Erythrocyte Incubation as a Method for Free-Dye Presence ...

Article pubs.acs.org/molecularpharmaceutics

Erythrocyte Incubation as a Method for Free-Dye Presence Determination in Fluorescently Labeled Nanoparticles Patrizia Andreozzi,† Chiara Martinelli,†,‡ Randy P. Carney,§ Tamara M. Carney,§ and Francesco Stellacci*,§ †

Fondazione IRCCS Istituto Neurologico Carlo Besta at IFOM (Fondazione Istituto FIRC di Oncologia Molecolare)European Institute of Oncology (IEO) Campus, Via Adamello 16, 20139 Milan, Italy ‡ IFOM (Fondazione Istituto FIRC di Oncologia Molecolare)European Institute of Oncology (IEO) Campus, Via Adamello 16, 20139 Milan, Italy § Institute of Materials, École Polytechnique Fédérale de Lausanne (EPFL), 1015 Lausanne, Switzerland S Supporting Information *

ABSTRACT: The field of nanotheranostics encompasses the integration of nanosized carriers in cancer imaging, diagnosis, and therapy. The use of nanomedicines for theranostic application typically depends on direct visualization of the nanocarriers. Normally fluorescent probes are attached to nanocarriers for biodistribution measurement through fluorescence imaging. However continued, noninvasive assurance that the fluorescent probe remains bound to the carrier has proven elusive. Mature erythrocytes, also known as red blood cells, are incapable of endocytosis. As a consequence, when incubated with fluorescently labeled particles, they do not show any signal coming from the membrane or the cytoplasm. Yet, these cells readily take up free BODIPY fluorescent dyes into their membranes. Here we show that incubation of nanoparticles with erythrocytes is a rapid and reliable method for the detection of unbound dye present within a nanoparticle sample, as the detection of a fluorescent signal coming from the cells can only be due to unbound dye present in the sample. We test the method on both sulfonate and PEG terminated gold nanoparticles, and we determine the minimum concentration of detectable dye for a specific gold nanoparticle sample. KEYWORDS: amphiphilic nanoparticles, cell internalization, cell uptake, unbound dye, nanoparticle drug delivery, thiolated dyes

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INTRODUCTION The development of biocompatible nanomaterials for cell imaging, tracking, and drug delivery is currently of great interest.1−4 The application of these nanomaterials for the diagnosis and study of the development of cancer has resulted in a surge for the field of nanotheranostics.5−7 A variety of inorganic core nanomaterials surrounded by a biocompatible ligand shell have been synthesized as diagnostic imaging tools and transport vessels for drug delivery or gene therapy.8 These applications typically require bright fluorescence signals for noninvasive live cellular imaging and biodistribution tracking.5,8 Although some types of nanomaterials, such as quantum dots (QDs), are inherently fluorescent, they remain limited in application due to their toxicity and difficulty in surface functionalization.8 Other systems, such as fluorescent core− shell silica nanoparticles (NPs),9,10 overcome issues of biocompatibility and ease of functionalization, yet remain unsuitable for some applications due to their large size, surface chemistry, or hemolytic activity.11,12 Instead, a large class of nanotheranostic tools are inherently nonfluorescent and employ fluorescent dyes as noncovalently bound labels at their surface.2,8,13,14 In the case of plasmonic particles, the core © 2012 American Chemical Society

material, e.g., gold, can quench, in part or totally, the fluorescence of the dye.15,16 When the dye is released from the surface of the nanomaterial, it regains fluorescence.13 The actual functionalization scheme of dye-labeled nanomaterials depends on the chemistry of the core material, but typically a bifunctional surface molecule, known as a ligand, is used. These ligands reserve one terminal end for immobilization onto the nanomaterial (such as thiol functionalization for gold or silver NPs, or carboxylic acid functionalization for iron oxide NPs) with the opposite terminus available for biomolecule functionality, such as the attachment of fluorescent dye molecules. On gold NPs, for example, dye molecules can be prethiolated and place exchanged onto the NP surface.17,18 Although ligands are normally considered moderately stable with respect to detachment over time, it is currently challenging Special Issue: Theranostic Nanomedicine with Functional Nanoarchitecture Received: Revised: Accepted: Published: 875

September 19, 2012 November 20, 2012 November 28, 2012 November 28, 2012 dx.doi.org/10.1021/mp300530c | Mol. Pharmaceutics 2013, 10, 875−882

