_5+ was employed as standard ligand because its structure is most similar to the D-enantiomeric amino acid peptide D3. Earlier experiments showed a reduction in plaque load and inflammation and improvement of cognitive function in an AD transgenic mouse model when treated with D3.27 The ligand density per nanoparticle was determined by separating unbound ligands from conjugates through centrifugation and quantifying ligand concentrations in the supernatant. The quantification of bound Trp-D3-Cys per nanoparticle shows that the maximum number of ligands per particle is approximately 164 ± 12 (Figure 1b), corresponding to 177 ± 13 pmol ligands per cm2 colloid. This finding is comparable to that of Petersen et al., who studied the conjugation of oligonucleotides to laser-generated AuNPs and found a maximum ligand load of 164 ± 7 pmol/cm2.35 Compared to chemically synthesized particles, the loading is notably higher. For instance, the nanoconjugates of Gao et al. carried only 10

The above scheme depicts the three selected D3 derivatives with C-terminally appended −Cys, −(Lys)3-Cys, and −(Lys)5Cys, differing in length and cationic charge. All basic residues are written in blue. Below, the sequence of Aβ1−42 is shown, indicating again cationic residues in blue (R-5, K-16, K-28), anionic ones in red (E-22, D-23), and hydrophobic areas in black (central hydrophobic cluster, C-terminal nucleation site). Colloidal stability was analyzed as a function of ligand to nanoparticle ratio via UV/vis extinction spectroscopy and zeta potential measurements (Figures 1, S8, and S10). By 7584


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Figure 2. Titration of Aβ to D3-functionalized nanoparticles changes zeta potentials and number-weighted hydrodynamic diameters of AuNPs (a). The green box indicates the sample employed in (b). Interaction of ligand-free and D3-functionalized nanoparticles with the Alzheimer peptide Aβ, as determined by zeta potential and dynamic light scattering experiments (b). Fluorescence quenching of FITC-labeled Aβ (c). Titration of D3-functionalized gold (top) and platinum nanoparticles (bottom) to Aβ-FITC and compared to the effects of bare nanoparticles and free D3. The applied ligand to NP ratio was 125:1, and the Aβ-FITC concentration was 0.125 μM.

Table 1. Dissociation Constants and Cooperativity Factors for the Interaction of NP/D3 Conjugates, Bare NP, or Free D3 with FITC-Labeled Aβa

D3_5+ with Aβ-FITC Au/D3_5+ with AβFITC Au with Aβ-FITC Pt/D3_5+ with AβFITC Pt with Aβ-FITC

dissociation constant, KD,Lig [nmol/L, applied D3]

dissociation constant, KD,NP [nmol/L, applied NP]

cooperativity factor, n

correl. coefficient, R2 (Hill fit)

1906 ± 587 271 ± 25

2.17 ± 0.20

0.63 ± 0.07 2.32 ± 0.42

0.97 0.98

973 ± 148

2.05 ± 0.11 7.78 ± 1.81

5.24 ± 2.01 1.36 ± 0.33

0.99 0.96

27.2 ± 6.0

1.70 ± 0.42

0.96

Dissociation constants and cooperativity factors were derived from Hill fitting of fluorescence quenching data. Ligands were present on the nanoparticle surface at a ratio of 125:1. a

Aβ-inhibiting LPFFD−peptide ligands per AuNP at even larger particle diameters (d = 22 nm) than the particles presented here.20 As can be seen from Figure 1b, although the number of total bound ligands per nanoparticle increases by increasing the ligand dose, conjugation becomes less efficient. This indicates that electrosteric ligand−ligand interactions may occur, limiting the nanoparticle coverage with ligand until monolayer coverage is reached.34 The applied ligand dose in further experiments was 418 ligands per NP to ensure maximum coverage of the NP surface. However, at this high ligand dose, some of the ligands may be free in solution. To better differentiate between the

effect of free and NP-immobilized ligands, the ratio was reduced to 125 ligands per NP when indicated. Physicochemical Interaction with Synthetic Aβ 1−42 Species. After having characterized how cationic peptide ligands affect nanoparticle stability and how ligand load can be adjusted, the interaction with Aβ was assessed. As a first measure, changes in the physicochemical properties of AuNP/ D3_5+ conjugates upon Aβ incubation were analyzed. For this purpose, bare nanoparticles and conjugates were incubated with 10 μM monomerized Aβ. As observed from TEM, interparticle distances were altered in the presence of Aβ compared to the Aβ-free reference, indicating that the peptide accumulates in 7585


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Figure 3. Analysis of the effect of pure nanoparticles, pure ligands, and ligand-coated nanoparticles on the molecular weight distribution of aggregated synthetic Aβ. The procedure includes incubation of samples with Aβ (a, left), density gradient centrifugation (a, middle), and immunologic detection of different Aβ species (a, right). Quantification of Aβ was performed with an Aβ-specific ELISA in each of the 23 fractions of the density gradient (from fraction 1/top to 23/bottom of the tube) after incubation periods of 90 min (b) and 24 h (c) at 37 °C, respectively. Note that Aβ species are present in all fractions after 90 min incubation, while the major portion of Aβ is present in fractions 17− 20 after 24 h incubation. Reductions of distinct Aβ species by free ligands and ligand-coated NPs after 90 min and 24 h are marked with arrows. Gold concentrations were 600 μg/mL in (b) and 200 μg/mL in (c), D3 concentration was 40 μM, and Aβ concentration was 40 μM (corresponding to the monomer). Each data point is an average of four samples ± SD.

