SERS - ACS Publications - American Chemical Society

Letter

A Novel Liposome-Based Surface Enhanced Raman Spectroscopy (SERS) Substrate William Lum, Ian R. Bruzas, Zohre Gorunmez, Sarah A. Unser, Thomas L. Beck, and Laura B. Sagle J. Phys. Chem. Lett., Just Accepted Manuscript • Publication Date (Web): 23 May 2017 Downloaded from http://pubs.acs.org on May 27, 2017

Just Accepted “Just Accepted” manuscripts have been peer-reviewed and accepted for publication. They are posted online prior to technical editing, formatting for publication and author proofing. The American Chemical Society provides “Just Accepted” as a free service to the research community to expedite the dissemination of scientific material as soon as possible after acceptance. “Just Accepted” manuscripts appear in full in PDF format accompanied by an HTML abstract. “Just Accepted” manuscripts have been fully peer reviewed, but should not be considered the official version of record. They are accessible to all readers and citable by the Digital Object Identifier (DOI®). “Just Accepted” is an optional service offered to authors. Therefore, the “Just Accepted” Web site may not include all articles that will be published in the journal. After a manuscript is technically edited and formatted, it will be removed from the “Just Accepted” Web site and published as an ASAP article. Note that technical editing may introduce minor changes to the manuscript text and/or graphics which could affect content, and all legal disclaimers and ethical guidelines that apply to the journal pertain. ACS cannot be held responsible for errors or consequences arising from the use of information contained in these “Just Accepted” manuscripts.

The Journal of Physical Chemistry Letters is published by the American Chemical Society. 1155 Sixteenth Street N.W., Washington, DC 20036 Published by American Chemical Society. Copyright © American Chemical Society. However, no copyright claim is made to original U.S. Government works, or works produced by employees of any Commonwealth realm Crown government in the course of their duties.

Page 1 of 24

1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60

The Journal of Physical Chemistry Letters

A Novel Liposome-Based Surface Enhanced Raman Spectroscopy (SERS) Substrate William Lum, Ian Bruzas, Zohre Gorunmez, Sarah Unser, Thomas Beck and Laura Sagle* Department of Chemistry, College of Arts and Sciences, University of Cincinnati, 301 West Clifton Court, Cincinnati OH 45221-0172 *Corresponding author Tel: +1 513 556 1034; Fax: +1 513 556 9239. E-mail: [email protected]

1 ACS Paragon Plus Environment


1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60

Abstract Although great strides have been made in recent years towards making highly enhancing surface enhanced Raman spectroscopy (SERS) substrates, the biological compatibility of such substrates remains a crucial problem. To address this issue, liposome-based SERS substrates have been constructed in which the biological probe molecule is encapsulated inside the aqueous liposome compartment, and metallic elements are assembled using the liposome as a scaffold. Thus, the probe molecule is not in contact with the metallic surfaces. Herein we report our initial characterization of these novel nanoparticle-on-mirror substrates, both experimentally and theoretically, using finite-difference time-domain (FDTD) calculations. The substrates are shown to be structurally stable to laser irradiation, the liposome compartment does not rise above 45 °C, and they exhibit an analytical enhancement factor of 8x106 for crystal violet encapsulated in 38 liposomes sandwiched between a 40 nm planar gold mirror and 80 nm gold colloid.


Page 2 of 24

Page 3 of 24

1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60


Surface enhanced Raman spectroscopy (SERS) offers the ability to probe single molecules with unprecedented detail for processes ranging from biological sensing1 and chemical transformation2 to optical waveguides3 and electronics.4 For biophysical studies in particular, SERS has enormous potential due to exquisite sensitivity, intrinsically fast timescales (1 million times faster than fluorescence)5,6 and narrow peak bandwidth which enables multiplexed labeling for site-specific information.7,8 Unfortunately, SERS signal is highest at the junction between metallic surfaces9, thus, the status quo for measuring biological species using SERS is to sandwich the probe molecule between aggregated silver colloids10,11 which often leads to denaturation and inactivation.12,13 Many recent biophysical studies have been carried out in this manner and have allowed dynamic insight into protein chromophore charge conversion14, conformational equilibrium15, and ligand binding.16 However, questions still remain concerning the biological significance of these measurements, since the reported SERS spectra were often quite different than the Raman spectra of the same proteins in solution.17 A common strategy towards improving biocompatibility of SERS substrates is to limit the interactions between the metallic substrate and the biological probe molecule. Some recent examples include silver colloids coated in iodide18, a flow-through cell in which the probe and substrate are added together for a brief time19,20, and the measurement of freely diffusing biological molecules using tip-enhanced Raman spectroscopy (TERS).21 While these studies show evidence of improved biocompatibility, the SERS enhancement and measurement capabilities for weaker interactions and longer timescales is often compromised. Thus, SERS substrates which



1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60

possess high enhancement, improved reproducibility, and at the same time allow for the measurement of biological species in a cell-like environment would be ideal. A possible solution is to use liposomes for probe molecule encapsulation, and build metallic, SERS enhancing elements around the liposome scaffold. Liposomes have been commonly used as a cellular mimic in single molecule fluorescence studies22,23 and have been shown to retain biological and chemical activity.24 In addition, liposomes create compartmentalization with little leakage25,26 and have long term stability to laser irradiation.22,25 Since lipids present in cellular membranes can be readily incorporated into liposomes, they have also been shown to reproduce specific biological interactions present in cells.27 Moreover, liposomes offer a wide range of functional groups on their surface, which have been utilized in the drug delivery community to interface lipids with polymers and metal nanoparticles.28 Some examples include gold-coated liposomes for photothermal drug delivery and imaging applications.29,30 However, the SERS properties of such liposome-metal composites remains completely unexplored.