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to ensure the continued attachment of fluorescent ligands to the nanomaterial, or the removal of free dye from the functionalization experiment. The collection and recharacterization of the nanomaterial just prior to (or after) application is generally necessary to ensure dye viability, an impractical and sometimes impossible restriction. Contamination of a sample with free dye from the labeling procedure is an issue that materializes persistently, yet not homogeneously in all experiments. It is therefore necessary to develop a control experiment for quickly detecting free dye molecules in the presence of dyefunctionalized nanotheranostic carriers. Commonly, centrifugation is used to purify bound dye−particle assemblies from unreacted free dye. Yet this method provides no feedback that the free dye has been truly removed. The method of detection we propose is to control that the free dye has been removed after purification by centrifugation. This experiment should be run in parallel with in vitro experiments on other types of cells on the dye-labeled sample in use. Erythrocytes, also known as red blood cells (RBCs), serve the primary purpose of delivering oxygen to the tissues in vertebrates. Mature mammalian RBCs characteristically lack nuclei and other intracellular organelles such as mitochondria, distinguishing them from most other mammalian eukaryotic cells. In addition, these cells do not have phagocytic receptors on their surface and therefore cannot employ endocytosis, the uptake of proteins or foreign material at the cell surface.19 For this reason RBCs are commonly used as a model for nonphagocytic/nonendocytic cells. 19 Many studies have indicated that small molecules (such as fluorescent dyes) have the ability to partition into the erythrocyte membrane.20,21 Boron-dipyrromethene (BODIPY) containing fluorescent dyes, originally developed for imaging application due to their high brightness, environment-independent fluorescent yields, and sharp excitation and emission peaks, are known to readily insert at trace levels into the RBC plasma membrane.21 This renders erythrocytes as a unique system that is unable to take up most nanomaterials yet can take up small amphiphilic molecules into their plasma membranes. We propose to exploit this property to develop RBCs as a quick screen to detect free dye in an aliquot of dye-functionalized nanomaterials. Previous work has explored the interaction of nanomaterials with RBCs for a variety of applications, including nanopore generation,22 passive membrane penetration mechanisms,23 enhanced-contrast blood flow imaging,24 and hemocompatibility.25−28 Nanoparticles have also been adhered to the outer surface of RBCs to improve their in vivo circulation time.29,30 Yet most of the recent interest at the intersection of RBCs and nanotheranostics has been devoted to the separation and use of RBC ghost cells. These naked RBC membranes can be isolated after osmotic hemolysis of RBCs in diluted, alkaline buffer.31 Returning the swelled membranes to hypertonic conditions, the RBCs shrink and crenate, leaving behind only empty membranes, or ghost cells. The emerging field of erythrocyteinspired delivery32 has primarily utilized RBC ghosts to encapsulate proteins, genetic material, cancer chemotherapeutics, and nanomaterials, dramatically increasing their circulation in the body and bioavailability.32−35 These types of cells are easily identifiable under confocal microscopy compared to normal RBCs and were excluded from quantification in this study, due to possible complexities arising from NP−ghost cell interactions. Here we present a protocol for the detection of free dye in a sample of fluorescently labeled nanoparticles. We test the

method on sulfonate-terminated NPs commonly used by our group for cell penetration studies. These NPs are soluble in water and can be easily functionalized with BODIPY dye by known place-exchange procedures. We additionally test the procedure on perfluorinated-PEG coated gold particles functionalized with FITC dye. After the assurance that the coincubation of nanoparticles with RBCs yields no qualitative changes to the RBC morphology, we systematically measure the fluorescence of the RBC membrane using confocal laser scanning microscopy (CLSM) and flow cytometry (FC) upon incubation with different concentrations of NPs and free dye. The underlying idea is simple. For particles that do not enter RBCs, incubation with nanoparticles leads to no fluorescence. Only when free dye is present, fluorescence signal from the RBCs is detected. Hence nanoparticles’ RBC incubation can be a rapid detection method for free dye in nanoparticle samples.

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MATERIALS AND METHODS Experimental Materials. 6-(((4,4-Difluoro-5-(2-thienyl)4-bora-3a,4a-diaza-s-indacene-3-yl)styryloxy)acetyl)aminohexanoic acid, succinimidyl ester (BODIPY 630/650-X, SE) was purchased from Life Technologies and thiolated through the succinimidyl ester group to yield BODIPY-SH. 11Mercaptoundecane sulfonate (MUS) and all NPs were synthesized in house.14 All other chemicals were purchased from Sigma-Aldrich. All solvents were reagent grade and were purged with nitrogen gas at least 1 h prior to use. All cell culture reagents were purchased from Life Technologies. All the procedures followed the Strain Laboratory Animal Care (Directive 86/609/EFC) at the live animal facility at IFOM. Source and Isolation of RBCs. Freshly drawn mouse blood was collected in EDTA coated tubes to minimize coagulation. Whole blood was then centrifuged at 800 RCF for 5 min and the plasma and buffy coat were removed. The packed red blood cells were washed in DPBS (Dulbecco’s phosphate buffered saline solution) three times and resuspended in a final volume of 10 mL of DPBS. This stock solution was stored at 4 °C. Nanoparticle Synthesis and Fluorescent Labeling. 0.9 mmol of gold salt (HAuCl4) was dissolved in 150 mL of ethanol under nitrogen flow. 0.9 mmol of thiolated ligand mixture (11-mercaptoundecane sulfonate, MUS, and MUS:octanethiol (OT), 2:1 mol:mol) was dissolved in 10 mL of methanol and subsequently added to the gold solution. After 5 min stirring, 150 mL of a saturated sodium borohydride (NaBH4) in ethanol solution was added dropwise to the gold/ thiol mixture. The solution was then stirred for 3 h under nitrogen flow on ice and placed in a refrigerator overnight to precipitate. The particles were collected via vacuum filtration with quantitative filter paper, washed with ethanol, methanol, and acetone, and dried under vacuum. 20 mg aliquots of NPs were dissolved in Milli-Q purified water and dialyzed with centrifugal dialysis membranes (10,000 Da MWCO) in order to remove free ligands, salts, and synthetic side products (as confirmed by 1H NMR). A stock solution of BODIPY-SH dye in DI-H2O/DMF (2:1 vol:vol) was used to place exchange an 80-fold molar excess onto the NPs (dissolved in 750 μL of DIH2O) over 2 days. The reaction was purified of excess dye by centrifugation. Three drops of the dye−NP solution was diluted to 2 mL with acetone in an eppendorf tube and centrifuged at 13,000 rpm for 5 min. Free Dye and Nanoparticle Incubation with RBCs. The RBC stock solution was diluted 1:100 in DPBS. For the free 876

