Fluorescent Nanodiamond Silk Fibroin Spheres: Advanced Nanoscale

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Fluorescent Nanodiamond Silk Fibroin Spheres: Advanced Nanoscale Bioimaging Tool asma khalid, Alexander Mitropoulos, Benedetto Marelli, David Allan Simpson, Phong Tran, Fiorenzo G Omenetto, and Snjezana Tomljenovic-Hanic ACS Biomater. Sci. Eng., Just Accepted Manuscript • DOI: 10.1021/acsbiomaterials.5b00220 • Publication Date (Web): 11 Sep 2015 Downloaded from http://pubs.acs.org on September 21, 2015

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ACS Biomaterials Science & Engineering

Fluorescent Nanodiamond Silk Fibroin Spheres: Advanced Nanoscale Bioimaging Tool Asma Khalid1, Alexander N. Mitropoulos2, BenedettoMarelli2, David A. Simpson1, Phong A. Tran3,4, Fiorenzo G. Omenetto2* and Snjezana Tomljenovic-Hanic1* 1. School of Physics, University of Melbourne, Parkville, VIC 3010, Australia. 2. Department of Biomedical Engineering, Tufts University, Medford MA 02155. 3. School of Biomolecular Engineering, University of Melbourne, Parkville, VIC 3010, Australia. 4. Institute of Health and Biomedical Innovation, Queensland University of Technology, Kelvin Grove, QLD, Australia. *Email: [email protected], [email protected] KEYWORDS: nanodiamonds, silk fibroin, fluorescence, bioimaging, mobility

ABSTRACT. High resolution bioimaging is not only critical to the study of cellular structures and processes but it also has important applications in drug delivery and therapeutics. Fluorescent nanodiamonds (NDs) are excellent candidates for long term bioimaging and tracking of biological structures at the nanoscale. Encapsulating NDs in natural biopolymers like silk fibroin (SF) widens their biomedical applications. Here we report the synthesis, structural and optical characterization of ND incorporated SF nanospheres. The photo-luminescence from optical defects within the NDs is found to increase when encapsulated in the SF spheres. The encapsulated NDs are applied in-vitro to investigate the intra-cellular mobility compared to bare NDs. The diffusion rate of encapsulated NDs is shown to improve due to SF coating. These ND-SF spheres are envisioned as highly suitable candidates for bio-injectable imaging and drug release carriers for targeted drug delivery applications. 1 ACS Paragon Plus Environment


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1. INTRODUCTION Nanodiamonds (NDs) are emerging as promising material for a variety of biomedical applications. Their biocompatibility, large surface area to volume ratios and superior optical properties make them attractive for drug delivery carriers1 and intra-cellular imaging tools.2-3 NDs are biologically compatible,4 noncytotoxic and are nonreactive inside the cells.5 The negatively-charged nitrogen vacancy (NV-) centre in NDs is a photostable emitter

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with a

bright broad fluorescence band extending from the visible to near infrared.7 The NVfluorescence is well separated from the cells autofluorescence and is highly resistant to photo-bleaching or photo-blinking. Unlike ultraviolet,8 these wavelengths of excitation and emission (in visible to near infrared wavelength range) for NV- do not interact with cells. These unique characteristics of NV- centres distinguish them from commercially used fluorescent proteins,9 dyes10 and quantum dots.11 ND’s biocompatibility combined with excellent optical properties of NV- emitter make NDs highly desirable for long term imaging and monitoring inside of cells.1-2, 12-13 However, the use of NDs without any surface modification in many bio-applications is limited. The cleaved edges and rough surfaces of NDs2 result in trapping of these nanoparticles in endosomes after cellular uptake.12 Moreover, the direct surface functionalization of NDs for specific biorecognition is not straight forward, which is an obstacle for the use of NDs in biological applications. These limitations are addressed by using biocompatible surface modification of NDs. Current research in the field of NDs has focused on the effects of lipid12, 14 and polymer15-16coatings to improve the dynamics such as mobility, diffusion and bio-suitability of NDs inside biological cells. Coating NDs with polymeric shells16 has resulted in improved colloidal stability, non-toxic delivery of DNA, proteins and drugs into the cells.12 The coating and encapsulation of NDs therefore allows

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probing of both short-term and long term dynamics of single NDs in living cells without compromising the mobility and biocompatibility of these nanoparticles in cells for extended periods of time.12, 16-17 In this manuscript, the novel combination of NDs and silk fibroin (SF) is reported, by encapsulating NDs within SF nanospheres. SF is a biocompatible natural protein which is degradable and exhibits low inflammatory response in-vivo.18-19 The surface of SF is easily functionalized with various other proteins19 making it highly suitable for applications involving protein and gene delivery,20 advanced targeted therapeutics and drug delivery.21-22 Coating of NDs with SF also ensures biologically safe and non-toxic surface modification of NDs for biomedical applications. Moreover, SF also holds special significance due to its biodegradability and bioresorbability.23 SF is a mechanically robust biocompatible material that degrades at a controlled rate in-vivo and in-vitro. The final breakdown products of SF degradation are the corresponding amino acids, which are easily absorbed in-vivo. This is one of the reasons for the extensive use of SF in the field of biomedicine.18-23 The degradability of SF can be engineered from weeks to months by modifying the crystallinity and beta sheet content,22 simply by processing the fibroin or using post-processing methods. The all aqueous SF solution at room temperature allows for the incorporation of labile biological components without loss of function and can preserve their bioactivity over extended periods.24 Furthermore, the side groups of the SF peptide can be easily functionalized for target specific delivery making them more favourable for drug delivery applications.21 SF is not only biocompatible and biodegradable but also possesses excellent optical properties. It is more than 90 % transparent to the photoluminescence produced by NDs7 and possesses a low background signal7 as compared to other biomaterials. In the current work, we have used SF as an encapsulating polymer due to its versatile nature. The robust