Page 4 of 24

Page 5 of 24

1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60


Herein, we have constructed liposome-based, highly enhancing SERS substrates and carried out initial characterization of these novel substrates. These liposome-based constructs are shown to be stable to laser irradiation, not susceptible to plasmonic heating above 42°C, exhibit high SERS enhancement of 8 x 106, and notably improved reproducibility over traditional aggregated structures. Since the probe molecules are encapsulated inside the liposome, freely diffusing in solution, and not in direct contact with metallic surfaces, these substrates show great promise towards improved

Scheme 1: Flow chart of substrate fabrication. Probe encapsulated liposomes are made from Egg-PC, DPPC, and Biotin PE lipids dried into a cake and rehydrated with probe containing buffer. Extrusion through differently sized membranes create small unilamellar liposomes containing the buffer with probe. Next, size exclusion chromatograph separates free probe from liposomes containing the probe and the fresh, biotinylated liposomes are added to gold covered glass coverslips functionalized with streptavidin. Last, streptavidin functionalized gold colloids are added to the biotinylated liposome-covered gold surface to complete the assembly.



1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60

biocompatibility, in which the probe molecules behave as if they reside in a cellular environment. The substrates were constructed using a self-assembly process described in Scheme 1 and the Supporting Information. Briefly, a self-assembled monolayer of biotin-PEG-thiol is created on a glass coverslip containing 40 nm of gold, followed by addition of Streptavidin protein. Next, unilamellar liposomes composed of a 1:2:0.03 ratio of phosphatidylcholine (PC):dipalmitoylphosphatidylcholine (DPPC):Biotincapped phosphatidylethanolamine (PE) were added to the streptavidin-coated surface at an optimal concentration for single construct studies. Last, 80 nm streptavidin-coated gold colloids are then added to conjugate with the biotinylated lipids and complete the assembly. The components of the liposome construct were characterized and optimized both separately and together. The size of the liposome component was measured using dynamic light scattering and transmission electron microscopy (TEM), see Figure S1 in the Supporting Information section. In addition, the size and shape of the gold colloids was measured using TEM, and confirmed to be ~80 nm in diameter. Systematic studies verified that the observed SERS signal was due to individual constructs containing a single gold colloid. First, the density of liposomes on the gold surface was measured through fluorescence microscopy of liposomes containing 5 mol% fluorescently-labeled NBD-PE lipids, see Figure S2a of the Supporting Information. Next, the density of gold colloids bound to lipids were measured by drying out the constructs and using scanning electron microscopy (SEM), see Figure S2b in the Supporting Information section. Lastly, correlated studies measuring the localized surface plasmon resonance (LSPR) and SERS spectrum of the same individual


Page 6 of 24

Page 7 of 24

1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60


construct were carried out using a standard SERS microscopy setup described in detail in the Supporting Information section. The correlated LSPR/SERS measurements were followed by SEM measurements of the same spots after drying, see Figure 1. Such measurements confirmed that SERS was indeed carried out on individual constructs containing individual gold colloids. The SERS spectra shown in Figure 1 were confirmed to be that of crystal violet, with the two strongest peaks at 1174 cm-1 and 1382 cm-1 assigned as a symmetric CCC stretch/bend combination and a CH bending mode of CV respectively.31,32 Optimization of the constructs involved investigating the role of liposome size in SERS enhancement and was aided by finite-difference time-domain (FDTD) calculations.33 Calculated SERS enhancement values were generated by taking the maximum |E/E0|4 value only for grid points inside the liposome component. Indeed, both experimental measurements and FDTD calculations confirm the smaller liposome does exhibit significantly higher SERS enhancement, and greater coupling with the planar gold surface, see Figure S3 in the Supporting Information section. This data implies that the observed SERS signal is not simply a result of dye molecules residing close to either the surface of the gold colloid or gold mirror.



1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60

Figure 1: Correlated LSPR/SERS/SEM measurements to verify single constructs with individual gold colloids were being measured. (Top) SERS spectra of 5 mM crystal violet in 38 nm gold colloid-liposome substrates illuminated with a 1.2 mW, 633 nm laser. The insets in these graphs are the LSPR spectra obtained under dark field illumination of the same constructs. (Bottom) SEM of constructs consisting of single gold colloids correlated with the LSPR and SERS spectra shown above. To investigate the stability of these substrates to laser irradiation, experiments were carried out using the self-quenching properties of toluidine blue dye. As shown in Figure 2a, when the liposomes are intact, the dye itself quenches, reducing the fluorescence signal. Upon disruption of the liposome and release of the dye, the


Page 8 of 24

Page 9 of 24

1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60


fluorescence is no longer self-quenched and an increase in intensity is observed. When illuminating the liposome constructs with the same laser intensity and time required for SERS measurements, no fluorescence intensity changes were observed, indicating the liposomes are not bursting during the SERS measurements. Control experiments were carried out by increasing the laser power and adding detergent to burst the liposomes and indeed, an increase in fluorescence intensity was observed in both cases, see Figure 2a.