dx.doi.org/10.1021/mp300530c | Mol. Pharmaceutics 2013, 10, 875−882


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Figure 1. Fluorescent dye place-exchange reaction. Gold NP structure during place-exchange reaction. The fluorescent BODIPY-SH molecules are mixed with monolayer protected NPs in molar excess for 2 days at room temperature.

oil-immersion objective (HCX PL APO 63X Lbd Bl, Leica Microsystems) was employed for analysis. MetaMorph 7.7 software (2011 Molecular Devices) was used for image quantification analysis according to previously described methodology.36 Briefly, CLSM images were exported to MetaMorph 7.7 software (2011 Molecular Devices), and regions of interest (ROIs) were hand-drawn around the circumference of the cell using the transmitted image for guidance (Figure 1 in the Supporting Information). The intracellular fluorescence intensity was calculated by subtraction of the background and normalized per unit area. Between 50 and 300 cells were quantified and averaged per image. Flow cytometry data were acquired with a Becton Dickinson LSR II SORP analyzer using the 640 nm red laser and optical filter 670/14 nm. Data were collected in triplicate (15,000 events each) with the following settings: forward scatter (FSC) 320, side scatter (SSC) 280, and red filter 610. Data were analyzed using the same gates on the SSC, FSC plots in FlowJo 7.6 software, and the median fluorescence intensities and standard deviations were exported and averaged for each treatment.

dye experiments, 6 serial dilutions, each decreasing by 10× concentration from an initial 10 mM stock of BODIPY-SH, were prepared in DPBS. For the RBC incubations, 600 μL of the diluted RBC stock solution was mixed with 15 μL of each BODIPY-SH dilution, plated in 8-well ibidi μ-slide chambers (ibidiTreat) and incubated at 37 °C for 45 min. After incubation the solutions were transferred to microcentrifuge tubes and spun at 800 RCF for 3 min. The supernatant was removed, and the RBC pellets were washed three times with 200 μL of fresh DPBS. The final washed samples were either returned to ibidi μ-slide chambers for CLSM acquisition or used directly with FC. Unwashed samples were used directly after incubation. For nanoparticle incubations, dried NPs were prepared in an aqueous solution to a final concentration of 5 μM as previously reported.36 Freshly filtered particle solutions (0.22 μm2 PTFE syringe filters, in DPBS) were directly incubated with RBCs. Timing, concentration, and washing were performed identically to the aforementioned dye experiments. The cells were left in the last wash and immediately imaged by CLSM or used directly with FC. Labeled Nanoparticles and Free Dye Incubation with RBCs. The RBC stock solution was diluted 1:100 in DPBS. Labeled NPs were prepared at four different concentrations (0.2 mg/mL, 0.1 mg/mL, 0.05 mg/mL, 0.01 mg/mL) in DPBS. We assume the molecular weight of the particles is ∼1 MDa; therefore mg/mL and μM can be used interchangeably. Free BODIPY solutions at differing BODIPY/NP ratios (10, 1, 0.1, 0.01) were prepared immediately before addition to the NP solutions. The BODIPY/NP mixtures were added to the RBC solutions and monitored by CSLM from time 0 to time 60 min. All experiments were performed at 37 °C. CLSM/FC Acquisition and CLSM Image Quantification. Confocal microscopy was performed on a Leica TCS SP5 confocal microscope equipped with blue (Argon, 488 nm) and red (633 nm HeNe Laser) excitation laser lines. A 63/1.4 NA

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RESULTS AND DISCUSSION For this study, homoligand sulfonate-terminated gold nanoparticles (MUS NPs) or heteroligand sulfonate-methyl terminated nanoparticles (MUS:OT 2:1) were used as sample nanotheranostic carriers that can be dye labeled. Their synthesis and composition are described in the Materials and Methods. MUS NPs have been used in our research group as a control NP against MUS:OT surface-structured NPs, and both types of NPs are well characterized and their typical behavior with cells is well documented.14,36,37 These NPs can be functionalized by known place-exchange procedures to incorporate secondary ligands such as thiolated dyes into their protecting shell (Figure 1).18 We have previously used BODIPY (630/650) dye functionalized in house with a thiol linker to make BODIPY877



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Figure 2. Free dye incubations with red blood cells. A dilution series of free BODIPY-SH dye was incubated with RBCs for less than 1 h. The cells were washed of free dye and imaged by confocal microscopy (top panel). BODIPY fluorescence was quantified from the CLSM images (blue bars) and by flow cytometry (red bars). The dynamic range of detectable free BODIPY-SH dye was established between 1 mM and 1 μM.

