Dynamics and Morphology of Nanoparticle-Linked Polymers

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Dynamics and Morphology of Nanoparticlelinked Polymers Elucidated by NMR Yongqian Zhang, Charles G. Fry, Joel A. Pedersen, and Robert J Hamers Anal. Chem., Just Accepted Manuscript • DOI: 10.1021/acs.analchem.7b03489 • Publication Date (Web): 16 Oct 2017 Downloaded from http://pubs.acs.org on October 18, 2017

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Analytical Chemistry

Dynamics and Morphology of Nanoparticle-linked Polymers Elucidated by NMR Yongqian Zhang,1 Charles G. Fry,1 Joel A. Pedersen,1,2 Robert J. Hamers1 1

University of Wisconsin-Madison, Department of Chemistry, 1101 University Ave., Madison, WI, 53706 University of Wisconsin-Madison, Environmental Chemistry and Technology Program, 1525 Observatory Dr., Madison, WI, 53706 2

KEYWORDS: diamond nanoparticle, NMR, solid/liquid interface, nanoparticle-polymer interaction, polymer dynamics ABSTRACT: Nanoparticles are frequently modified with polymer layers to control their physical and chemical properties, but little is understood about the morphology and dynamics of these polymer layers. We report here an NMR-based investigation of a model polymer-modified nanoparticle using multiple NMR techniques including 1H NMR, DOSY, TOCSY, and T2 relaxometry to characterize the dynamics of the nanoparticle-polymer interface. Using 5 nm detonation nanodiamond covalently linked to poly(allylamine) hydrochloride as a model system, we demonstrate the use of NMR to distinguish between free and bound polymer and to characterize the degree to which the segments of the nanoparticle-wrapping polymer are mobile (loops and tails) vs. immobile (trains). Our results show that the polymer-wrapped contain a large fraction of highly mobile polymer segments, implying that the polymer extends well into solution away from the nanoparticle surface. Flexible, distal polymer segments are likely to be more accessible to extended objects such as cell membranes, compared with polymer segments that are in close proximity to the nanoparticle surface. Thus, these flexible segments may be particularly important in controlling subsequent interactions of the nanoparticles. While reported here for a model system, the methodology used demonstrates how NMR methods can provide important insights into the structure and dynamics at nanoparticle-polymer interfaces, leading to new perspectives on the behavior and interactions of polymer-functionalized nanoparticles in aqueous systems.

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Synthetic nanoparticles are often functionalized with molecular ligands or polymer coatings to control properties such colloidal stability1-3 and nanoparticle interaction with biomolecules,4-5 as well as to facilitate incorporation into nanoparticle composites.1,6-7 The increasing interest in understanding nanoparticle interactions in environmental and biological systems motivates a desire to understand the molecular-level structure and dynamics of molecule-nanoparticle interfaces and interactions. Previous studies have found that nanoparticles functionalized with positively charged ligands and/or polycation wraps are toxic to many singlecell8-10 and multicellular organisms.11-14 Recent studies suggest that cationic, amine-based polymers produce larger biological responses than simpler amineterminated molecular ligands.14 These results suggest that the morphology and dynamics of polymers and ligands at

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nanoparticle interfaces may play an important role in controlling their biological interactions.12 For example, molecular functional groups located in close proximity to a nanoparticle surface might be expected to induce significantly different interactions with cell membranes compared with equivalent groups located farther away and therefore in less hindered environments. A key challenge in understanding nanoparticle interfaces is the dearth of analytical tools able to selectively probe chemically and structurally heterogeneous interfaces in aqueous media.12 This problem is exacerbated by the fact that the organic ligands and polymers used for functionalization of nanoparticles often remain in solution, either in dynamic exchange with the nanoparticles15 or present as residual contaminants.16-17 NMR methods are well suited to investigate polymer dynamics and have been used previously to investigate

