Lighting Up Plasmonic Nanostar Colloids for Metal Enhanced

LIGHTING UP PLASMONIC NANOSTAR COLLOIDS FOR METAL ENHANCED FLUORESCENCE. UNDER TWO-PHOTON NIR EXCITATION. Lixia Zhou ...
0 downloads 0 Views 3MB Size
Download PDF
Article pubs.acs.org/JPCC

Cite This: J. Phys. Chem. C XXXX, XXX, XXX−XXX

Lighting up Plasmonic Nanostar Colloids for Metal-Enhanced Fluorescence under Two-Photon Near-Infrared Excitation Lixia Zhou, Kuan-Jen Chen, Jacob D. Ramsey, C. Kyle Almlie, and Sean M. Burrows* Department of Chemistry, Oregon State University, 153 Gilbert Hall, Corvallis, Oregon 97331, United States

J. Phys. Chem. C Downloaded from pubs.acs.org by UNIV OF KENTUCKY on 08/23/18. For personal use only.

S Supporting Information *

ABSTRACT: Literature reports detail the theory for strong electric-field enhancements sustained and localized on gold nanostars (NSs). The enhanced fields are expected to alter the fluorescence of nearby fluorophores. However, there are very few literature reports on experimental evidence for enhanced fluorescence from dyes bound to gold NSs. Furthermore, the literature lacks experimental studies on metal-enhanced fluorescence under near-infrared (NIR) two-photon excitation. In our work, a nanoassembly was made with a NS at the core, then coated with avidin followed by biotinylated-DNA-linkers that hybridized to different Cy3-labeled DNA strands. We will discuss experimental results on physical insights including: (1) how the nanoassembly synthesis conditions and dye orientation affected the fluorescence enhancement and (2) how the NSs affected the organic fluorophore’s photostabilityboth under NIR two-photon excitation. We observed enhancement factors ranging from 1.2 to 3.5 depending on the nanoassembly synthesis conditions. A biotin displacement experiment supported the fluorescence enhancement results. Our work contributes new knowledge and empirical insights that will enable advances in the field of colloidal dye NS-based applications.

■

INTRODUCTION Metals can cause a dye’s fluorescence to enhance or quench depending on the type of materials, nanostructures, and dyes. Metal-enhanced fluorescence (MEF) is a phenomenon that results from photophysical interactions between fluorophores and the nanostructure of metallic surfaces.1−6 These photophysical interactions depend on the distance between the metal and fluorophores, as well as the ability of the metal’s scattering field and plasmon to couple with the fluorophore’s excitation and emission spectra.7,8 A plasmon is the oscillation of the metal’s free electrons upon interaction with incident light. The resulting enhanced electromagnetic fields can interact with a fluorophore’s absorption and emission properties to alter the fluorescence intensity.1 One theoretical approach to predict electromagnetic field enhancement around a metal is the finite-difference time domain (FDTD) calculations.9 There have been many examples of FDTD calculations for nanoparticles that showed electric-field-enhancement factors over one and corresponding experiments that showed MEF.10−12 Many MEF experiments have been performed on different materials. For example, silver island films13−15 are capable of enhancement factors over 2. Others have found that nanoparticles10,16−18 such as nanospheres, nanorods, or nanoshells achieve enhancement factors ranging from 1.1 to 90. Another study of fluorescence on nanobipyramids19 showed a 1.4-fold fluorescence enhancement. Although many metal nanostructures have been studied for their MEF phenomenon, new nanostructures are continually emerging. Thus, there is a need to elucidate the photophysical © XXXX American Chemical Society

relationships between each new type of nanomaterial and a dye. In particular, investigating the MEF properties on different materials is needed to give researchers better ideas on how to incorporate them into their research or application needs. One emergent nanostructure is gold nanostars (NSs). Gold NSs are nanoparticles with sharp spikes protruding from the center core. Better understanding of the MEF phenomenon will facilitate future applications of NSs into a variety of fluorescence-based applications such as biosensors and chemical imaging. NSs are known to have strong enhanced electric fields.2,20−25 One FDTD calculation revealed that the sharp spikes on the individual NS generate electric-field-enhancement factors up to and over 200.20 Such a strong electric-field-enhancement suggests that gold NSs are able to enhance the fluorescence of nearby fluorophores. One study of the fluorescence phenomena of dyes on NSs showed strongly quenched fluorescence at short distances.26 The dyes (Rhodamine B, Atto 590, and Atto 610) were only able to achieve full fluorescence recovery when the NSs and dyes were separated by about 15 nm.26 However, another study showed strong fluorescence enhancement of Alexa Fluor 750 and 790 on gold NS substrate.27 The following reasons outline how colloidal gold NSs interacting with ultrafast near-infrared (NIR) laser excitation Received: July 10, 2018 Revised: July 30, 2018 Published: July 31, 2018 A

DOI: 10.1021/acs.jpcc.8b06609 J. Phys. Chem. C XXXX, XXX, XXX−XXX

Article

The Journal of Physical Chemistry C Table 1. Oligonucleotide Sequences name a

3pBT(n)-cpDNA 5pBT(n)-cpDNAa Cy3DNAa

sequence 1

sequence 2

5′-TGAGGTAGTAGGTTGTATAGTT+T(30) regions-CnO-biotin-3′ 5′-biotin-CnO+T(0) regions-TGAGGTAGTAGGTTGTATAGTT-3′ 5′-AACTATACAACCTACTACCTCA-3′-Cy3

none 5′-biotin-CnO+T(30) regions-TGAGGTAGTAGGTTGTATAGTT-3′ none

Duplex formation between the BT(n)-cpDNA and Cy3DNA were predicted to have melting temperatures over 50 °C and Gibbs free energies around −25 kcal/mol; thus, the duplexes were favored to form at room temperature. The thermodynamic results of ΔG and Tm were obtained from online freeware36−38 that takes the DNA sequences as inputs. a

