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Adsorption and Unfolding of a Single Protein Triggers Nanoparticle Aggregation Sergio Dominguez-Medina, Lydia Kisley, Lawrence J. Tauzin, Anneli Hoggard, Bo Shuang, A.Swarnapali D. S. Indrasekara, Sishan Chen, Lin-Yung Wang, Paul J Derry, Anton Liopo, Eugene R Zubarev, Christy F. Landes, and Stephan Link ACS Nano, Just Accepted Manuscript • DOI: 10.1021/acsnano.5b06439 • Publication Date (Web): 11 Jan 2016 Downloaded from http://pubs.acs.org on January 13, 2016

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Adsorption and Unfolding of a Single Protein

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Triggers Nanoparticle Aggregation

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Sergio Dominguez-Medina,1†# Lydia Kisley,1†∆ Lawrence J. Tauzin,1 Anneli Hoggard,1 Bo

4

Shuang,1 A. Swarnapali D. S. Indrasekara,1 Sishan Chen,1 Lin-Yung Wang,1 Paul J. Derry,1

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Anton Liopo,1 Eugene R. Zubarev,1,2 Christy F. Landes,1,3* Stephan Link1,3*

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1 Department of Chemistry, Rice University, Houston, TX 77251

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2 Department of Materials Science and NanoEngineering, Rice University, Houston, TX 77251

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3 Department of Electrical and Computer Engineering, Rice University, Houston, TX 77251

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† These authors contributed equally to this work

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*Corresponding author's email: [email protected], [email protected]

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ABSTRACT

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The response of living systems to nanoparticles is thought to depend on the protein corona,

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which forms shortly after exposure to physiological fluids and which is linked to a wide array of

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pathophysiologies. A mechanistic understanding of the dynamic interaction between proteins and

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nanoparticles and thus the biological fate of nanoparticles and associated proteins is, however,

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often missing mainly due to the inadequacies in current ensemble experimental approaches.

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Through the application of a variety of single molecule and single particle spectroscopic

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techniques in combination with ensemble level characterization tools, we have been able to

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identify different interaction pathways between gold nanorods and bovine serum albumin

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depending on the protein concentration. Overall, we found that local changes in protein

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concentration influence everything from cancer cell uptake to nanoparticle stability and even

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protein secondary structure. We envision that our findings and methods will lead to strategies to

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control the associated pathophysiology of nanoparticle exposure in vivo.

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KEYWORDS: Protein corona, nanorods, super-localization microscopy, correlation

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spectroscopy, surface plasmon

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The interaction of nanoparticles with living organisms has received growing attention in the

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past decade1–3 due to the development of novel therapeutic and diagnostic tools,4–6 and the

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growing concerns regarding the safety of nanomaterials in vivo.7–11 It is understood that

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nanoparticles adsorb proteins in biological fluids forming a “protein corona”, which influences

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the nanoparticle physicochemical properties and subsequent interactions with the physiological

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environment.12–27 Both the possibility that in vivo administration of therapeutic nanoparticles can

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disturb the structural integrity of proteins and the current knowledge of the direct link between

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denatured proteins and severe neurodegenerative diseases suggest that nano-therapeutic

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platforms potentially pose a health risk.2,28–30 Therefore understanding the interaction of proteins

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with nanoparticles on a molecular level is paramount for the efficient and safe application of

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engineered nanoparticles.

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The quantification and identification of the protein corona composition for different types of

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nanoparticles with varying surface chemistries have been undertaken.12,14,30–34 Ex-situ ensemble

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techniques such as gel electrophoresis coupled with mass spectrometry have been widely used to

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characterize of chemical identity and abundance of constituent corona proteins after separation

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from the nanoparticles.12,29 Less disruptive, complementary optical spectroscopy approaches can

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be used to study the protein-nanoparticle complex in-situ and were able to determine the

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thermodynamics of protein binding as well as to provide insight into the interactions between

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protein coated nanoparticles and cells.15,35–38 For instance, the micromolar affinity of serum

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albumin to colloidal nanoparticles resulting in a protein monolayer15 at physiological

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concentrations and enhanced colloidal stability in plasma39 has been revealed by a combination

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of UV-VIS and fluorescence correlation spectroscopic techniques. Payne and co-workers have

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used super-resolution optical microscopy imaging under in vitro conditions, and demonstrated Page 3 of 33

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that what cellular receptors actually screen are the adsorbed proteins on the nanoparticles and not

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the initially synthesized nanoparticles themselves.40,41 Spectroscopic methods are furthermore

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ideally suited to characterize changes in protein structure when bound to a nanoparticle surface.

