Antibody Conjugated, Raman Tagged Hollow GoldâSilver

Subscriber access provided by Binghamton University | Libraries

Article

Antibody conjugated, Raman tagged hollow gold-silver nanospheres for specific targeting and multimodal Dark Field/ SERS/ Two Photon-FLIM imaging of CD19(+) B lymphoblasts Timea Nagy-Simon, Andra-Sorina Tatar, Ana-Maria Craciun, Adriana Vulpoi, Maria-Ancuta Jurj, Adrian Florea, Ciprian Tomuleasa, Ioana Berindan-Neagoe, Simion Astilean, and Sanda Boca ACS Appl. Mater. Interfaces, Just Accepted Manuscript • Publication Date (Web): 02 Jun 2017 Downloaded from http://pubs.acs.org on June 3, 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.

ACS Applied Materials & Interfaces 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 50

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

ACS Applied Materials & Interfaces

Antibody conjugated, Raman tagged hollow goldsilver nanospheres for specific targeting and multimodal Dark Field/ SERS/ Two Photon-FLIM imaging of CD19(+) B lymphoblasts Timea Nagy-Simon#, Andra-Sorina Tatar#,†, Ana-Maria Craciun#, Adriana Vulpoi§, MariaAncuta Jurj‡, Adrian Florea∥, Ciprian Tomuleasa‡, Ioana Berindan-Neagoe‡, Simion Astilean#, †, Sanda Boca#,* # Nanobiophotonics and Laser Microspectroscopy Center, Interdisciplinary Research Institute on Bio-Nano-Sciences, Babes-Bolyai University, T. Laurian 42, 400271, Cluj-Napoca, Romania † Faculty of Physics, Babes-Bolyai University, Kogalniceanu 1, 400084 Cluj-Napoca, Romania § Nanostructured Materials and Bio-Nano-Interfaces Center, Interdisciplinary Research Institute on Bio-Nano-Sciences, Babes-Bolyai University, T. Laurian 42, 400271, Cluj-Napoca, Romania ‡ Research Center for Functional Genomics and Translational Medicine, Iuliu Hatieganu University of Medicine and Pharmacy, Cluj-Napoca, Romania ∥ Department of Cell and Molecular Biology, Iuliu Hatieganu University of Medicine and Pharmacy, Cluj-Napoca, Romania

ACS Paragon Plus Environment

1


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

Page 2 of 50

KEYWORDS Hollow gold-silver nanospheres, antiCD19, B lymphoblasts, multimodal imaging, SERS, TPE FLIM

ABSTRACT In this research paper we propose a new class of contrast agents for the detection and multimodal imaging of CD19(+) cancer lymphoblasts. The proposed agents are based on NIR responsive hollow gold-silver nanospheres conjugated with antiCD19 monoclonal antibodies and marked with Nile Blue (NB) SERS active molecules (HNS-NB-PEG-antiCD19). Proof of concept experiments on specificity of the proposed complex for the investigated cells was achieved by Transmission Electron Microscopy (TEM). The microspectroscopic investigations via Dark Field (DF), Surface Enhanced Raman Spectroscopy (SERS), and Two Photon Excited Fluorescence Lifetime Imaging Microscopy (TPE-FLIM) corroborate with TEM and demonstrate successful and preferential internalization of the antibody-nanocomplex. The combination of the microspectroscopic techniques enables contrast and sensitivity that competes more invasive and time demanding cell imaging modalities, while depth sectioning images provide real time localization of the nanoparticles in the whole cytoplasm at the entire depth of the cells. Our findings prove that HNS-NB-PEG-antiCD19 represent a promising type of new contrast agents with great possibility of being detected by multiple, non invasive, rapid and accessible microspectroscopic techniques and real applicability for specific targeting of CD19(+) cancer cells. Such versatile nanocomplexes combine in one single platform the detection and imaging of cancer lymphoblast by DF, SERS and TPE-FLIM microspectroscopy.