Figure 2: Measuring liposome stability and temperature changes upon laser irradiation. a) The fluorescence intensity of the 650 nm toluidine blue o emission peak with 1.2 mW 633nm excitation was monitored over time. No increase in intensity suggest liposomes stay intact throughout illumination. Triton X was added to trigger release of self-quenched dye, resulting in increase in peak intensity. At a higher power (3.2 mW), an increase in intensity was also observed suggesting liposome dye leakage. At a lower power (0.4 mW) the intensity is steady, as expected. b) Correlation of 570 nm sulforhodamine g (SRG) emission peak intensity to the calibration curve of KV6-ELP and SRG. Inset compares the raw spectra of two calibration spectra to the spectrum under SERS measurement conditions. Measurements indicate the substrate only rises to ~42 °C when SERS 9 measurements are taken. ACS Paragon Plus Environment


1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60

In order to carry out measurements with biological probe molecules in the liposome component of the substrates, the temperature should not rise above 60°C, since most enzymes are completely denatured at this temperature.34–38 Experiments were carried out to ensure that the temperature inside the liposomes did not rise above 60°C through the co-encapsulation of an elastin-like polypeptide39 (ELP), with a lower critical solution temperature (LCST) of ~45 °C, and oppositely charged fluorescence dye, sulforhodamine g (SRG). Upon raising the temperature above the LCST, the fluorescence signal shows a marked decrease in intensity due to the collapse of the ELP-dye complex causing self-quenching, see Figure 2b. It is verified that this change is due to the collapse of the ELP-dye complex, since liposomes containing no ELP and just dye do not show similar behavior, see Figure S4 in Supporting Information. Indeed, when illuminating single liposome constructs containing the dye-ELP mixture with laser irradiation of the same intensity and time required for SERS measurements, only a small decrease in fluorescence was observed. These measurements indicate that the temperature inside the liposomes remains ~42 °C, and thus, the constructs should render biological probe molecules fully functional. The SERS enhancement of the liposome-based substrates were first evaluated experimentally. Analytical SERS enhancement factors were calculated by measuring the laser spot size in the x, y, and z directions to be 1.6um x 0.8um x 16um respectively using the 520 cm-1 band of a thin (100 µm) silicon wafer. Next, it was assumed that single liposome constructs were being measured, as verified above, and that the dye encapsulation efficiency was 50%, based on previous studies with crystal violet dye in liposomes of similar composition.40,41 Last, the normal Raman spectrum of 5 mM


Page 10 of 24

Page 11 of 24

1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60


crystal violet in aqueous solution was measured under the same conditions for comparison. The SERS enhancement factors for the three liposome constructs made herein are 8x106, 3x105, 3x104 for the 38 nm, 65 nm and 104 nm liposome sizes respectively. These values are in the right range for a weakly coupled nanoparticle-onmirror substrate, roughly a factor of 103 less than strongly coupled systems.42 The strongly coupled systems in which the nanoparticle is less than 10 nm from the gold mirror exhibit SERS enhancements in the range of108 to 1010.43–47 One interesting thing to note is that the SERS spectra obtained herein do not contain peaks from the lipid or protein components of the constructs, despite the fact that these components reside closer to the metallic surfaces. This observation is explained by the fact that Raman cross sections of crystal violet are ~10,000 times higher than that of the protein or lipid components, thus only SERS from the crystal violet probe is observed.48,49 Measurements of the same constructs containing liposomes without crystal violet show SERS signal from the streptavidin and possibly lipid components when considerably longer acquisition times are used, see Figure S5. Thus, the liposome-based constructs are stable enough to allow for the long-time measurement of biological species. In order to verify that our measurements are in fact measuring dye freely diffusing inside the liposome, a control experiment was carried out to assess the possibility that dye could be bound to the membrane components or leak out of the liposome, coming into direct contact with either the gold surface or gold colloid. These experiments were carried out by forming unilamellar liposomes which were then soaked in a dye solution similar in concentration to that used for dye encapsulation, see Figure S6 in the Supporting Information. Similar to constructs prepared with encapsulated dye,



1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60

the liposome-dye solution was then passed through a size exclusion column to remove excess, unbound dye, and assembled onto the gold surface. Although some SERS intensity was observed in these constructs, it was considerably less than that observed when the dye is encapsulated within the liposome component. Thus, a large part of the dye molecules generating our measured SERS signal appear to reside inside the liposome component of our constructs. Another control experiment was carried out assessing the ability of the liposomes to flatten themselves, trapping dye in a small gap between the gold colloid and planar gold surface. An investigation of possible liposome collapse was carried out by varying the distance between the gold colloid and planar gold surface using polymers and measuring the LSPR spectra of the ‘nanoparticle on mirror’ assemblies, see Figure S7. Through such systematic studies, it has been shown that one can directly calculate the distance between the gold colloid and gold surface from the LSPR spectra.42 Indeed, results indicate the distances between the gold colloid and the 40 nm planar gold surface is calculated to be 27 ± 10 nm when sampling 30 individual constructs. Thus, the 38 nm liposomes are not noticeably flattened or reduced in size to create a smaller, more SERS-amenable gap. Indeed, these substrates appear to be unique in generating high SERS enhancement for probe molecules that are not in contact with a metallic surface and further, reside at least 8 nm from any metallic surface. In order to address the mechanism of SERS enhancement, FDTD calculations were carried out on the colloid-liposome constructs depicted in Figure 3a using the FDTD++ open source software.50 The Drude plus two-pole Lorentz Oscillators Model was used to fit the Johnson and Christy data51 to model the dispersive properties of gold