Figure 3. Fluorescently labeled nanoparticles and free dye incubations with HeLa and red blood cells. Both MUS (a, b, g, h) and MUS:OT NPs (e, f, k, l) were dye-labeled and incubated at 37 and 4 °C with either HeLa or RBCs. Free BODIPY-SH dye was also incubated for 1 h with HeLa and RBCs at the same concentration as found on the NPs (0.1 mM). The striking difference lies in the lack of penetration from MUS:OT NPs in RBCs compared to HeLa cells. The previously proposed energy-independent mechanism acting to internalize MUS:OT NPs in HeLa cells, even at 4 °C (f), is not found with RBCs (l). The free dye however does not change between HeLa and RBC incubation (d compared to j). This renders erythrocyte incubation a useful tool to check for the presence of free dye in labeled NP solutions.

SH.14,36 The functionalization of the NPs with BODIPY-SH is monitored through fluorescence microscopy of the monolayerprotected nanoparticles or through measurement of the supernatant solution’s absorbance following dye exchange.18

The reduced concentration of dye in the supernatant compared to the initial stock solution incubated with the NPs is used to calculate the concentration of dye attached to the NP, using a calibration curve measured for the BODIPY-SH (Figure 2 in 878



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were previously with HeLa, both in this study (Figure 3e,f) and in previous ones.36 The free dye however (Figure 3c,d compared to Figure 3i,j) retained identical fluorescence regardless of HeLa or RBC incubation. This is a key result for the development of this method, as it renders erythrocytes as a cell line where the behavior of the free dye and that of the NP are clearly different and easily distinguishable in fluorescence imaging. So far we have assumed that a lack of fluorescent signal from clean, dye-labeled NPs in RBCs means that they do not penetrate RBCs as they do with other cell types. It should be stressed that this article is not the place to discuss if these MUS:OT particles truly do not interact with RBCs. Based on the experiments and controls we used in this manuscript to test these NPs with RBCs, we have observed that they appear not to enter these cells in a similar manner as previous cell lines, yet we have to state that further tests are needed to confirm this point. Nevertheless, the key idea that we have developed here is that when a nanoparticle sample is incubated with RBCs, a fluorescent signal can only be detected if free dye is present. To test this assumption we used a batch of dye-labeled NPs that we suspected could contain traces of free dye, possibly due to incomplete washing procedures or release of dye molecules over time from the ligand shell. Figure 4 shows that after

the Supporting Information). Under standard conditions, this place exchange results in nanoparticles that contain very few dye molecules per NP; a good approximation is to assume ∼10 dye molecules per particle. In order to first test the interaction of free BODIPY-SH dye with RBCs, a dilution series of BODIPY-SH solutions, ranging from the concentrated stock (1 mM) to a 1:103 (1 μM) dilution, was incubated with RBCs. The median concentration of 0.1 mM represents the quantified amount of BODIPY-SH functionalized to NPs at the concentration used for standard cell−NP incubations (0.1 μM), such as the dye-labeled NPs in Figure 3. The fluorescence intensities were measured and quantified by both CLSM and FC of the BODIPY-SH serial dilutions and are presented in Figure 2. At the minimum concentration of 1 μM BODIPY-SH dye, the fluorescence intensity is comparable to the control experiment with no dye. Overall, this dilution series represents the dynamic range of the detection of free BODIPY-SH dye with erythrocytes. Following RBC incubation with free BODIPY-SH dye, the dye was added via place-exchange reaction onto the surface of the NPs (Figure 1). The reaction conditions used were such that 10 dye molecules per MUS:OT and MUS nanoparticles were supposed to be present in the ligand shell (2 day mixing of 80-fold molar excess of dye to NPs).36 The particles were thoroughly cleaned by repeated centrifugation in a polar solvent (see Materials and Methods). The dye-labeled particles (0.1 μM) were incubated with HeLa cells at 37 and 4 °C. Figure 3a shows that at 37 °C MUS particles produce a fluorescence signal, while at 4 °C (Figure 3b) no signal is present as at this temperature no energy dependent mechanism occurs, in accordance with our previous studies using these same NPs.14,36 These images alone provide a good indication that no/little dye is released from the nanoparticles, compared to free BODIPY-SH incubation with HeLa cells in which a diffuse fluorescence pattern is obtained at both 37 and 4 °C (Figure 3c,d). In the case of MUS:OT particles the images are somewhat different, as at both 37 °C (Figure 3e) and 4 °C (Figure 3f) a diffuse fluorescence background can be seen. Using transmission electron microscopy,14 and more recently photothermal heterodyne imaging,38 we have been able to interpret that, for MUS:OT particles, there exists an additional energy-independent penetration mechanism, initiated by a fusion mechanism with the cell membrane.39 However, it is clear that resorting to additional microscopy techniques that are independent of fluorescent labeling for each experiment like the ones in Figure 3e and Figure 3f is both time and resource consuming. There is a need for a quick and reliable method to determine if free dye is present in a nanoparticle sample. The shelf life of monolayer protected NPs after synthesis is on the order of weeks to months, and so previously tested particle batches are continuously resynthesized. This need for free dye testing exists not only for wholly new types of nanoparticles but also for these batches of particles that have fully been previously tested but newly synthesized. As reliable as a cleaning procedure can be, it is not reasonable to expect a 100% success rate. To address this problem, given that free dye freely penetrates in RBCs (Figure 3i,j), we devised the method that we are illustrating here. We took the dye-labeled particles used to produce the images in Figure 3 and incubated them for 1 h with RBCs. As shown in Figure 3g,h,k,l no fluorescence signal was observed for both types of particles. Notably, the dye-labeled MUS:OT NPs were not internalized into RBCs at 37 or 4 °C (Figure 3k,l), as they