will contribute to the field of MEF. First, there are conflicting reports on the ability of NSs to enhance or quench fluorescence. Second, there are very few, if any, studies of MEF from dyes on a NS surface or in a colloid under twophoton excitation. Third, our NS synthesis approach yields sharp spikes with highly reproducible plasmon resonances in the NIR range 718 ± 27 nm (N = 27).28 NIR radiation has the potential to stimulate the NS’s plasmon and excite the fluorophore such that they interact to enhance emission. One plausible reason for this is the NS’s plasmon band and fluorophores’ two-photon absorption are both in the NIR range. Fourth, NIR excitation will benefit research involving applications in media that are highly scattering.18,21,29−35 Fifth, MEF research on colloidal NSs is not well-studied. NSs in colloids are advantageous over NSs on substrates because colloids simplify sample preparation steps such as deposition and washing, typical of MEF on substrates. Moreover, the mass transportation limit on surfaces can be circumvented with colloids. In this work, we present on the groundwork for two-photoninduced fluorescence enhancement of Cy3 near gold NSs in colloids. To investigate the NS’s influence on fluorescence, Cy3-labeled DNA (Cy3DNA) strands were tethered onto gold NSs through avidin and biotinylated DNA linkers (BT(n)cpDNA). The enhancement factors were determined based on a ratio of Cy3 signal on the NSs to that of controls in the absence of NSs. To further validate MEF, a biotin displacement experiment was performed. The fluorescence signals of Cy3 bound to and freed from the NSs was compared. Next, various experimental working conditions were optimized to achieve the highest possible fluorescence enhancement and evaluate the reproducibility. Finally, the photostability of Cy3 was studied to learn more about how gold NSs influence the fluorescence signals of nearby fluorophores. To the best of our knowledge, this is the first study of two-photon-induced MEF from colloidal gold NS.

ascorbic acid were added simultaneously, with pipets vertical. The reaction was carried out for 30 seconds under stirring at 700 rpm. Next, 100 μL of 2% SDS was added for another 30 s. To quench the reaction, the solution from the previous step was centrifuged for 15 min at 3000g (gravitational) and 20 °C. The supernatant was removed to leave a 2 mL suspension of NSs. Finally, the suspension was sonicated for 8 min and filtered by 0.22 μm nitrocellulose membrane purchased from Corning.21,28 Oligonucleotides. Oligonucleotides were purchased from Integrated DNA Technologies (IDT, Coralville, IA) and used as received. The DNA came as stock solutions prepared in IDTE buffer at a pH of 8.0. Table 1 lists the DNA sequences for the Cy3-labeled DNA strand (Cy3DNA) and the biotinylated DNA strands with polythymine and capture sections (BT(n)-cpDNA). The capture DNA sequence (cpDNA) on 3pBT(n)-cpDNA or 5pBT(n)-cpDNA were complementary to the Cy3DNA. The distance between Cy3 and the NS’s surface was controlled by the number (n) of thymine bases (n = 0 and 30) on the BT(n)-cpDNA. The predicted melting temperature for the duplex of Cy3DNA and BT(n)-cpDNA was above 50 °C, and the Gibbs free energy was about −25 kcal/mol; thus, the hybridized strands were expected to be stable at room temperature. Nanoassembly Steps for Metal Enhanced Fluorescence Experiments. Avidin was purchased from Thermo and resuspended in sterile 18 MΩ cm H2O. Tris buffer (pH 10) and 2 M magnesium chloride (MgCl2) were obtained from Fisher Scientific. TWEEN 20 was purchased from Acros. Phosphate-buffered saline (PBS, pH 7.4) was obtained from Gibco. All chemicals were used as received. Working solutions of DNA were prepared by diluting the stock solutions with a house buffer containing: 10 mM Tris buffer, 2.5 mM MgCl2, and 0.005% TWEEN 20 in 0.1× PBS. The pH of the house buffer was about 8. To functionalize NSs, the biotin−avidin system was chosen for two reasons: (1) the binding affinity39 between avidin and biotin is approximately 1015 and (2) to validate MEF by performing competitive biotin displacement studies. Scheme 1 shows how the NSs were functionalized. To coat avidin on the NSs, a mixture of NSs and avidin was reacted for 20 hours at 4 °C to form avidin-coated NSs (NS-A). Meanwhile, 5pBT(n)-cpDNA linkers were hybridized with Cy3DNA for at least 30 min, but no more than 1 day, to form a complex called 5pBT(n)Cy3. A similar procedure was carried out for the 3pBT(n)-cpDNA linkers to form 3pBT(n)Cy3 complexes. To complete the nanoassembly, the 5pBT(n)Cy3 was added into the NS-A solution to form Cy3-coated NSs (NS-A-5pBT(n)Cy3). Similarly, the 3pBT(n)Cy3 and NS-A reacted to form NS-A-3pBT(n)Cy3. Two-Photon Fluorescence Instrument. In this work, a custom-built two-photon fluorimeter, previously described,32 delivered laser light to excite and collect fluorescence spectra

■

EXPERIMENTAL SECTION Nanostar Synthesis. Chloroauric acid (HAuCl4) was purchased from Sigma-Aldrich, 12 M HCl was purchased from EMD Millipore, silver nitrate (AgNO3) was purchased from Alfa Aesar, ascorbic acid was purchased from Macron Chemical, and sodium dodecyl sulfate (SDS) was purchased from Avantor. NanoXact silica nanospheres (80 nm) were purchased from nanoComposix (10 mg/mL). All chemicals were used as received. The NSs were synthesized by a seed-mediated method carried out at 5 °C.28 A detailed description to synthesize the seed particles and NSs has been previously published.28 For the NS synthesis, first, 10 mL of 0.25 mM HAuCl4, 10 μL of 1 M HCl, and 100 μL of seed gold nanospheres (∼12 nm) were mixed together. Second, the solution was brought to 5 °C and then 100 μL of 3 mM AgNO3 and 50 μL of 100 mM B