Page 12 of 24

Page 13 of 24

1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60


in the appropriate wavelength region, and a grid size of 1.55 nm was used.52,53 Since our measurements used unpolarized light in an epi configuration, in calculating the LSPR spectra and SERS enhancements, all possible linear polarizations of light were evaluated. However, the TM polarization, which is normal to our gold surfaces, as expected, exhibit the largest absorption of the metal and dominated the field enhancements.54–56 Thus, only this configuration is shown in the electric field and charge distribution diagrams, see Figure 3c (and Figure S9 in the Supporting Information for these diagrams for different polarizations). Experimental scattering spectra (an average of 10 individual constructs) and calculated scattering spectra for the TM polarization show good agreement, with a large resonance ~570 nm, see Figure 3b. This is mainly attributed to the dipolar plasmon mode of the colloid coupled to the gold surface. Diagrams depicting the E-fields in and around the constructs are shown in Figure 3c. Interestingly, these diagrams show a substantial amount of field inside the liposome itself, with the main ‘hot spot’ residing close to the gold colloid on the side that faces the gold mirror. SERS enhancements (|E/E0|4) were calculated for the constructs by averaging the different polarizations and gold colloid binding positions (90 and 45 degrees) and only evaluating the grid points residing inside the liposome component. These numbers reflect an electromagnetic SERS enhancement, |E/E0|4 avg being 8.8x106, which is in very good agreement with experimental results (for a more in depth description of the SERS enhancement calculation, please see Supporting Information). It should be noted that CV dye has minimal fluorescence background at 633 nm and previous studies have taken the analytical experimental enhancement factors calculated



1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60

in such manner, where the normal Raman of the dye is divided, as the nonresonant enhancement factor.49,57 The E-field diagrams highlight the fact that the high SERS enhancements observed here may be a result of the liposomes trapping probe molecules between the surface of the colloid and mirror, i.e. the main ‘hot spot’ shown in such diagrams. This is notably different than some current nanoparticle on mirror SERS studies in which probe molecules are either placed directly on the gold mirror or on the outer surface of the gold colloid.44–46,58 Alternatively, other studies have attached probe molecules to the outside of the gold colloid prior to nanoparticle-on-mirror assembly, but, even though the nanoparticles were within 3 nm from the mirrors, the


Page 14 of 24

Page 15 of 24

1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60


Figure 3. FDTD calculations depicting how different polarized light interacts with the gold colloid-liposome constructs. a) Schematic of the constructs showing the dimensions of each component. b) Calculated scattering spectra of all polarizations overlaid with the average experimental scattering spectra of 10 individual constructs. c) Cross sectional E-field diagrams (|E/E0|) for constructs with different nanoparticle binding orientations. Black arrows point to the liposomes. Diagrams depicting the cross sectional surface charge distributions of every element in the constructs, except for the underlying titanium and silica layers. SERS enhancement factors reported were only in the 106 range. This is most likely due to normalization with respect to all dye molecules residing on the entire surface of the colloid, not just those residing in the ‘hot spots’.46,59,60 Next to the electric field diagrams in Figure 3, diagrams displaying charge on all surfaces in the constructs are depicted. Interestingly, these diagrams show that not only is the gold colloid and gold surface displaying charge, but the liposomes themselves also contain charged surfaces. This is most likely due to the high refractive index of the lipids, and may allow for an optical waveguide effect which confines high electromagnetic fields in the liposome component, much like a higher refractive index 15 ACS Paragon Plus Environment


1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60

silicon interface, consequently leading to high SERS enhancement.61,62 Additionally, the high refractive index of the liposome ‘spacer’ between the gold mirror and colloid may also create interference effects at these interfaces which have been shown to increase fields and SERS enhancement.63,64 Lastly, even though the dipole modes of both the gold colloid and planar gold surface are excited at 633 nm, the liposome component of the constructs (where the probe molecule resides) shows a clear higher order antibonding dipole mode. Excitation of such higher energy modes and their coupling with dipole modes has also been shown to increase SERS enhancements in core-shell and nanoraspberry-type structures.65–68. Further studies are currently underway to elucidate the role of the liposome in the observed SERS enhancement. As shown in Figure 3, the calculated E-field diagrams indicate the ‘hot spots’ inside the liposome component are fairly delocalized when compared to more typical SERS substrates involving aggregated structures.69 Consequently, this could lead to more reproducible SERS signal. Measurements were carried out to investigate the SERS signal reproducibility of the novel liposome-based substrates and the results are shown in Figure 4. For these substrates, dye was encapsulated into the liposome component, rather than adsorbing dye to the outside surface of a metallic structure. Measurements were carried out on at least 100 individual structures and the standard deviation calculated from fitting the data to a Gaussian distribution. The relative standard deviation for the 38 nm colloid-liposome constructs was calculated as 28%, which is considerably lower than that reported for aggregated silver colloids and single gold nanostars (see Figure S8 in the Supporting Information). Despite the low standard deviation of the substrates, it is still higher than modern SERS substrates such as film


Page 16 of 24

Page 17 of 24

1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60


over nanospheres, which can be as low as 10%.70 In our case, the main sources of irreproducibility are most likely due to variations in liposome size, as evidenced in Figure S3, and colloid placement on the liposome surface. Indeed, the FDTD calculations reveal that changing the placement of the colloid from 90 ° to 45 ° can result in a change in the SERS enhancement factors of ten times, see Figure S9. Because these changes are so significant, it is likely that only binding orientations close to 90 ° are being selected out in measuring single constructs, since the SERS signal from other orientations would be difficult to measure.

