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Development of Bright and Biocompatible Nanoruby and its Application to Background-free Time-gated Imaging of G-protein Coupled Receptors Varun K. A. Sreenivasan, Wan Aizuddin W Razali, Kai Zhang, Rashmi R Pillai, Avishkar Saini, Denitza Denkova, Marina Santiago, Hannah Brown, Jeremy Thompson, Mark Connor, Ewa M. Goldys, and Andrei V Zvyagin ACS Appl. Mater. Interfaces, Just Accepted Manuscript • DOI: 10.1021/acsami.7b12665 • Publication Date (Web): 12 Oct 2017 Downloaded from http://pubs.acs.org on October 13, 2017

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ACS Applied Materials & Interfaces

Development of Bright and Biocompatible Nanoruby and its Application to Background-free Time-gated Imaging of G-protein Coupled Receptors Varun K. A. Sreenivasanχ*, Wan Aizuddin W Razaliψχ, Kai Zhangρχ, Rashmi R. Pillaiχ, Avishkar Sainiϕ, Denitza Denkovaχ, Marina Santiagoϐ, Hannah Brownϕ, Jeremy Thompsonϕ, Mark Connorϐ, Ewa M. Goldysχ, Andrei. V. Zvyaginϖχ χ

Department of Physics and Astronomy and Centre for Nanoscale BioPhotonics, Macquarie University, NSW 2122, Australia;

ϕ

Robinson Research Institute and Centre for Nanoscale BioPhotonics, Adelaide Medical School, University of Adelaide, SA 5005, Australia; ϐ

Department of Biomedical Sciences, Macquarie University, NSW 2122, Australia;

ϖ

Institute of Molecular Medicine, Sechenov First Moscow State University, Moscow 119991 and

Institute of Biology and Biomedicine, Lobachevsky Nizhny Novgorod State University, Nizhny Novgorod 603022, Russia Keywords: nanoruby, opioid, GPCR, time-gated microscopy, single-particle Corresponding Author

ACS Paragon Plus Environment

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*Single Molecule Science, Level 3 Lowy, University of New South Wales, NSW 2052, Australia. E-mail: [email protected] Present Addresses ψ

Current Affilitation: Faculty of Applied Science, Universiti Teknologi MARA Cawangan

Pahang, 26400 Bandar Tun Abdul Razak Jengka, Pahang, Malaysia ρ

Current Affilitation: State Key Laboratory of Supramolecular Structure and Materials, Jilin

University, Changchun 130012, P. R. China


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Abstract

At the forefront of development of fluorescent probes for biological imaging applications are enhancements aimed at increasing their brightness, contrast, and photostability, especially towards demanding applications of single molecule detection. In comparison with existing probes, nanorubies exhibit unlimited photostability and a long emission lifetime (~ 4 ms), which enable continuous imaging at single-particle sensitivity in highly scattering and fluorescent biological specimens. However, their wide application as fluorescence probes has so far been hindered by the absence of facile methods for scaled-up high-volume production and molecularly-specific targeting. The present work encompasses large scale production of colloidally stable nanoruby particles, demonstration of their biofunctionality and negligible cytotoxicity, as well as validation of its use for targeted biomolecular imaging. In addition, optical characteristics of nanorubies are found to be comparable or superior to state-of-the-art quantum dots. Protocols of reproducible and robust coupling of functional proteins to the nanoruby surface are also presented. As an example, NeutrAvidin-coupled nanoruby show excellent affinity and specificity to µ-opioid receptors in fixed and live cells, allowing wide-field imaging of G-protein coupled receptors with single particle sensitivity.