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small ligands on nanoparticles17-19 and the adsorption/desorption behavior of solution-phase polymers interaction with nanoparticles such as silica20-21 and CdSe.22 However, very few NMR studies have investigated the dynamics of polymers that are covalently linked to nanoparticle surfaces, thereby constraining the polymer morphology and dynamics. The dearth of prior studies is due in part to the relatively low sensitivity of NMR, which makes it difficult to study nanoparticle systems at low concentrations. Nevertheless, covalently linked nanoparticle-polymer systems are of great interest in understanding the fundamental molecular factors that control the biological interactions of engineered nanoparticles, and NMR techniques can provide a wealth of chemical information about the structure and dynamics of these systems. Here, we demonstrate the synergistic application of multiple liquid-based NMR techniques including diffusion-ordered spectroscopy (DOSY), total correlation spectroscopy (TOCSY) in 2-dimensional and selective 1dimensional variants, and spin-spin (T2) relaxation measurements using the Carr Purcell Meiboom Gill (CPMG) pulse sequence to provide insight into the morphology and dynamics of polymers covalently bonded to nanoparticles. As a model system we use poly(allyl amine hydrochloride) (PAH) covalently linked to 5 nm diameter diamond nanoparticles (DNPs). This specific system was chosen because (1) covalently functionalized DNPs exhibit extraordinary chemical stability24-25 and (2) PAH is representative of a wide class of polycationic polymers that stabilize nanoparticles but also can render them potentially toxic.11-14, 26-30 Thus, this combination of nanoparticle core and surface functionalization provides an important model system for understanding fundamental mechanisms of nanoparticle toxicity as well as being a model system for establishing the capabilities and limitations of NMR to provide information about the local chemical environment of polymers bonded to nanoparticles. In additional to definitively identifying spectroscopic features associated with polymers linked to the nanoparticles, we demonstrate that NMR methods can be used to categorize and quantify protons within polymer segments that are motion-constrained and those that are highly mobile. Taken together, these measurements show that polymer-functionalized nanoparticles have a large fraction of the protons in highly mobile environments. This in turn shows that the polymer does not wrap around nanoparticles in a tightly, multi-layer manner but forms many “loops” and “tails” that protrude into solution. These NMR-based approaches provide important new insights into the dynamics and morphology at polymer-nanoparticle interfaces.

Figure 1: Steps in covalent functionalization of nanodiamond with PAH. PAH is depicted in red. Molecules and diamond are not drawn to scale. 92 93 94 95 96 97 98 99 100 101 102 103 104 105 106 107 108 109 110 111 112 113 114 115 116 117 118 119 120 121 122 123 124

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EXPERIMENTAL SECTION

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Material. All reagents were purchased from SigmaAldrich, unless noted otherwise. Ultrapure water (resistivity ≥18 MΩ·cm) was used for all experiments.

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Figure 1 shows an overview of the process used for functionalization of the nanoparticles. Preparation of 2-(10-undecen-1-yl)-1,3-dioxolane. The 2-(10-undecen-1-yl)-1,3-dioxolane molecule was prepared by dissolving 10-undecenal (5.9 mmol, 95% purity), anhydrous ethylene glycol (1.2 mol, 99.8% purity) and a catalytic amount of p-toluenesulfonic acid (0.5 mmol, 98.5% purity) in 20 mL of anhydrous toluene. The resulting mixture was refluxed at 110 °C under N2 for 1 h. 31 The product was purified using rotary evaporation followed by vacuum distillation, giving a yield of 81% of the protected aldehyde. The product was stored under 3 Å molecular sieves. The chemical structure of the 2-(10undecen-1-yl)-1,3-dioxolane product was confirmed by 1H and 13C NMR. 1H NMR (CDCl3) δ: 5.76 (ddt, 1H), 4.91 (dd, 2H), 4.79 (t, 1H), 3.83 (m,4H), 1.98 (q,2H), 1.60 (dt, 2H), 1.30 (m, 12H). 13C NMR (CDCl3) δ: 139.1 (1H), 114.2 (1H), 104.7 (1H), 64.9 (2H), 34 (1H), 33.9 (1H), 29.6 (2H), 29.4 (1H), 29.2 (1H), 29.0 (1H), 24.2 (1H). High-resolution electrospray ionization mass spectrometry for C13H24O2 (M + H+) yielded found a mass-to-charge ratio m/z=213.1847, nearly identical to the expected value of m/z=213.1849. Detailed spectra and IR results are reported in the Supporting Information (Figure S1-2). Hydrogenation of 5 nm core detonation DNP surfaces. Detonation nanodiamond (5 nm average primary particle size, Nanostructured & Amorphous Material Inc.) was annealed at 500 °C in pure H2 (1 atmosphere, flow rate at 50 standard cm3/min) for 5 h to hydrogen-terminate the diamond surface and to disaggregate the particles.32 Diffuse reflectance infrared Fourier transform spectroscopy (DRIFTS) was used to verify H-termination of DNPs; spectra are shown in SI, Figure S3. Preparation of DNPs functionalized with 2-(10undecen-1-yl)-1,3-dioxolane. The H-functionalized DNPs were mixed with excess 2-(10-undecen-1-yl)-1,3dioxolane in argon and irradiated under a 254 nm UV