between the conjugates (Figure S7). Furthermore, increased hydrodynamic sizes (dynamic light scattering, DLS) and reduced overall electrophoretic mobilities (zeta potential) suggest that Aβ attaches to the conjugate surface. When titrated with different amounts of Aβ, an Aβ-concentrationdependent increase in NP size and a decrease in zeta potential can be observed (Figure 2a). Importantly, Aβ also modified the properties of ligand-free reference particles (Figure 2b). Thus, unspecific interactions between the ligand-free nanoparticle and Aβ may have led to the formation of an Aβ “protein corona”, which is a phenomenon well-described in the literature.36,13 With regard to Aβ-affinity, dissociation constants between Aβ and bare NP, free ligands, and ligand-coated nanoparticles were derived from titration experiments (Figure 2c). Different concentrations of NP, ligand, or NP−ligand conjugates were titrated into a stock solution of fluorophore-labeled Aβ. A decrease in fluorescence was interpreted as quenching due to the formation of complexes between Aβ-FITC and NP, D3, or AuNP/D3 conjugates. The results were fitted with the Hill function as described in the Methods section, and dissociation constants with regard to D3 and the nanoparticles were derived, respectively. Dissociation constants for the free ligand were found to be in the low-micromolar range, comparable to those results obtained for the unmodified D3 via SPR measurements.37,25 With regard to the ligand, the dissociation constant of AuNP-immobilized D3 appears 7 times lower than that of the free ligand (Table 1). Notably, also bare AuNPs strongly quench the fluorescence of Aβ-FITC. The resulting dissociation constant in the nanomolar range is almost identical to the dissociation constant achieved with D3-coated AuNPs. Two simultaneous effects may contribute to the observed quenching effects of AuNPs. First, a distance-dependent quenching of fluorophores in the vicinity of plasmonic NPs

based on radiative and nonradiative effects has been described, and reduction of Aβ-FITC fluorescence can thus indicate the binding of the labeled Aβ to the NP surface.38 Second, especially for the case of FITC, its fluorescence emission (520 nm) overlaps with the surface plasmon resonance peak of AuNP (517 nm). Hence, a significant amount of energy can be reabsorbed by the particle (energy transfer quenching), which in turn reduces the quantum yield of the fluorophore.39 To evaluate nanoparticle quenching independent of plasmonic effects, additional experiments were performed with laser-generated platinum nanoparticles (PtNPs) and PtNP/D3_5+ conjugates. As shown in Figure S9, lasergenerated PtNPs feature a similar size and a negative zeta potential comparable to AuNPs. Furthermore, the addition of thiolated D3 results in Pt/D3 conjugates with overall negative or positive charge, dependent on the ligand to NP ratio (Figure S10). In contrast to AuNPs, PtNPs do not feature a surface plasmon resonance peak in the UV/vis range (Figure S9c). Thus, the dominant quenching may be “aggregation-induced quenching”,40 if high local concentrations of Aβ-FITC adsorb on the particle surface. As shown in Figure 2 and Table 1, bare PtNPs quench the fluorescence of Aβ-FITC less efficiently than AuNPs possibly due to the missing absorbance/emission spectral overlap and the absence of energy-transfer quenching. However, upon functionalization with the D3_5+ ligand, quenching becomes more efficient and the dissociation constant decreases by a factor of 3.5 compared to bare PtNP. Since the only difference in the two samples is the presence/ absence of D3, the increased affinity must result from the ligand coating. With respect to the ligand dissociation constant, the value is even lower for PtNP-immobilized D3 than for the free ligand, indicating that NP immobilization may induce 7586


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Figure 4. Atomic force micrographs of preformed synthetic Aβ fibrils after incubation with pure nanoparticles, pure ligands, and ligand-coated nanoparticles directly after addition of samples, after 24 h, and after 7 d. Gold concentration was 125 μg/mL, D3 concentration 25 μM, and Aβ concentration 50 μM (corresponding to the monomer).

Kogan et al. showed that ligand functionalization with the CLPFFD peptide increased the adhesion of nanoparticles to Aβ fibrils. Compared to unfunctionalized AuNPs, peptide-coated AuNPs were attached at 4 times higher concentrations to the fibrils compared to unfunctionalized NPs, as analyzed by separation of NP-bound fibrils and free AuNP through centrifugation and subsequently analyzing the gold content in the supernatant and pellet.42 In summary, the results indicate that although unspecific Aβ binding to bare nanoparticles occurs, the affinity of bare nanoparticles for Aβ can be increased through the immobilization of D3 on their surface. With regard to the ligand functionality, the immobilization on the nanoparticle surface does not impair but rather enhances their molecular recognition of Aβ. Density Gradient Centrifugation−ELISA Assays. Density gradient centrifugation (DGC) experiments followed by evaluation through reversed-phase high-performance liquid chromatography (RP-HPLC) allow the quantitative monitoring of a molecular weight distribution that is produced by the Aβ aggregation process.43 Here, DGC is combined with ELISA to