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1. Introduction Functional biological imaging of molecular processes represents a powerful experimental approach in the life sciences, particularly in cell biology and pharmacology.1 Recent advances in fluorescence microscopy have enabled real-time imaging, with spatial and temporal resolutions improved down to tens of nanometers and several milliseconds, respectively.2-3 However, singlemolecule optical microscopy instrumentation remains largely restricted to advanced optical laboratories2, 4-5 and operates within the narrow margins of specific applications, such as singlereceptor tracking on the surface of neurons.6-8 One of the key challenges in the observation, analysis and quantification of biomolecular events, particularly on a single molecule level, is the intense autofluorescence background produced by cells and biological tissue as well as the strong emission from other intentionally introduced fluorophores highlighting cellular processes of simultaneous interest. Appropriate selection of fluorescent probes and excitation/emission configurations can help obviate unwanted effects of the autofluorescence background. For example, the choice of fluorescent probes with excitation/emission spectral bands in red and near-infrared (e.g. carbon nanotubes, far-red dyes, quantum dots and genetically engineered proteins)9-11 allows suppression of biological specimen autofluorescence, which is commonly characterized by excitation/emission in UV-blue-green spectral range, and hence improves the imaging sensitivity. However, near-infrared (NIR) probes requires specialized detectors and optics tailored to minimize aberrations and losses, especially for single molecule sensitivity. Multiphoton12 and up-conversion13-14 microscopy employing the excitation in the NIR spectral range (800 - 1000 nm) also allows minimization of the sample autofluorescence at the expense of high excitation intensities (10-100 W.cm-2 in upconversion up to MW.cm-2 in multiphoton microscopy) and long, seconds or more, acquisition times.


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Amongst the imaging methods to discriminate faint single molecule labels from these interfering signals, time-gated imaging offers enhancement of imaging contrast by orders of magnitude.15-16 Time-gated imaging is carried out by synchronizing the pulsed excitation and detection time-windows, with a delay (gate) time between the termination of the excitation laser pulse and the onset of image acquisition. The gate allows short (nanosecond) lifetime background to decay before signal from the photoluminescent probes with longer (e.g. millisecond) emission lifetimes ߬௣௟ is collected. Ruby nanocrystals that we demonstrated earlier exhibit large action cross-sections in combination with millisecond lifetimes and it is uniquely suited as a label for time-gated imaging.17 Nanorubies exhibit unfading photoluminescence10,

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in contrast to organic

fluorophores which photobleach within seconds and semiconductor quantum dots that exhibit intermittent emission (blinking), limiting the duration and reliability of imaging and tracking experiments. The photostability of nanoruby enables continuous imaging, limited only by instrumentation.10, 17 Photoluminescence of nanoruby originates from Cr3+ color-centers located within the physically and chemically robust host material of α-phase alumina.18 These colorcenters are isolated from biological and chemical microenvironments, including potentially confounding factors such as pH variation, which, for example,

affects the fluorescence

properties of fluorescent organic dyes. The emission band of nanoruby in the deep red spectral range is narrow (693 ± 2 nm), and this enables simple spectral unmixing with other colocalized fluorophores. The fluorescence of nanoruby can be optimally excited using standard 405, 532 and 561 nm laser sources.16 These properties make nanorubies well-suited for long-term, highsensitivity imaging applications. The adoption of nanoruby has thus far been limited due to limited availability, difficulties in large scale production, poorly characterized surfaces, lack of


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biofunctionalization and bioconjugation methods, uncharacterized cytotoxicity, and scarcity of demonstrated applications in the life sciences. Taking advantage of these unique properties of nanoruby,16 we now introduce a nanorubybased imaging platform for conjugation to receptors of interest, by reporting its scaled-up production, surface engineering and the attachment of functional targeting biomolecules. Firstly, we demonstrate the facile production of large quantities of nanorubies by high-energy ball milling at a high yield,19 followed by simple postprocessing steps to obtain bright, photostable and colloidally stable nanoruby within the size range of 40-70 nm. These nanorubies were nontoxic to cultured cells and to very susceptible early mouse embryos. We report protocols for nanoruby surface functionalization using silica and silane-based reagents. The silica-coating allowed stable conjugation of a streptavidin analogue, which facilitated further molecular targeting in fixed and live cells. Using this approach, we demonstrate specific labeling and ultrasensitive imaging of µ-opioid receptors, an important class of G-protein coupled receptors.