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mechanical properties, the controllable degradation, and unique optical properties make SF superior to other degradable polymers (collagen or polylactic acid) and provide a solid basis for utilizing SF as a biomedical material for many applications.25 Coating the fluorescent NDs with a biopolymer like SF has the ability to improve their biological properties like cellular uptake,12 biocompatibility, and optical properties.7 This has already been shown in our previous work7 where the ND-SF films showed a 2-4 time enhancement in the NV-emission properties, and the implantable ND-SF films were found to be non-inflammatory and non-toxic in-vivo. SF coated NDs are reported to exhibit superior emission properties due to the higher refractive index of SF as compared to air.7 The coating fully preserves the unique optical properties of the NV- centres that are crucial for bioimaging applications. Submicron SF spheres are increasingly studied and investigated recently as efficient carriers for local and target specific drug release.21-22 However the incorporation of NDs into these spheres for imaging administered biomedicine applications has not been examined to date. SF structures, which are mostly used in conjunction with rapidly photobleaching fluorescent dyes,21 would benefit from NDs in terms of improved bioimaging ability at the nanoscale. The ND-SF combination will enable optical structures that can adapt to the surrounding environment in a non-invasive and non-inflammatory manner and at the same time can convert chemical and structural changes into optical signals. In this manuscript, the fabrication, surface and optical characterization, and dynamics of ND-SF spheres incorporated into living cells are investigated. The optical properties of NDSF spheres and emission comparison with NDs and SF alone were investigated with near field confocal imaging. The fluorescence properties and mobility of ND-SF spheres in fibroblast cells were recorded in-vitro using wide field imaging. 4 ACS Paragon Plus Environment

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2. MATERIALS AND METHODS 2.1. SF processing. The protocol for purifying the SF protein is well established and documented in the literature.25 SF was obtained from Bombyx mori cocoons and protocols outlined by Kaplan et al were followed. Briefly, the purification of SF initially involved the removal of sericin, by boiling the cocoons in 0.02 M aqueous solution of sodium carbonate (Sigma Aldrich, USA) to remove the undesired sericin molecule. The fibroin bundle was washed in deionized water, dried overnight, and then solublized in 9.3 M aqueous lithium bromide (Sigma Aldrich, USA) at 60°C for three hours. The solution was dialyzed against deionized water (dialysis cassettes Slide-a-Lyzer, Pierce, MWCO 3.5K) and enabled the production of 6 % w/v SF solution. The solution is then centrifuged to remove any large aggregates. 2.2. Synthesis of spheres. Spheres were fabricated with the co-flow technique.26 As shown in the schematic of Figure 1, the continuous phase consisting of Polyvinyl Alcohol (PVA) was flowed over a discrete phase containing ND and SF solutions to produce ND-SF spheres. A 16-gauge needle (inner diameter=1.2 mm) was used for the outer continuous phase channel while for the inner discrete phase channel, a 30-gauge needle (inner diameter=152 µm) was used. The needle diameter, flow rates selected for the two channels and concentration of SF solution are the important parameters that directly control the sphere diameter.26 For sphere fabrication, NDs with an average size of 45 nm and irradiated with NV- centres were used. A 2 mg/mL NDs solution and a 10 mg/mL (1 %) SF solution was used. A concentration of 10 mg/mL for SF was achieved by diluting 362 µL of the as prepared SF solution (58 mg/mL) in 500 µL of NDs as prepared solution (2 mg/mL) and 1138 µL of de-ionized water. The resulting 2 mL of ND-SF mixture was injected through the smaller diameter inner needle connected to the co-flow device. The 5 % PVA solution was



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injected through the larger diameter outer needle connected to the co-flow device. The syringe pumps for the inner and outer needles were set to flow the solutions at rates of 40 µL/h and 4 mL/h respectively. In this study, the smallest inner gauge needle, slowest discrete phase flow rate, and lowest SF concentration22,

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synthesize the spheres. Spheres collected and suspended in PVA were cast on a clean flat PDMS surface. Once it dried, the PVA was washed by centrifuging the spheres in water twice with 11000 RCF for 10 minutes at 4˚C. The washed spheres were then redispersed in 1 mL of ultrapure water or phosphate buffer saline (PBS) for optical and surface characterization. 2.3. Scanning electron microscopy (SEM). Each sample was sputter coated with palladium/gold before imaging.The spheres were sputter coated with 5 nm of gold using an Electron Microscopy Sciences 300T dual head sputter coater. Sputtering was conducted at 20 mA with an average deposition rate of 4 nm/min. Images of each batch of microspheres were examined under a Zeiss EVO MA 10 (Carl Zeiss SMT, UK) Scanning Electron Microscope (SEM) at 3keV. For higher magnification images, a Supra 55VP FESEM (Zeiss) was used using the SE2 detector at 4kV. 2.4. Confocal imaging. Confocal imaging of the NDs, ND-SF and SF only spheres was undertaken on a custom build confocal microscope.7 The samples on Si substrate were mounted and illuminated with a 532 nm excitation from a frequency doubled Nd:YAG continuous wave laser through a 100×, 0.95NA objective. The fluorescence from the samples was filtered using a long pass (560 nm, Semrock) and band pass (650-750 nm, Semrock) filter and coupled to a fibre based avalanche photo detector (APD, Perkin Elmer). A Physik Instrument peizo scanning stage was used to scan the sample over a 100x100 µm2 area. Fluorescence spectra of particles of interest were collected using a 300 lines/mm imaging


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spectrometer (Acton Research) over the wavelength range from 550-850 nm, in which case the fluorescence band-pass filter and 560 nm long pass filter were removed for the spectral acquisitions. 2.5. NPs internalization in fibroblast cells. The as prepared ND-SF spheres were incubated in PBS at 37˚C and uniformly rotated for 7 days. The reduction in the sphere diameter due to SF degradation was checked with SEM at day 1, 3 and 7 during incubation. The degraded SF was removed every 24 hours and the spheres were resuspended and incubated in fresh PBS. The spheres (ND-SF and SF only) with appropriate size distribution at day 3 after SF degradation were selected for cell culture study. Mouse embryo fibroblast (ATCC) were used for the in-vitro imaging experiments. The cells were placed in the glass bottom cell chambers and incubated in a cell culture incubator at 37˚C. When the cells reach approximately 70-80 % confluency, spheres or NDs were introduced and incubated for 3-4 hours to allow for cellular uptake of the particles. After this time period, PBS was used to wash the cells on the glass coverslips in the wells to remove free particles. Fresh culture medium was added to each well for subsequent wide field imaging of adherent cells on the glass coverslips. 2.6. Wide field imaging. The wide-field imaging was performed on a commercial fluorescence microscope (Nikon, Eclipse Ti-U) adapted for wide-field imaging. Optical excitation was provided by a 532 nm Verdi laser operating with a typical power density of 30 W/mm2. A x5 beam expander was used to expand the excitation beam to 10 mm before focusing the excitation light onto the back aperture of the 100×, 1.45 NA (Nikon) oil immersion objective via a dichroic mirror (Semrock-Di02-R561-25x36).27 The resulting fluorescence was filtered using two band-pass filters 650-750 nm (Semrock) before being imaged onto a sCMOS camera (Andor, Neo) with a f=300 mm tube lens. The operating field