cooperative effects leading to increased affinities of the individual ligands. With regard to cooperative effects, the cooperativity factor derived from the Hill fit indicates whether Aβ moieties already adsorbed on the particle/ligand sample promote (n > 1) or inhibit (n < 1) further Aβ adsorption. For samples containing NPs, positive cooperativity (n > 1) can be derived from the Hill fit, while negative cooperativity (n < 1) is observed for the free ligand. Hence, NP/D3-bound Aβ seems to increase the binding strength as further Aβ adsorbs, possibly due to synergistic effects of multiple ligands on the nanoparticle surface. In contrast, the affinity for Aβ progressively decreases once that Aβ is bound to the free ligand, possible due to saturation of the ligand’s binding site. Douglas et al. have analyzed the interaction of unfunctionalized AuNPs with human blood plasma proteins. While anti-cooperative binding has been reported for the interaction of AuNP with albumin, fibrinogen, histone, and globulin proteins, positive cooperativity was observed for insulin. The high cooperativity value was attributed to the NP-induced formation of ordered insulin fiber structures.41 Concerning functionalized nanoparticles, 7587


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inhibition of de novo fibril formation and destruction of already formed fibrils. Quantification of sample density on the surface indicated that free ligands and NP−ligand conjugates strongly reduced Aβ fibril density compared to the reference samples (at the same time) and also compared to the initially preformed Aβ fibril density (Figure 5). It is noted that Aβ monomers could

quantify Aβ aggregate size distributions and to take snapshots during different stages of Aβ aggregation. Thus, it is even possible to create a picture of the kinetic development of smaller and larger aggregates. Since D3 was selected against Aβ monomers to small oligomers through a mirror-image phage display, 26 it was especially interesting to monitor the occurrence of these most neurotoxic species in the absence and presence of AuNPs. Aggregation was initiated by dissolving 40 μM Aβ in 10 mM phosphate buffer at pH 7.4. Incubation with gentle agitation for either 90 min or 24 h at 37 °C preceded the analysis by density gradient centrifugation at 7 °C. The resulting harvested 23 fractions were further analyzed via ELISA. Thus, the relative concentration of Aβ was determined in each fraction. The size of Aβ assemblies was derived from the position in the gradient in comparison to the positions obtained for a set of lead proteins with known molecular mass (Figure 3a). Early Phase (after 90 min Incubation). After 90 min, Aβ formed small- and medium-size oligomers; the presence of AuNPs did not alter this distribution. The free ligand D3_5+ significantly lowered formation of medium-size oligomers, but produced some higher oligomers in fractions 15−20. With D3 immobilized on AuNPs, the same amount of ligand retarded the development of oligomers and produced at the same time very large aggregates. The same effects were reproduced in two consecutive sets of experiments. Late Phase (after 24 h Incubation). After 1 day, protofibrils are formed and the lateral association begins to form mature fibrils. Now the picture completely changed: almost no oligomers are present any more, and large species prevail (fractions 16−23) in the Aβ control sample. Molecular weight distributions in the presence of free or AuNP-supported ligands are now very similar and show a strong reduction of Aβ from fractions 16−23. We would like to emphasize that naked/pure AuNPs did not alter the course of Aβ aggregation over the entire 24 h period. On the contrary, at an early stage free and immobilized D3 ligands both effectively prevented formation of the most neurotoxic small Aβ oligomers, indicating that the ligand’s selectivity was retained in the AuNP conjugates. As Aβ aggregation proceeds, both free and AuNP-immobilized ligands eliminate large aggregates (see also Supporting Information Figure S22), in sharp contrast to the AuNP control.63 Atomic Force Microscopy. Atomic force microscopy allows the facile visualization of Aβ fibrils, oligomers, and nanoparticles, giving insight into fibril length, height, and density. Lin et al. reported the formation of fibrillary structures on the AFM substrate itself attributed to surface-mediated adsorption and self-assembly of monomers.44 Thus, morphologies observed in AFM images may not always accurately reflect the peptide structures in solution.44 For AFM experiments, Aβ fibrils were preformed at a monomer concentration of 100 μM according to a protocol from Stine et al.45 After 24 h, ligands, nanoparticles, and AuNP−ligand conjugates were added to the preformed fibrils, and their effects were assessed. AFM snapshots of the samples were taken 0, 1, and 7 days after sample addition (Figure 4). Notably, fibril formation is a dynamic process and fibrils can be formed as long as monomers of Aβ remain in the sample. It was observed that even after the preincubation period the density of Aβ species in the reference samples (Aβ only and Au/Aβ) increased over time. Hence, all effects observed in the samples containing Aβ, AuNP, and/or ligands can result from a combination of two effects, namely,

Figure 5. Analysis of atomic force micrographs of preformed synthetic Aβ fibrils after incubation with pure nanoparticles, pure ligands, and ligand-coated nanoparticles. The surface coverage is evaluated as a function of incubation time (a). Temporal evolution of fibril height (b) and length (c) after different incubation periods with ligand-coated nanoparticles.

not be detected by AFM; neither were spherically appearing oligomers and nanoparticles distinguished when determining Aβ density on the substrate. However, height and length of the fibrils were also greatly reduced over time by free ligands and ligand-carrying nanoparticles. It is therefore concluded that free and immobilized ligands both destroyed fibrils and inhibited fibril formation, while Aβ aggregation proceeded in the presence of ligand-free nanoparticles. This was independently confirmed through transmission electron microscopy (TEM) analysis of aggregated Aβ fibrils alone and after the addition of the AuNP/D3 conjugates (Figure S23). After 7 days, preformed fibrils were almost completely dissolved by free and immobilized ligands, which retain their disaggregating ability on the particle surface. In previous studies, Thakur et al. analyzed DHLA-capped quantum dots and Aβ1−40 fibrillation. After 7 days of incubation, the overall number of fibrils and fibril heights were strongly reduced in the presence of quantum dots.12 Moreover, Palmal et al. analyzed the fibril morphology of Aβ1−40 in the presence of curcumin-functionalized gold nanoparticles via TEM. While average fibril lengths of 2.3 μm 7588