2. Results and Discussion 2.1. Production of nanoruby by high energy ball milling Several physical (e.g. ball milling, pyrolysis,20 laser ablation,17,

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combustion22-23) and

chemical (e.g. sol-gel24-25 resin synthesis26) methods have been developed to produce nanoparticles of α-alumina and ruby, with or without additional processing steps. Most of these studies focused on large-scale preparations for engineering applications and did not address the purity, colloidal stability or surface functionality of the as-produced nanoparticles, which are crucial for applications in life sciences. Other methods, including a method reported by us,17 resulted in production of colloidally stable nanorubies, but with a low yield. This work aimed at


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both increasing a production yield by high-energy ball milling (HEBM) and improving colloidal properties of nanorubies by simple processing steps. The HEBM production of nanoalumina and related materials has been previously demonstrated, achieving low levels of contamination as well as a high production volume and yield with the use of appropriate milling media and process control agents (PCAs).19,

23, 27-30

Yttria-stabilized zirconia (hereafter, referred to as zirconia) has been identified as the best commercially available milling medium to attain the lowest contamination for milling alumina,19, 27

because of the matching Mohs hardness indices of 9 for zirconia and alumina (following

diamond at 10). PCAs increase the effectiveness of HEBM by influencing the balance between three competing processes: comminution, aggregation and de-aggregation.31 PCAs, including sugar,32 salt,32 surfactants,33-34 gas,35 fatty acids

36

and organic solvents

33

have been reported to

improve milling for various materials. We explored different PCAs to optimize the milling conditions for ruby. Salt (sodium chloride) and sugar (glucose) shown to be effective in stabilizing nanoparticles2223, 32

did not result in appreciable size-reduction of ruby particles (data not shown). In contrast,

three wet PCAs (water, a non-ionic surfactant Pluronic F-127, and low pH) previously shown to increase milling efficiency19,

31, 33

were more promising (Figure 1a). Further characterization

showed milling at low pH results in a rapid decrease of the particle size to < 100 nm, with a concomitant increase in the fraction of particles smaller than 100 nm. In comparison, Pluronic was not effective as a PCA. Milling in water resulted in a marginal size reduction, where the fraction of particles under 100 nm was less than 50% even after 5 h milling. Nanoruby particles produced at low pH and in water had sizes of 80 and 140 nm with polydispersity indices of 0.18 ± 0.03 and 0.24 ± 0.05, and zeta-potentials >> 30 mV.


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Figure 1. Effects of milling media on nanoruby properties. (a) Time evolution of colloidal properties of nanorubies milled in water, low pH and Pluronic, showing (top) hydrodynamic diameter, (middle) yield or the number fraction of particles of size less than 100 nm and (bottom) zeta-potential. Milling in Pluronic, water and low pH-based milling were carried out once, twice and four times, respectively. Comparison of (b, c) crystallinity, (d) contamination and (e, f) morphology of nanorubies milled in water or at low pH. (b) X-ray diffraction spectra of nanorubies milled for 300 min in water or at low pH. Peaks matching α-alumina are marked blue. Peaks at 18.4°, 20.3° and 53° originate from aluminum hydroxide in the form of bayerite (18.6° and 20.4°) and gibbsite (18.4° and 53°). (c) Williamson-Hall plot of nanorubies milled in


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water and low pH. Open and solid circles demarcate samples milled for 60 and 300 min. Dashed (60 min) and solid lines (300 min) are linear fit to the data. (d) Fractional mass of zirconia expressed in percentage. Error bars represent standard deviation of two independent repeats for low pH milling and 3 independent repeats for milling in water. The low pH milling was characterized in detail only once beyond 180 min. (e, f) TEM images of nanoruby samples milled for 300 min in (e) water or (f) low pH. Scale bars, 100 nm.

The size-reduction of alumina particles resulting from HEBM at a low pH in comparison with that in water has been attributed to a combination of two factors.31 First is the formation of aluminum hydroxide and amorphous alumina on the particle surface.19, 31, 37 We confirmed this formation by detecting 18.4°, 20.3° and 53° peaks in the x-ray diffraction (XRD) spectrogram (Figure 1b), representative of hydroxide. The formed hydroxide layer rendered the nanoruby surface plastic and deformable, reducing the milling energy transferred to the crystals and contributing towards the initiation and propagation of fractures. The dissolution of the unwanted hydroxide layer was promoted by low pH (4 - 5),31, 38 thus exposing the crystallite surface to direct the impact during the milling. Note: Figure 1b shows the presence of hydroxide nanorubies even at low-pH milled, because of formation of gibbsite platelets upon extraction into water after milling. These platelets are visible as hexagonal crystals39 in transmission electron microgram (TEM) in Figure 1f. XRD peak at 20.3° also suggests the presence of bayerite in this sample, but these were not observed in TEM. Such crystals were absent when ruby samples were milled in water (Figure 1e), because hydroxides in the form of bayerite (XRD peaks at 18.4°, 20.3°) remained bound to the nanoruby particles at neutral pH. The second factor, as explained in Ref