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The structural integrity of the corona proteins - which domains of the protein bind and which

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ones potentially change their structure28,42–48 - has been studied by using far and near UV circular

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dichroism (CD),28,42,43,49,50 IR,43,50 and Raman44 spectroscopy.

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While the protein-nanoparticle community has begun to transition from the mere identification

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of adsorbed proteins to the understanding of the physiological responses that can arise due to the

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presence of the protein corona, it is still not fully understood how protein adsorption affinities,

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relative concentrations, and surface chemistries are ultimately linked to the corona composition

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and possible structural changes of the proteins. A mechanistic bridge between these aspects is

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lacking due to the low spatio-temporal resolution and ex-situ requirements of most current

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techniques.18 Ensemble analytical techniques that assume quasi equilibrium conditions with

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exchange between free and bound proteins have furthermore led to controversial results.12,19,30–

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33,51

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Here we have used a combined approach of ensemble and single protein/single nanoparticle

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methods to answer the following questions about the nanoparticle protein corona: Is

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thermodynamic equilibrium a valid concept for the protein corona as local changes in protein

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concentrations occur? How does protein adsorption influence protein chemistry? How is

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nanoparticle stability altered?

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protein/nanoparticle interactions influence physiological outcomes? Mechanistic understanding

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of the physicochemical properties of bovine serum albumin (BSA) adsorbed to cationic-ligand

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functionalized gold nanorods (AuNRs) is used as the experimental system. While an ensemble

Finally, how might equilibrium and non-equilibrium

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protein adsorption isotherm implicates the adsorption of several proteins to the nanoparticle

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surface, single-protein/single-nanoparticle interactions visualized through super-localization

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imaging reveal that in fact only one protein is irreversibly adsorbed at a time. The conflicting

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findings are resolved through surface plasmon coupled circular dichroism (CD) spectroscopy,

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and demonstrated that the local changes in protein concentration affect the colloidal stability of

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nanoparticles, protein secondary structure and eventually cellular uptake of nanoparticles.

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RESULTS AND DISCUSSION

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When nanoparticles are pre-incubated in bovine serum albumin (BSA) at lower than

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physiological concentrations, properties such as uptake by MCF-7 cancer cells, BSA adsorption,

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and

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mercaptoundecyltrimethylammonium bromide coated AuNRs (MUTAB-AuNRs) incubated in

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10 % fetal bovine serum (FBS) are uptaken by MCF-7 cancer cells (Figure 1a, b).52 However, by

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pre-incubating the MUTAB-AuNRs in 1 % w/v BSA the uptake is increased three-fold (Figure

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1b). This increase is not due to MUTAB-AuNRs adhered to the outside of the cells as the cells

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were washed several times with PBS following a previously published protocol.52 As the most

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abundant serum protein, BSA is present in higher concentrations in FBS than 1 % w/v. Pre-

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incubation of MUTAB-AuNRs with 1 % w/v BSA, however, seems to influence their interaction

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with cancer cells and cause increased uptake. An important question to ask is therefore if and

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how a BSA-based protein corona is concentration dependent. This question will be addressed

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next with a combination of in situ ensemble and single molecule and single particle

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spectroscopic techniques.

nanoparticle

aggregation

are

strongly

influenced

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(Figure

1).

Cationic

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Figure 1 | Pre-incubation of MUTAB-AuNRs with low concentrations of BSA strongly influences AuNR cell uptake, BSA protein adsorption, and NR aggregation. a, Cartoon representation of (left, blue) pre-incubation of MUTAB-AuNRs with low concentration of BSA compared to (right, orange) MUTAB-AuNRs exposed to high concentration of proteins in serum without preincubation. b, Number of MUTAB-AuNRs uptaken per MCF-7 cell for MUTAB-AuNRs preincubated with 1% BSA before suspending in 10 % FBS and Eagle’s minimum essential media (EMEM), compared to MUTAB-AuNRs only suspended in 10% FBS and EMEM. Cells suspended in 10% FBS and EMEM without MUTAB-AuNRs are shown as a control. c, Increases in MUTAB-AuNR fluctuation correlation decay time caused by BSA adsorption. Luminescence correlation decays for MUTAB-AuNRs alone (gray), incubated with low concentrations of BSA (blue) vs. in higher serum-level BSA concentrations (orange). Autocorrelation curves with respective decay fits shown as solid lines. Under low BSA concentration the fit did not follow a predictable model; dashed line is instead shown as a guide for the eye. d, UV/vis spectra under similar conditions as c suggest MUTAB-NR aggregation occurs when incubated at low concentrations of BSA.