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lamp for 18 h.24 The 2-(10-undecen-1-yl)-1,3-dioxolanefunctionalized DNPs were cleaned via centrifugation (5 times, 5 min each, 14100×g) in ethanol followed by ultrapure water, and dried in a vacuum oven at 110 °C. The resulting powder was characterized using DRIFTS (SI, Figure S4).

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Preparation of DNPs covalently functionalized with PAH. Poly(allylamine hydrochloride) (PAH), Mr = 17500 g/mol was used without further purification. The 2-(10undecen-1-yl)-1,3-dioxolane-functionalized DNPs from the previous step were deprotected into aldehydefunctionalized DNPs in 1.5 M HCl at 60 °C for 16 h and were then mixed with PAH solution (0.1 mM) and sodium cyanoborohydride (50 mM) in 0.1 M Na2HCO3/Na3CO3 buffer solution at pH=10 and stirred overnight. The aldehyde-functionalized DNPs were characterized using DRIFTS (Figure S5), with successful deprotection established by the disappearance of acetal modes at 1142 and 1135 cm-1 and the concurrent appearance of new, strong C=O mode at 1761 cm-1. To clean the nanoparticles, ultrapure water was added to the PAH-functionalized DNPs and the suspension was ultrasonicated for 5 min and centrifuged (14,100×g). The supernatant (containing residual molecules from previous steps) was discarded. These washing and centrifugation steps were then repeated a minimum of 3 times, and the resulting pellet was re-suspended in 1 mM HCl.

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Because PAH-functionalized DNPs undergo aggregation in suspension, we also used centrifugation to produce size-selected subsets that could be easily resuspended. Centrifugation at 14,100×g was sufficient to pelletize the larger aggregates; the residual supernatant was then ultracentrifuged (288,000×g) to pelletize the smaller aggregates. The DNPs in each size-selected pellet were easily re-suspended. We used the larger aggregates (i.e., DNPs suspended from the pellet after centrifugation at 14,100×g) for DOSY and TOCSY characterization because these particles gave the highest concentration of the sample, which is optimal for NMR characterization; the smallest aggregates were suspended in D2O for T2 relaxometry measurements. The hydrodynamic size and the zeta potential distributions of the smallest aggregates were measured using dynamic light scattering and electrophoretic light scattering using a Malvern ZetaSizer ZS. (SI, Figure S6). Dynamic light scattering measurements of the aggregates yielded hydrodynamic diameters of 17.2 nm ± 1.8 nm for the small aggregates and 177±7 nm for the larger aggregates (data shown in SI Figure S6-S7). The surface functionalization of both sizes is the same because they both underwent the same functionalization procedures.

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NMR Measurements: 1H NMR, TOCSY, DOSY, and T2 relaxometry experiments were all performed using a Bruker Avance III 600 MHz spectrometer. DOSY measurements were carried out at 298 K using the Jerschow-Müller convection-compensated pulse sequence 33 using a liquid He-cooled cryoprobe that has a zgradient coil with a maximum gradient strength of 0.498 T·m-1. For all experiments, spinning was turned off. The

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recycling time (d1) was always 3-5 times longer than the longest T1 of interest. Diffusion gradient length δ and diffusion time Δ were optimized for the broad peaks during the experiments. For the free PAH polymer solution, gradient length δ (p30) was 550 ms and diffusion time Δ (d20) was 0.15 s. For the suspension of PAHfunctionalized DNPs, the gradient length δ (p30) was 2000 ms and diffusion time Δ (d20) was 0.48 s.