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ACS Nano were observed in the control after 7 days of incubation, fibril lengths were restricted to 20 nm for 10 min at 18 000 rpm (24000g, Hettich centrifuge, angle rotor 1612). The nanoparticle concentration after centrifugation was determined spectrophotometrically (ε = 8930 L mg−1 cm−1 at 380 nm). A calibration curve relating the absorbance at 380 nm to the gold mass concentration is included in the Supporting Information (Figure S2). UV/vis extinction spectroscopy was performed using a Thermo Scientific Evolution 201 spectrophotometer. Samples were measured in a 1.5 mL quartz cuvette with 10 mm path length covering a spectral range from 190 to 900 nm. Nanoparticle size was analyzed via analytical disc centrifugation (ADC, DC 24,000, CPS Instruments), TEM, and DLS. For ADC, 100 μL of PVC calibration standard (0.239 μm, CPS Instruments) or sample was loaded into the spinning centrifuge at 24 000 rpm, which contained a continuous sucrose gradient (8 to 20 wt %). The lower limit of detection was 4 nm. DLS and nanoparticle zeta potential measurements were performed using a Zetasizer Nano ZS (Malvern). The sample was filled into a disposable capillary cell (V = 750 μL), and the temperature equilibrated for 120 s. Each sample was measured three times, and each measurement consisted of at least 20 runs. For pH-dependent stability analysis, AuNPs were titrated with 1 M NaOH or HCl, and the pH was recorded with a PCE-PHD pH meter and a microelectrode from Sartorius at 23 °C before zeta potential measurements. The standard Hückel model56 was applied to interpret the zeta potential results, applicable for small nanoparticles and low ionic strength media. Average particle diameters (xC) are obtained by fitting the size distribution obtained from ADC, TEM, or DLS to logNormal distributions. Nanoparticle number concentrations were calculated according to formula 1:57

ELISA assay and AFM images demonstrate that the AuNPimmobilized ligands are able to inhibit Aβ aggregation as well as dissolve existing fibrils with comparable performance to free ligands and that they retain their selectivity for small oligomers. Aβ-binding, ThT, and cell assays even indicate significantly improved effects through ligand immobilization and presenting multiple ligands in local vicinity on the particle surface. In the future, we will proceed to heteroavidic nanoparticles that combine two and more different functional units and examine the effect of hybridization on the AuNP surface toward Aβ aggregation and cell lesion. A cooperative effect may be expected if such units bind to distinctly different regions on the Aβ target and if they offer complementary functionalities, e.g., molecular recognition and β-sheet breakage. In a generic approach, the synergistic immobilization of functional ligands on nanoparticle surfaces may serve for the development of new biomedical tools offering numerous advantages over free ligands.

METHODS/EXPERIMENTAL Nanoparticle Synthesis and Characterization. Gold nanoparticles were generated by pulsed laser ablation in liquids.30 A nanosecond Nd:YAG laser (Rofin PowerLine E 20) was employed at a fundamental wavelength of 1064 nm, with a pulse duration of 8 ns, and at a repetition rate of 15 kHz. The corresponding laser pulse energy was 367 μJ. Briefly, a gold foil (99.99% Allgemeine Gold and Silberscheideanstalt AG, Pforzheim) was placed in a self-designed, stirred Teflon batch reactor containing 30 mL of aqueous solution (18.2 MΩ·cm@25 °C) with 1 mmol/L NaCl (99.9% VWR). The laser beam was focused on the target via an F-Theta lens ( f = 100 mm) with a liquid layer of 3 mm and spirally scanned on the target with a scanner optic (SCANcube10, Scanlab). After 15 min, approximately 22.5 mg of gold was ablated by the laser, resulting in colloid concentrations of 750 μg/mL and an extrapolated productivity of 90 mg/h using 5.5 W laser power. The

NNP,tot = 7592

βNP mNP

=

3βNP 4ρπrNP3

(1) DOI: 10.1021/acsnano.6b02627 ACS Nano 2016, 10, 7582−7597

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ACS Nano where NNP,tot [1/mL] is the nanoparticle number concentration, ρ [μg/cm3] is the material density (i.e., 19.3 g/cm3 for AuNP), rNP [cm] is the nanoparticle radius, and bNP [μg/mL] is the nanoparticle mass concentration. Stability upon freezing was analyzed by coating ultrapure AuNPs with O-(2-mercaptoethyl)-O′-methylhexa(ethylene glycol) (mPEGSH, M = 356.48 g/mol, purity 98% (Figures S11−S14). Nanoparticle Conjugation with Ligands. Nanoparticles were freshly mixed with thiol ligands before use in further experiments. Applied ligand dosages were calculated according to formula 2: NLigands =