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of view was 60×60 µm2. The temperature of the wide-field microscope was maintained at 37 ˚C for imaging using an environmental chamber (Clear State Solutions). Once the NPs were internalized in to the fibroblast cells, the culture medium was replaced with PBS to avoid autofluorescence from the culture medium. 2.7. Mobility analysis. The fluorescence images recorded with wide field imaging, taken at a rate of 7 frames/s for 60 s were converted to a movie in MATLAB. The successive frames in MATLAB were cropped and backgrounded subtracted with noise filters. The particle locator and centroid finding functions were employed to trace the trajectories of each particle to sub-pixel accuracy. Finally the locations were translated to displacements and scaled to microns. 3. RESULTS AND DISCUSSION The manuscript reports the synthesis and characterization of ND-SF spheres. This work investigates and compares the optical, structural and dynamical properties of ND-SF spheres with NDs and/or SF only spheres. Section 3.1 briefly describes the method and schematic of sphere fabrication. In section 3.2, the structure of the ND-SF spheres is examined and their size range is estimated with scanning electron microscopy (SEM). Section 3.3 investigates the optical emission properties of spheres through near field fluorescence microscopy. Wide field fluorescence measurements performed in-vitro are discussed in section 3.4 and the section 3.5 presents a comparison and measurement of the mobility of spheres and NDs invitro. 3.1. Synthesis of spheres. SF solution was extracted, as reported in previous literature25 and ND-SF or SF only spheres were fabricated using all aqueous solutions at room temperature without involving any harsh chemicals or high temperature and pressure conditions. The synthesis of spheres was done through already developed co-flow technique. 8 ACS Paragon Plus Environment

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The co-flow device was used because it is capable of producing consistent and reproducible diameter spheres (with SF only) compared to other methods such as bulk mixing of solutions that induces particle formation. The quantities required for specific applications can be easily attained using one device which is easily controllable using hydrodynamic properties of fluids used. The particle size is repeatable as the method can control the sphere diameter based on the flow rates, fluid concentrations, fluid viscosities, and the dimensions of the device. Using the current method without NDs, consistent sized SF only spheres are attainable since molecules are transported in a relatively predictable manner and the set parameters do not change once the flows have started.26 The concentrations of SF and NDs for ND-SF or SF spheres and NDs alone were kept constant for all three types of samples. Figure 1 shows the schematic of the co-flow technique, where a continuous phase (PVA 5 %) is flowed over a discrete phase (ND and SF solution). For the discrete phase, the inner needle of 30 G was fitted and a flow rate of 40 µL/h was selected, controlled by a syringe pump, to produce smallest sphere size achievable with the equipment.



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Figure 1. Schematic diagram of the co-flow method for the synthesis of ND-SFspheres. 3.2. Surface analysis and degradation of SF. A 5 µL volume of ND-SF spheres or ND particles suspended in water was drop-cast onto clean Si substrates and were allowed to dry on a SEM sample stub. The samples were then sputter coated with gold under a vacuum and the morphologies of nanoparticles were imaged using the SEM at 3kV. Figure 2 (a) and (b) show the SEM images of the ND-SF spheres at two different magnifications. Under SEM, the sample was found to contain nano-sized spheres. These NDs incorporated SF spheres existed within a range of 400 to 600 nm in diameter, which was estimated with MATLAB. The range for the diameters was measured to vary from 190±5 nm for the smallest sphere to 850±17 nm, with the maximum number of particles in the 400-600 nm range. The mean for the wide distribution was found (using MATLAB) to be 540 nm.


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Figure 2. SEM image of ND-SF spheres at magnifications of (a) 25k and (b) 250k. (c) After 7 days of incubation, spheres degraded leaving NDs alone imaged at 100k magnification. (d) NDs alone on the substrate scanned at 100k magnification. The ND-SF spheres were then incubated in PBS at 37˚C and constantly rotated and monitored with SEM. At day 7, a 5 µL volume of solution was drop-cast onto a Si substrate. SEM image of the post-PBS ND-SF spheres, at day 7 during incubation, is shown in Figure 2 (c), where the spheres have clearly degraded leaving behind the NDs. The diameter of these residual nanoparticles was calculated using MATLAB and existed in a range of 17±2 nm to 60±8 nm for 18 nanoparticles. The average of the distribution was found to be 30 nm. A size comparison was made between bare NDs and sphere degraded NDs using SEM. A 5 µL volume of fresh NDs solution in water was drop-cast on Si substrate and analysed with SEM at a 100k× magnification as shown in Figure 2 (d). NDs were found to be agglomerated together onto the Si surface. For NDs alone, the size distribution for 20 particles existed from 22±4 nm for the smallest ND to 70±3 nm for the largest less agglomerated bare NDs. The average size was calculated to be 47 nm. Hence the size range of SF degraded NDs coincides with that of bare NDs. The degradation of the SF in PBS is attributed to bulk swelling of the SF causing silk molecules to detach from the surface reducing the diameter of the spheres overtime. The degradation is accelerated by the phosphate ions in PBS because the positive and negative charges more effectively interfere with the beta-sheet structure of the silk molecules causing them to swell at different rates compared to water. As mentioned in Section 2.5, the silk molecules that detached from the spheres existed in the solution and were removed upon rinsing every 24 h. 3.3. Confocal imaging. The optical fluorescence measurements were performed with a custom confocal microscope. For the as prepared ND-SF spheres with a size range of 400-



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600 nm (measured in Section 3.2), a comparison of the fluorescence was made with bare NDs. The concentration of NDs for both spheres and NDs only samples was maintained constant. Finally SF only spheres were synthesized to measure the fluorescence contribution of SF structure to the ND-SF spheres. SF spheres were fabricated following the same concentration of SF and flow rates of both channels as were adapted for fabricating ND-SF spheres. Suspensions of NDs and spheres (ND-SF and SF alone) in water were drop cast on three individual Si substrates and the samples were air dried overnight. Fluorescence scans of 100×100 µm2 were taken, emission count rates recorded and spectra acquired for the three samples. A CW 532 nm green laser was used to illuminate the three samples at a constant low excitation power of 10 µW. Figure 3 shows 10×10 µm2 fluorescence zoomed scans for NDs, ND-SF spheres and SF only spheres in Figure 3 (a), 3 (b) and 3 (c) respectively.