31

, is related to the amphoteric property of alumina (ruby) which has an isoelectric point


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between 9 – 11 (Figure S1c). Therefore, milling at low pH conditions increases the colloidal stability, and decreases aggregation or cold welding. Taken together, these two factors resulted in a net increase in the effectiveness of the comminution. The Williamson-Hall analysis of the XRD peaks corresponding to α-alumina indicated that milling at low pH resulted in significantly smaller crystals than milling in water. Increasing intercepts in Figure 1c between 60 and 300 min of milling indicates the reduction of the crystallite size from 290 ± 150 nm to 100 ± 80 nm, when milled in water, and from 45 ± 80 nm to 13 ± 11 nm, when milled at low pH. The uncertainty in the estimation of crystallite sizes originates from the high polydispersity of the particle size distribution and the aspect ratios of raw nanoparticles.

2.2. Reduction of zirconia and gibbsite contamination and improving colloidal stability by AAA process The choice of PCA also affected the chemical composition of the milled product (Figure 1d) as measured by Energy dispersive x-ray spectroscopy (EDS). We have shown that EDS provides an accurate measurement of chemical composition of milled nanomaterials and agrees with mass spectroscopy data19. EDS revealed the presence of significant quantities of zirconium, likely in the form of zirconia, which increased with milling time. We used a previously demonstrated method of sulfuric acid-based etching to reduce the zirconia contamination.19 In order to suppress the accompanying aggregation (Figure 2a,c), we developed a washing procedure to remove anionic sulfate groups adsorbed onto the positively charged nanoruby surface at neutral pH.19, 40 To this aim, we increased the pH of the colloid during washing to 12, exceeding the isoelectric point of nanoruby (Figure S1c), to render it negatively charged. In the subsequent


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washing step, we used hydrochloric acid to lower the pH to 2, close to the pKa2 of sulfuric acid (1.92). Upon dispersing acid-alkali-acid (AAA) processed nanorubies in water, the zeta-potential returned to a high value of 51 mV and remained stable for months (Figure 2a, 20 d post treatment). We confirmed the reproducibility of the AAA method by testing samples milled for 120, 180 and 300 min, yielding an 8-fold reduction of the zirconia content in all samples (Figure 2b).

Figure 2. Purification and colloidal stabilization of as-milled nanorubies. (a) Hydrodynamic size and zeta-potential of nanoruby during AAA procedure, measured in water. Error bars represent standard deviation. (b) Zirconia contamination in nanoruby milled for 120, 180 and 300 min, before and after AAA processing. The contamination is expressed in weight percent zirconia/(zirconia+alumina). TEM images of nanoruby (c) after sulfuric acid treatment and (d) after AAA processing. Scale bars, 100 nm. (e) XRD spectra of nanoruby at the start (top, black) and after AAA (bottom, red) procedure. Blue lines on the top indicate peaks that match these of corundum. Pink lines mark peaks originating from hydroxides. (f) Williamson-Hall plot of


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nanorubies obtained using data in (e). Lines represent linear fit to the data, with R2 = 0.15 and 0.69 at the start and after AAA, respectively. While the detailed characterization was only carried out once, the AAA procedure was carried out thrice independently and yielded similar TEM micrographs and DLS values.