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High-resolution measurements of the spin-spin (T2) relaxation times were performed using the Carr-PurcellMeiboom Gill (CPMG) pulse sequence. The 90° (pw90) and 180° (pw180) pulse lengths were 8 and 16 µs, respectively. The recycling time (d1) was 30 s, which is 5 times longer than the longest T1 of interest for all experiments. The number of scans was 16 per spectrum and CPMG echo time τ equaled 1 ms. The evolution time was calculated as (2τ + pw180)·(# of echo loops).

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RESULTS AND DISCUSSION

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The PAH-linked DNPs were covalently functionalized as depicted in Figure 1. After covalently grafting poly(allylamine) hydrochloride to the nanoparticles, we used x-ray photoelectron spectroscopy (XPS) to verify that the DNPs were successfully functionalized with the PAH polymer. The DNPs show a substantial increase in the N(1s) signal after PAH functionalization, indicating that the PAH polymer had successfully grafted to the DNPs. Quantitative analysis of the N(1s) and C(1s) XPS data yields a coverage of 30.9 ± 1.4 N atoms/nm2 after functionalization. This density is larger than the number density of N atoms typically produced by grafting of simple N-containing ligands (~4 atoms/nm2)34 and is even higher than the number density of C atoms at the closepacked diamond (111) surface (18.15 atoms/nm2) due to the three-dimensional nature of the PAH polymer. The XPS spectra and details of the analysis are in Supporting Information (Figure S8).

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Identification of surface-bound vs. free polymer via H NMR. Previous studies have shown that free and nanoparticle-bound PAH molecules induce significantly different biological responses.14, 35 However, quantifying free vs. bound polymers is difficult using most analytical methods.16 To evaluate the ability of NMR to distinguish between free and nanoparticle-bound PAH polymer, we first used 1H NMR to characterize the PAH-functionalized DNPs via conventional 1H NMR and via diffusion-ordered spectroscopy (DOSY). 1

Figure 2 shows 1H-NMR spectra of the hydrogenterminated DNPs, the free PAH polymer in solution, and the PAH-functionalized DNPs, with chemical shift assignments in Figure 2b. For the free polymer in solution the 1.56 ppm peak corresponds to the methylene protons (–CH2–) on the PAH polymer backbone (labeled “Hc”), the 2.06 ppm peak corresponds to the backbone methine protons (Hb), and the 3.10 ppm peak corresponds to the methylene protons adjacent to the amine group (Ha). These peaks exhibit apparent line broadening. After grafting to the diamond nanoparticles (Figure 2c), these peaks are shifted to 0.88 ppm, 1.30 ppm and 1.63 ppm and



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broaden further, with peak widths increasing from 57 ± 9 Hz for the free polymer to 144 ± 13 Hz for PAH bound to the DNPs. The diamond core provides increased shielding for molecules near the surface and therefore shifts the three proton peaks from 3.10, 2.06 and 1.56 ppm to slightly higher values.

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The large linewidths from these shifted new peaks suggest but do not prove that they arise from protons associated with PAH molecules bound to the DNPs. To identify the atoms in the PAH structure associated with these peaks, we explored the use of 1H-13C heteronuclear single-quantum correlation (HSQC) methods. HSQC measurements were successful on the free polymer, allowing definitive assignments of the peaks (SI, Figure S9). However, when this same method was applied to PAH-functionalized nanodiamond, the greater linebroadening resulted in very low HSQC signal intensity (SI, Figure S10) insufficient to definitively assign the broadened peaks.

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establish whether the PAH polymer is linked to the nanoparticles. Figure 3 shows a series of 1H spectra acquired as a function of the magnetic field gradient strength, GZ. As the magnetic field gradient increases, the signal intensity decreases because molecules diffuse out of the sampled region. In Figure 3, the signals of the sharp peaks attenuate much more quickly than do the broad peaks, indicating that the species giving rise to sharp peaks diffuse much faster than those giving rise to the broad peaks. To quantify the diffusion coefficients, we integrated the area under each of the broad peaks for each series of 1H spectra. Based on the Stejskal-Tanner equation, 37 the peak area I depends on the diffusion coefficient (D), the gyromagnetic ratio γ, and the magnetic field gradient GZ via equation 1:

(1)

I0 can be treated as an arbitrary scaling factor in our analysis.