c LigandsNAv NNP,tot

=

complete ligand binding at the same ligand concentration (125 ligands per particle, conjugation efficiency 100%). Preparation of Aβ. Aβ1−42 peptide (Bachem, Germany) was monomerized overnight in 1,1,1,3,3,3-hexafluoro-2-propanol (Sigma, Germany), lyophilized, redissolved in DMSO, and stored at −20 °C. Affinity Experiments. Affinity experiments were conducted in order to assess protein binding on the nanoparticle surface. FITClabeled Aβ (FITC-β-Ala-amyloid β (1−42), purity 88%, M = 4974.57 g/mol, Bachem) was employed. The titration experiments were performed in a 384-well plate (Nunclon 384 flat-bottom black polystyrol, Sigma). A stock solution of gold colloid (c = 500 μg/mL) with ligand (c = 100 μM) was prepared. A 1:1 dilution series of AuNPs was created with a total volume of 45 μL per well. Equal amounts of labeled protein stock (c = 0.25 μmol/L Aβ-FITC; V = 45 μL) were added to each well, thus keeping the protein concentration and sample volume constant. Measurements were conducted in a Tecan InfiniTe 200 at room temperature, with 100% amplification. Integration time was 20 μs, delay time was set to 0 s, and the number of flashes was 25. The excitation wavelength was 485 nm for FITC-labeled proteins (emission at 520 nm). As described previously,38,41 AuNP quench fluorescence with respect to the fluorophore’s proximity to the AuNP surface was determined. In this respect, normalized quenching is defined as the ratio between the apparent fluorescence in the presence of gold nanoparticles (F0 − F) vs the fluorescence of the pure fluorophore (F0). By assuming that protein binding occurs at equilibrium (AuNP + protein ⇌ AuNP − protein), the quenching data can be fit with the Hill formula 3: y = Bmax

(3)

in which x is the nanoparticle concentration [nM], y is the fluorescence quenching, Bmax is the maximum quenching, KD [nM] is the dissociation constant describing the equilibrium, and n is the cooperativity factor. Thioflavin T Fluorescence Assay. ThT fluorescence measurements were performed on a 384-well plate (Nunc GmbH, Germany) in a Tecan InfiniTe 200. Fluorescence intensity was measured at 37 °C, 446 nm excitation wavelength (bandwidth 9 nm), and 490 nm emission wavelength (bandwidth 20 nm). Each data point was averaged over 40 lamp flashes. Each measurement cycle was started by shaking the sample carrier orbitally for 30 s at medium intensity to avoid settling of larger aggregates. Each sample (60 μL) was composed of 10 μM Aβ1−42 in 10 mM phosphate-buffered saline, 3.03 μM ThT (Sigma, Germany), and 60 μM of the test compound. After 5 days of incubation in a thermomixer (650 rpm) at 37 °C, ThT fluorescence was determined. For graph representation, emission values of 4-fold samples were averaged. Each test compound was measured separately, in both 10 mM phosphate-buffered saline (PBS) and 10 mM PBS with ThT to exclude any potential interactions between ligand and ThT. Kinetics of Aβ Aggregation. A mixture of 2 μL of Aβ1−42 peptide (300 μM stock solution), 6 μL of PBS (10×), 2.9 μL of ThT (62.7 μM stock solution), and 49.1 μL of bidistilled water was vortexed. The volume of 60 μL was pipetted in a 384-well plate (Nunc GmbH, Germany). ThT fluorescence intensity was determined every hour at 37 °C (λEx = 446 nm, λEm = 490 nm). The sample carrier was agitated 30 s before each measurement. Each point is the average of quadruplication. Circular Dichroism. Aβ1−42 was dissolved in hexafluoroisopropanol to a concentration of 500 μM. For the final peptide concentration of 10 μM, the solution was diluted with 10 mM potassium phosphate buffer (pH 7.0). Each sample was composed of 10 μM Ab1−42, 10 mM potassium phosphate buffer (pH 7.0), 10 μM ligand, and 50 μg/mL AuNP. Samples were incubated for 1 day in a thermomixer (650 rpm) at room temperature; spectra were recorded after 0 h, 1 h, and 1 day. Circular dichroism measurements were carried out on a J-815 spectropolarimeter (Jasco) at 20 °C in a 0.1 cm cell between 200 and 400 nm (data pitch: 1 nm, response: 1 s., sensitivity: standard, scanning speed: 100 nm/min, accumulation 1). Peptide secondary

4πrNP3ρc LigandsNAv 3βNP

xn (KD)n + x n

(2)

where NLigands [#/NP] is the number of ligands per nanoparticle, NAv [1/mol] is the Avogadro constant, cLigands [mol/L] is the ligand concentration, NNP,tot [1/mL] is the nanoparticle number concentration, ρ [μg/cm3] is the material density (i.e., 19.3 g/cm3 for AuNP), rNP [cm] is the nanoparticle radius, and bNP [μg/mL] is the nanoparticle mass concentration. To quantify the amount of bound ligands, nanoparticles were conjugated with tryptophan-labeled peptide (Trp-D3-Cys) and centrifuged at 100000g for 1 h at 7 °C (Optima Max-XP, Beckman Coulter). Nanoparticles with bound ligands formed a red-black pellet, whereas unbound ligands remained in the supernatant. Ligands in the supernatant were quantified spectrophotometrically (ε(Trp-D3-Cys) = 3850 M−1 cm−1@280 nm, Figure S15), and the number of bound ligands per nanoparticle was calculated as the difference from the initially applied number of ligands and the number of unbound ligands. To test the functionality of ligand-coated nanoparticles, particles were routinely incubated with 418 ligands per particle, leading to full coverage of the particle surface (approximately 125 bound ligands, conjugation efficiency 30%). In order to increase the proportion of bound ligands, nanoparticle concentration was tripled to allow 7593