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Luminescence correlation spectroscopy provides strong support that the BSA protein corona

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that forms on MUTAB-AuNRs is concentration dependent (Figure 1c). At high protein

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concentrations (20 µM), the increase in measured hydrodynamic radius matches the dimensions

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of native BSA53 and hence implies the formation of a protein monolayer, consistent with earlier

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reports of BSA monolayer formation on nanoparticles (Table S1).15,54,55 In contrast, at lower

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BSA concentrations (2 nM) the luminescence correlation decay shifts to longer average decay

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times, indicating that surprisingly the nanoparticle-protein complex has become even larger than

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what is observed under the high BSA concentration conditions. Most importantly, the data could

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not be fit to an equilibrium adsorption model, suggesting non-equilibrium behavior, consistent

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with nanoparticle aggregation as verified further below.

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Further support for the strong deviation between low and high BSA concentrations is found in

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the UV-VIS spectra (Figure 1d). At high BSA concentration the extinction spectrum of the

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MUTAB-AuNRs is slightly red-shifted, due to a change in dielectric environment and consistent

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with equilibrium formation of a stable BSA monolayer. However, at low BSA concentrations the

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decreased intensity and resonance shift of the plasmon resonance indicate that the MUTAB-

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AuNRs aggregate in the presence of low BSA concentrations.

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The combination of our results on cancer cell uptake, protein adsorption, and nanoparticle

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colloidal stability strongly suggest that local protein concentrations must be considered when

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assessing corona chemistry and its influence on nanoparticles’ ultimate physiological fate, as

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suggested previously.11,20 Of particular interest in Figure 1 is the case of low protein

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concentrations. Low protein-to-nanoparticle ratios can, for example, occur under controlled pre-

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incubation conditions after injection into a living organism and upon accumulation in cellular

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compartments. Thus, we examine more closely the structure of protein and nanoparticles when

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they interact at low concentrations and suggest how these processes lead to the ultimate fate of

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the protein-nanoparticle colloid.

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BSA adsorbs at low concentrations to cationic ligand-coated gold nanomaterials of varying

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geometries (Figure 2). A 2 nM concentration of Alexa647-labeled BSA (3 dyes per protein) is

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flowed over individual nanomaterials of varying surface chemistries (cationic MUTAB, cationic

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amine poly(ethylene glycol) [NH2-PEG], and anionic citrate) and different geometries

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(nanospheres [~50 nm diameter], nanowires [width: ~70 nm; length: 1 - 10 µm], and nanorods

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[width: 20 nm, length: 58 nm]). BSA binding to these different nanostructures is visualized via

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an increase in fluorescence intensity due to adsorption of the fluorescently labeled protein. Due

Figure 2 | Single molecule imaging of BSA interactions with nanomaterials of varying geometries (sphere, wire, rod) and surface chemistries (cationic MUTAB, cationic NH2-PEGSH, anionic citrate). Images show nanomaterial highlighted with dashed red outline (left) before and (right) after 2 nM Alexa-BSA is introduced. Images are binned, ranging from 10-100 frames. These results show that BSA binding occurs independent of the nanoparticle geometry. Furthermore, a different surface chemistry also facilitates binding as long as the surface charge remains positive. No significant adsorption is seen for negative charged citrate capped nanoparticles at this BSA concentration.

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to total internal refection excitation, only individual molecules at the surface are registered while

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freely diffusing proteins only contribute to a slightly larger average background signal.56 We find

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that BSA binds to the cationic-ligand functionalized nanomaterials at this low concentration. On

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the other hand, negligible binding of BSA to the citrate capped nanospheres is observed under

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the same conditions. Much higher µM concentrations are needed for this ligand.35 To understand

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the mechanistic details of protein adsorption at low concentrations to cationic nanomaterials, we

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will concentrate on the interactions between BSA and MUTAB-NRs as example nanoparticle

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supports and serum proteins, respectively. Further analysis of other nanoparticle chemistries and

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serum proteins are presented in the Supplementary Information (SI).