Figure 4: Reproducibility measurements of 100 particles plotting the normalized 1175 cm-1 mode intensity of 5 mM crystal violet encapsulated in 38nm gold colloidliposome substrates. Peak intensities were normalized by dividing by the maximum value for each dataset. The relative standard deviation was calculated from a Gaussian fit of the data and found to be 28% for the gold colloid-liposome constructs. In conclusion, we report the construction of a novel SERS substrate and its initial characterization. The results indicate this substrate exhibits high SERS enhancement of 8 x 106, improved reproducibility over typical SERS substrates such as aggregated silver colloids, and the potential to exhibit greatly improved biocompatibility. Interestingly, unlike the SERS mechanism observed in many systems where a SERS 17 ACS Paragon Plus Environment


1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60

‘hot spot’ is created through a small gap between metallic particles, the SERS ‘hot spots’ for this system falls in the weakly coupled nanoparticle on mirror regime, are more delocalized, and exhibit long range decay. Future experimental and computational studies will involve the elucidation of the liposome role in electromagnetic field enhancements. In addition, further optimization of these substrates and encapsulation of biological molecules in the liposome component are currently underway.

Supporting Information Available: Additional information on experimental and computational materials and methods, instrumentation, and characterization are available free of charge via the Internet at http://pubs.acs.org.

Notes The authors declare no competing financial interest.

Acknowledgments:

This work was supported by University of Cincinnati start-up

funds. TLB acknowledges support from NSF grant CHE-1565632. The authors also gratefully acknowledge Prof. Jeff McMahon for helpful discussions and guidance on the FDTD calculations.

References 18 ACS Paragon Plus Environment

Page 18 of 24

Page 19 of 24

1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60


(1)

Kneipp, J.; Kneipp, H.; Kneipp, K. SERS--a Single-Molecule and Nanoscale Tool for Bioanalytics. Chem. Soc. Rev. 2008, 37, 1052–1060.

(2)

Xie, W.; Walkenfort, B.; Schlücker, S. Label-Free SERS Monitoring of Chemical Reactions Catalyzed by Small Gold Nanoparticles Using 3D Plasmonic Superstructures. J. Am. Chem. Soc. 2013, 135, 1657–1660.

(3)

Gu, Y.; Xu, S.; Li, H.; Wang, S.; Cong, M.; Lombardi, J. R.; Xu, W. Waveguide-Enhanced Surface Plasmons for Ultrasensitive SERS Detection. J. Phys. Chem. Lett. 2013, 4, 3153–3157.

(4)

Matsushita, R.; Kiguchi, M. Surface Enhanced Raman Scattering of a Single Molecular Junction. Phys. Chem. Chem. Phys. 2015, 1.

(5)

Singhal, K.; Kalkan, A. K. Surface-Enhanced Raman Scattering Captures Conformational Changes of Single Photoactive Yellow Protein Molecules under Photoexcitation. J. Am. Chem. Soc. 2010, 132, 429–431.

(6)

Nicholas D. Spencer, J. H. M. Encyclopedia of Chemical Physics and Physical Chemistry: Fundamentals; IOP Publishing Ltd.: London, U.K.; 2001.

(7)

Chen, Z.; Tabakman, S. M.; Goodwin, A. P.; Kattah, M. G.; Daranciang, D.; Wang, X.; Zhang, G.; Li, X.; Liu, Z.; Utz, P. J.; et al. Protein Microarrays with Carbon Nanotubes as Multicolor Raman Labels. Nat. Biotechnol. 2008, 26, 1285–1292.

(8)

Cao, Y. C.; Jin, R.; Mirkin, C. a. Nanoparticles with Raman Spectroscopic Fingerprints for DNA and RNA Detection. Science. 2002, 297, 1536–1540.

(9)

Michaels, A. M.; Brus, L. Ag Nanocrystal Junctions as the Site for Surface-Enhanced Raman Scattering of Single Rhodamine 6G Molecules. J. Phys. Chem. B 2000, 104, 11965–11971.

(10)

Delfino, I.; Bizzarri, A. R.; Cannistraro, S. Single-Molecule Detection of Yeast Cytochrome c by Surface-Enhanced Raman Spectroscopy. Biophys. Chem. 2005, 113, 41–51.

(11)

Habuchi, S.; Cotlet, M.; Gronheid, R.; Dirix, G.; Michiels, J.; Vanderleyden, J.; De Schryver, F. C.; Hofkens, J. Single-Molecule Surface Enhanced Resonance Raman Spectroscopy of the Enhanced Green Fluorescent Protein. J. Am. Chem. Soc. 2003, 125, 8446–8447.

(12)

Banerjee, V.; Das, K. P. Structure and Functional Properties of a Multimeric Protein alphaACrystallin Adsorbed on Silver Nanoparticle Surface. Langmuir 2014, 30, 4775–4783.