DOI: 10.1021/acs.jpcc.8b06609 J. Phys. Chem. C XXXX, XXX, XXX−XXX

Article

The Journal of Physical Chemistry C

sure the SiNsp did not quench the Cy3 fluorescence (data not shown). Because the SiNsp with A-BT(n)Cy3 control was able to account for the presence of nanoparticles, it was a better control than either a simple buffer solution or the A-BT(n)Cy3 in buffer. The background for the A-BT(n)Cy3 in 0.1× PBS control was a complex between avidin and unlabeled biotinylated DNA in buffer solution. The background for the A-BT(n)Cy3 in SiNsp colloid control was the avidin complexed with unlabeled biotinylated DNA in the SiNsp colloid. The background-corrected spectrum of Cy3 in control was the difference between the signal of Cy3 in control solutions and their respective backgrounds (more details are described in section S1.2 of the Supporting Information). The enhancement factor was the ratio of the summed counts of background-corrected Cy3 on NSs to that in controls of 0.1× PBS or SiNsp (more details are described in section S1.3 of the Supporting Information). Nanostar and Nanoassembly Characterization. Bare NSs were imaged with a transmission electron microscope (TEM). The TEM is an FEI Titan 80-200 TEM/scanning TEM (FEI, Hillsboro, OR, USA). The voltage was set to 200 kV and the magnifications were 19.5k× and 51k×. To confirm that avidin and the BT(n)Cy3 were bound to the NSs, we ran UV/vis, NanoSight, and dynamic light scattering (DLS) experiments. For each technique, we analyzed bare NSs, avidin-coated NSs (NS-A), and Cy3coated NSs (NS-A-BT(n)Cy3). UV/vis spectrophotometry measured changes in the peak plasmon wavelength as the surface chemistry around the NSs changed. NanoSight provided size and concentration information. The DLS validated the size results from the NanoSight. A UV-1800 spectrometer (Shimadzu, Kyoto, Japan) measured the plasmon band of bare and chemically coated NS. The wavelength range was scanned from 200 to 1000 nm with a pitch of 2 nm. Scan speed was set to fast. Functionalized NSs were observed as shifts in the peak plasmon wavelength because avidin and biotinylated DNA were added. The NSs’ size and concentration were determined with a NanoSight (NS 500, NanoSight Ltd, Salisbury, UK). The experiments were all performed at 25 °C. More details for the experimental settings are in section S2 of the Supporting Information. DLS analysis was performed on a Zetasizer Nano ZS (Malvern Instruments Ltd., Worcestershire, UK). DLS validated the NanoSight size data and measured the NSs’ zeta potential before, in the middle of, and after they were functionalized. The bare and chemically coated NS suspensions were measured directly without further dilution. For the detection of NS size and zeta potential with DLS, a disposable folded capillary cell containing the sample was measured three times at 25 °C. Optimizing Nanoassembly Reaction Conditions for Metal Enhanced Fluorescence Experiments. The following describes experiments that revealed the optimal working conditions and evaluated the reproducibility for fluorescence enhancement. First, we evaluated the following reaction conditions: (i) the concentration of NS, (ii) the mole ratio of BT(30)Cy3 to avidin, and (iii) the amount of avidin to NS. In total, there were 26 different reaction conditions. We identified and repeated four reaction conditions that gave the largest enhancement factors.

Scheme 1. Procedure to Functionalize Gold NSs with Avidin and then the Biotinylated DNA Linker-Cy3 Complexa

a

(NS = bare NS; NS-A = avidin-coated NS; BT(n)-cpDNA = biotinylated DNA linker; Cy3DNA = Cy3-labeled DNA strand; BT(n)Cy3 = biotinylated DNA linker-Cy3 complex; NS-A-BT(n)Cy3 = NS-coated with avidin and biotinylated DNA linker-Cy3 complex).

from the samples. Briefly, a Spectra-Physics Mai Tai laser fixed at 742 nm with an average power of 75 mW excited the NS− fluorophore complex. Signal was collected in an epiconfiguration and directed to a fiber-optic-coupled spectrometer (Acton, SP2300). The spectrometer was outfitted with a 300 grooves/ mm grating blazed at 500 nm. The exit port was outfitted with a Princeton Instrument electron-multiplying charge-coupled device (CCD) camera. Light-field software controlled the spectrometer and camera with the following settings: slit width was 1000 μm and grating center wavelength was set to 570 nm for Cy3. Six frames (spectra) were collected with an exposure time of 500 ms and 10 exposures per frame, for each sample measured. The analog conditions were listed as low-noise, 100 kHz, and high analog gain. The readout was full frame with a readout time of 77.485 ms. To keep the thermal noise low, the CCD chip was cooled to −70 °C. Metal Enhanced Fluoresence Controls and Analysis Methods. To extract the Cy3 signal, we needed to measure the background signal from all-sample components except the dye. To account for background signal from each nanoparticle and bound constituent, we evaluated the signal from gold NSs coated with avidin and DNA strands that lacked Cy3 (referred to as unlabeled NSs). The signal from unlabeled NSs served as the background for the signal of NSs coated with avidin and Cy3-labeled DNA (referred to as Cy3-labeled NSs). The background-corrected spectrum of Cy3 on NSs was the difference between the signal from Cy3-labeled NSs and the background signal from unlabeled NSs (more details are described in section S1.1 of the Supporting Information). To determine the fluorescence enhancement factor, we compared the background-corrected Cy3 signal on NSs to that in controls. We relied on two Cy3 controls: (1) an avidin and BT(n)Cy3 complex (A-BT(n)Cy3) in buffer solution (0.1× PBS control) and (2) A-BT(n)Cy3 in a suspension of 80 nm silica nanospheres (SiNsp control). We chose the A-BT(n)Cy3 in buffer solution as a control because it contained all of the experimental components except the NSs. The A-BT(n)Cy3 was a better control sample than BT(n)Cy3 in buffer solution alone because A-BT(n)Cy3 better approximates the fluorophore’s avidin environment on gold NSs. We chose SiNsps as another control because they had a similar size as gold NSs and thus created a colloidal environment without plasmons or enhanced electric fields around the fluorophores. We made C

DOI: 10.1021/acs.jpcc.8b06609 J. Phys. Chem. C XXXX, XXX, XXX−XXX

Article

The Journal of Physical Chemistry C

agreed well with the NanoSight measurements of 81.1 ± 8.7 nm (N = 9). The predicted final structures and orientation of dye with respect to the nanosurface are shown in Scheme 2. The