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Single molecule analysis of the fluorescence imaging data reveals that only one BSA protein

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binds irreversibly to each MUTAB-AuNR (Figure 3). BSA adsorption onto single MUTAB-

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AuNRs is identified by co-localization57 of dye fluorescence and intrinsic AuNR luminescence

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(Figure 3a, Figure S1, S2, S3, Video S1). Scanning electron microscopy (SEM) image analysis

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confirms that 99% of the analyzed BSA adsorption events occur at single MUTAB-AuNRs

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(Figure S4) with little non-specific BSA interaction with the substrate because of surface

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passivation (Figure S5).

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Although in its native form smaller than a AuNR, we find no evidence for multiple BSA

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adsorption by analyzing fluorescence time transients (Figure 3b,c). Protein adsorption is

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observed as an increase in fluorescence intensity, followed by a step-wise decrease as each of the

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dye molecules on the protein photobleach (Figure 3b). Photobleaching step and intensity

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distribution analyses are commonly used in single molecule experiments to detect single vs.

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oligomer states of proteins,58,59 including proteins with multiple dyes.60,61 The number of

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photobleaching steps identified by an 46

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established step-finding algorithm

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thus a direct measure of the BSA

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molecules per AuNR (Figure 3b, line).

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Analysis for 86 AuNRs yields an

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average of 3.2 ± 1.2 photobleaching

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steps (Figure 3c), matching the labeling

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density

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(Molecular Probes, A34785). Further

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controls

involve

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presence

of

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photobleaching

of

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(Figure

reversible

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desorption/re-adsorption (Figure S6), or

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dye fluorescence quenching close to the

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AuNRs (Video S2). In particular,

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Figure 3d confirms that despite the

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labeling with 3 dyes the BSA protein

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molecule

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multichromophoric system capable of

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simultaneous off/on blinking of all dyes

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due to energy transfer.

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indicated

3d),

does

by

the

is

supplier

eliminating

the

simultaneous

not

multiple

behave

dyes protein

as

a

Figure 3 | Single BSA proteins adsorb to single MUTABAuNRs. a, Co-localization of single MUTAB-AuNRs, confirmed by SEM (inset; scale bar, 50 nm), with fluorescently labeled BSA. Fluorescence images are 74 frames binned with the same intensity; scale bar 500 nm. AuNRs are visible through their weak intrinsic photoluminescence using long exposure times before introducing BSA b, Intensity transient (black) of BSA adsorbed to a MUTAB-AuNR with state and step identification (teal dashed line) of photobleaching steps. Simultaneous photobleaching was discounted by probability analysis (Figure S6). c, Distribution of the number of photobleaching steps for 86 transients. d, Simultaneous photobleaching steps are unlikely. For a labeling density of 3 fluorophores per BSA molecule, the average photobleaching step size as a percent of the maximum intensity should be ~33%, which is indeed confirmed by the observed average step size of 30 ± 10%. A value of > 66% indicates simultaneous photobleaching events. Out of the 213 photobleaching steps analyzed, < 4% of steps had values greater than 60%.

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proteins to single MUTAB-AuNRs is

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irreversible on a time scale of 8 hours as

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demonstrated by a constant percentage of

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AuNR-BSA

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continuously rinsing with buffer solution

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without proteins (Figure 4). It is not

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clear, however, if this observation of only

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one

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immobilized MUTAB-AuNR extends to

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the solution case. The exposed AuNR

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surface area is about twice as large in

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solution, but the protein-to-nanoparticle

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ratio is only ~26, as ~75 pM nanorods are

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exposed to 2 nM BSA. It is not trivial to

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quantify this ratio for MUTAB-AuNRs

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immobilized on a substrate, but we estimate this number to be orders of magnitude larger given

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that 2 nM BSA (~108 BSA proteins per µl) is flowed over single nanorods spaced out at least 1

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µm from each other at a flow rate of 5 µl/min for 15 min. Regardless of the experimental

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geometry (i.e. free MUTAB-AuNRs in solution vs. immobilized on a surface), there is space for

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many dozens of proteins to bind to each AuNR (width = 20 nm, length = 58 nm; resulting

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surface area of ~ 3000 nm2), assuming a native BSA conformation that can be approximated by

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an equilateral triangular prism with 2 triangular facets of ~ 28 nm2 and 3 smaller rectangular