(13)

Hegyi, H.; Gerstein, M. The Relationship between Protein Structure and Function: A Comprehensive Survey with Application to the Yeast Genome. J. Mol. Biol. 1999, 288, 147–164.

(14)

Habuchi, S.; Ando, R.; Dedecker, P.; Verheijen, W.; Mizuno, H.; Miyawaki, A.; Hofkens, J. Reversible Single-Molecule Photoswitching in the GFP-like Fluorescent Protein Dronpa. Proc. Natl. Acad. Sci. USA 2005, 102, 9511–9516.

(15)

Brazhe, N. A.; Abdali, S.; Brazhe, A. R.; Luneva, O. G.; Bryzgalova, N. Y.; Parshina, E. Y.; Sosnovtseva, O. V.; Maksimov, G. V. New Insight into Erythrocyte through in Vivo SurfaceEnhanced Raman Spectroscopy. Biophys. J. 2009, 97, 3206–3214.

(16)

Costas, C.; López-Puente, V.; Bodelón, G.; González-Bello, C.; Pérez-Juste, J.; Pastoriza-Santos, I.; Liz-Marzán, L. M. Using Surface Enhanced Raman Scattering to Analyze the Interactions of Protein Receptors with Bacterial Quorum Sensing Modulators. ACS Nano 2015, 9, 5567–5576. 19 ACS Paragon Plus Environment


1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60

(17)

Otto, A. What Is Observed in Single Molecule SERS, and Why? J. Raman Spectrosc. 2002, 33, 593– 598.

(18)

Xu, L. J.; Zong, C.; Zheng, X. S.; Hu, P.; Feng, J. M.; Ren, B. Label-Free Detection of Native Proteins by Surface-Enhanced Raman Spectroscopy Using Iodide-Modified Nanoparticles. Anal. Chem. 2014, 86, 2238–2245.

(19)

Feng, M.; Tachikawa, H. Surface-Enhanced Resonance Raman Spectroscopic Characterization of the Protein Native Structure. J. Am. Chem. Soc. 2008, 130, 7443–7448.

(20)

Bailey, M. R.; Pentecost, A. M.; Selimovic, A.; Martin, R. S.; Schultz, Z. D. Sheath-Flow Microfluidic Approach for Combined Surface Enhanced Raman Scattering and Electrochemical Detection. Anal. Chem. 2015, 87, 4347–4355.

(21)

Cowcher, D. P.; Deckert-Gaudig, T.; Brewster, V. L.; Ashton, L.; Deckert, V.; Goodacre, R. Detection of Protein Glycosylation Using Tip-Enhanced Raman Scattering. Anal. Chem. 2016, 88, 2105–2112.

(22)

Pirchi, M.; Ziv, G.; Riven, I.; Cohen, S. S.; Zohar, N.; Barak, Y.; Haran, G. Single-Molecule Fluorescence Spectroscopy Maps the Folding Landscape of a Large Protein. Nat. Commun. 2011, 2, 493.

(23)

Oberholzer, T.; Luisi, P. L. The Use of Liposomes for Constructing Cell Models. J. Biol. Phys. 2002, 28, 733–744.

(24)

Debs, R. J.; Düzgüneş, N.; Brunette, E. N.; Fendly, B.; Patton, J.; Philip, R. Liposome-Associated Tumor Necrosis Factor Retains Bioactivity in the Presence of Neutralizing Anti-Tumor Necrosis Factor Antibodies. J. Immunol. 1989, 143, 1192–1197.

(25)

Boukobza, E.; Sonnenfeld, A.; Haran, G. Immobilization in Surface-Tethered Lipid Vesicles as a New Tool for Single Biomolecule Spectroscopy. J. Phys. Chem. B 2001, 105, 12165–12170.

(26)

Tokarz, M.; Akerman, B.; Olofsson, J.; Joanny, J.-F.; Dommersnes, P.; Orwar, O. Single-File Electrophoretic Transport and Counting of Individual DNA Molecules in Surfactant Nanotubes. Proc. Natl. Acad. Sci. U. S. A. 2005, 102, 9127–9132.

(27)

Chakrabarti, A. C.; Breaker, R. R.; Joyce, G. F.; Deamer, D. W. Production of RNA by a Polymerase Protein Encapsulated within Phospholipid Vesicles. J. Mol. Evol. 1994, 39, 555–559.

(28)

Al-Jamal, W. T.; Kostarelos, K. Liposome-Nanoparticle Hybrids for Multimodal Diagnostic and Therapeutic Applications. Nanomedicine 2007, 2, 85–98.

(29)

Troutman, T. S.; Barton, J. K.; Romanowski, M. Biodegradable Plasmon Resonant Nanoshells. Adv. Mater. 2008, 20, 2604–2608.

(30)

Jin, Y.; Gao, X. Spectrally Tunable Leakage-Free Gold Nanocontainers. J. Am. Chem. Soc. 2009, 131, 17774–17776.

(31)

Camamares, M. V.; Chenal, C.; Birke, R. L.; Lombardi, J. R. DFT, SERS, and Single-Molecule SERS of Crystal Violet. J. Phys. Chem. C 2008, 112, 20295–20300.