The four reaction conditions were (i) 1 mL of NSs with 80 pmol avidin and 160 pmol 3pBT(30)Cy3; (ii) 1 mL of NSs with 80 pmol avidin and 240 pmol 3pBT(30)Cy3; (iii) 1 mL of NSs with 80 pmol avidin and 160 pmol 5pBT(30)Cy3; and (iv) 1 mL of NSs with 40 pmol avidin and 120 pmol 5pBT(30)Cy3. For statistical analysis, each of the four reaction conditions were repeated four times. Each replicate reflected independent avidin and BT(30)Cy3 coatings from the same batch of NS. Data were analyzed with the student t-test to reject the null hypothesis at the 95% confidence level. For these experiments, the NSs’ concentration was kept at an optical density of 0.20 ± 0.002 (N = 3). Gold NSs were functionalized with avidin and BT(30)Cy3 by following the same procedures described in the Nanoassembly Steps for Metal Enhanced Fluorescence Experiments, Experimental Section. Effect of Gold Nanostar on the Photostability of Nearby Fluorophores under Two-Photon Excitation. To study the photostability of Cy3 in the presence of NSs, the fluorescence of Cy3 in 0.1× PBS and on gold NSs were examined over 30 min (data not shown). Precursor experiments over a 4 h period showed that the fluorescence signal remained stable after 30 min. The Cy3 was exposed to laser light during the entire time period. Cy3-coated gold NSs (NS-A-3pBT(30)Cy3) were prepared as previously described in Scheme 1. As a no NS control, Cy3 in 0.1× PBS was prepared by adding avidin and 3pBT(30)Cy3 in 0.1× PBS. The photostability study had the same acquisition settings as described in the Two-Photon Fluorescence Instrument, Experimental Section.

Scheme 2. Predicted Nanoassembly and Dye Orientation of NS-A-3pBT(n)Cy3 and NS-A-5pBT(n)Cy3a

a Just one avidin and BT(n)Cy3 complex are shown to simplify the schematic.

3pBT(n)Cy3 orients Cy3 far away from the nanosurface. The 5pBT(n)Cy3 orients Cy3 22 nucleic acids closer to the nanosurface than the 3pBT(n)Cy3. Both 3pBT(n)Cy3 and 5pBT(n)Cy3 were studied to learn the effect of dye orientation on NS-induced MEF. The BT(0)-cpDNA and BT(30)-cpDNA were evaluated because they demonstrated sufficient fluorescence enhancement factors for the BT(n)Cy3-NS complex (data not shown). Cy3 was chosen because of its low quantum yield around 0.24.40 We chose a dye with low quantum yield for two reasons. First, previous studies of MEF on NSs with highquantum-yield dyes did not show fluorescence enhancement.26 Second, reports on other nanomaterials recommend lowquantum-yield dyes for enhancement purposes.15 Metal Enhanced Fluorescence from Nanostar Colloids. Figure 2 shows the MEF from freely diffusing NSs

■

RESULTS AND DISCUSSION Nanostar Morphology and Nanoassembly Scheme. Figure 1 shows the NSs’ morphology from a TEM image.

Figure 2. Two-photon MEF from Cy3 on ∼80 nm gold NSs (NS, blue) compared to fluorescence on 80 nm Si nanospheres (SiNsp, red). NS and SiNsp were coated with avidin and 3pBT(30)Cy3. Samples were excited at 742 nm.

compared to the control of SiNsp with avidin and 3pBT(30)Cy3. Fluorescence of Cy3 on NSs was 1.68 ± 0.01 (N = 6) times the signal in SiNsp colloids. We observed higher Cy3 fluorescence signals in a NS colloid compared to that in the 0.1× PBS control with avidin and 3pBT(30)Cy3 (data not shown). These results provide evidence that NSs were able to enhance the Cy3 fluorescence. Theoretical literature suggests that one nanostructural cause for NS MEF of Cy3 is the many sharp spikes in close proximity to each other that localize the enhanced electric fields on each NS.20−24,41,42 A plausible mechanism for observed fluorescence enhancement is that the NS acts as a “donor” as the plasmon

Figure 1. TEM image of NSs. Spikes ranged in length from 6 to 46 nm with an average tip-to-tip distance of 73 ± 47 nm (N = 156).

Some NSs had very short spikes, some had very long spikes, and some had both short and long spikes. To measure the overall tip-to-tip diameter of the NSs, a representative sample group containing NSs with spikes that ranged in length from 6 to 46 nm were selected. From the NSs’ TEM images, the overall tip-to-tip distance was found to be 73 ± 47 nm (N = 156) from ImageJ measurements. The TEM measurements D

DOI: 10.1021/acs.jpcc.8b06609 J. Phys. Chem. C XXXX, XXX, XXX−XXX

Article

The Journal of Physical Chemistry C

details in section S3 of the Supporting Information). In contrast, the fluorescence decreased by (0.74 ± 0.14) × 106 counts upon biotin addition in a NS colloid. These controls provide additional evidence to validate that the enhanced fluorescence was due to the NS and not an artifact. Confirmation of Nanostar Surface Functionalization. To gauge the success of forming the nanoassembly, we made DLS, NanoSight, and spectrophotometric measurements. These measurements were made before, during, and after the functionalization of NSs with avidin and then BT(n)Cy3. The NanoSight and DLS measurements revealed a change in size from bare to chemically coated NSs. The results provided evidence that the NS’s surface was functionalized. The size of NSs, NS-A, and NS-A-BT(30)Cy3 from NanoSight and DLS are shown in Figure 4.

absorbs photons in the NIR and then couples with the excitation and emission properties of nearby “acceptor” fluorophores. In a follow-up study, we will study how different extents of spectral overlap between the plasmon and fluorophore’s excitation and emission spectra alter the dye’s fluorescence intensity. We observed similar fluorescence enhancement from NSs with 5pBT(30)Cy3 as we did with 3pBT(30)Cy3. Because there was no dramatic difference among the enhancement factors for the different dye orientations, we were not able to draw any conclusions on the effect of dye orientation on MEF. Competitive Biotin Displacement Study to Validate Metal Enhanced Fluorescence on Nanostars. To further confirm MEF of Cy3 by the NSs, the Cy3-biotin-linker was displaced from the NS with pure unlabeled biotin. Unlabeled pure biotin was added to a solution of NS-A-5pBT(0)Cy3. The displacement was driven by the higher binding affinity of biotin for avidin over the Cy3-biotin-linker. The house buffer was added to a separate solution to control for dilution of NSA-5pBT(0)Cy3. For both solutions, the fluorescence of Cy3 was observed over time. Measurements were made before (time zero) and after the biotin or buffer was added. Figure 3 shows the Cy3 fluorescence signal stabilized after 20 min when either the buffer or biotin was added. Fluctuation

Figure 4. Change in size as the NSs were functionalized with different nanoassembly constituents. The size changed from bare NSs to NSs with avidin to NSs with avidin and BT(30)Cy3. Sizes were measured with NanoSight (blue) and DLS (red). 5p = biotin on 5-prime end of the DNA linker and 3p = biotin on the 3-prime end of DNA linker. N was the number of replicate samples measured (N = 3).