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sides of ~ 12 nm2.53 The only hypothesis that explains how the strong, irreversible adsorption of

protein

complexes

adsorbing

while

to

each

Figure 4 | Irreversible adsorption of BSA to MUTABAuNRs. a, Representative images of BSA/MUTABAuNRs identified based on the stronger BSA signal (circles) before (No BSA), after (t = 0 min) adding a 2 nM BSA solution, and after 500 min of rinsing with buffer. False identification (No BSA) is likely due to 1 MΩ) and

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placed overnight in a water bath at 35° C. Excess MUTAB was removed by centrifugation at

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7,500 rpm for 10 minutes and replaced with Millipore H2O for storage. We estimate that the final

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concentration of free MUTAB in solution after this centrifugation step was less than 0.01 mg/ml.

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The MUTAB-AuNR solutions were positively charged (ζ = 35 ± 5 mV) at pH 7.2 according to

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zeta-potential measurements (Malvern Zen 3600). Further characterization of the MUTAB-

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AuNRs in the buffer conditions of the experiments (20 mM HEPES, 20 mM NaCl in Molecular

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Biology Grade H2O) by UV/vis spectroscopy (Figure S17-S18) and by transmission electron

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microscopy (Figure S19) are included in the SI.

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Commercially available citrate-coated gold nanospheres (AuNPs) with a nominal diameter of

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50 nm were purchased from BBI solutions (Cardiff, UK). Sizing performed by the manufacturer

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using transmission electron microscopy (TEM) showed that their actual size is 48 ± 4 nm (Batch

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# 16659, ~ 75 pM concentration based on the mass of gold per milliliter used for the synthesis).

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For the control experiments shown in Figure S7-9, citrate-AuNPs were functionalized with

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MUTAB using the same procedure as described above for the AuNRs. The zeta-potential of the

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AuNPs went from negatively charged (ζ = -35 ± 5 mV) in the presence of citrate, to positively

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charged once functionalized with MUTAB (ζ = 40 ± 5 mV), confirming the ligand exchange

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reaction occurred. Page 18 of 33

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Gold nanowires (Au nanowires, diameter ~ 70 nm, length 1 ~ 10 µm) were synthesized using a

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modified three-step seeding synthesis originally developed for AuNRs.71 Seed particles were

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prepared by a rapid reduction of HAuCl4 (gold precursor) using NaBH4 (reducing agent). These

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particles were then grown in a solution containing HAuCl4, ascorbic acid, and CTAB. The

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presence of ascorbic acid reduces the gold salt from its Au(III) to a stable Au(I) state. Further

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addition of seeds to this solution induces an autocatalytic reaction of Au(I) that enlarges the seed

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particles. While this method would typically yield nanowires with an aspect ratio smaller than

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10, aspect ratios larger than 100 can be achieved when the reaction takes place in an acidic

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environment (pH ~2) and less seeds are added to the growth solution. CTAB on the as-

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synthesized Au nanowires was replaced with MUTAB. The CTAB-Au nanowires sedimented at

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the bottom of the vial after 1-2 hours at room temperature without mixing or shaking. Carefully,

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CTAB was removed out of solution without removing the nanowires. An equal volume of

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MUTAB (1 mg/ml in Millipore H2O) was added to fill the volume left by removed CTAB. The

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solution was mixed using a micro-pipette and placed overnight in a water bath at 35° C. The

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MUTAB-Au nanowires were then re-suspended in solution by gently shaking the vial, and then

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left untouched for 1-2 hours at room temperature to allow for sedimentation. Finally, excess

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MUTAB was removed out of solution (carefully without removing the nanowires), and an equal

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volume of Millipore H2O was added to fill the volume left by excess MUTAB.

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Cell Culture. BSA (7.5% in DBPS), penicillin-streptomycin (10,000 U-10 mg/mL), HCl

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(Trace Metals Grade) and HNO3 (Trace Metals Grade) were purchased from Sigma-Aldrich.

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MCF-7 cells were acquired from ATCC (HTB-22.) EMEM (with EBSS and L-Glutamine) was

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purchased from Lonza. FBS (heat-inactivated) was purchased from Seradigm. Lactate

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dehydrogenase (LDH) activity assay kit was purchased from Thermo Scientific. All chemicals

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were used without further purification.