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Figure 3. Series of DOSY plots of H spectra as a function of the magnetic field gradient strength (GZ) of both free PAH polymer and PAH-functionalized DNPs in D2O. 300 301 302 303 304 305 306 307 267 268 269 270 271

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Figure 2. (a) The H spectra of the hydrogen-functionalized DNPs (H-DNPs), (b) the free PAH ligand (0.1 mM) and (c) the PAH-functionalized DNPs (1 mg/mL) in D2O. All spectra use D2O peak as the reference peak.

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Identification of surface-bound vs. free polymer via DOSY. Since HSQC measurements were not able to definitively establish that the shifted broad peaks in Figure 2c arise from PAH bound to nanoparticles, we used an alternative approach based on measuring the diffusion coefficients of corresponding protons by DOSY.36 DOSY measures the diffusion coefficients of the targeted species, thereby providing a direct way to

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To extract the diffusion coefficient, we integrated the broad peaks at 1.56 ppm, 2.06 ppm, 3.10 ppm from Figure 2b and at 0.88 ppm, 1.30 ppm, 1.63 ppm from Figure 2c, and plotted ln(I) vs. . Figure 4a and 4b show the resulting plots for free PAH polymer and the PAHfunctionalized DNPs. The data points at the lowest gradient strength in Figure 4b include fast diffusion components from an impurity (small sharp peaks on top of broad peaks near 1.30 ppm and 1.63 ppm in Figure 2b) and were therefore excluded from the linear plots. The slopes of the resulting fits from Figure 4a and 4b yield the diffusion coefficient of the chemical moiety associated with each peak. Figure 4c shows the resulting diffusion coefficients plotted as a function of their chemical shifts. The data show that the translational diffusion coefficients associated with the three broad peaks observed for free PAH polymer are in the range from 5 × 10-7 to 6 × 10-7 cm2s-1, while those associated with the three broad peaks observed from PAH surface-bound DNPs are much smaller, in the range of 2 × 10-8 to 3 × 10-8 cm2s-1.

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Figure 4. Linear fits of natural logarithm of integration of three broad peaks in (a) free PAH polymer and (b) PAHfunctionalized DNPs vs the square of gradient strength using the Stejskal-Tanner equation adjusted to the JerschowMuller convection-compensated experiment. Error analysis were obtained using Least-Squares fitting. (c) Comparison of diffusion coefficients of each three broad peaks from free PAH polymer and PAH-functionalized DNPs as a function of their chemical shifts.

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where kB is the Boltzmann’s constant, η is the dynamic viscosity of the water, and T is absolute temperature (298 K). The calculated dH equals to 176 ± 38 nm and is consistent with the hydrodynamic diameter measured by dynamic light scattering, which is 177 ± 7 nm. Hydrodynamic diameters obtained from NMR measurements and those obtained from dynamic light scattering results are reported in the Supporting Information (Figure S7). The 1H-NMR and DOSY-NMR data together establish that the three peaks at 0.88, 1.30, and 1.63 ppm in Figure 2c are associated with a 10-fold decrease in diffusion coefficient relative to the peaks attributed to free PAH polymer. These data further establish that these peaks arise from PAH polymer that has covalently linked to the DNP surface and allow us to conclude that the amount of free, unbound polymer is below the limit of detection. Identification of polymer segments of PAHfunctionalized DNPs. One of the important aspects of this study is to determine whether covalent functionalization with a cationic polymer leads to nanoparticles in which the polymer coating wraps tightly around the particle, or whether there are conformationally flexible segments extending into the surrounding aqueous solution. A tightly wrapped polymer would be expected to be rigid, eliminating or reducing the motional averaging effects that ordinarily are important to achieving narrow 1H NMR linewidths, and would therefore be expected to give rise exclusively to broadened peaks. However, sharp peaks are observed in Figure 2c between 1.90 ppm and 3.60 ppm in the 1H spectrum of PAH-functionalized DNPs that were not present in the free PAH spectrum. The presence of these peaks in the spectrum of PAH-functionalized DNPs but not in the spectrum of the free PAH suggesting that either (1) covalent grafting of PAH to DNPs leads to polymer segments with high local mobility,38 or (2) they arise from the linker molecule used to initially functionalize the DNPs before grafting of the PAH. We hypothesize that the sharp peaks result from highly mobile segments of DNP-attached polymers and not from the residual free species in solution for two primary reasons: (1) the intensities of these peaks do not decrease even after repeated ultra-sonication and centrifugation, and (2) these sharp peaks do not overlap with any of those in the spectra of the linker ligands used throughout the functionalization process. Spectra of the linker ligands and the PAH-functionalized DNPs are presented in the Supporting Information (Figure S11). To test this hypothesis, we used Total Correlation Spectroscopy (TOCSY) to establish whether there are through-bond correlations between the sharp and broad resonances.