Figure 3. Cy3 fluorescence over time after buffer or biotin were added to NS-A-5pBT(0)Cy3 (run in triplicate). Trials with biotin added (freed Cy3 from NSs) had lower fluorescence than those with buffer added (bound Cy3 on NSs). The biotin displacement studies provide evidence to validate two-photon MEF by gold NSs.

Prior to functionalization of NSs with avidin and 5pBT(30)Cy3, DLS measurements determined that the bare NSs were 56.4 ± 11.5 nm (N = 3). After 0.4 μM avidin was added to the NSs, the size increased to 79.7 ± 20.1 nm (N = 3). Addition of 5pBT(30)Cy3 to the avidin-coated NSs further increased the size to 158.1 ± 43.3 nm (N = 3). The increase in size with the addition of avidin and 5pBT(30)Cy3 provided evidence of successful functionalization of the gold NSs. A similar trend in size change was observed from NanoSight size measurements (Figure 4). The NanoSight results provided complementary evidence that the NSs were coated with 0.4 μM avidin and 5pBT(30)Cy3. DLS and NanoSight data from coating NSs with avidin and 3pBT(30)Cy3 showed similar changes in size as described for coating 0.4 μM avidin and 5pBT(30)Cy3. Upon adding 0.8 μM avidin, the size of the NSs increased to 238.4 ± 75.0 nm (DLS, N = 3) and then decreased to 105.0 ± 25.3 nm (DLS, N = 3) after addition of 3pBT(30)Cy3. We suspect the avidincoated NSs (NS-A) aggregated together, resulting in large particles (>200 nm); then addition of 3pBT(30)Cy3 broke the NS-A aggregates up and caused the size to decrease. The change in size for each coating stage was indicative of successful attachment of avidin and 3pBT(30)Cy3 on NS. Analysis of the zeta potential during different stages of nanoassembly provided additional evidence that the NSs were successfully coated with avidin and BT(30)Cy3 (Table 2). The

of the fluorescence in the first 20 min after dilution with buffer was attributed to equilibration of the new concentration. After 20 min, the NSs bound to Cy3 had (5.41 ± 0.14) × 106 counts per 500 ms (N = 6), but the freed 5pBT(0)Cy3 had (4.68 ± 0.04) × 106 counts per 500 ms (N = 6). The Cy3 fluorescence was dimmer after biotin displacement because the Cy3 was no longer near the NS surface and thus unable to interact with the NS’s plasmon or scattering properties. These results provide additional evidence for two-photon-induced MEF by NSs. For statistical rigor, we repeated these experiments three times. Each replicate was prepared from the same batch of avidincoated NSs and represented independent coatings with 5pBT(0)Cy3. To further prove the NS’s ability to enhance fluorescence, Cy3-biotin-linker displacement control experiments were performed that consisted of either house buffer or nonplasmonic Si nanospheres. Similar to previous studies, the Cy3biotin linker was bound to avidin in solutions of 0.1× PBS or Si nanospheres (SiNsp). Figures S5 and S6 show the Cy3 fluorescence signal in 0.1× PBS and SiNsp suspensions, respectively, with the addition of either the buffer or biotin. Adding biotin to the control samples did not decrease the fluorescence signals compared to the buffer dilution (see E

DOI: 10.1021/acs.jpcc.8b06609 J. Phys. Chem. C XXXX, XXX, XXX−XXX

Article

The Journal of Physical Chemistry C Table 2. DLS Zeta Potentials of Bare NSs, NSs with Avidin, and NSs with Avidin and BT(30)Cy3a sample (N = 3)

NS

NS + 0.4 μM avidin

zeta potential (mV)

−32.6 ± 9.9

14.3 ± 3.3

NS + 0.8 μM avidin NS + 0.4 μM avidin + 5pBT(30)Cy3 NS + 0.8 μM Avidin + 3pBT(30)Cy3 13.8 ± 3.9

−25.2 ± 6.1

−18.2 ± 5.8

a

5p = biotin on the 5-prime end of DNA linker and 3p = biotin on the 3-prime end of DNA linker.

zeta potential changed from −32.6 ± 9.9 mV (N = 3) for bare NSs to 14.3 ± 3.3 mV (N = 3) after 0.4 μM avidin addition. The zeta potential further changed to −25.2 ± 6.1 mV (N = 3) after addition of the 5pBT(30)Cy3. The fact the zeta potential changed throughout the functionalization process confirmed successful coatings. Similar trends in zeta potential change were observed for the addition of 0.8 μM avidin and 3pBT(30)Cy3 to the NSs (Table 2). Figure 5 provides further evidence of the NS functionalization by comparison with the different UV/vis spectra of bare