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ACS Nano structures give rise to distinct CD patterns in the far-UV region (αhelix: minimum in ellipticities at 208 and 222 nm, β-sheet: minimum at 218 nm).58 Density Gradient Centrifugation and Enzyme-Linked Immunosorbent Assay. Lyophilized aliquots of 36 μg of Aβ1−42 were dissolved in 100 μL of 10 mM sodium phosphate buffer, pH 7.4, and 100 μL of sample was added, containing ligand-coated nanoparticles, nanoparticles, or the ligand alone, respectively. As a reference, 100 μL of Aβ solution was diluted with 100 μL of water as control. The solutions were incubated at 37 °C for 90 min or 24 h. To separate differently sized Aβ species, samples were subjected to density gradient centrifugation. The gradient was prepared in ultracentrifugation tubes (Quick Seal Polyallomer tubes, 8.9 mL, Beckman Coulter) by successively overlaying iodixanol solutions (Optiprep, 60% w/v, Sigma) using the following concentrations and volumes: 1155 μL 50%, 1155 μL 40%, 1155 μL 30%, 3455 μL 20%, 1150 μL 10%, 442 μL 5% (v/v) iodixanol in 10 mM sodium phosphate buffer at pH 7.4. A 200 μL amount of sample was layered on top. Centrifugation was carried out at 7 °C for 4.5 h in a Beckman Optima MAX-XP centrifuge (fixed angle rotor MLA-55) at a relative centrifugal force of 287000g. Finally, 23 fractions of 352 μL were harvested from each tube from top to bottom and further analyzed via an enzyme-linked immunosorbent assay. ELISA was carried out in 96-well plates (Nunc, flat-bottom, MaxiSorp, 350 μL, Thermo Fisher Scientific). Each fraction was applied to four wells in a 1:100 dilution. Four wells containing water were treated as blanks. The plate was incubated overnight at 4 °C with shaking at 400 rpm. ELISA was performed according to the manufacturer’s protocol of the secondary antibody with minor adaptions. Briefly, the plate was washed with 200 μL of PBS containing 0.05% Tween 20 (PBS-T) per well. After removing PBS, each well was blocked with 0.25% milk powder solution and incubated at room temperature with shaking at 400 rpm. After 35 min, the milk powder solution was removed and the well washed with PBS-T, before adding 100 μL/well of a 1:5000 dilution of the primary antibody, 6E10 (Covance). The plate was incubated for 2 h at room temperature with shaking at 400 rpm, emptied out, and washed again with PBS-T. Next, 100 μL/well of a 1:6000 dilution of the second antibody, GAMPO (Dianova), was added and incubated for 2 h at room temperature, with shaking at 400 rpm. After emptying out the plate and washing with PBS-T, 100 μL of developer solution from the QuantaBlu fluorogenic peroxidase substrate kit (Thermo Scientific, Rockford, IL, USA) was added per well and incubated at room temperature with shaking at 400 rpm. After 30 min, 100 μL/well stop solution was added to the developer solution, and fluorescence was measured using a microplate reader (Tecan InfiniTe 200, Crailsheim, Germany). The excitation wavelength was 325 nm with gain 100, averaging of 50 flashes, and 20 μs integration time. Emission was recorded from 419 to 421 nm, and the average over the three wavelengths and four wells was calculated to determine the Aβ content in each DGC fraction. Atomic Force Microscopy. For AFM experiments, Aβ fibrils were preformed from monomerized, lyophilized Aβ in 10 mM HCl at a monomer concentration of 100 μM at 37 °C (shaking at 650 rpm) for 24 h according to a protocol from Stine et al.45 After the incubation period, samples of pure AuNP, pure D3_5+, and AuNP/D3_5+ conjugates were added (1:1 volume ratio). Final concentrations were 50 μM Aβ, 125 μg/mL AuNP, and 25 μM D3. The samples were further incubated for 7 days at 37 °C (shaking at 650 rpm). AFM analysis was performed directly after addition of pure AuNP, pure D3, and AuNP−D3 conjugates to Aβ, after 24 h and after 7 days. Briefly, 10 μL aliquots were deposited on freshly cleaned mica substrates (Plano GmbH). After incubation for 5 min, the sample was dried under gentle argon flow and scanned in ScanAsyst mode using a MultiMode microscope (Bruker) equipped with a Nanoscope V controller. Force constant cantilevers (0.4 N/m) with sharpened pyramidal tips (ScanAsyst-Air tips, Bruker) were used for scanning. After engagement the peak force set point was typically 0.02 V and the scan rates were about 1 Hz. Image processing was performed with NanoScope Analysis software (version 1.5, Bruker). Fibril length and height were evaluated with Gwyddion software (http://gwyddion.net/