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Microscope glass coverslips. Glass coverslips (22 x 22 mm2, VWR) were cleaned by

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sonication for 10 minutes in acetone, 10 minutes in water/detergent mixture, and 10 minutes in

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deionized water. The coverslips were then immersed in a bath of base (6:1:1 water, 30% H2O2,

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NH4OH) for 90 seconds at 80 °C followed by rinsing with copious amounts of deionized water.

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The glass was then cleaned for 2 minutes in O2 plasma (Harrick Plasma). Cleaned coverslips

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were placed under vacuum for long term storage. Details of the passivation of the surface for

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correlation spectroscopy experiments and single protein binding experiments can be found in the

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caption of Figure S5.

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Luminescence correlation spectroscopy. Luminescence of MUTAB-AuNRs was recorded

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with a home-built confocal microscope described elsewhere.72 The instrument is based on an

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inverted epifluorescence microscope (Observer.D1, Zeiss) equipped with a 638 nm diode laser

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(CUBE, Coherent). The laser light was collimated and delivered to the back aperture of an oil

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immersion objective (Apochromat 64X, NA = 1.4, Zeiss). The objective focused the light onto a

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small liquid cell placed on top of a glass coverslip containing 45 µl of sample solution. The

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emitted light was collected by the same objective and passed through a dichroic mirror (ZT

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532/638 rpc, Chroma Technolgoy), a long pass filter (XLP-647, CVI Melles-Griot), and a 50-

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µm-diameter pinhole (Thorlabs) placed at the focal plane of the microscope. The luminescence

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signal was detected by a single element diode detector (PDM 50ct, Micro-Photon-Devices) and

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processed by a home-built photon counting module (LabView). All measurements were recorded

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mW/cm2 at the sample. This excitation power ensured minimal heating and negligible auto-

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fluorescence background of unlabeled BSA (Figure S20). For protein concentration dependent

377

measurements as summarized in Table S1, ~75 pM MUTAB-AuNR solutions with varying

378

concentrations of BSA (Catalog #A7906, Sigma-Aldrich; BSA concentrations: 0.1 nM, 0.5 nM,

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1 nM, 1.5 nM, 2 nM, 3 nM, 4 nM, 5 nM; 10 nM; BSA:AuNR ratios: 1.33, 6.66, 13.33, 20, 26.66,

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40, 53.33, 66.66, 133.33) were prepared by mixing equal volumes of both solutions in buffer.

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The dimensions of the observation volume were determined using a calibration sample with a

382

known diffusion coefficient at 20 °C (Figure S21).

383

Data acquisition and analysis for correlation spectroscopy experiments. The luminescence

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intensity was recorded in data sets of 40 s at a temporal resolution of 10 µs. Each data set was

385

autocorrelated using an automated routine in Matlab. At least seven data sets were averaged per

386

measurement and the experiment was repeated independently three times at each protein

387

concentration. The autocorrelation functions G(τ) were analyzed in Matlab using a one species,

388

three dimensional diffusion model with an additional exponential term that accounts for nanorod

389

rotation:73

390

1 G (τ ) = N

 τ 1 +  τD

  

−1

  r 2 τ 1 +  0    z 0  τ D 

   



1 2

τ −  τR  ⋅ 1 + Ae  

   

391

where denotes the average number of particles in the three-dimensional Gaussian-shaped

392

observation volume, with radial and axial dimensions r0 and z0, respectively. The translational

393

diffusion time τD is related to the translational diffusion coefficient of the particles, Dtr =

394

r02/(4τD). An additional exponential term of amplitude A and rotational diffusion time τR was

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used to account for nanorod rotational diffusion. The translational diffusion coefficient was used Page 21 of 33

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to estimate the equivalent hydrodynamic radius of the nanoparticles using the Stokes-Einstein

397

relationship Rh = kT/6πηDtr. Changes in viscosity due to the presence of BSA were calculated

398

using the intrinsic viscosity of this protein5 (4.2 cm3/g) assuming a linear relationship in the range

399

of concentrations used. Results of correlation spectroscopy experiments before and after BSA

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binding at different concentrations are shown in Table S1.