Figure 3. Zoomed fluorescence scans, 10×10 µm2 for (a) NDs, (b) ND-SF spheres and (c) SF only spheres on the Si substrate. The particles similar to the ones labelled with the cross hair in the figures were checked for stability and emission count rates. To confirm that each analysed particle is a single ND, Gaussian fitting was done (on a 10×10 µm2 fine-scan) along x- and y-axis to locate the best position of the particle. The NDs whose fluorescence can be distinguished down to the diffraction limit of the emitted light (340 nm) are considered as single particles and were the ones selected for emission characterization of NDs alone sample. For the ND-SF spheres, the smallest spheres were selected and the same technique was applied to determine the position of the sphere on a fine12 ACS Paragon Plus Environment

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scan area of the sample. After a few seconds of fluorescence bleaching (from SF), the location of the ND-SF sphere was Gaussian fitted to the diffraction limit, indicating the presence of a single fluorescing ND inside the sphere.



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Figure 4. (a) Emission rates for four NDs and (b) ND-SF spheres. The NDs with or without SF are found to fluoresce with high photostability. (c) Low and decaying emission intensity of two SF only spheres. The dashed line indicates the background from Si substrate. Emission counts were recorded for 10-12 fluorescent NDs with and without SF encapsulation. The count traces for four of these smallest particles are shown in Figure 4. Figure 4 (a) shows the intensity record for the weakest to the brightest intensity ND from the 100×100 µm2 scan on the sample. The brightness lies in a range of 0.06-2 Mcounts/s, which corresponds to different sizes of NDs, the orientation and location of the NV- centre inside the ND, as well as the presence of single or multiple NV- centers provide a range of intensities for different NDs. The stable traces in the figure show that the NDs are fluorescing with high photostability, without any blinking or reduction in counts over the measured time. Figure 4 (b) shows the evolution of emission counts for ND-SF spheres. As mentioned above, the smallest spheres on the scan of Figure 3 (b) were selected to record the intensity traces because the spheres are clustered close to each other in a similar way as they appear in the SEM images of Figure 2. However there was a possibility of more than one ND being present inside the spheres. Figure 4 (b) shows four of the selected emitters where the count rate ranges from 0.25 Mcounts/s for the least bright to 4 Mcounts/s for the brightest ND-SF sphere. The brighter emission is consistent with our previous studies on the optical emission with and without silk coating.7 To check the background fluorescence of SF and its contribution to the fluorescence of ND-SF spheres, the SF only spheres were fabricated and characterized optically with the confocal system. Figure 3 (c) shows one of the SF spheres in a 10×10 µm2 scan window with a very low contrast as compared to the background. The count traces for 8 SF spheres on the confocal scan were analysed and two of these SF only spheres are shown in Figure 4 (c). The


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intensity was not only faint but exponentially decayed to the background value of 2 kcounts/s from the Si substrate. The brighter emission from ND-SF spheres on Si substrate was also calculated numerically. A 3-dimensional Finite-Difference Time-Domain (FDTD) calculation using RSoft was performed to measure the ratio of emission from ND-SF spheres on the Si substrate with respect to emission of NDs alone deposited on the substrate. The calculations were done for two extreme polarizations (orthogonal and parallel) of NV- dipole with respect to the surface of the substrate. The simulations were run for three different sphere diameters in 100-600 nm diameter spheres, as shown in Table 1. The calculations for varying sphere diameters provided emission enhancement in a range of 0.64-2.86 for the orthogonal polarization and 1.51-17.89 for the parallel polarization of NV- dipole with respect to the substrate. The experimentally observed enhancement ratio existed in a range of 2-4 times for the smallest ND-SF spheres in the confocal scans. Although the location of the ND inside the sphere and the polarization of the NV- dipole were unknown for the experimentally obtained data, the enhancement range exists inside the numerically obtained ratios. Sphere size (nm) ND SF Si

ND closest to the substrate

ND in the centre of the sphere

ND at the edge of the sphere

Orthogonal Parallel Orthogonal Parallel Orthogonal Parallel 100 1.57 1.51 1.20 2.13 0.88 3.13 300 2.86 3.75 0.97 14.19 0.64 6.18 600 2.05 1.76 1.58 17.89 0.75 10.49 Table 1: Emission enhancement ratios for ND-SF spheres with respect to bare NDs obtained

numerically for three different sphere sizes, ND’s location inside the sphere and two polarizations of NV- dipole. The orthogonal and parallel polarizations are defined with respect to the Si substrate.



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The numerical results illustrate that the enhancement in emission is proportional to the sphere size. Table 1 reveals that the range of enhancement obtained with the orthogonal polarization matches to a greater extent with the experimentally obtained results. The discrepancy in the enhancement is more pronounced for the parallel polarization which is less likely to be collected, as it is propagating parallel to the substrate’s surface. This contradiction comes as we collect the radiation emanating from the sample with a 0.95 NA air objective, making an angular aperture of ∼72˚ (half angle for the light cone collected by the objective) with respect to the normal on the substrate’s surface. Therefore, with the current setup we can only collect efficiently within the cone which includes polarized light from the orthogonal polarization. The light emitted from the parallel polarization radiating along the substrate doesn’t all get collected due to the angular aperture of the 0.95 NA objective used for experimental studies. It is important to mention here that the comparisons between the measured and calculated rates are made on the basis of unity quantum efficiency from the NV centres. However, there is an on-going debate in the literature28 as to the actual quantum efficiency of these defects. Therefore, these measurements serve as a qualitative rather than a quantitative comparison. To confirm the presence of NDs inside the SF spheres, photoluminescence spectra were acquired for the samples. Spectra were taken for the particles shown in Figure 3, where the emission was collected in 550-850 nm detection window. Several bright spots were selected separately and fluorescence spectra were collected using a spectrometer. Figure 5 shows the spectra at room temperature from each of the NDs, ND-SF spheres and SF only spheres. In Figure 5, the peak at 637 nm is the NV- zero phonon line and the phonon sideband ranging from 650-750 nm. Spectra were background corrected by subtracting the background noise


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from the Si substrate. The spectrum from SF sphere was found to be very low in intensity in the near infrared region with peaks at 608, 641 and 668 nm.