were accessible to the biotin-linker. Third, if there are too many avidins on the NS’s surface, then, the Cy3-biotin-linkers may get too closely packed. In this case, the fluorescence can be quenched from the Cy3 interacting with other nearby dyes or nucleic acids.22 Hence, we evaluated the following reaction conditions: (i) the concentration of NSs (Tables S2 and S3), (ii) the mole ratio of BT(30)Cy3 to avidin (Tables S4 and S5), and (iii) the amount of avidin to NSs (Tables S6 and S7). Tables S2−S7 contain the fluorescence enhancement results under different reaction conditions. Analysis of Tables S2−S7 (with 26 nanoassembly reaction conditions in total) revealed some similarities and differences among the various reaction conditions (more details are in section S4 of the Supporting Information). To evaluate the reproducibility of the optimal nanoassembly reaction conditions to achieve the largest MEF enhancement factor, 4 out of the 26 reaction conditions were chosen to repeat because they achieved the largest enhancement factors (based on results from Tables S2−S7). The four working conditions are listed in the Experimental Section. Each replicate came from the same batch of NS but consisted of independent coatings of avidin and BT(30)Cy3. Table 3 reveals that for NS-A-3pBT(30)Cy3, the largest enhancement was 2.25 ± 0.97 (N = 4) when 80 pmol of avidin was added in 1 mL of the NS colloid with a 2−1 mole ratio of 3pBT(30)Cy3 to avidin. In contrast, the best enhancement for the other dye orientation, NS-A-5pBT(30)Cy3, was 1.50 ± 0.31 (N = 4) when 40 pmol avidin was added in 1 mL of the NS colloid with a 3−1 mole ratio of 5pBT(30)Cy3 to avidin. Cy3 Fluorescence Photostability on Gold Nanostars. Figure 6 shows the normalized fluorescence signal of Cy3 on

Figure 5. UV/vis spectrum of bare NSs, NSs with avidin, and NSs with avidin and BT(30)Cy3. 5p = biotin on the 5-prime end of DNA linker and 3p = biotin on the 3-prime end of DNA linker.

NSs, NSs with avidin, and NSs with avidin plus BT(30)Cy3. The bare NS’ plasmon band peaked near 668 nm prior to functionalization. After adding 0.4 μM avidin, the peak plasmon wavelength red-shifted about 47 nm. Addition of the 5pBT(30)Cy3 did not exhibit a further shift in the plasmon band; however, the absorption peak from Cy3 appeared around 550 nm after adding 5pBT(30)Cy3 to NS-A. A similar trend of peak plasmon wavelength change was observed from the functionalization NSs with avidin and 3pBT(30)Cy3. Optimizing Nanoassembly Reaction Conditions for Metal Enhanced Fluorescence Experiments. To achieve better fluorescence enhancement, the nanoassembly parameters were optimized. First, the NS’s concentration must be controlled because the NS’s plasmon and scattering spectra can interact with both the excitation light and the fluorescent light from the dye. Second, the ratio of biotin to avidin was important because when avidin was on the NS’s surface, it was not safe to assume that all four of avidin’s biotin-binding sites

Figure 6. Fluorescence intensity of Cy3 when continuously excited at 742 nm for 30 min. Blue stars and line are for Cy3 on gold NSs and red circles and line are for Cy3 in 0.1× PBS solution. Data provide evidence that gold NSs improve the photostability of Cy3.

Table 3. Average Fluorescence Enhancement Factor of Cy3 on Gold NSs [UV/Vis Optical Density is 0.2 ± 0.002 (N = 3)] under Different Reaction Conditionsa CNS

BT(30)Cy3

avidin in 1 mL NS (pmol)

molBT(30)Cy3/molavidin

(2.63 ± 0.08) × 109 particles/mL

3pBT(30)Cy3

80 80 80 40

2 3 2 3

5pBT(30)Cy3

enhancement factor 2.25 1.54 1.16 1.50

± ± ± ±

0.97 0.34 0.19 0.31

a

N was the number of replicate samples measured (N = 4). F

DOI: 10.1021/acs.jpcc.8b06609 J. Phys. Chem. C XXXX, XXX, XXX−XXX

Article

The Journal of Physical Chemistry C NSs and in 0.1× PBS overtime. In the two-photon excitation photostability study, the intensity of Cy3 on gold NSs remained constant over 30 min. In contrast, without the NSs, the intensity of Cy3 in 0.1× PBS reduced by 6.2 ± 0.3% (N = 3). Each replicate was an independent coating of the biotinylated DNA linker-Cy3 complex onto avidin-coated NSs that came from the same batch. From these studies, we found that the photostability of Cy3 improved after being coated on gold NS (p-value < 0.0001). Next, we examined Cy3’s photostability on gold NS when excited by a single-photon mechanism. Again, the photostability of Cy3 was better when coated on gold NSs than without them (see Figure S7). The intensity of Cy3 on gold NSs was reduced by (43.3 ± 0.2)% (N = 3) of its original intensity; the intensity of Cy3 in 0.1× PBS was reduced by (48.6 ± 0.4)% (N = 3) of its original intensity. More details for Cy3’s photostability with single-photon excitation are in section S5 of the Supporting Information. Collectively, these results show that gold NSs help to improve the photostability of Cy3.

■

AUTHOR INFORMATION

Corresponding Author

*E-mail: [email protected]. ORCID

Sean M. Burrows: 0000-0001-8296-0459 Notes

The authors declare no competing financial interest.

■

■

ACKNOWLEDGMENTS The authors acknowledge Oregon State University for the support of this research, especially from the Research Office General Research Grant (The Oregon State University Electron Microscopy Suite). The electron microscopy facility used to acquire TEM images is supported by the National Science Foundation via the Major Research Instrumentation (MRI) Program under grant no. 1040588. We also gratefully acknowledge the financial support for the acquisition of the TEM instrument from the Murdock Charitable Trust and the Oregon Nanoscience and Microtechnologies Institute (ONAMI). We would like to thank and extend our great appreciation to the Koley Group (Oregon State University) for the use of their UV/Vis spectrophotometer. In addition, we would like to thank the Harper Group (Oregon State University) for the use of their NanoSight and DLS instruments.

CONCLUSIONS The ability to achieve two-photon-induced MEF of Cy3 on gold NSs in a colloid was demonstrated. Enhancement factors ranged from 1.2 to 3.5, with many around 1.5 in all working conditions performed in our studies. Further evidence of the fluorescence enhancement came from the biotin displacement study. Finally, similar to nanobipyramids and other nanomaterials, the NSs were able to improve the photostability of a nearby fluorophore because of surface interactions between NSs and the fluorophores.19 These results are an important step toward opening opportunities to apply gold NS colloids into fluorescence-related techniques. The enhancement factors reported here must be improved before any major applications will be possible. This will require studies to elucidate if moving the dye closer to or farther from the surface will improve the enhancement. More details on the fluorescence enhancement factors with different number of thymine bases from 0 to 50 will be the subject of the follow-up work. In that work, we will further investigate whether the dye orientation with respect to the nanosurface has any effect on the fluorescence enhancement. In addition, the relationship between the NS’s plasmon spectrum and the fluorophore’s absorption and emission spectra will be illuminated to better understand the enhancement mechanism. Other future research should involve testing the hypothesis: if the plasmon absorbs ultrafast NIR, then the NS acts as a “donor” because the plasmon absorbs photons and sustains an enhanced twophoton electric field in the NIR and couples with the excitation and emission properties of the nearby “acceptor” fluorophores. Moreover, work from both a theoretical and empirical standpoint ought to be done to tease out the photophysical processes between NSs and dyes.