Figure 4. Washing removal of free dye on labeled nanoparticles. A sample of BODIPY-SH labeled MUS nanoparticles containing a small amount of unbound dye was incubated with RBCs (a). The same sample was washed to remove the unbound dye and reincubated with RBCs (b). After washing, confocal microscopy reveals the absence of any remaining free dye.

incubation with RBC a diffuse fluorescence signal is clearly visible. We washed the same particle batch by centrifugation in acetone and incubated again with RBC for 1 h. The result this time was that of no diffuse fluorescence signal. We concluded that the incubation with RBCs could be a good test for the presence of free dye, and for testing the success of subsequent washing steps to remove any free dye. In order to test this in a controlled manner, a washing experiment was designed. A typical place-exchange procedure for dye-labeled NPs begins with the mixing of the thiolated free dye with NPs. An equilibrium develops between dye molecules that attach to the surface of the NP and dye molecules that remain in solution. The entire sample is centrifuged so that dye-labeled NPs are pelleted and the remaining free dye is left behind in the supernatant. After one spin, the pelleted dyelabeled NPs were divided from the supernatant containing the remaining free dye, and both were separately incubated with 879



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Figure 5. Dye-labeled NP incubations with red blood cells. Following the dye place-exchange reaction, the supernatants (a−c) and dye-labeled NP pellets (d−f) of successive washes were incubated with RBCs. All of the unbound dye is removed in the first wash.

absorb to RBCs with a similar pattern to BODIPY. Afterward we demonstrate that we can detect free dye in a sample of unpurified dye-labeled AU-F8-PEG NPs, and that we can remove this by washing procedures (Figure 5 in the Supporting Information). Due to the similarity in behavior with RBCs between the two NP−dye samples, we believe that this method is quite general for the case of metallic NPs with ligand shells that can be modified by place exchange, provided that the type of dye used interacts with the membrane of erythrocytes. The method we are proposing is to use RBCs as a test for free-dye presence in nanoparticle batches. Of course any method has its own detection limit. Here we assume that, for a decently cleaned nanoparticle batch, free dye will come only from the NPs themselves, hence the NP concentration will determine the concentration of the free dye. To determine the detectable levels of free dye we incubated RBCs with particles freshly mixed with thiolated free dye. In one case we tested freshly prepared mixtures of free dye with MUS:OT particles and in another with dye-labeled MUS:OT particles. In the latter case we tested that the batch used initially gave no fluorescence signal. Comfortably, we found that already after 1 h of incubation with free dye, when using MUS:OT particles, little to no fluorescence signal was observed in RBCs, indicating a higher affinity of the dye to the particles than to penetration in the cells (Figure 4 in the Supporting Information). This means that a simple release of dye from the particles will result in readsorption of the dye onto another particle, and thus significant release will happen only in the case of imperfect cleaning. To find the detection limit we then limited ourselves to RBC incubations with dye-labeled MUS:OT particles freshly mixed

RBCs. The results of this experiment are presented in Figure 5. The controls are RBCs alone (Figure 5a) and unwashed NP/ dye mixed solution (Figure 5d). Following the first wash, the supernatant contains all of the free dye (Figure 5b), while the pelleted NPs are free of any unbound dye (Figure 5e). To further confirm this, the washing step was repeated and again the supernatant solution (Figure 5c) and the NP pellet (Figure 5f) were incubated separately with RBCs. Both were then free of unbound dye. To ensure that the NPs were not losing dye over time while suspended in solution, the same NPs used for Figure 5 were further incubated with RBCs in successive experiments over a period of 7 days. At no time was free dye detected (Figure 3 in the Supporting Information), indicating that the dye remains bound to the NP surface, at least above the detection limit of our methodology. Significant NP aggregation did occur after 24 h. Additionally nanoparticle batches as much as 2 months old were tested with RBCs and HeLa cells, and they showed no fluorescence with the former and the usual diffuse fluorescence images with the latter (data not shown). In order to assess the generality of this method for dye labeled particle systems we have repeated the experiments in this paper with another type of gold nanoparticle with a different surface composition and labeled with a different type of dye molecule. We used AU-F8-PEG NPs that were coated with a perfluorinated-PEG type of amphiphilic ligand. These NPs have been previously shown to retain water solubility despite the increased hydrophobic nature of the ligand, and as such could have interesting application in biological systems.40 The AU-F8-PEG NPs were labeled with another hydrophobic dye, fluorescein isothiocyanate (FITC), that we first show can 880