The increase in colloidal stability after the AAA procedure was also observable in TEM micrographs (Figure 2c, d), where particles appear well-separated. Hexagonal gibbsite platelets visible in Figure 1f are also absent in Figure 2c, d. The AAA process also led to narrowing of the peaks in XRD spectra, with R2 of the Williamson-Hall fit increasing from 0.16 to 0.69 (Figure 2e, f). The mean crystal grain size was estimated to be 60 ± 40 nm, matching that measured by dynamic light scattering (DLS, 60 ± 20 nm). XRD also confirmed that colloidal particles were mostly single crystals. We attribute the size uncertainty of 20-40 nm to the anisotropic sheet-like structure of nanoruby deduced by comparing TEM and atomic force microscopy images (Figure S2, S3 – Supporting Information figure). We estimated that 3 to 5-h milling, followed by the AAA processing yielded, approximately, 200 mg of high-quality colloidally-stable nanoruby sample from 3-g micron-sized ruby as starting material. Since smaller nanoparticles in the size range of 10 to 70 nm are the most useful for molecular labeling applications, the data presented hereafter were obtained using the low-pH-milled, AAAwashed nanorubies, unless otherwise stated.

2.3. Photoluminescence of nanorubies compared with quantum dots The as-produced nanorubies exhibited a sharp ≈4-nm wide photoluminescence spectral peak centered at 693 nm, with no broad peaks characteristic of γ phase observable (Figure 3a, Figure


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S4).17 Our spectral acquisition window of 5 nm did not allow resolving two closely spaced zero-phonon lines originating from the transitions of tri-valent chromium ions in the α-alumina crystal matrix.10 The excitation spectra (not shown) also resembled that of bulk ruby, with two broad peaks centered at 402 nm and 557 nm.16

Figure 3. Photoluminescence properties of nanoruby. (a) Photoluminescence emission of nanoruby centered at 693 nm falls within the biological tissue transparency window (white region), where the cumulative absorption of hemoglobin and water is at minimum.41 (b) Specific fluorescence intensity (per mass) of two nanoruby samples NR2 and NR8 before and after AAA processing, respectively. (c) Probability and (d) cumulative distributions of the single particle


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brightness of NR2, NR8 and 705 QDot. (e) Time-gated (left) and normal (right) photoluminescence images of nanorubies in cells featuring optical background progressively graded as Low, Medium and Very High. Note, the pixel values of Very High normal image (bottom right panel) divided by 8. The corresponding time-gated image is noisy, because it was acquired with same low-gain acquisition parameters to maintain a large dynamic range. (f) Ratio of signal to backgrounds of nanoruby spots correlated in time-gated and normal images in (e). At very high background levels, nanoruby spots could not be found in normal imaging, thus the value practically approaches infinity.

We compared the brightness, defined as the detectability over background, of nanoruby with that of state-of-the-art commercial quantum dots (refer to Experimental Section for details on the quantification of brightness). An earlier study based on colloidal measurements showed that 60-nm nanorubies exhibited photoluminescence signals comparable to those of 20-nm commercial quantum dots.10 However, inferences from ensemble measurements required assumptions about the density and size of nanoparticles. Here, we measured single particle photoluminescence of quantum dots (705 QDot) and nanorubies of two types, containing either 0.15% or 0.8% Cr3+ (NR2 or NR8 respectively) deposited on coverslips by spin-coating. We confirmed that the spin-coating yielded well-separated single particles by replicating the procedure on TEM grids (Figure S2). Since it is possible that spin-coating on TEM grids might not fully represent spin-coating on a coverslip, we compared the brightness of spin-coated nanorubies to that of nanorubies confirmed to be singles by atomic force microscopy (AFM) images (Figure S3 and Supporting Information (SI)).


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Photoluminescence images of NR2, NR8 and 705 QDot used for single particle quantification are shown in Figure S2. Histograms (probability distributions) and cumulative distributions of the single particle brightness were obtained by analyzing 800 particles of QDot, 1245 particles of NR2 and 265 particles of NR8 (Figure 3c, d). The cumulative distributions show that NR8 particles are approximately 30% as bright as 705 QDot, and NR2 particles are approximately 30 - 40% as bright as NR8, under continuous (normal) excitation/emission imaging conditions. Colloidal photoluminescence measurements also indicated that NR8 was approximately 3-fold brighter than NR2 (Figure 3b), where the enhanced brightness following the AAA process is attributed to the removal of contamination. While quantum dots are brighter than nanorubies under these conventional, non-time-gated, imaging circumstances, the long emission lifetime (~4 ms, Figure S4) can be exploited to dramatically improve the imaging contrast, particularly in samples with high optical background. Figure 3e compares the time-gated and conventional images of nanorubies in a cellular environment with varying levels of optical background. Time-gating resulted in an improvement of the signal-to-background ratio (2 fold to large unquantifiable values), depending on the optical background, as quantified in Figure 3f. Most notably, nanorubies, which were not detectable in the presence of very high background, became easily detectable by switching to the time-gated mode of imaging.