). ImageJ software (National Institute of Health, Bethesda, MD, USA) was employed to determine the substrate coverage with fibrils. Cellular Aβ Assay/Western Blots. In order to test the influence of the bare or ligand-coated AuNPs on naturally expressed Aβ species, we generated and subcloned a CHO cell line that stably expresses APP751 including the familial Indiana mutation (V717F) termed Hennes20. A similar cell line, termed 7PA2, has been shown to secrete Aβ oligomers and N-terminal-elongated monomers.28,29 We directly compared the Hennes20 and 7PA2 cell lines and found a slightly stronger secretion of these Aβ species in Hennes20 (data not shown). The experiments were performed according to the protocol described by Podlisny et al.28 Briefly, Hennes20 cells were cultured in DMEM with 10% fetal bovine serum supplemented with 100 U/mL penicillin and 100 μg/mL streptomycin. A total of 5 × 105 cells were seeded into 10 cm cell culture dishes, and after 2 h the settled cells were treated with freshly prepared pure or ligand-coated nanoparticles or controls as indicated in the figures. After 4 days the medium was replaced with serum-free medium, and fresh preparations of nanoparticles or control compounds were added to the cells. After an additional 72 h the conditioned medium was harvested and cleared by centrifugation (20000g for 5 min at 4 °C). Aβ species were immunoprecipitated from the CM overnight at 4 °C with NHS-sepharose-coupled mAB-IC16, recognizing amino acid 2−8 of Aβ.59,60 After two washes with PBS, samples were electrophoresed on 10−20% tricine peptide gels (Biorad, Hercules, CA, USA) and transferred to 0.2 μM nitrocellulose (NC) membranes at 400 mA for 2 h. Filters were boiled for 10 min in PBS and blocked overnight at 4 °C with 5% skimmed milk (Oxoid, Thermo Scientific, Bonn, Germany) in PBS-T. After three washes in PBS-T, each for 10 min, the membranes were probed with monoclonal 4G8 antibody (Signet, Dedhem, MA, USA; 1:500 in PBS-T) that recognizes amino acid 17−24 of the Aβ peptide. Bound antibody was detected with horseradish peroxidase conjugated goat anti-mouse Ig (diluted 1:25 000 in PBS-T) (Thermo Scientific, Bonn, Germany) and the Amersham ECL Western blotting detection reagent (GE, Buckinghamshire, UK). Aβ signals were quantified by densitometry using the ImageJ software (National Institutes of Health, Bethesda, MD, USA). For detection of APP and caspase-3 cells were lysed in 750 μL of lysis buffer (50 mM Tris pH 8, 150 mM NaCl, 1% NP40, 5 mM EDTA) including the Complete protease inhibitor cocktail (Roche, Mannheim, Germany). The protein concentrations of the lysates were quantified using the DC protein detection kit (Biorad). A 30 μg amount of lysates was separated on a NuPAGE 4−12% Bis-Tris gel (Invitrogen, Carlsbad, CA, USA) and transferred to a 0.2 μm NC membrane. After blocking with PBS-T/5% skimmed milk, APP and caspase-3 were detected by CT15 polyclonal rabbit antiserum61 (diluted 1:3500 in TBS-T) and caspase-3 antibody (diluted 1:1000 in TBS-T; #9662, Cell Signaling, Danvers, MA, USA). Detection of tubulin with a monoclonal anti-α-tubulin antibody (T9026, SigmaAldrich, St. Louis, MO, USA; diluted 1:15 000 in PBS-T) served as internal control. After incubation with 1:25 000 dilutions of goat antimouse IRDye 680RD and goat anti-rabbit IRDye 800CW (LI-COR, Lincoln, NE, USA), signal intensities were analyzed on a LI-COR Odyssey CLX using the corresponding Image Studio version 2.1 software (LI-COR Biosciences, NE, USA). Light and Confocal Microscopy. Cells (2 ×104) were seeded on cover glasses (13 mm diameter) that were placed in 24-well plates. After settling of the cells, they were treated with the compounds. At day 4, the cells were washed with PBS and fixed with PBS/4% paraformaldehyde for 30 min at room temperature. After two washing steps with PBS and one with pure water, cells were mounted with ProLong Gold including DAPI (Invitrogen, Hercules, CA, USA). Images were collected with a Zeiss Axiovision Apotome.2 (Zeiss) using the differential interference contrast (DIC) filter. Additionally, images were collected with a Leica SP8 gSTED confocal microscope, using three channels. AuNPs were excited at 532 nm (emission: 425− 510 nm), and DAPI-stained cell nuclei were excited at 405 nm (emission: 534−590 nm) using a white light laser source. The DIC filter was employed for visualization of the cell membrane. 7594


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ASSOCIATED CONTENT S Supporting Information *

The Supporting Information is available free of charge on the ACS Publications website at DOI: 10.1021/acsnano.6b02627. Supplementary material on the Synthesis and characterization of nanoparticles and NP-ligand conjugates; Characterization of NP-ligand conjugates; Characterization of thiol-modified D3 ligands; Aβ secondary structure analysis; Density Gradient Centrifugation − ELISA Experiments; TEM analysis and the Cellular Aβ assay (PDF)

AUTHOR INFORMATION Corresponding Authors

*E-mail: [email protected] (ligands). *E-mail: [email protected] (nanoparticles). Author Contributions △

C. Streich and L. Akkari contributed equally.

Notes

The authors declare no competing financial interest.