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composition of blood (Figure 6 in the Supporting Information).

with thiolated free dye. The BODIPY-SH dye/NP/erythrocyte mixture was incubated at 37 °C for 1 h and imaged with confocal microscopy under the same conditions as the previous experiments. Given that we sought to determine detection limits and validity of this method we (1) used the most common NP concentrations adopted in various past experiments with our particles and (2) used free dye concentrations that resulted in reasonable amount of free dye molecules for particles. For example we assumed that any decent cleaning step would remove an amount of dye that is in excess of 10 dye molecules per particle. We incubated RBCs with various concentrations of particles and dye molecules in excess or defect of number of particles (e.g., from 10 dye molecules per NP, down to 100 NPs per 1 dye molecule). Each incubation was imaged with confocal microscopy in order to quantify the amount of remaining free dye staining the RBCs. The result is a 3D plot shown where the fluorescence intensity is plotted as a function of NP concentration and the number of BODIPY-SH molecules per NP (Figure 6). This plot, while specific to the

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CONCLUSION The control experiment we propose in this work is of general applicability and relevance for the field of nanotheranostics. We show that incubation of nanoparticles with erythrocytes is a good tool to determine if free dye is present. As only free dye can penetrate (in an efficiently and detectable way) RBCs, the presence of fluorescence signal inside these cells is a clear indicator of free dye presence in a dye-labeled NP sample. Furthermore, we establish a procedure for developing design rules for the minimum concentration of NPs that should be tested with RBCs to ensure the detection of free dye molecules.

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ASSOCIATED CONTENT

S Supporting Information *

Additional information on how to quantify the dye using MetaMorph software, calibration curves for the BODIPY dye, more washing control experiments, data using another type of NP−dye system, and whole blood incubations. This material is available free of charge via the Internet at http://pubs.acs.org.

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AUTHOR INFORMATION

Corresponding Author

*EPFL, Institute of Materials, EPFL STI IMX SUNMIL, Station 12, 1015 Lausanne, Switzerland. E-mail: francesco. stellacci@epfl.ch. Tel: +41 21 6937872. Fax: +41 21 6935270. Notes

The authors declare no competing financial interest.

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ACKNOWLEDGMENTS We gratefully acknowledge Dr. Paola Alberici (IFOM, Milan) for providing fresh blood samples. We also thank the imaging facility (IFOM, Milan), in particular Dr. Sara Barozzi, Dr. Amanda Oldani, Dr. Massimiliano Garrè, and Dr. Dario Parazzoli, for providing excellent technical assistance and helpful discussion. P.A. would like to acknowledge support from CariPisa, C.M. from AIRC-IG11723, and R.P.C. from the Swiss National Foundation NRP 64 program. Finally we would like to thank Lucia Pasquato and Silvia Bidoggia from the University of Trieste for providing the AU-F8-PEG-FITC particles.

Figure 6. Extent of particle dye labeling as a function of particle and dye concentrations. A series of 4 concentrations of NPs (blue, 0.01 μM; green, 0.05 μM; purple, 0.1 μM; and red, 0.2 μM) and 4 ratios of BODIPY-SH/NP (0.01, 0.1, 1, and 10) were used to measure the extent of BODIPY labeling of NPs in the presence of RBCs. This diagram provides the minimum nanoparticle concentrations in order to detect free dye for a given number of dye molecules per particle.

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MUS:OT NPs we used for this study, provides a convenient tool to quantify the extent of free dye detection as a function of NP concentration. We can for example conclude that to detect the presence of one free dye molecule per 10 NPs it is necessary to test a concentration of at least 0.2 μM, while a concentration of 0.1 μM is necessary for 10 dye molecules per NP. From this plot we can also conclude that when using concentrations lower than 0.05 μM a simple cleaning procedure is sufficient to ensure that no free dye artifact can affect the results of an incubation study of NPs with cells. Finally, a successful theranostic approach should not lose its usefulness upon the increased complexity of in vivo environments. To determine if our method could serve as a viable tool even in living organisms we tested the protocol with whole blood, i.e., freshly drawn blood containing a mixture of erythrocytes, leukocytes, platelets, and plasma. We found that dye-labeled particles remained suspended in solution without significant aggregation and that free dye was readily adsorbed by erythrocytes, even when they represent less than 50% of the