Figure 5. Background subtracted spectra for a representative ND (black trace), ND-SF sphere (red trace) and SF only sphere (blue trace). Collection time was 30s. The true counts for NDs alone and SF only spheres are shown at the left y-scale and those for ND-SF spheres are shown to the right. For the left y-axis, the emission counts for ND-SF sphere are multiplied by 0.2 to show the three spectra distinctly in the same plot. Figure 5 contains spectra that are representative of the three samples, each for NDs alone, ND-SF spheres and SF only spheres on the Si substrate. The spectra were collected for 10, 10 and 8 individual NDs, ND-SF spheres and SF only spheres respectively. While the emission counts for different spectra (along the y-axis) may vary for the distribution of particles, the general trend (as also discussed in Section 3.3) was that the ND-SF spheres showed brightest counts and SF only spheres showed the lowest counts.



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3.4. Wide field imaging. To check the emission properties and hence the imaging efficiency of ND-SF spheres inside biological structures, the in-vitro imaging of spheres was performed. As mentioned in Section 3.2, the diameters for the as prepared spheres were larger and the majority existed in a range of 400-600 nm, possibly caused by the varying diameters of bare NDs (22-70 nm), as mentioned in Section 3.2. The addition of NDs could also have changed the viscosity of the fluid in unexpected ways, ultimately resulting in a broad size range of the as prepared ND-SF spheres. However, incubating the spheres with constant shaking in PBS resulted in their size reduction according to the degradation mechanism (mentioned in Section 3.2). A consistent decrease in the sizes of the spheres was noticed at day 1, 3 and 7 during incubation of spheres at 37 ˚C. At day 3, the sphere distribution reduced to size range of 85±8 to 400±9 nm, which is suitable to be uptaken by the skin cells. The distribution averages around 200 nm. The bare NDs and size reduced ND-SF or SF spheres were introduced to three different cell chambers and incubated for 6-8 hours to allow for cellular uptake. Fibroblast cells (3T3 fibroblast, ATCC) were used for this experiment. 3T3 cells were chosen for this study because they are routinely used to test cellular response to materials29 and cellular uptake, as compared to the most commonly used HeLa cells30-31 which are routinely used to evaluate cellular responses of biomaterials. Wide-field imaging was used to take sequential images of the entire field of view for the spheres and NDs cultured cells. The technique provided the ability to track the stability and brightness of multiple spheres inside cells. Wide field imaging27 was performed with an inverted fluorescence microscope. The in-vitro imaging was done for NDs alone, ND-SF spheres and SF only spheres at 37 ˚C inside an incubated chamber.


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A white light bright field microscopic image of the cells was captured and used to determine the imaging plane. Figure 6 (a), (b) and (c) show white light microscopic images of fibroblast cells containing NDs, ND-SF and SF only spheres respectively. The corresponding wide field fluorescence scans for the three samples are shown to the right of each white light image in Figure 6 (d), (e) and (f).



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Figure 6. (a)-(c) Bright field microscopic images of the fibroblast cells cultured with NDs, ND-SF and SF only spheres respectively. (b), (d), (f )Wide-field fluorescence images for 60×60 µm2 areas on the samples illuminated uniformly with the laser beam. Fluorescence imaging of the spheres was performed several microns above the surface of the coverglass where the fluorescent spheres came into focus. Care was taken to ensure the focal point did not exceed the average height of the cultured cells. High contrast fluorescence was observed for NDs and ND-SF spheres, as apparent in Figure 6 (d) and (e), which was distinguished from the cells’ autofluorescence and background noise. However with SF only spheres, a weak fluorescence of the particles and significantly low contrast with respect to the background was noticed as can be seen in Figure 6 (f). The NDs and spheres appeared to be uptaken by the cells and showed no indication of cytotoxicity. Bright fluorescence allowed efficient tracking of the emission intensities over time. The fluorescence of NDs was compared with that of ND-SF spheres in cells. The evolution of emission counts with time was recorded for a number of cells cultured with NDs, ND-SF or SF only spheres. Figure 7 (a) and (b) show the integrated emission counts collected for four of these cells internalized with NDs and ND-SF spheres respectively. The photostability of the emission is clearly obvious from the time traces with the counts ranging from 0.12 to 0.30 Mcounts/s for NDs and 0.27-1.20 Mcounts/s for ND-SF spheres. A comparison in counts between Figures 7 (a) and (b) for NDs alone and ND-SF spheres respectively indicates on average a 3 fold brighter emission from ND-SF spheres. Each sphere may contain multiple NDs but the NDs being smaller in size are uptaken in greater number by the cells, as shown in Figure 6 (a).


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Figure 7. The fluorescence intensities versus time for (a) NDs and (b) ND-SF spheres inside four individual cells.The average counts are labelled for each cell. (c) Wide field fluorescence



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intensity plotted as a function of time for the SF only spheres inside two different cells. The emission rate varies between 13-25 kcounts/s for different cells containing SF spheres. As reported by Khalid et al in Ref7 the enhancement in the emission of ND-SF comes from the encapsulation of the NDs by SF’s high refractive index, n=1.54 at the wavelength of NV- emission, λ=637 nm. The fluorescence intensities of SF only spheres were recorded to investigate SF contribution to fluorescence. The traces of Figure 7 (c) reveal a ∼100 times less fluorescence of the internalized SF only spheres and rapidly decreasing counts with time. The magnitude of intensity indicates that the fluorescence of SF structures contribute only 8% to the fluorescence of the spheres. The major contribution 92% comes from the modified emission properties of encapsulated NDs. The brighter emission from ND-SF spheres suspended in PBS was also calculated numerically using FDTD method. The emission ratio was calculated for the ND-SF sphere’s emission rate with respect to the emission rate of NDs alone suspended in water. The NVdipole’s polarizations were not considered as the nanoparticles are freely suspended in the solution during in-vitro imaging. The calculations showed that for sphere diameters of 100400 nm, the emission enhances in a range of 1.19-1.58 when the ND is positioned at the centre of the sphere and 1.15-1.45 when the ND exists close to the edges of the sphere. The experimentally observed enhancement exists in a range of 2.2-4 times for different SF coated ND spheres in-vitro with diameters from 85 to 400 nm. The greater experimental enhancement possibly comes from the collective fluorescence response of multiple spheres and NDs in the individual cells. A background check for the autofluorescence of the fibroblast cells was done as shown in Figure 8. Figure 8 (a) shows white light image of cells alone without the addition of any


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foreign fluorescent agent, while (b) and (c) show the corresponding wide field fluorescence images at different time intervals. A bright autofluorescence was seen in the first few frames which bleached rapidly after a few seconds. The frame after delays of 2 s and 20 s are shown in the Figure 8 (b) and (c). Hence, the wide-field microscopy data shows that NDs fluoresce with high photostability on their own and with enhanced brightness when encapsulated inside SF spheres. This fluorescence is long lived without any signs of bleaching or blinking as compared to the fluorescence of SF only spheres or autofluorescence of the cells.