(32)

Angeloni, L.; Smulevich, G.; Marzocchi, M. P. Resonance Raman Spectrum of Crystal Violet. J. Raman Spectrosc. 1979, 8, 305–310.


Page 20 of 24

Page 21 of 24

1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60


(33)

Taflove, A.; Hagness, S. C.; Piket-May, M. Computational Electromagnetics: The Finite-Difference Time-Domain Method; Artech House: Norwood, MA, 2005.

(34)

Hill, C. G.; Root, T. W. An Introduction to Chemical Engineering Kinetics & Reactor Design; John Wiley & Sons: New York; 2014; 232–233.

(35)

Purich, D. L. Enzyme Kinetics: Catalysis & Control; Elsevier: London, U.K.; 2010; 425–429.

(36)

Johansen, P.; Senti, G.; Martinez Gomez, J. M.; Wuthrich, B.; Bot, A.; Kundig, T. M. Heat Denaturation, a Simple Method to Improve the Immunotherapeutic Potential of Allergens. Eur. J. Immunol. 2005, 35, 3591–3598.

(37)

Pohl, F. M.; Thomae, R.; Karst, A. Temperature Dependence of the Activity of DNA-Modifying Enzymes: Endonucleases and DNA Ligase. Eur. J. Biochem. 1982, 123, 141–152.

(38)

Pereira, E. B.; De Castro, H. F.; De Moraes, F. F.; Zanin, G. M. Kinetic Studies of Lipase from Candida Rugosa: A Comparative Study between Free and Immobilized Enzyme onto Porous Chitosan Beads. Appl. Biochem. Biotechnol. 2001, 91–93, 739–752.

(39)

MacEwan, S. R.; Chilkoti, A. Elastin-like Polypeptides: Biomedical Applications of Tunable Biopolymers. Biopolymers 2010, 94, 60–77.

(40)

Koneracká, M.; Kopčanský, P.; Sosa, P.; Bageľová, J.; Timko, M. Interliposomal Transfer of Crystal Violet Dye from DPPC Liposomes to Magnetoliposomes. J. Magn. Magn. Mater. 2005, 293, 271– 276.

(41)

Boualem, K.; Subirade, M.; Desjardins, Y.; Saucier, L. Development of an Encapsulation System for the Protection and Controlled Release of Antimicrobial Nisin at Meat Cooking Temperature. J. Food Res. 2013, 2, 36–45.

(42)

Mock, J. J.; Hill, R. T.; Degiron, A.; Zauscher, S.; Chilkoti, A.; Smith, D. R. Distance-Dependent Plasmon Resonant Coupling between a Gold Nanoparticle and Gold Film. Nano Lett. 2008, 8, 2245–2252.

(43)

Huang, S.; Ming, T.; Lin, Y.; Ling, X.; Ruan, Q.; Palacios, T.; Wang, J.; Dresselhaus, M.; Kong, J. Ultrasmall Mode Volumes in Plasmonic Cavities of Nanoparticle-On-Mirror Structures. Small 2016, 12, 5190–5199.

(44)

Lombardi, A.; Demetriadou, A.; Weller, L.; Andrae, P.; Benz, F.; Chikkaraddy, R.; Aizpurua, J.; Baumberg, J. J. Anomalous Spectral Shift of Near- A Nd Far-Field Plasmonic Resonances in Nanogaps. ACS Photonics 2016, 3, 471–477.

(45)

Rodriguez-Lorenzo, L.; aLvarez-Puebla, R. A.; Pastoriza-Santos, I.; Mazzucco, S.; Stephan, O.; Kociak, M.; Liz-Marzan, L. M.; De Abajo, F. J. G. Zeptomol Detection through Controlled Ultrasensitive Surface-Enhanced Raman Scattering. J. Am. Chem. Soc. 2009, 131, 4616–4618.

(46)

Mubeen, S.; Zhang, S.; Kim, N.; Lee, S.; Krämer, S.; Xu, H.; Moskovits, M. Plasmonic Properties of Gold Nanoparticles Separated from a Gold Mirror by an Ultrathin Oxide. Nano Lett. 2012, 12, 2088–2094.

(47)

Li, A.; Isaacs, S.; Abdulhalim, I.; Li, S. Ultrahigh Enhancement of Electromagnetic Fields by Exciting Localized with Extended Surface Plasmons. J. Phys. Chem. C 2015, 119, 19382–19389.

(48)

Manoharan, R.; Baraga, J. J.; Feld, M. S.; Rava, R. P. Quantitative Histochemical Analysis of Human 21 ACS Paragon Plus Environment


1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60

Artery Using Raman Spectroscopy. J. Photochem. Photobiol. B Biol. 1992, 16, 211–233. (49)

Le Ru, E. C.; Blackie, E.; Meyer, M.; Etchegoint, P. G. Surface Enhanced Raman Scattering Enhancement Factors: A Comprehensive Study. J. Phys. Chem. C 2007, 111, 13794–13803.

(50)

McMahon, J.M. FDTD++ 1.7; Algorithms in Motion LLC. 2016..

(51)

Johnson, P. B.; Christy, R. W. Optical Constants of the Noble Metals. Phys. Rev. B 1972, 6, 4370– 4379.

(52)

McMahon, J. M.; Gray, S. K.; Schatz, G. C. Nonlocal Optical Response of Metal Nanostructures with Arbitrary Shape. Phys. Rev. Lett. 2009, 103, 97403.