REFERENCES

(1) Ferrari, M. Cancer Nanotechnology: Opportunities and Challenges. Nat. Rev. Cancer 2005, 5, 161−171. (2) Ghosh, P.; Han, G.; De, M.; Kim, C.; Rotello, V. Gold Nanoparticles in Delivery Applications. Adv. Drug Delivery Rev. 2008, 60, 1307−1315. (3) Farokhzad, O. C.; Langer, R. Impact of Nanotechnology on Drug Delivery. ACS Nano 2009, 3, 16−20. (4) Youan, B. B. C. Impact of Nanoscience and Nanotechnology on Controlled Drug Delivery. Nanomedicine (London, U.K.) 2008, 3, 401−406. (5) Lammers, T.; Kiessling, F.; Hennink, W. E.; Storm, G. Nanotheranostics and Image-Guided Drug Delivery: Current Concepts and Future Directions. Mol. Pharmaceutics 2010, 7, 1899−1912. (6) Peer, D.; Karp, J. M.; Hong, S.; FarokHzad, O. C.; Margalit, R.; Langer, R. Nanocarriers as an Emerging Platform for Cancer Therapy. Nat. Nanotechnol. 2007, 2, 751−760. (7) Davis, M. E.; Chen, Z.; Shin, D. M. Nanoparticle Therapeutics: An Emerging Treatment Modality for Cancer. Nat. Rev. Drug Discovery 2008, 7, 771−782. 881



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(8) Xie, J.; Lee, S.; Chen, X. Y. Nanoparticle-Based Theranostic Agents. Adv. Drug Delivery Rev. 2010, 62, 1064−1079. (9) Liong, M.; Lu, J.; Kovochich, M.; Xia, T.; Ruehm, S. G.; Nel, A. E.; Tamanoi, F.; Zink, J. I. Multifunctional Inorganic Nanoparticles for Imaging, Targeting, and Drug Delivery. ACS Nano 2008, 2, 889−896. (10) Mullner, M.; Schallon, A.; Walther, A.; Freitag, R.; Muller, A. H. E. Clickable, Biocompatible, and Fluorescent Hybrid Nanoparticles for Intracellular Delivery and Optical Imaging. Biomacromolecules 2010, 11, 390−396. (11) Lin, Y. S.; Haynes, C. L. Impacts of Mesoporous Silica Nanoparticle Size, Pore Ordering, and Pore Integrity on Hemolytic Activity. J. Am. Chem. Soc. 2010, 132, 4834−4842. (12) Zhao, Y. N.; Sun, X. X.; Zhang, G. N.; Trewyn, B. G.; Slowing, I. I.; Lin, V. S. Y. Interaction of Mesoporous Silica Nanoparticles with Human Red Blood Cell Membranes: Size and Surface Effects. ACS Nano 2011, 5, 1366−1375. (13) Hong, R.; Han, G.; Fernandez, J. M.; Kim, B. J.; Forbes, N. S.; Rotello, V. M. Glutathione-Mediated Delivery and Release Using Monolayer Protected Nanoparticle Carriers. J. Am. Chem. Soc. 2006, 128, 1078−1079. (14) Verma, A.; Uzun, O.; Hu, Y. H.; Hu, Y.; Han, H. S.; Watson, N.; Chen, S. L.; Irvine, D. J.; Stellacci, F. Surface-Structure-Regulated CellMembrane Penetration by Monolayer-Protected Nanoparticles. Nat. Mater. 2008, 7, 588−595. (15) Dulkeith, E.; Morteani, A. C.; Niedereichholz, T.; Klar, T. A.; Feldmann, J.; Levi, S. A.; van Veggel, F. C. J. M.; Reinhoudt, D. N.; Moller, M.; Gittins, D. I. Fluorescence Quenching of Dye Molecules near Gold Nanoparticles: Radiative and Nonradiative Effects. Phys. Rev. Lett. 2002, 89. (16) Sapsford, K. E.; Berti, L.; Medintz, I. L. Materials for Fluorescence Resonance Energy Transfer Analysis: Beyond Traditional Donor-Acceptor Combinations. Angew. Chem., Int. Ed. 2006, 45, 4562−4588. (17) Daniel, M. C.; Astruc, D. Gold Nanoparticles: Assembly, Supramolecular Chemistry, Quantum-Size-Related Properties, and Applications toward Biology, Catalysis, and Nanotechnology. Chem. Rev. 2004, 104, 293−346. (18) Templeton, A. C.; Cliffel, D. E.; Murray, R. W. Redox and Fluorophore Functionalization of Water-Soluble, Tiopronin-Protected Gold Clusters. J. Am. Chem. Soc. 1999, 121, 7081−7089. (19) Rothen-Rutishauser, B. M.; Schurch, S.; Haenni, B.; Kapp, N.; Gehr, P. Interaction of Fine Particles and Nanoparticles with Red Blood Cells Visualized with Advanced Microscopic Techniques. Environ. Sci. Technol. 2006, 40, 4353−4359. (20) Sheetz, M. P.; Singer, S. J. Biological-Membranes as Bilayer Couples - Molecular Mechanism of Drug-Erythrocyte Interactions. Proc. Natl. Acad. Sci. U.S.A. 1974, 71, 4457−4461. (21) Holowka, D.; D′auria, L.; Van Der Smissen, P.; Bruyneel, F.; Courtoy, P. J.; Tyteca, D. Segregation of Fluorescent Membrane Lipids into Distinct Micrometric Domains: Evidence for Phase Compartmentation of Natural Lipids? PLoS ONE 2011, 6, 17021. (22) Delcea, M.; Sternberg, N.; Yashchenok, A. M.; Georgieva, R.; Baumler, H.; Mohwald, H.; Skirtach, A. G. Nanoplasmonics for DualMolecule Release through Nanopores in the Membrane of Red Blood Cells. ACS Nano 2012, 6, 4169−4180. (23) Wang, T. T.; Bai, J.; Jiang, X.; Nienhaus, G. U. Cellular Uptake of Nanoparticles by Membrane Penetration: A Study Combining Confocal Microscopy with Ftir Spectroelectrochemistry. ACS Nano 2012, 6, 1251−1259. (24) Ahn, S.; Jung, S. Y.; Seo, E.; Lee, S. J. Gold NanoparticleIncorporated Human Red Blood Cells (Rbcs) for X-Ray Dynamic Imaging. Biomaterials 2011, 32, 7191−7199. (25) Asharani, P. V.; Sethu, S.; Vadukumpully, S.; Zhong, S.; Lim, C. T.; Hande, M. P.; Valiyaveettil, S. Investigations on the Structural Damage in Human Erythrocytes Exposed to Silver, Gold, and Platinum Nanoparticles. Adv. Funct. Mater. 2010, 20, 1233−1242. (26) Dobrovoiskaia, M. A.; Clogston, J. D.; Neun, B. W.; Hall, J. B.; Patri, A. K.; McNeil, S. E. Method for Analysis of Nanoparticle Hemolytic Properties in Vitro. Nano Lett. 2008, 8, 2180−2187.