2

Page 3 of 50

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. INTRODUCTION In present-day medicine, state-of-the-art translational research is focused on the development of new agents for specific targeting and molecular imaging of cancer cells 1. Current cell-based diagnostic tools such as immunocitochemistry require large numbers of malignant cells, which are usually present in advanced stages of the disease. Early cancer detection, the ability to predict cancer progression, and the ability to monitor response to the therapy is no longer a utopia as it can be clinically achieved grace to the use of molecular and prototypical agents such as fluorophores, quantum dots or nanoparticles 2. When compared to the bulk material, nanosized gold and silver particles present distinct physical and chemical properties, which favored their applicability in various areas such as electronics, chemistry, biology or medicine 3. In medicine, the application of noble metal nanoparticles has known an extensive growth during the past years and studies at nano-bio interface include the use of nanoparticles as contrast agents for microscopy imaging, vehicles for drug delivery or targeted probes for cancer cell detection and therapy 4. Noble metal nanoparticles have unique optical characteristics which are linked to their surface plasmon resonances (SPR). By varying the particle size, morphology and inherent structure, the SPR of noble metal nanoparticles can be tuned from visible to the biologically approachable and tissue transparency near-infrared (NIR) domain. Noble metal nanoparticles amplify the electromagnetic field in their immediate vicinity, property that was explored for their use as efficient Surface enhanced Raman Scattering (SERS) substrates for various (bio)molecules. Reporters with inherent large Raman cross sections were used for preparing the so-called SERS nanotags which, depending on the particle type and the choice of the laser excitation line, proved applicability in intracellular and in vivo imaging and efficacy that often surpassed that of the commonly used


3


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

fluorophores

5,6

Page 4 of 50

. Moreover, compared to other metallic nanoparticles, gold nanoparticles have

excellent biocompatibility and offer ease of conjugation with targeting molecules such are peptides, aptamers or antibodies, providing molecular recognition and differentiation between malignant and normal cells 7,8. From the class of noble metal nanoparticles, hollow gold-silver nanoparticles (HNS) satisfy all the above requirements for being used in cell imaging applications, especially when the experiments involve cells that are physiologically grown in suspension as is the case of leukemia and lymphoma cells. The main biomedical applications of HNS include biosensing, bioimaging, photothermal cancer therapy, and drug or gene delivery 9. Regarding bioimaging, photoacoustic visualisation is the widest used imaging technique that exploits HNS as contrast agents

10

, but other methods such as CT and OCT imaging

11

or microspectroscopic techniques

such as SERS are also applied. Despite numerous reports on the SERS activity of HNS both in solution as well as on solid substrates 12,13, very little data has yet to assess the intracellular SERS imaging and detection capability of such nanostructures. Lee et al. have reported antibodyconjugated hollow gold nanospheres as multimodal sensing agents for both dark-field and SERS imaging of HER2/neu markers overexpressed in single MCF7 breast cancer cells

14

. The same

group has demonstrated a highly efficient multiplex SERS imaging of biomarkers (EGF, ErbB2 and IGF-1) expressed in three types of breast cancer cell lines (MDA-MB-468, KPL4 and SKBR-3) using silica-encapsulated hollow gold nanospheres conjugated with specific antibodies 15. Quantitative analysis of EGFR, ErbB2 and IGF-1R marker proteins distribution has been reported; meanwhile the cancer cell phenotypes could also be successfully identified from their SERS-mapping images. The therapeutic potential of HNS is somewhat better exploited and documented, especially focusing on the photothermal effect induced upon laser irradiation 16.


4

Page 5 of 50

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


Recently, a new imaging method evidenced amongst the microspectroscopic techniques that are currently used for cell and tissue imaging applications. Two-photon excitation (TPE) microscopy is an emerging non-linear technique with versatile possibilities and numerous advantages

17

. First of all, as two-photon excited non-linear processes imply the simultaneous

absorption of lower energy photons, longer excitation wavelengths allow researchers to effectively exploit the tissue “optical window” in the 700–1000 nm spectral domain. Therefore, deep penetration of the excitation light can be achieved facilitating imaging of thick biological specimens. On the other hand, in TPE, most of the generated signal is limited to the focal region which results in greatly improved axial depth discrimination and improvement in the image contrast, compared with conventional microscopy. There are several reports that exploit the two photon luminescence (TPL) of gold nanoparticles for cellular imaging, but very few exploit the promising combination with timeresolved fluorescence technique, allowing the distinction between signals that have overlapping emissions, but different fluorescence decay times. Durr et al. were the first to use gold nanorods for molecularly specific nonlinear imaging of skin cancer cells in a three-dimensional tissue phantom

18

. Later, Zhao et al. proved that gold nanorods can be successfully employed as dual

photo-sensitizing and imaging agent in TPE photodynamic therapy, enabling image-guided therapy assays 19, while Wu et. al showed that their aspect ratio should be in resonance with the excitation wavelength in order to obtain high quality cell images under TPE

20

. Other research

groups have demonstrated that branched nanoparticles exhibit remarkable TPE luminescence being less toxic than gold nanorods, which makes them a valuable alternative for cell-targeted treatments

21

. Despite the already demonstrated feasibility of using different types of gold

nanoparticles as contrast agents in TPE imaging assays, the combination of TPE with the