(53)

McMahon, J. M.; Gray, S. K.; Schatz, G. C. Calculating Nonlocal Optical Properties of Structures with Arbitrary Shape. Phys. Rev. B 2010, 82, 35423.

(54)

Boardman, A. D. Electromagnetic Surface Modes; John Wiley & Sons: New York; 1982.

(55)

Chen, S. Y.; Mock, J. J.; Hill, R. T.; Chilkoti, A.; Smith, D. R.; Lazarides, A. A. Gold Nanoparticles on Polarizable Surfaces as Raman Scattering Antennas. ACS Nano 2010, 4, 6535–6546.

(56)

Homola, J. Present and Future of Surface Plasmon Resonance Biosensors. Anal. Bioanal. Chem. 2003, 377, 528–539.

(57)

Kleinman, S. L.; Ringe, E.; Valley, N.; Wustholz, K. L.; Phillips, E.; Scheidt, K. A.; Schatz, G. C.; Van Duyne, R. P. Single-Molecule Surface-Enhanced Raman Spectroscopy of Crystal Violet Isotopologues: Theory and Experiment. J. Am. Chem. Soc. 2011, 133, 4115–4122.

(58)

Benz, F.; Chikkaraddy, R.; Salmon, A.; Ohadi, H.; De Nijs, B.; Mertens, J.; Carnegie, C.; Bowman, R. W.; Baumberg, J. J. SERS of Individual Nanoparticles on a Mirror: Size Does Matter, but so Does Shape. J. Phys. Chem. Lett. 2016, 7, 2264–2269.

(59)

Hill, R. T.; Mock, J. J.; Urzhumov, Y.; Sebba, D. S.; Oldenburg, S. J.; Chen, S. Y.; Lazarides, A. A.; Chilkoti, A.; Smith, D. R. Leveraging Nanoscale Plasmonic Modes to Achieve Reproducible Enhancement of Light. Nano Lett. 2010, 10, 4150–4154.

(60)

Rycenga, M.; Xia, X.; Moran, C. H.; Zhou, F.; Qin, D.; Li, Z.-Y.; Xia, Y. Generation of Hot Spots with Silver Nanocubes for Single-Molecule Detection by Surface-Enhanced Raman Scattering. Angew. Chemie Int. Ed. 2011, 50, 5473–5477.

(61)

Wen, J.; Wang, W. J.; Li, N.; Li, Z. F.; Lu, W. Light Enhancement by Metal-Insulator-Metal Plasmonic Focusing Cavity. Opt. Quantum Electron. 2016, 48, 150.

(62)

Lin, S.; Zhu, W.; Jin, Y.; Crozier, K. B. Surface-Enhanced Raman Scattering with Ag Nanoparticles Optically Trapped by a Photonic Crystal Cavity. Nano Lett. 2013, 13, 559–563.

(63)

Shoute, L. C. T.; Bergren, A. J.; Mahmoud, A. M.; Harris, K. D.; McCreery, R. L. Optical Interference Effects in the Design of Substrates for Surface-Enhanced Raman Spectroscopy. Appl. Spectrosc. 2009, 63, 133–140.

(64)

Jayawardhana, S.; Rosa, L.; Juodkazis, S.; Stoddart, P. R. Additional Enhancement of Electric Field in Surface-Enhanced Raman Scattering due to Fresnel Mechanism. Sci. Rep. 2013, 3, 2335.

(65)

Hastings, S. P.; Swanglap, P.; Qian, Z.; Fang, Y.; Park, S. J.; Link, S.; Engheta, N.; Fakhraai, Z. Quadrupole-Enhanced Raman Scattering. ACS Nano 2014, 8, 9025–9034. 22 ACS Paragon Plus Environment

Page 22 of 24

Page 23 of 24

1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60


(66)

Qian, Z.; Li, C.; Fakhraai, Z.; Park, S. J. Unusual Weak Interparticle Distance Dependence in Raman Enhancement from Nanoparticle Dimers. J. Phys. Chem. C 2016, 120, 1824–1830.

(67)

Lal, S.; Grady, N. K.; Goodrich, G. P.; Halas, N. J. Profiling the near Field of a Plasmonic Nanoparticle with Raman-Based Molecular Rulers. Nano Lett. 2006, 6, 2338–2343.

(68)

Prodan, E.; Radloff, C. J.; Halas, N. J.; Norlander, P. A Hybridization Model for the Plasmon Response of Complex Nanostructures. Science. 2003, 302, 419–422.

(69)

Henry, A. I.; Bingham, J. M.; Ringe, E.; Marks, L. D.; Schatz, G. C.; Van Duyne, R. P. Correlated Structure and Optical Property Studies of Plasmonic Nanoparticles. J. Phys. Chem. C 2011, 115, 9291–9305.

(70)

Dick, L. A.; McFarland, A. D.; Haynes, C. L.; Van Duyne, R. P. Metal Film over Nanosphere (MFON) Electrodes for Surface-Enhanced Raman Spectroscopy (SERS): Improvements in Surface Nanostructure Stability and Suppression of Irreversible Loss. J. Phys. Chem. B 2002, 106, 853– 860.



1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60

50x50mm (300 x 300 DPI)

ACS Paragon Plus Environment

Page 24 of 24

SERS - ACS Publications - American Chemical Society

Recommend Documents