2.4. Functionalisation of nanoruby using silane reagents The described optimization of the nanoruby sample production protocols resulted in its excellent aqueous colloidal stability. However, large aggregates formed once the sample was suspended in saline containing physiological concentrations of salt (Figure 4b). When


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suspended in anionic buffers, including PBS, nanoruby also acquired a negative charge due to the adsorption of phosphate ions,40 as illustrated in Figure 4d. This presents a problem, as labeling applications require stable colloids in physiologically relevant solvents.

Figure 4. Silane-based biofunctionalization of nanoruby, showing that the bonding was unstable and easily hydrolyzed. (a) FTIR spectra of NR (AAA-processed) before and after biofunctionalization with three different concentrations of sil-PEG. Black arrows indicate the expected peaks. (b) Hydrodynamic size and (c) zetapotential of nanoruby before, immediately after and three weeks after functionalization with sil-PEG. Error bars are standard deviation of measurements from two functionalization repeats (d, e) Schematic diagram of surface charge (+ and – symbols) and slipping plane (dashed line) of (d) NR and (e) NR-PEG in water and PBS. Detailed colloidal characterization was carried out twice, but results were routinely reproducible.


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Introducing steric stabilization using polyethylene glycol (PEG) and other block polymers is a common strategy to increase colloidal stability when the electrostatic stabilization is insufficient.42 Therefore, we tested the stabilization of nanorubies by their functionalization with polyethylene glycol containing reactive silane-groups (sil-PEG). Firstly, we confirmed that nanorubies, like nanoalumina 19, 43-44 and silica, were reactive to silane groups (Figure S1). Figure 4a indicates the presence of C-H groups on PEG-functionalized nanoruby (NR-PEG) at three concentrations of sil-PEG reagent. The PEG-ylation enhanced the colloidal stability of nanorubies in PBS (Figure 4b) and in cell culture media (data not shown), albeit an observable increase of the hydrodynamic size in water, possibly due to the expansion of the fluid slipping plane, as illustrated in Figure 4d, e. A comparison of the zeta-potentials before and after the PEG functionalization further confirmed the successful functionalization. We also verified that this technique allowed to introduce reactive functional groups, such as biotin, to form NR-PEG-biotin (data not shown). Following functionalization, the nanoruby colloids were stable in PBS for up to three weeks (Figure 4b, c). The observed deterioration of the functionalization properties was attributed to the hydrolysis of Al-O-Si bonds,45 as indicated by a reversing trend of the zeta-potential and size (in both water and PBS) towards those of unfunctionalized nanorubies.

2.5. Biocompatibility of nanorubies We tested the potential toxicity of nanoruby in two biological systems: in mouse pituitary tumor cell line (AtT-20 cells) and in early murine embryos. AtT-20 cells were incubated overnight with unprocessed nanorubies milled in water at three different concentrations, and the viability was measured using MTS assay. The results shown in Figure 5 indicate no signs of


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cytotoxicity up to the maximum assessed concentration of 100 µg.mL-1. This corroborated our earlier observation on the lack of toxicity of nanoalumina.19 The result also indicated that zirconia, which constituted ≈20% of this sample, exerted no detectable stress to cells. Next, we tested cytotoxicity of PEG-functionalized AAA-processed nanorubies (NR-PEG) to early murine embryos, which are highly sensitive to environmental conditions.46 No statistically significant difference in the embryonic development was observed after 2 or 5 days of the nanoruby incubation (Figure 5). To clarify whether the observed inertness of nanorubies was due to their extracellular localization and reduced extent of interaction with developing embryos, we imaged the nanoruby-treated embryos (Figure S5) and confirmed the embryos contained internalized nanorubies. Thus, nanorubies appeared to be non-toxic at the concentrations below 100 µg.mL-1, even in the presence of up to 20% zirconia contamination.