■

enhancement factors on gold nanostars), NanoSight measurements, plasmon-free control experiments for biotin competition study of Cy3 in: 0.1× PBS and Si nanospheres, optimization of nanoassembly working conditions (optimal nanostar concentration, optimal mole ratio of 3p and 5p BT30Cy3 to avidin, optimal mole ratio of avidin to nanostars, and summary of the optimal nanoassembly working conditions), and photostability of Cy3 on gold nanostars under single-photon excitation (PDF)

■

REFERENCES

(1) Lakowicz, J. R. Principles of Fluorescence Spectroscopy; Springer: USA, 2006. (2) Jans, H.; Huo, Q. Gold Nanoparticle-Enabled Biological and Chemical Detection and Analysis. Chem. Soc. Rev. 2012, 41, 2849− 2866. (3) Zhang, Y.; Dragan, A.; Geddes, C. D. Wavelength Dependence of Metal-Enhanced Fluorescence. J. Phys. Chem. C 2009, 113, 12095− 12100. (4) Hao, Q.; Qiu, T.; Chu, P. K. Surfaced-Enhanced Cellular Fluorescence Imaging. Prog. Surf. Sci. 2012, 87, 23−45. (5) Dong, J.; Zhang, Z.; Zheng, H.; Sun, M. Recent Progress on Plasmon-Enhanced Fluorescence. Nanophotonics 2015, 4, 472−490. (6) Lin, H.-H.; Chen, I.-C. Study of the Interaction between Gold Nanoparticles and Rose Bengal Fluorophores with Silica Spacers by Time-Resolved Fluorescence Spectroscopy. J. Phys. Chem. C 2015, 119, 26663−26671. (7) Chen, Y.; Munechika, K.; Ginger, D. S. Dependence of Fluorescence Intensity on the Spectral Overlap between Fluorophores and Plasmon Resonant Single Silver Nanoparticles. Nano Lett. 2007, 7, 690−696. (8) Tam, F.; Goodrich, G. P.; Johnson, B. R.; Halas, N. J. Plasmonic Enhancement of Molecular Fluorescence. Nano Lett. 2007, 7, 496− 501. (9) Buil, S.; Laverdant, J.; Berini, B.; Maso, P.; Hermier, J.-P.; Quélin, X. FDTD simulations of localization and enhancements on fractal plasmonics nanostructures. Opt. Express 2012, 20, 11968.

ASSOCIATED CONTENT

S Supporting Information *

The Supporting Information is available free of charge on the ACS Publications website at DOI: 10.1021/acs.jpcc.8b06609. Method for emission spectra data analysis (backgroundcorrected spectrum of Cy3 on gold nanostars, background corrected spectrum of Cy3 in 0.1× PBS or Si nanospheres, and determination of the fluorescence G

DOI: 10.1021/acs.jpcc.8b06609 J. Phys. Chem. C XXXX, XXX, XXX−XXX

Article

The Journal of Physical Chemistry C

Probes in Cells and Tissues. J. Am. Chem. Soc. 2012, 134, 12157− 12167. (31) Qiu, J.-P.; Liang, R.-F.; Peng, X.; Li, Y.-H.; Liu, L.-X.; Yin, J.; Qu, J.-L.; Niu, H.-B. Experimental Study on Multicolor Two-Photon Excited Fluorescence Microscopy. Acta Phys. Sin. 2015, 64, 048701. (32) Larkey, N. E.; Almlie, C. K.; Tran, V.; Egan, M.; Burrows, S. M. Detection of miRNA Using a Double-Strand Displacement Biosensor with a Self-Complementary Fluorescent Reporter. Anal. Chem. 2014, 86, 1853−1863. (33) Larkey, N. E.; Brucks, C. N.; Lansing, S. S.; Le, S. D.; Smith, N. M.; Tran, V.; Zhang, L.; Burrows, S. M. Molecular Structure and Thermodynamic Predictions to Create Highly Sensitive microRNA Biosensors. Anal. Chim. Acta 2016, 909, 109−120. (34) Larkey, N. E.; Zhang, L.; Lansing, S. S.; Tran, V.; Seewaldt, V. L.; Burrows, S. M. Förster resonance energy transfer to impart signalon and -off capabilities in a single microRNA biosensor. Analyst 2016, 141, 6239−6250. (35) Weissleder, R. A Clearer Vision for in vivo Imaging. Nat. Biotechnol. 2001, 19, 316−317. (36) Dimitrov, R. A.; Zuker, M. Prediction of Hybridization and Melting for Double-Stranded Nucleic Acids. Biophys. J. 2004, 87, 215−226. (37) Markham, N. R.; Zuker, M. DINAMelt Web Server for Nucleic Acid Melting Prediction. Nucleic Acids Res. 2005, 33, W577−W581. (38) Markham, N. R.; Zuker, M. UNAFold: Software for Nucleic Acid Folding and Hybridization. Methods in Molecular Biology; Springer, 2008; Vol. 453, pp 3−31. (39) Kumari, A.; Koyama, T.; Hatano, K.; Matsuoka, K. Synthetic Assembly of Novel Avidin-Biotin-GlcNAc (ABG) Complex as an Attractive Bio-Probe and Its Interaction with Wheat Germ Agglutinin (WGA). Bioorg. Chem. 2016, 68, 219−225. (40) Malicka, J.; Gryczynski, I.; Fang, J.; Kusba, J.; Lakowicz, J. R. Photostability of Cy3 and Cy5-Labeled DNA in the Presence of Metallic Silver Particles. J. Fluoresc. 2002, 12, 439−447. (41) Khoury, C. G.; Norton, S. J.; Vo-Dinh, T. Investigating the Plasmonics of a Dipole-Excited Silver Nanoshell: Mie Theory Versus Finite Element Method. Nanotechnology 2010, 21, 315203. (42) Khoury, C. G.; Norton, S. J.; Vo-Dinh, T. Plasmonics of 3-D Nanoshell Dimers Using Multipole Expansion and Finite Element Method. ACS Nano 2009, 3, 2776−2788.