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Super-localization single molecule microscopy. Samples containing single immobilized

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MUTAB-AuNRs (Figures 2, 3) and MUTAB-AuNPs (Figure 2) were prepared by dropcasting

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~40 µl of ~12.5 pM of nanoparticle solutions (1:6 dilution of the concentration used for

404

luminescence correlation spectroscopy experiments) on microscope coverslips passivated with

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unlabeled BSA (see caption of Figure S5 for details on the passivation procedure). The droplet

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was then dried for 10 minutes at room temperature, and the coverslips were rinsed with 1 ml of

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Millipore H2O and gently dried with nitrogen gas. A home-built total internal reflection wide

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field microscope with 637 nm excitation (Coherent OBIS-FP 637 LX) as described elsewhere74

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was used with a custom flow chamber (1 mm height, elliptical opening of 12×5 mm; 43018C,

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Grace BioLabs) placed over the sample containing immobilized MUTAB-nanoparticles. A

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solution of 2 nM Alexa647-labeled BSA (A34785, Molecular Probes, 3 dyes/protein) was flowed

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over the sample at 5 µl/min using a Genie Plus flow system (Kent Scientific). This flow rate does

413

not impart strong forces on protein-binding site interactions and results in effectively negligible

414

net directed diffusion due to flow.75 The sample was allowed to equilibrate for 15 min with

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protein flow before measuring, ensuring that an excess of proteins is exposed to each

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nanoparticle (2 nmol BSA/l x 5 ul/min x 15 min x NA = 9 x 1010 BSA molecules compared to

417

~10 particles/5 µm2 in Figure S3 x 60 mm2 area of sample = 12 x 105 particles/sample). Data was

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rate of 7.5 Hz, and electron multiplying gain of 300. Analysis was possible at the super-

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localization level, as the fluorescent BSA can be selectively observed only when adsorbed to the

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interface, and was unobservable by motion blur when freely diffusing (D ∼ 60 µm2/s). Increases

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in signal to noise ratio of each frame, identification of adsorbed BSA, and super-localization

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analysis of the location by radial symmetry fitting was performed using MATLAB 2011b as

424

described previously.57,68 Samples containing single immobilized MUTAB-Au nanowires

425

(Figure 2, 5c, S14) were prepared using the same drying procedure, but because the

426

concentration was unknown, the solution was repeatedly diluted until single isolated nanowires

427

were observed on the microscope coverslip under the optical microscope.

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Circular dichroism (CD) spectroscopy. Ensemble CD measurements were taken on a J-815

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circular dichroism spectrometer (JASCO) with a 1 mm pathlength quartz cuvette at 20 ± 0.1 °C

430

using a Peltier temperature controller. Equal volumes of ~1 nM nanoparticle solutions (MUTAB-

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AuNRs or AuNPs) and 0.1 mg/ml BSA solutions were mixed ~30 minutes before each

432

measurement. Wavelength ranges of 190-260 nm and 500-750 nm were collected every 1 nm

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under standard sensitivity, bandwidth of 1.00 nm, and rate of 50 nm/min. Four scans were

434

collected for each trial and averaged. The spectra were baseline-corrected and presented as mean

435

residue ellipticity. The solvent for CD measurements was 1 mM phosphate buffer (pH 7.2) due

436

to its low absorbance in the far UV. This solvent did not affect the stability of MUTAB-AuNRs,

437

MUTAB-AuNPs or BSA.

438

SEM imaging. SEM images were taken using a FEI Quanta 400 ESEM FEG, operated at 10

439

kV under low vacuum conditions. Analysis of aggregate size was performed in ImageJ.

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MUTAB-AuNR uptake by MCF-7 cells. Page 23 of 33

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(a)

Pre-incubation of MUTAB-AuNRs under different media conditions before

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incubation with MCF-7 cells. For the experiments shown in Figure 1a, 1b, MUTAB-

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AuNRs were incubated in two different media conditions: 1) 1 mL of MUTAB-AuNRs

444

were mixed with 133 μl of 7.5% (w/v) BSA (final concentration of BSA ~1 % w/v) and

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incubated overnight. This formed a BSA-corona onto the MUTAB-AuNRs before exposure

446

to serum. Next, the BSA-MUTAB-AuNRs were diluted (1:10) in EMEM with 10% FBS

447

(with 1% PCN) and incubated overnight. 2) 1 mL of MUTAB-AuNRs were directly diluted

448

(1:10) in EMEM with 10% FBS (with 1% PCN) and incubated overnight. Both MUTAB-

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AuNR solutions were found to be stable in EMEM over the timescale of the experiments,

450

as observed via UV/Vis spectroscopy.