Assuming the functionalized DNPs have a spherical shape in solution, the hydrodynamic diameter dH of the PAH-functionalized DNPs was calculated from the measured translational diffusion coefficient (D) using the Stokes-Einstein equation:

(2)



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Figure 5. (a) 2D TOCSY spectrum of PAH-functionalized DNPs in D2O with a mixing time of 120 ms, showing all through-bond peak correlations. (b) Expanded view of broad peaks (a). (c) Selective-1D TOCSY compared with standard proton spectrum of PAH-functionalized DNPs in D2O (bottom) to confirm the through-bond correlation between sharp and broad peaks. The selective excitation is labeled in box at 0.88 ppm with a mixing time of 120 ms.

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Figure 5a shows a 2D-TOCSY 1H spectrum of the PAHfunctionalized DNPs. Crosspeaks at the intercept of two peaks indicate that the two protons are connected through chemical bonds via spin-spin coupling. From Figure 5a, the presence of crosspeaks among all four sharp resonances indicates that they are all connected through chemical bonds. As discussed below, the 2D-TOCSY pulse sequence that was used reduces the correlation amplitudes of broad peaks compared with those of sharp peaks. To show the correlations of the broad peaks more clearly, Figure 5b shows an expanded region from Figure 5a, highlighting the region encompassing the broad peaks and with the contour enhanced to accentuate correlations of smaller amplitude. Chemical bond correlations can be observed at a low contour level in Figure 5b between the

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broad peaks at 1.30 ppm and 1.63 ppm from crosspeak A. Correlation between the broad peak at 1.30 ppm and the sharp peak at 2.43 ppm (that is bond-correlated with the other three sharp peaks) is also observed from crosspeak B in Figure 5b. These results show that through-bond correlations exist between the broadened and narrow lines, thereby establishing that the narrow and broadened peaks arise from the same molecule. To confirm that the above interpretation is correct and that the observed correlations are not artifacts due to thermal noise, we performed selective-1D TOCSY experiments. Selective-1D TOCSY probes specific through-bond correlations among protons. In a 2DTOCSY experiment, all resonances in the spectra are excited simultaneously to obtain all the correlations within the molecule; whereas in a selective-1D experiment, a specific resonance of interest is selectively excited to show the correlations originating only from that peak. In general, broad peak signals and their correlations from macromolecules are greatly attenuated or eliminated in 2-D TOCSY because relaxation occurs during the pulse sequence; sharp signals are less affected.36, 39 Therefore, observing through-bond correlations from the broad peaks in a 2D-TOCSY is difficult. We resolved this challenge using selective-1D TOCSY and exciting the broad resonances; if throughbond correlations among the broad peaks and the sharp peaks exist, then sharp peaks are expected to appear in the TOCSY spectra. Figure 5c shows that this is indeed the case, confirming that the sharp and broad peaks come from the same molecule. The top spectrum results from selective excitation of the broad peak at 0.88 ppm (indicated by the box). After a 120 ms mixing time the spectrum displays all four peaks in the 2.0 to 3.5 ppm range, indicating that magnetization was transferred from the protons at 0.88 ppm to all four of these J-coupled protons through chemical bonds. These peaks are the same as those observed in a standard 1H spectrum without selective excitation (bottom spectrum). The presence of the four sharp peaks due to selective excitation of the broad peak at 0.88 ppm establishes through-bond correlation among sharp and broad peaks. Taken together, the 2D and selective-1D TOCSY data establish that the four main sharp peaks at 2.03, 2.42, 2.82, and 3.50 ppm and the broad peaks at 0.88, 1.30, and 1.62 ppm are all connected through chemical bonds. The mixing time used in the 2D and selective-1D TOCSY experiments allow us to observe correlations through four or more bonds in some cases. The sharp peaks at 2.04, 2.43 and 3.50 ppm correspond to the three protons on the PAH polymer backbone. The “singlet” at 2.83 ppm is assigned as an unresolved multiplet from amine protons on the PAH polymer, as evidenced by associated crosspeaks in the TOCSY spectrum in Figure 5a. The appearance of sharp peaks indicates that some regions of the PAH polymer retain high local mobility even after the polymer is grafted to the DNPs. In this model, the linewidths are then expected to be directly related to the distance between the protons and the DNP surfaces:


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sharp peaks correspond to the flexible motion of polymer segments that are further away from the DNP surfaces, while the broad peaks arise from polymer segments that have restricted motion due to their closer proximity to the DNP surfaces. Quantification of mobile vs. immobile polymer segments. The above analysis indicates that the polymer segments (and the associated protons) can be categorized into two classes based on their linewidth. Segments that are rigidly bound to the surface (sometimes referred to as “trains” 40-42) yield broad peaks, while those that are farther away from the surface and in high mobile environments (“loops” and “tails”) yield sharp peaks. We believe these differences may be important to understanding how nanoparticles interact with biological systems. For instance, NPs with a higher ratio of mobile segments can result in a much larger hydrodynamic diameter than NPs with polymer segments tightly bound to the surface (motion-restricted segments). Moreover, these details of the polymer shell can potentially determine the types of interactions (such as penetration, adsorption and disruption) that NPs encounter with the lipid membrane, which are often used as a model system in biological studies. 43-47 To achieve a deeper insight into and more accurate picture of the morphology of the ligand shell on these nanoparticles, we calculated the ratio of ligand segments that are in a motion-restricted environment vs. those in a highly mobile and more solvated environment. This was achieved using the Carr-Purcell-Meiboom Gill (CPMG) pulse sequence48-49 to measure spin-spin (T2) relaxation times. In this method, an initial 90o pulse is applied, followed by a series of 180o spin echo pulses; the intensities of the peaks are measured as a function of the number of repeated echoes separated by a relaxation delay period, the full length of which corresponds to a total evolution time. The peak intensities typically decay exponentially with increasing evolution time. The T2 relaxation time and the associated population of spins can then be calculated from measurements made at different evolution times via the relationship:

M y = M 0e

−τ T2

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Figure 6. (a) A series of H spectra result from the CPMG experiment with increasing evolution times (b) T2 exponential decay and associated curve-fits of peaks of interest vs time.

(3)

where My is the transverse magnetization, M0 is the equilibrium magnetization (proportional to the population of each corresponding proton species), and τ is the delay period. In cases where two discrete populations exist, protons near the particle surface (“bound”) are motion-restricted and have short relaxation times, while protons further away from the particle surface are less motion-restricted, having mobility similar to the free polymer in solution, and have longer relaxation times.50

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We refer to the T2 relaxation times of protons within these “bound” and “free” segments as T2b and T2f, respectively, and their corresponding fractional populations as Pb and Pf. Figure 6a shows a series of 1H spectra acquired with increasing evolution times. We plotted the integrated areas of peaks of interest and plotted these areas vs. evolution time to provide the inverse of the relaxation times T2 and their corresponding populations at τ=0. In Figure 6b, T2 decay curves of all peaks that are associated with the polymer shell (identified in TOCSY section) were fit to an exponential decay curve shown to obtain the data in Table 1. From these data, we determine that the fraction of protons in the bound environment (Pb) from the polymer shell is



1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60

547 548 549 550 551 552 553 554 555 556 557 558 559 560 561 562 563

55±3 %. This result indicates that 55 % of the polymer shell is in a bound environment on the DNP surfaces, and the rest is in the form of loops or tails extending into the adjacent aqueous solution. As a point of comparison, we also quantified the hydrodynamic diameter by Dynamic Light Scattering (DLS). Those measurements showed a hydrodynamic diameter of 17.2 ± 1.8 nm, a value significantly larger than the 5 nm diameter of the nanoparticle cores. However, DLS is not able to determine whether the hydrodynamic diameter is larger than the nanoparticle physical diameter because of nanoparticle aggregation or because loops and ends of the polymer chains extend in the aqueous medium. Out data suggest that at least part of the larger diameter measured by DLS arises from the extension of the polymer chains into solution. Figure 7 shows a pictorial illustration (not drawn to scale) that is consistent with the available data.