Figure 5. Viability of AtT-20 cells and murine embryo development rate upon nanoruby incubation. Values are expressed as percentage of the maximum with (vehicle) control. The viability of AtT-20 cells was assessed 24 h post nanoruby (or vehicle) incubation. The embryo development was assessed 2 d or 5 d post nanoruby (or vehicle) treatment. Error bars represent standard deviation for test on AtT-20 cells and embryos carried out in 4 and 5 experimental replicates, respectively. Data from embryo experiments follow normal distribution with marginal


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variations in the variance, and one-way ANOVA analysis did not show any statistical difference (P2.5 h of incubation using a plate reader (PHERAstar FS, BMG Labtech). The absorption background was measured by using values from wells containing MTS reagent, but without cells. We verified that nanorubies did not measurably contribute to the 490-nm absorption. Four experimental repeats showed no difference between treatments.

4.7. Biocompatibility in early mouse embryos All experiments were approved by The University of Adelaide Animal Ethics Committee (M2015-072) and were conducted in accordance with the Australian Code of Practice for the Care and Use of Animals for Scientific Purposes. CBA x C57Bl/6 F1 hybrid mice were housed within Laboratory Animal Services (University of Adelaide, Australia) under controlled temperature, photoperiod (14h l: 10h d) and with water and feed ad libitum. Three-week old female mice were superovulated with 5 IU equine chorionic gonadotropin (eCG; Folligon, Intervet) administered i.p., followed 46 h later by 5 IU human chorionic gonadotrophin (hCG/Pregnyl; Merck). Females were then mated with males of the same strain O/N. Copulation plugs were checked to confirm mating and 23 h later, mice were culled via cervical dislocation and one cell presumptive zygotes (PZs) were collected from the ampulla of the reproductive tract into Cook Research Wash Media (Cook Medical). All embryo culturing was performed in Cook Research Cleave media and embryos were cultured in humidified gas incubators (6% CO2, 5% O2 & 89% N2 at 37 °C) in groups of 10 in


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20-µL drops overlaid with paraffin oil. To examine the toxicity of nanorubies, embryos were cultured from PZ to blastocyst (4 d) in the presence of 1, 10 and 100 µg.mL-1 nanoruby samples. Nanorubies used were functionalized with sil-PEG as described above to render them colloidally stable in culture media. 2-cell embryo development was assessed at 24 h (cleavage rate) and blastocysts assessed at 118 h, and graded (early, expanded or hatching). Data were arc-sine transformed prior to analysis. One-way ANOVA statistical analysis with appropriate post-hoc was conducted on all treatment groups using Graph Pad Prism Version 7 for Windows and significance was assumed at p < 0.05. Data were presented as mean + SEM. The blastocyst rate was calculated as a percentage of 2 cell embryos.

4.8. Conjugation of azide-functionalised nanoruby with NeutrAvidin Firstly, NeutrAvidin was tagged with a copper-free click chemistry counterpart, dibenzocyclooctyne. Under gentle vortexing, 4-fold molar excess of dibenzocyclooctyne-NHS ester (DBCO-NHS, Nanocs) prepared in dimethyl sulfoxide was added to a solution containing 0.2-mM NeutrAvidin (Life Technologies) in PBS. The reaction was continued for 1 h at RT. Unreacted excess of DBCO-NHS was removed using a desalting column. 0.8 mg of azide-functionalized nanoruby solution was mixed with 16 µL of 0.12-mM alkynated NeutrAvidin (or non-alkynated for control) and incubated under stirring for 3 h at RT. Unbound and loosely bound NeutrAvidin was removed by rinsing 5 times in PBS. As-produced conjugates were stored in PBS for cell labeling experiments within 1 week. Longer storage resulted in aggregation followed by precipitation.