(27) Cótica, L. F.; Freitas, V. F.; Dias, G. S.; Santos, I. A.; Vendrame, S. C.; Khalil, N. M.; Mainardes, R. M.; Staruch, M.; Jain, M. Simple and Facile Approach to Synthesize Magnetite Nanoparticles and Assessment of Their Effects on Blood Cells. J. Magn. Magn. Mater. 2012, 324, 559−563. (28) Creangă, D. E.; Culea, M.; Nădejde, C.; Oancea, S.; Curecheriu, L.; Racuciu, M. Magnetic Nanoparticle Effects on the Red Blood Cells. J. Phys. Conf. Ser. 2009, 170, 012019. (29) Chambers, E.; Mitragotri, S. Long Circulating Nanoparticles Via Adhesion on Red Blood Cells: Mechanism and Extended Circulation. Exp. Biol. Med. 2007, 232, 958−966. (30) Fan, W.; Yan, W.; Xu, Z.; Ni, H. Erythrocytes Load of Low Molecular Weight Chitosan Nanoparticles as a Potential Vascular Drug Delivery System. Colloids Surf., B 2012, 95, 258−265. (31) Schwoch, G.; Passow, H. Preparation and Properties of Human Erythrocyte-Ghosts. Mol. Cell. Biochem. 1973, 2, 197−218. (32) Hu, C.-M. J.; Fang, R. H.; Zhang, L. Erythrocyte-Inspired Delivery Systems. Adv. Healthcare Mater. 2012, 1 (5), 537−547. (33) Antonelli, A.; Sfara, C.; Manuali, E.; Bruce, I. J.; Magnani, M. Encapsulation of Superparamagnetic Nanoparticles into Red Blood Cells as New Carriers of MRI Contrast Agents. Nanomedicine (London, U.K.) 2011, 6, 211−223. (34) Hu, C. M. J.; Zhang, L.; Aryal, S.; Cheung, C.; Fang, R. H.; Zhang, L. F. Erythrocyte Membrane-Camouflaged Polymeric Nanoparticles as a Biomimetic Delivery Platform. Proc. Natl. Acad. Sci. U.S.A. 2011, 108, 10980−10985. (35) Brahler, M.; Georgieva, R.; Buske, N.; Muller, A.; Muller, S.; Pinkernelle, J.; Teichgraber, U.; Voigt, A.; Baumler, H. MagnetiteLoaded Carrier Erythrocytes as Contrast Agents for Magnetic Resonance Imaging. Nano Lett. 2006, 6, 2505−2509. (36) Carney, R. P.; Carney, T. M.; Mueller, M.; Stellacci, F. Dynamic Cellular Uptake of Mixed-Monolayer Protected Nanoparticles. Biointerphases 2012, 7. (37) Jewell, C. M.; Jung, J. M.; Atukorale, P. U.; Carney, R. P.; Stellacci, F.; Irvine, D. J. Oligonucleotide Delivery by Cell-Penetrating ″Striped″ Nanoparticles. Angew. Chem., Int. Ed. 2011, 50, 12312− 12315. (38) Leduc, C.; Jung, J. M.; Carney, R. P.; Stellacci, F.; Lounis, B. Direct Investigation of Intracellular Presence of Gold Nanoparticles Via Photothermal Heterodyne Imaging. ACS Nano 2011, 5, 2587− 2592. (39) Van Lehn, R.; Autokorale, P.; Carney, R. P.; Voitchovsky, K.; Stellacci, F.; Alexander-Katz, A. Bilayer Penetration by Surface Monolayer Protected Gold Nanoparticles. Unpublished. (40) Gentilini, C.; Evangelista, F.; Rudolf, P.; Franchi, P.; Lucarini, M.; Pasquato, L. Water-Soluble Gold Nanoparticles Protected by Fluorinated Amphiphilic Thiolates. J. Am. Chem. Soc. 2008, 130, 15678−15682.

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