451

(b)

Page 24 of 33

Nanoparticle incubation with MCF-7 Cells. MCF-7 cells were seeded into two 6-well

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plates with each well containing 2 ml of 100,000 cells/ml solution and cultured for three

453

days in EMEM with 10% FBS and 1% PCN (same as incubation conditions for MUTAB-

454

AuNRs). On the third day, the media was removed and each well was carefully washed

455

three times with PBS to remove adhered proteins. Two wells were filled with 2 ml media

456

containing either 1% BSA, 10% FBS in EMEM with 1% PCN, or 10% FBS in EMEM with

457

1% PCN. The remainder of the wells contained 2 ml of the corresponding BSA-MUTAB-

458

AuNR solutions, each in triplicate. The plates were incubated at 37° C in 4.5% CO2 for 6

459

hours. After incubation for 6 hours, 50 μl of media from each well was removed in

460

triplicate using a pipette and kept for the LDH assay. The wells were washed with PBS

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three times to remove adhered proteins and nanorods. The cells were then detached via

462

incubation in 0.05% trypsin with EDTA and counted using a standard hemacytometer

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(Figure S22). After counting, the cells were transferred to a solution of 0.1% Triton X-100

464

and placed in a -20 °C freezer overnight.

465

(c)

LDH cytotoxicity assay of MCF-7 cells incubated with BSA-MUTAB-AuNRs. A LDH

466

assay was performed by adding 50 μl of formazan dye (Thermo Scientific) to each well

467

containing media from the nanoparticle-cell incubation experiments (Figure S23). The

468

plates were placed in an incubator at 37 °C with 4.5% CO2 for 30 minutes and read on a

469

BioTek Uniread 800 plate reader. The purpose of this assay was to measure the cytotoxicity

470

of the MUTAB-AuNRs under the different incubation conditions with BSA and FBS,

471

compared to MCF-7 cells alone.

472

(d)

ICP-MS measurement of BSA-MUTAB-AuNRs uptaken by MCF-7 cells. The lysed

473

cells were thawed and centrifuged at 8000 g for 10 minutes. The supernatant was removed

474

and the pellets were digested using aqua regia (3:1 solution of HNO3 and HCl). Following

475

digestion, the pellets were diluted to 10 ml using 5% HCl in milli-Q water. The samples

476

were measured for Au concentration using a Perkin-Elmer Nexion 300 ICP-MS. The

477

concentration of Au was determined for each cell using the cell count collected after

478

detaching the cells (Figure S24).

479

Calculation of the number of AuNRs uptaken by each cell. The average number of AuNRs

480

uptaken per cell was calculated by first determining the number of AuNRs in the different

481

solutions with cells from the concentration of Au determined via ICP-MS. The number of

482

nanorods was then divided by the average number of cells. The addition of BSA to the MUTAB-

483

AuNRs increases the uptake by MCF-7 cells by 320% compared to the conventional incubation

484

conditions in EMEM with 10% FBS. Page 25 of 33

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485

CONFLICT OF INTEREST

486

The authors declare no competing financial interests.

Page 26 of 33

487 488

ACKNOWLEDGEMENTS

489

This work was funded by the National Science Foundation (Grants CBET-1134417 and CHE-

490

1151647 to C. Landes; Grant CBET-1438634 to S. Link and C. Landes; Grant DMR-1105878 to

491

E. Zubarev), the Welch Foundation (Grants C-1787 to C. Landes and C-1664 to S. Link), and the

492

National Institute of Health (Grant GM94246-01A1 to C. Landes). L. Kisley and A. Hoggard

493

acknowledge the National Science Foundation Graduate Research Fellowship (Grant 0940902).

494

S. Dominguez-Medina acknowledges partial support from a Lodieska Stockbridge Vaughn

495

Fellowship from Rice University. We would like to thank Professors Peter Wolynes, Naomi

496

Halas, and Carsten Sönnichsen for their valuable feedback.

497 498

SUPPORTING INFORMATION AVAILABLE

499

Details regarding experimental & computational methods and data analysis. This material is

500

available free of charge via the Internet at http://pubs.acs.org.

501 502

AUTHOR PRESENT ADDRESSES

503

# - CEA-Grenoble; 17 Rue des Martyrs; F-38054 Grenoble Cedex 9, France

504

∆ - University of Illinois at Urbana-Champaign; School of Chemical Sciences; 600 S. Mathews

505

Ave.; MC-712, Box C-3; Urbana, IL 61801

506

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