4.9. Opioid receptor labeling in live and fixed cells using biotin-functionalised nanorubies


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AtT-20 cells stably expressing hemagglutinin-tagged wild-type human µ-opioid receptor (HAhMOR)56 or FLAG-tagged wild-type mouse µ-opioid receptor (FLAG-mMOR, negative control)57 were plated in an 8-chambered slide (Thermofisher Scientific). Cells were grown overnight in an incubator maintained at 37 °C and 5% CO2 in DMEM supplemented with 10% FBS, Penicillin/Streptomycin and 100 µg.mL-1 Hygromycin (selection antibiotic for HA-hMOR) or 300 µg.mL-1 G418 (FLAG-mMOR). For labeling in fixed cells, cells were fixed with 3.7% paraformaldehyde in PBS for 20 min and blocked with Leibovitz’s L-15 medium (L15, Thermofisher Scientific) containing 1% bovine serum albumin (hereafter referred to as serumfree media) for 1 h at RT. For labeling in live cells, cells were firstly equilibrated to RT for 15 min and incubated in serum-free media for 20 min at RT. Biotinylated anti-HA antibody (Covance # BIOT-101L), diluted 500 times in serum-free media, was added to the cells and incubated at RT for 1 h. This antibody was tested in house for specificity (data not shown). The cells were rinsed three times with serum-free media. ExtrAvidin-FITC (Sigma) was diluted 1000 times in serum-free media and added to the cells, followed by incubation for 1 h at RT. The cells were rinsed three times with serum-free media. Biotinylated nanorubies (with or without silica coating) prepared at concentrations ranging from 10-50 µg.mL-1 were prepared in serum-free media, added to the cells, and incubated at RT under gentle rocking. Cells were washed once with serum-free media and twice with PBS. For fixed cell labeling, samples were mounted for imaging. For live cell labeling, cells were fixed for 20 min using 3.7% paraformaldehyde solution prepared in PBS. Cells were rinsed twice with PBS and mounted for imaging.

4.10. Opioid receptor labeling in live and fixed cells using NeutrAvidin-conjugated nanorubies


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The labeling step until the incubation with biotinylated anti-HA antibody was same as described above. After the antibody incubation step, cells were washed three times using serumfree media. Nanoruby-NeutrAvidin conjugates were added to cells within a concentration range of 30-150 µg.mL-1, and incubated for 10 mins at RT. Cells were washed once with serum-free media and twice with PBS and then prepared for imaging, as described above.


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Acknowledgement Authors acknowledge the help of Mel McDowall with the biocompatibility studies in embryo; Ben Johnston and ANFF – for laser engraving grids on coverslips; Ekaterina Grebenik for discussions on ion adsorption on nanoruby; and the MQ Microscopy facility for electron microscopy and elemental analysis.

Abbreviations AAA, acid-alkali-acid (processing); AFM, atomic force microscopy; DLS, dynamic light scattering; EDS, energy dispersive x-ray spectroscopy; HEBM, high-energy ball milling; NA, NeutrAvidin; NIR, near infrared; NR2/NR8, nanoruby with 0.15% or 0.8% Cr3+; PCA, process control agent; PEG, polyethylene glycol; PVP, polyvinylpyrrolidone; QDot, quantum dot; TEM, transmission electron microscopy or microgram; XRD, x-ray diffraction;


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Supporting Information. A PDF file containing additional figures, experimental methods and results as cited in the manuscript is available online, free of charge. Author Contributions V.K.A.S, M.S., J.T., M.C., E.M.G., and A.V.Z. designed the study. V.K.A.S, W.A.W.R, A.S., D.D., E.M.G., and A.V.Z. wrote the paper. V.K.A.S, A.V., W.A.W.R, K.Z. and R.R.P. carried out experiments and analyzed the data. A.S., and H.B. carried out embryo toxicity experiments and analyzed the data. D.D. carried out AFM measurements and analyzed the data. All authors have given approval to the final version of the manuscript. Conflicts of Interest V.K.A.S, W.A.W.R, E.M.G., and A.V.Z are named on Australian patent application 2017901011 related to this work. Funding Sources V.S. received the Macquarie University Research Fellowship 9201200881 and the Australian Research Council Discovery Early Career Research Fellowship DE160100888. This work was partially supported by the Australian Research Council CE14010003 award to E.M.G and Russian Science Foundation (7-19-01416) awarded to A.V.Z. AFM measurements were supported by the Australian Research Council funding LE150100177.


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