566 567 568 569

Figure 7. Schematic illustration depicting mobile and immobile molecular segments of PAH polymer covalently linked to DNPs. Figure is not drawn to scale 590 591 592

564 565

Table 1. T2 relaxation times and populations of peaks associated with specific chemical shifts. The peaks are classified as corresponding to T2 relaxation times as bound (T2b) and free (T2f) segments, with populations Pb and Pf.

570

593 594 595 596 597 598 599 600

1

Hδ

T2b

(ppm)

(ms)

1 T2f Bound Hδ Population (ppm) (ms) Pb

Free Population

601

Pf

603

960±20

0.90

47±21

300±30

2.04

73±7

1.30

16±2

2300±200

2.43

120±30 990±30

1.63

29±5

990±70

3.50

69±5

960±20

602

604 605 606 607 608 609

Total

3600±200

Sum

2900±40

Pb (%)

55±3

Pf (%)

45±3

610 611

621 622

ASSOCIATED CONTENT

623

Supporting Information

624

627

Details of nanoparticle functionalization and characterization using XPS, NMR and dynamic light scattering. The Supporting Information is available free of charge on the ACS Publications website.

628

AUTHOR INFORMATION

629

Corresponding Author

614

CONCLUSIONS

615 573 574 575 576 577 578 579 580 581 582 583 584 585 586 587 588 589

616

The use of complementary NMR-based methods provides important new insights into the morphology and dynamics of polymer-modified nanoparticles. DOSY allows robust identification of surface-tethered polymer vs. solution-phase polymer by making use of the significantly hindered translational diffusion rates of polymers tethered to nanoparticles. TOCSY provides definitive identification of the origins of the broadened peaks, while T2 measurements allow quantification of the proton relaxation and allow protons to be classified according to their spin-spin relaxation rates. For our model system of PAH covalently linked to 5 nm nanodiamond, our results show that even when covalently bonded to nanodiamond, 45 % of the polymer retains a high degree of mobility in the form of polymer loops and the free tails from the polymer ends. This result

is important to understanding of nanoparticle-biological interactions. While prior studies have shown that positively charged nanoparticles are more toxic than anionic nanoparticles,8-14 there is comparatively little understanding of how the detailed molecular structure controls biological response. Of particular importance is that nanoparticles functionalized with cationic polymers have been reported to induce larger biological response than nanoparticles functionalized with molecular ligands terminated with single positively charged amino groups.14 These studies, and related studies by Stellaci and coworkers45 show that in addition to the net charge, the spatial distribution of charge plays an important role. We hypothesize that cationic groups within polymer loops and tails more easily access and interact with functional groups bearing complementary charges (e.g., negatively charged phosphate groups of phospholipids) within cell membranes. Compared with free polymers in solution, linking polymers to nanoparticles may concentrate these group into smaller spaces, alter the way that they are displayed to cell membranes, and increase electrostatic interactions. The NMR techniques employed in this study provide a platform to investigate the dynamics and morphology for a variety of synthetic nanomaterials in solution, leading to a foundation for the mechanistic investigation of interactions between nanoparticle-ligand complexes in biological systems. We anticipate that these same methods could be applied to systems with higher complexity, such as binding of proteins, natural organic material, and other molecules that form “coronas” around nanoparticles. 51-54

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* Email: [email protected]

690 691

631

Funding Sources

632 634

This work was supported by the National Science Foundation Centers for Chemical Innovation Program Award # CHE1503408.

635

ACKNOWLEDGMENT

697

636

698

644

This work was supported by the National Science Foundation Center for Chemical Innovation Program grant CHE-1503408 for the Center for Sustainable Nanotechnology. The Bruker Avance 600 NMR instrument used in this work was supported by the National Institutes of Health grants S10OD012245. We thank Mark McCormick (3M Corporation) for valuable discussions regarding T2 relaxation experiments and Professor Christy Haynes for suggestions regarding the manuscript.

645

ABBREVIATIONS

709

646

710

649

NMR, Nuclear Magnetic Resonance; PAH, Poly(allylamine hydrochloride); DOSY, Diffusion-ordered Spectroscopy; TOCSY, Total Correlated Spectroscopy; DLS, Dynamic Light Scattering.

650

REFERENCES

651

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Dynamics and Morphology of Nanoparticle-Linked Polymers

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