Figure 8. (a) Bright field microscopic images from cells alone. The corresponding 60×60 µm2 wide field fluorescence images after (b) 2 s and (c) 20 s are shown.

3.5. Mobility analysis. The bare NDs and the ND-SF spheres in the fibroblast cell were monitored by tracking their movement in cells and recording their two-dimensional trajectories under a wide-field fluorescence microscope. The diffusion coefficients for 21 NDs were compared with that of 22 ND-SF spheres in cells and the calculations have demonstrated that the encapsulation of NDs within SF spheres makes the particles increasingly mobile in the cytoplasm of the cells. Higher diffusion coefficients of ND-SF



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spheres indicate that the silk coated NDs are potentially more suitable for long term imaging and tracking in biological environment.32 Mobility calculations were performed to compare the dynamics of ND-SF and NDs alone inside the cells. With wide-field fluorescence microscope, sequential images were recorded to track the movement of particles inside the cells. A tracking program in MATLAB was coded to detect, trace and find locations of individual nanoparticles (NDs or ND-SF spheres) in successive frames. The movement of single nanoparticles was tracked in the plane (along xand y- axes) over a time span of 60s. Mean squared displacement analysis was performed for the two dimensional trajectories and the diffusion coefficients were determined for the internalized NDs and ND-SF spheres. Figure 9 (a) and (b) show the fluorescence images of NDs inside the cells. Brownian motion particle trajectories were traced for each individual ND using MATLAB and displacement square data was plotted against time. Figure 9 (c) and (d) show the 2dimensional Brownian motion trajectories for the individual NDs labelled in Figure 9 (a) and (b) respectively. The displacement squared (r2) data plotted versus time for the circled NDs are shown in Figure 9 (e) and (f). The trajectory for each particle was traced with respect to its origin at (xn,yn) = (0,0) and t = 0. The r2 versus time plots were fit to a linear expression described in Equation 1 to measure the diffusion coefficient. The lines of best fit are also shown in the corresponding figures. Brownian motion trajectories were drawn for each individual ND using MATLAB and displacement square data plotted against time. The diffusion coefficient33 was measured using Equation 1 for 2-dimensional Brownian motion,

= 4 ∆ = 4∆ ,

(1)


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where ∆t is the time interval and D is the diffusion coefficient. The procedure of plotting Brownian motion and displacement square versus time was repeated for 21 NDs to find the corresponding diffusion coefficients. The D values existed in a range of (1.0±0.2)×10-4 to 0.50±0.05 µm2/s for NDs in-vitro. The mean value of the diffusion coefficient from the broad range calculated for 21 NDs is 0.054 µm2/s.

Figure 9. (a), (b) Wide field fluorescence image of fibroblast cells internalized with NDs. (c), (d) 2D Brownian motion trajectory for the circled NDs in (a) and (b) were traced. (e), (f) 25 ACS Paragon Plus Environment


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Displacement square versus time plot for the selected NDs. The red data in figures show the lines of best fit and the equations of the fitted line are shown in the inset. The broad distribution of the diffusion coefficents for NDs (blue), shown with the bar graph in Figure 10, might have resulted due to the differences in ND size as well as in the cellular environment. Generally lower diffusion coefficients < 0.1 µm2/s (with an exception of few) were recorded for NDs as illustrated by the bar graph of Figure 10 in the manuscript. A comparison of diffusion coefficients of bare NDs, measured for the current study, with the literature is shown in Table 2 below. Table 2. Comparison of diffusion coefficients of NDs and ND-SF spheres with already published work for non-coated and coated nanoparticles.

D values for non-coated NPs in cells (µm2/s) D values for encapsulated NPs in cells (µm2/s)

Current work Reference12 1.00×10-4-0.50 45 nm NDs: 3.00×10-3

Reference 34 163 nm NDs: 6.00×10-31×10-2

SF sphere 160 nm lipid encapsulation: coated ND: 6.00×10-4-1.02 4.00×10-20.12

Reference35 50 nm NDs: 1.00 µm2/s (rare)

Reference36 QDs: 0.30

Lipid molecule only: 4.00

The low mobility of NDs agrees with the previous data34-35 published for NDs of similar sizes inside the cells. In some cases the NDs are even reported to get trapped and most of the NDs become immobilized inside the cells after a short time,35 which might have resulted due to their rough edges and jagged surfaces. The reason for the low mobility of NDs in cells is that they get captured in endosomal or lysosomal vesicles.34


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Reports35, 37 have previously shown that NDs coated with lipid layers internalized into cells easily and showed improvement in diffusion despite the increase in size after coating. A recent study by Hui et al32 reports enhanced diffusion of the cholesterol-based lipids encapsulation of NDs. The diffusion coefficient for this study was enhanced by a factor larger than 10 as compared to bare NDs. Further research by Smith et al37 showed that NDs were efficiently transfected into cells after applying cationic liposome coating, however the diffusion coefficient of these particles in the cells was not reported.

Figure 10. Histogram plot for the comparison of diffusion coefficeints of NDs (blue) and ND-SF spheres (red). The intracellular dynamics of NDs with SF sphere encapsulation were analysed and the diffusion coefficient compared with already published work for lipid coated NPs.12, 36 Similar procedure was repeated and measurements were made with ND-SF spheres to compare the dynamics of SF coated ND-SF spheres with NDs alone in-vitro. Figure 11 (a) and (b) show the fluorescence images of ND-SF spheres inside the cells. Figure 11 (c) and (d) show the 2dimensional Brownian motion trajectories for the circled ND-SF spheres of Figure 11 (a) and



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(b) respectively. Figure 11 (e) and (f) plots the r2 versus time graph for the two circled NDSF spheres for 60 s and 40 s respectively. The diffusion coefficients were measured and were found to vary from (6.0±1.0)×10-4 to 1.025±0.075 µm2/s for 22 tracked individual ND-SF spheres. The mean value of the diffusion coefficient from the range is 0.124 µm2/s. The average increase in D for ND-SF spheres as compared to bare NDs is found to be 2.3±0.9. The statistics for the diffusion coefficients of bare NDs and ND-SF spheres were compared by plotting histograms (red) as shown in Figure 10.