5


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

Page 6 of 50

powerful Fluorescence Lifetime Imaging Microscopy (FLIM) techniques has only been reported by one research group so far. Specifically, Zhang et al. employed gold nanorods as fluorescent labels to successfully visualize kidney cells through TPE-FLIM. This approach enables better contrast and sensibility than conventional intensity imaging considering that the fluorescence lifetime of gold nanorods in less than 100 ps, being easily distinguished from other fluorescent labels or endogenous fluorophores 22. Previous work of the author demonstrated the pertinence of using multimodal imaging via scanning confocal Surface Enhanced Resonant Raman Spectroscopy (SERRS) and FLIM of cancer cells incubated in the presence of Methylene Blue loaded, Pluronic stabilized 3D nanoassemblies of gold nanoparticles with plasmon resonances in the visible domain

23

. Herein, we

propose a new class of contrast agents for in vitro detection and multimodal imaging of CD19(+) lymphoma cells via highly informative microspectroscopic techniques: Dark Field (DF), SERS, and TPE-FLIM. The multifunctional agents are based on NIR-responsive HNS conjugated with monoclonal antibodies and marked with spectroscopic active, Raman reporter molecules. The hollow gold-silver nanoparticles were prepared by galvanic replacement reaction using silver nanospheres as template, method that allows the precise control on the spectral position of the SPR. For application in SERS imaging, we conjugated the particles with Nile Blue (NB) cationic dye, a widely used Raman reporter molecule with affinity for the nanoparticles surface through electrostatic interaction and demonstrated high Raman activity. Further, the HNS-NB dyenanoparticle complex was covered by heterofunctional PEG which has a role in improving nanoparticle stability and biocompatibility but also to provide functional groups for nanoparticle functionalization with antiCD19 targeting antibody via EDC/NHS coupling. Specificity of the proposed complex for the investigated cells was demonstrated by TEM, which successfully


6

Page 7 of 50

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


showed that antibody conjugated nanoparticles preferentially internalize the cells in comparison with their nonconjugated homologous nanoparticles. Combined approaches based on less invasive TPE-FLIM measurements, SERS mapping and DF microscopy on lymphoma (SKW 6.4-human Epstein-Barr virus-transformed B cell) cells incubated with HNS-NB-PEG-antiCD19 corroborate with the TEM results and showed successful internalization of the antibody conjugated particles by cells. Supplementary, depth sectioning images show localization of the nanoparticles in the whole cytoplasm at the entire depth of the cells. The obtained results demonstrate that HNS-NB-PEG-antiCD19 are versatile nanocomplexes that combine in one single platform the detection and imaging by DF, SERS and TPE-FLIM microspectroscopy, being promising agents for specific targeting and multimodal microspectroscopic imaging of CD19(+) lymphoma cells and of cancer lymphoblast. Moreover, due to its non invasive character and operability in the NIR biological window we are confident that the presented methods have a real possibility for the near time future in vivo applicability including real time imaging for guided chemotherapy and tracking of the cancer cells response to drug treatment. Such approach could not only save the unwanted costs of the treatment but can considerably diminish patient discomfort and side effects.

2. MATERIALS AND METHODS

2.1. Materials Ascorbic

acid

(AA),

silver

nitrate

(AgNO3),

trisodium

citrate

(Na3C6H5O7),

Poly(vinylpyrrolidone) of MW 10000 (PVP), Hydrogen tetrachloroaurate (III) hydrate (HAuCl4.3H2O,

99.99%),

N-(3-Dimethylaminopropyl)-N′-ethylcarbodiimide

hydrochloride


7


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

Page 8 of 50

(EDC), N-Hydroxysulfosuccinimide sodium salt (S-NSH) were purchased from Sigma-Aldrich. Nile Blue A (NB) was obtained from Alfa Aesar. Anti-CD19 antibody [MB19-1] (FITC) was purchased from Abcam. Alpha-Methoxy-omega-mercapto poly(ethylene glycol) of 5000 Dalton (mPEG-SH) and alpha-Thio-omega-carboxy poly(ethylene glycol) of 3000 Dalton (SH-PEGCOOH) were purchased from Iris Biotech GmBh. The following materials were used for biological experiments: Gibco® RPMI 1640 Medium 1X (500 mL), Gibco® Minimim Essential Medium (500 mL), Gibco® Fetal Bovine Serum (500 mL), Gibco® Penicillin-Streptomycin (100 mL), paraformaldehyde (Sigma-Aldrich). All the aqueous solutions were prepared with ultrapure water (with a resistivity of 18.2 MΩ cm) from a Milli-Q purification system (Millipore, Merck). Materials for TEM: glutaraldehyde (Electron Microscopy Sciences, Hatfield, USA), osmium tetroxide (Sigma-Aldrich), acetone (Merck, Darmstadt, Germany), EMBed-812 embedding kit:

EMBed-812,

dodecenyl

succinic anhydride,

methyl-5-norbornene-2,3-

dicarboxylic anhydride, and 2,4,6-tris-(dimethylaminomethyl)phenol (EMS), 300 mesh copper grids (Agar Scientific Ltd., Stansted, UK), uranyl acetate (Merck), ethanol (Merck).