ACKNOWLEDGMENTS L. Gamrad is acknowledged for performing confocal microscopy at the imaging facility IMCES. M. Heidelmann from the Interdisciplinary Center for Analytics on the Nanoscale (ICAN, CENIDE) and J. Jakobi are acknowledged for performing TEM measurements. W. Pfeiffer and B. Hilterhaus are acknowledged for support with AFM measurements and data evaluation. H. Wöll is acknowledged for working on the peptide synthesizer and HPLC measurements. P. Schneider is acknowledged for performing CD spectroscopy. C.S. thanks the Faculty of Chemistry, University of Duisburg-Essen, and Evonik Industries for financial support. A.M.S. was supported by a grant from the Forschungskommission of the Heinrich-Heine University Düsseldorf- Medical Faculty (54/2013). C.D. has been supported by the Helmholtz Association through the Portfolio Topics “Technology and Medicine” and “Drug Research”. REFERENCES (1) Stockwin, L. H.; Holmes, S. Antibodies as Therapeutic Agents: Vive la Renaissance! Expert Opin. Biol. Ther. 2003, 3, 1133−1152. (2) Keefe, A. D.; Pai, S.; Ellington, A. Aptamers as Therapeutics. Nat. Rev. Drug Discovery 2010, 9, 537−550. (3) Fasting, C.; Schalley, C. A.; Weber, M.; Seitz, O.; Hecht, S.; Koksch, B.; Dernedde, J.; Graf, C.; Knapp, E.-W.; Haag, R. Multivalency as a Chemical Organization and Action Principle. Angew. Chem., Int. Ed. 2012, 51, 10472−10498. (4) Mammen, M.; Choi, S.-K.; Whitesides, G. M. Polyvalent Interactions in Biological Systems: Implications for Design and Use of Multivalent Ligands and Inhibitors. Angew. Chem., Int. Ed. 1998, 37, 2754−2794. (5) Tassa, C.; Duffner, J. L.; Lewis, T. A.; Weissleder, R.; Schreiber, S. L.; Koehler, A. N.; Shaw, S. Y. Binding Affinity and Kinetic Analysis of Targeted Small Molecule-Modified Nanoparticles. Bioconjugate Chem. 2010, 21, 14−19. (6) Hong, S.; Leroueil, P. R.; Majoros, I. J.; Orr, B. G.; Baker, J. R.; Banaszak Holl, M. M. The Binding Avidity of a Nanoparticle-Based Multivalent Targeted Drug Delivery Platform. Chem. Biol. (Oxford, U. K.) 2007, 14, 107−115. (7) Rosi, N. L.; Giljohann, D. A.; Thaxton, C. S.; Lytton-Jean, A. K. R; Han, M. S.; Mirkin, C. A. Oligonucleotide-Modified Gold 7595


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ACS Nano (59) Müller-Schiffmann, A.; März-Berberich, J.; Andreyeva, A.; Ronicke, R.; Bartnik, D.; Brener, O.; Kutzsche, J.; Horn, A. H. C.; Hellmert, M.; Polkowska, J.; Gottmann, K.; Reymann, K. G.; Funke, S. A.; Nagel-Steger, L.; Moriscot, C.; Schoehn, G.; Sticht, H.; Willbold, D.; Schrader, T.; Korth, C. Combining Independent Drug Classes into Superior, Synergistically Acting Hybrid Molecules. Angew. Chem., Int. Ed. 2010, 49, 8743−8746. (60) Müller-Schiffmann, A.; Herring, A.; Abdel-Hafiz, L.; Chepkova, A. N.; Schäble, S.; Wedel, D.; Horn, A. H. C.; Sticht, H.; de Souza Silva, M. A.; Gottmann, K.; Sergeeva, O. A.; Huston, J. P.; Keyvani, K.; Korth, C. Amyloid-β Dimers in the Absence of Plaque Pathology Impair Learning and Synaptic Plasticity. Brain 2016, 139, 509−525. (61) Sisodia, S. S.; Koo, E. H.; Hoffman, P. N.; Perry, G.; Price, D. L. Identification and Transport of Full-Length Amyloid Precursor Proteins in Rat Peripheral Nervous System. J. Neurosci. 1993, 13, 3136−3142. (62) Streubel, R.; Barcikowski, S.; Goekce, B. Continuous Multigram Nanoparticle Synthesis by High-Power, High-Repetition-Rate Ultrafast Laser Ablation in Liquids. Opt. Lett. 2016, 41, 1486−1489. (63) The quantification of the Aβ content in the density gradient fractions should be treated with care: (a) Typically, the last fraction contained a significant amount of very large aggregates, when AuNPs were present, which became detectable only after denaturation with 6 M guanidine hydrochloride and heating to 100 °C (fraction 23 in Figures 4b and S22). Due to the high density of AuNPs, the particles will sediment under the applied DGC conditions. It is thus assumed that all species of Aβ (independent of their size and shape) will when adsorbed on AuNPsbe detected only in the last fraction after denaturation.. (b) As shown in the Supporting Information (Figure S21), the linear range of quantitative Aβ detection is limited with ELISA, meaning that the antibody did not detect the whole amount of Aβ, but was already saturated with Aβ in some fractions. Thus one has to consider that the black reference line (Aβ control) would appear much higher and the measured inhibitory effects may be much larger. (c) Additionally, it is noted that the area under the curve is not the same for each sample. In this regard, it was not determined whether D3 bound to Aβ negatively influences Aβ detection. Considering that D3 binds to the N-terminus of Aβ similarly to the antibody used for ELISA, some epitopes of Aβ may already be blocked by D3 and bias the quantitative detection of Aβ.

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Characterizing the Effect of Multivalent Conjugates Composed of AÎ²

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