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Figure 11. (a), (b) Wide field fluorescence image of fibroblast cells internalized with ND-SF spheres. (c), (d) 2D Brownian motion trajectories for the spheres marked in (a) and (b) respectively. (e), (f) Displacement square versus time plots for the selected spheres. The red data shows the line of best fit and the equations of the fitted lines are shown in the inset of the figures. For consecutive frames, the spheres can be seen to have taken short but continuous movements for the entire period of observation as can be seen by comparing the Brownian 29 ACS Paragon Plus Environment


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motions of NDs with and without SF sphere encapsulation in Figures 9 and 11, subplots (c) and (d). The Figure 10 illustrates that the range of D for spheres is shifted to higher values despite their larger size range. The increase in mobility is found to be consistent with previous work.12 The second row of Table 2 compares the diffusion of ND-SF spheres to lipid coated NDs as reported in literature. Our results show good agreement with the already published work that proves the enhanced dynamics of nanoparticles in the presence of a bio-coating. In conclusion, the Section demonstrates that the encapsulation of NDs within SF spheres makes the particles increasingly mobile in the cytoplasm of the cells. Higher diffusion coefficients of ND-SF spheres indicate that the silk coated NDs are potentially more suitable for longer term imaging and tracking in biological environment. This would ultimately result in a better quantitative analysis of structure and interactions for biological structures and processes.38 4. CONCLUSIONS In this manuscript the fabrication and characterization of ND-SF spheres is reported. SF spheres encapsulating NDs were synthesized with the co-flow technique. The SEM analysis showed a 400-600 nm range for the diameter of spheres. These SF spheres which are otherwise resistant to degradability in aqueous suspensions at room temperature were found to degrade completely within a week when incubated at 37 ˚C (with slow continuous shaking). The presence of NDs inside SF spheres was confirmed through emission spectral analysis. Comparison of emission properties of ND-SF spheres with NDs revealed that the spheres were able to fluoresce with 2-4 times higher brightness as compared to NDs alone. The smallest spheres were selected for experimental emission analysis. The enhancement range was consistent with numerical calculations that indicated ∼0.6-17.9 times increased 30 ACS Paragon Plus Environment

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emission for spheres on substrate and ∼1.2-1.6 times increase for ND-SF spheres suspended in water. The photostability of NDs was uncompromised with SF coating. The dynamics of these spheres were analysed inside the cells and compared with NDs on their own, which clearly showed enhanced mobility and increased diffusion for SF encapsulated NDs. In conclusion, SF coating not only enhanced the optical emission properties of NDs but also provided a biodegradable coating to the fluorescing NDs which increased their mobility and dynamics inside the biological structures as compared to bare NDs. The stable fluorescence and enhanced brightness of ND-SF spheres has great advantage for bioimaging applications since one can efficiently isolate the emission of spheres from the background signals. This approach of spheres encapsulation is particularly useful in enhancing the image contrast and signal-to-noise ratio of single NDs in biological structures. The high image contrast and the enhanced sensitivity, combined with the perfect photostability of ND would allow long term bio-tracking and imaging. SF encapsulation would facilitate the transport of NDs as non-toxic, highly photostable imaging enabled carriers to deliver drugs, proteins and genetic material into the cells and across cellular membranes, without getting trapped in the cell walls or endosomes. Hence ND-SF spheres can be essentially used for longer-term photostable single particle tracking, and have therefore the potential of being reliable biomarkers. SUPPLEMENTARY MATERIAL I. Numerical analysis for emission enhancement: Numerical simulations were performed using an RSoft FullWAVE (3-dimensional Finite Difference Time Domain package). Emission ratios for ND-SF spheres were calculated with respect to NDs either on Si substrate or inside a PBS suspension. The orthogonal and parallel orientations of the dipole were considered when ratios were calculated with relative to the Si substrate. For enhancement 31 ACS Paragon Plus Environment


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comparison in-vitro, the ratios were calculated independent of dipole polarizationas there is no substrate. The optical emission characteristics of ND encapsulated NV− defects encapsulated in SF were computationally investigated and the study was carried out in two steps. In the ﬁrst step, ND on the Si substrate was considered. In the second step, a sphere encapsulation of SF was introduced in the model. The refractive indices of Si, SF and PBS used in the simulations were nSi = 3.875 + 0.0111i, nSF = 1.54 and nPBS= 1.34 respectively. II. Calculation of NP diameter: The diameters were calculated using the scale bar of SEM at a given magnification. The SEM image was exported to MATLAB using the “Imtool” image processing command. From the image toolkit bar, the “Measure distance” option was selected and the number of pixels (along x-axis) existing in the scale bar (of SEM) were calculated. The factor was used to convert the sizes to nm. For bare NDs alone, the size distribution for 20 particles existed from 22 nm for the smallest ND and 70 nm for the largest agglomerated NDs. The average size was estimated to be 47 nm and the majority existed between 30-60 nm as shown in the bar plot of Figure S1 (a) of supporting information. For the distribution of 28 as prepared ND-SF spheres, the diameters were measured to vary from 190 nm for the smallest sphere to 850 nm, with the maximum number of particles in the 400600 nm range, as shown in Figure S1 (b) in supporting information. The mean value for the distribution were found to be 540 nm. The relevant numbers and discussion has been added to the Section 3.2 of the manuscript. The ND-SF sphere size distribution during PBS incubation on day 3 was also calculated with MATLAB. Due to SF degradation, the size of the spheres reduced to a range of 85-400 nm, as shown in the Figure S2 in the supplementary information. The majority of particles can be clearly seen to have a size range between 85-