(10) Bardhan, R.; Grady, N. K.; Cole, J. R.; Joshi, A.; Halas, N. J. Fluorescence Enhancement by Au Nanostructures: Nanoshells and Nanorods. ACS Nano 2009, 3, 744−752. (11) Shi, X.; Ji, Y.; Hou, S.; Liu, W.; Zhang, H.; Wen, T.; Yan, J.; Song, M.; Hu, Z.; Wu, X. Plasmon Enhancement Effect in Au Gold Nanorods@Cu2O Core-Shell Nanostructures and Their Use in Probing Defect States. Langmuir 2015, 31, 1537−1546. (12) Wu, D.; Xu, X.; Liu, X. Electric Field Enhancement in Bimetallic Gold and Silver Nanoshells. Solid State Commun. 2008, 148, 163−167. (13) Zhang, Y.; Mandeng, L. N.; Bondre, N.; Dragan, A.; Geddes, C. D. Metal-Enhanced Fluorescence from Silver−SiO2−Silver Nanoburger Structures. Langmuir 2010, 26, 12371−12376. (14) Malicka, J.; Gryczynski, I.; Gryczynski, Z.; Lakowicz, J. R. Effects of fluorophore-to-silver distance on the emission of cyaninedye-labeled oligonucleotides. Anal. Biochem. 2003, 315, 57−66. (15) Lakowicz, J. R.; Shen, Y.; D’Auria, S.; Malicka, J.; Fang, J.; Gryczynski, Z.; Gryczynski, I. Radiative Decay Engineering. Anal. Biochem. 2002, 301, 261−277. (16) Shi, F.; Jia, Z.; Lv, X.; Zhang, H.; Zhou, J. Enhancement of the R6G Fluorescence by Gold Nanoparticle Depositions in Porous Silicon Bragg Reflectors. Phys. Status Solidi A 2015, 212, 662−665. (17) Zhang, J.; Thurber, A.; Tenne, D. A.; Rasmussen, J. W.; Wingett, D.; Hanna, C.; Punnoose, A. Enhanced Dye Fluorescence in Novel Dye-ZnO Nanocomposites. Adv. Funct. Mater. 2010, 20, 4358−4363. (18) Ray, A.; Lee, Y.-E. K.; Kim, G.; Kopelman, R. Two-Photon Fluorescence Imaging Super-Enhanced by Multishell Nanophotonic Particles, with Application to Subcellular pH. Small 2012, 8, 2213− 2221. (19) Navarro, J. R. G.; Lerouge, F.; Micouin, G.; Cepraga, C.; Favier, A.; Charreyre, M. T.; Blanchard, N. P.; Lermé, J.; Chaput, F.; Focsan, M.; et al. Plasmonic Bipyramids for Fluorescence Enhancement and Protection against Photobleaching. Nanoscale 2014, 6, 5138−5145. (20) Hao, F.; Nehl, C. L.; Hafner, J. H.; Nordlander, P. Plasmon Resonances of a Gold Nanostar. Nano Lett. 2007, 7, 729−732. (21) Yuan, H.; Khoury, C. G.; Hwang, H.; Wilson, C. M.; Grant, G. A.; Vo-Dinh, T. Gold Nanostars: Surfactant-Free Synthesis, 3D Modelling, and Two-Photon Photoluminescence Imaging. Nanotechnology 2012, 23, 075102. (22) Zhu, S.; Cortie, M.; Blakey, I. Effect of Multimodal Plasmon Resonances on the Optical Properties of Five-Pointed Nanostars. Nanomater. Nanotechnol. 2015, 5, 22. (23) Tsoulos, T. V.; Han, L.; Weir, J.; Xin, H. L.; Fabris, L. A Closer Look at the Physical and Optical Properties of Gold Nanostars: an Experimental and Computational Study. Nanoscale 2017, 9, 3766− 3773. (24) Rodríguez-Oliveros, R.; Sánchez-Gil, J. A. Gold nanostars as thermoplasmonic nanoparticles for optical heating. Opt. Express 2012, 20, 621−626. (25) Yguerabide, J.; Yguerabide, E. E. Light-Scattering Submicroscopic Particles as Highly Fluorescent Analogs and Their Use as Tracer Labels in Clinical and Biological Applications. Anal. Biochem. 1998, 262, 137−156. (26) Navarro, J. R. G.; Liotta, A.; Faure, A.-C.; Lerouge, F.; Chaput, F.; Micouin, G.; Baldeck, P. L.; Parola, S. Tuning Dye-to-Particle Interactions toward Luminescent Gold Nanostars. Langmuir 2013, 29, 10915−10921. (27) Theodorou, I. G.; Jawad, Z. A. R.; Jiang, Q.; Aboagye, E. O.; Porter, A. E.; Ryan, M. P.; Xie, F. Gold Nanostar Substrates for MetalEnhanced Fluorescence through the First and Second Near-Infrared Windows. Chem. Mater. 2017, 29, 6916−6926. (28) Ramsey, J. D.; Zhou, L.; Almlie, C. K.; Lange, J. D.; Burrows, S. M. Achieving Plasmon Reproducibility from Surfactant Free Gold Nanostar Synthesis. New J. Chem. 2015, 39, 9098−9108. (29) Helmchen, F.; Denk, W. Deep tissue two-photon microscopy. Methods 2005, 2, 932−940. (30) Li, L.; Ge, J.; Wu, H.; Xu, Q.-H.; Yao, S. Q. Organelle-Specific Detection of Phosphatase Activities with Two-Photon Fluorogenic H

DOI: 10.1021/acs.jpcc.8b06609 J. Phys. Chem. C XXXX, XXX, XXX−XXX