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200 nm, suitable for cell culture and hence the ones used for wide-field imaging in-vitro study. III. Confocal imaging and spectral analysis. For each of the NDs alone, as prepared ND-SF spheres and SF only spheres on Si substrates, 100×100 µm2 fluorescence scans were recorded with confocal microscopy and shown in Figure S3 of supporting information. Several particles (8-20) on the scans were analysed for their magnified fluorescence scans, emission rates and spectral analysis. The statistics were accumulated and a representative particle was selected from each sample and shown in Section 3.3. The Figure S3 also illustrates that the agglomeration of NDs can be prevented using SF sphere coating. IV. Background autofluorescence from cells alone. The background autofluorescence from the cells is shown in Figure S4 of the supporting information, where the counts photobleach rapidly, minimizing the background signal within a few seconds. AUTHOR INFORMATION Author Contributions The manuscript was written through contributions of all authors. All authors have given approval to the final version of the manuscript. ACKNOWLEDGEMENTS A. Khalid is supported by Melbourne Research Scholarships awarded by The University of Melbourne and acknowledges Melbourne Scholarships for providing ORES funding to visit Prof. F.G. Omenetto at Tufts University S. Tomljenovic-Hanic is supported by an ARC Australian Research Fellowship (DP1096288). The numerical part of research was undertaken with the assistance of resources provided at the NCI National Facility systems at



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The Australian National University through the National Computational Merit Allocation Scheme supported by the Australian Government. REFERENCES 1. Man, H. B.; Ho, D., Diamond as a nanomedical agent for versatile applications in drug delivery, imaging, and sensing. Physica Status Solidi a-Applications and Materials Science 2012, 209 (9), 1609-1618. DOI: 10.1002/pssa.201200470. 2. Mochalin, V. N.; Shenderova, O.; Ho, D.; Gogotsi, Y., The properties and applications of nanodiamonds. Nat Nano 2012, 7 (1), 11-23. 3. Perevedentseva, E.; Lin, Y. C.; Jani, M.; Cheng, C. L., Biomedical applications of nanodiamonds in imaging and therapy. Nanomedicine (London, England) 2013, 8 (12), 2041-60. DOI: 10.2217/nnm.13.183. 4. Xing, Y.; Dai, L. M., Nanodiamonds for nanomedicine. Nanomedicine 2009, 4 (2), 207-218. DOI: 10.2217/17435889.4.2.207. 5. Schrand, A. M.; Huang, H.; Carlson, C.; Schlager, J. J.; Ōsawa, E.; Hussain, S. M.; Dai, L., Are Diamond Nanoparticles Cytotoxic? The Journal of Physical Chemistry B 2007, 111 (1), 2-7. DOI: 10.1021/jp066387v. 6. Schirhagl, R.; Chang, K.; Loretz, M.; Degen, C. L., Nitrogen-Vacancy Centers in Diamond: Nanoscale Sensors for Physics and Biology. Annual Review of Physical Chemistry, Vol 65 2014, 65, 83-105. DOI: 10.1146/annurev-physchem-040513-103659. 7. Khalid, A.; Lodin, R.; Domachuk, P.; Tao, H.; Moreau, J. E.; Kaplan, D. L.; Omenetto, F. G.; Gibson, B. C.; Tomljenovic-Hanic, S., Synthesis and characterization of biocompatible nanodiamondsilk hybrid material. Biomedical Optics Express 2014, 5 (2), 596-608. DOI: 10.1364/boe.5.000596. 8. Youn, H.-Y.; McCanna, D. J.; Sivak, J. G.; Jones, L. W., In vitro ultraviolet–induced damage in human corneal, lens, and retinal pigment epithelial cells. Molecular Vision 2011, 17, 237-246. 9. Fu, Y.; Zhang, J.; Lakowicz, J. R., Metal-enhanced fluorescence of single green fluorescent protein (GFP). Biochemical and Biophysical Research Communications 2008, 376 (4), 712-717. DOI: 10.1016/j.bbrc.2008.09.062. 10. Mohan, N.; Tzeng, Y. K.; Yang, L.; Chen, Y. Y.; Hui, Y. Y.; Fang, C. Y.; Chang, H. C., Sub20-nm Fluorescent Nanodiamonds as Photostable Biolabels and Fluorescence Resonance Energy Transfer Donors. Advanced Materials 2010, 22 (7), 843-+. DOI: 10.1002/adma.200901596. 11. Cinteza, L. O., Quantum dots in biomedical applications: advances and challenges. NANOP 2010, 4. DOI: 10.1117/1.3500388. 12. Hui, Y. Y.; Zhang, B. L.; Chang, Y. C.; Chang, C. C.; Chang, H. C.; Hsu, J. H.; Chang, K.; Chang, F. H., Two-photon fluorescence correlation spectroscopy of lipid-encapsulated fluorescent nanodiamonds in living cells. Optics Express 2010, 18 (6), 5896-5905. DOI: 10.1364/oe.18.005896. 13. Krueger, A.; Lang, D., Functionality is Key: Recent Progress in the Surface Modification of Nanodiamond. Advanced Functional Materials 2012, 22 (5), 890-906. DOI: 10.1002/adfm.201102670. 14. Rehor, I.; Slegerova, J.; Kucka, J.; Proks, V.; Petrakova, V.; Adam, M. P.; Treussart, F.; Turner, S.; Bals, S.; Sacha, P.; Ledvina, M.; Wen, A. M.; Steinmetz, N. F.; Cigler, P., Fluorescent Nanodiamonds Embedded in Biocompatible Translucent Shells. Small 2014, 10 (6), 1106-1115. DOI: 10.1002/smll.201302336. 15. Rehor, I.; Mackova, H.; Filippov, S. K.; Kucka, J.; Proks, V.; Slegerova, J.; Turner, S.; Van Tendeloo, G.; Ledvina, M.; Hruby, M.; Cigler, P., Fluorescent Nanodiamonds with Bioorthogonally Reactive Protein-Resistant Polymeric Coatings. Chempluschem 2014, 79 (1), 21-24. DOI: 10.1002/cplu.201300339. 16. Zhang, Z. W.; Niu, B. H.; Chen, J.; He, X. Y.; Bao, X. Y.; Zhu, J. H.; Yu, H. J.; Li, Y. P., The use of lipid-coated nanodiamond to improve bioavailability and efficacy of sorafenib in resisting


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Table of content graphic

Fluorescent Nanodiamond Silk Fibroin Spheres: Advanced Nanoscale Bioimaging Tool Asma Khalid1, Alexander N. Mitropoulos2, BenedettoMarelli2, David A. Simpson1, Phong A. Tran3,4, Fiorenzo G. Omenetto2* and Snjezana Tomljenovic-Hanic1*

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Fluorescent Nanodiamond Silk Fibroin Spheres: Advanced Nanoscale

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