2.2. Synthesis and surface modification of HNS 2.2.1 Synthesis of HNS To synthesize HNS we used a galvanic replacement method, based on the reduction of gold tetrachloroauric acid (HAuCl4) onto silver nanoparticle templates. Quasi-spherical silver nanoparticles were synthesized by adapting the protocol by Li et al 24. Briefly, 700 µl of 10 mM AA were added to 47 ml of ultrapure water and brought to boiling under continuous magnetic stirring. 1 ml of aqueous solution of 1% sodium citrate and 0.25 ml of aqueous solution of 1% AgNO3 were added to 1.25 ml water and incubated for 5 min at room temperature under stirring,


8

Page 9 of 50

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


then injected to the boiling solution of AA. The boiling was continued for 15 min, after which the heat was stopped while keeping the stirring process for another 15 min. The pristine AgNP colloidal solution was incubated with an aqueous solution of PVP at a final polymer concentration of 0.4 mM for two hours. The unbound polymer was removed by centrifugation and resuspension of the particles in ultrapure water. To tune the extinction of the obtained HNS to a desired wavelength, different colloidal samples were prepared by varying the amount of HAuCl4 in the galvanic replacement reaction. Briefly, 5 ml of AgNP-PVP were boiled under continuous magnetic stirring and incremental volumes (50, 62.5, 75, 100 and 125 µl) of 2.5 mM HAuCl4 were rapidly injected in the different samples. A rapid gradual color change (from yellow to blue) was observed in the mixed solutions. The nanoparticles were purified from the unreacted chemicals by centrifugation at 12290 ×g for 10 min. The AgCl precipitate was separated by centrifugation for 1 h at low speed (31 ×g) at 4 °C.

2.2.2 Preparation of antibody-SERS HNS nanoprobes Purified HNS were mixed with a solution of Nile Blue (NB) Raman reporter at 10-5 M concentration, in a 10:1 volume ratio and incubated for short term. After the reporter tagging step, a two-step PEGylation protocol followed. First, COOH-PEG-SH (10-3 M) was conjugated onto nanoparticle surface by the addition of the polymer to the nanoparticle solution and their overnight incubation at 4oC. Second, mPEG-SH (10-4 M) was added and left to react at room temperature (RT) for another 2 hours. Both PEGs carry a thiol group (-SH) at one end which forms covalent bonding with the NP surface. Excess polymer was removed by centrifugation (10 minutes, 20 oC, 12290 ×g) and resuspension of the particles in PBS. The terminal -COOH groups of the PEGylated HNS were activated using a modified EDC-NHS protocol 25. Briefly, a mixture


9


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

Page 10 of 50

of EDC and S-NHS (2:1 molar ratio) was added to the NP solution and left to react for 2 hours at RT, followed by purification by centrifugation (10 minutes, 20 oC, 12290 ×g) and resuspension in PBS. AntiCD19 monoclonal antibody was added to the activated particles in a final concentration of 2.5 ug/ml. The mixture was incubated for 2 hours at RT, followed by an overnight incubation at 4 oC. Antibody conjugated particles were purified by centrifugation (10 minutes, 20 oC, 12290 ×g) and resuspension in PBS. After the removal of the unbound protein, the particles were stored at -20 ºC prior use. The quantification of the amount of antiCD19 antibody conjugated to nanoparticles was performed by the fluorescence emission at 520 nm of the FITC molecules stoichiometrically conjugated to the antibody (excitation at 495 nm) as measured from the supernatant solution, and by fitting the value on a calibration curve.

2.3. Characterization of the colloidal samples UV-Vis-NIR extinction spectra of the colloidal samples were recorded using a Jasco V-670 UV-Vis-NIR spectrometer in 2 mm path length quartz cuvettes at 1 nm spectral resolution. Fluorescence emission spectra were measured on a Jasco FP6500 spectrofluorimeter. For Transmission Electron Microscopy (TEM) the colloids were added dropwise onto a carbon film covered copper grid. The morphology of the nanoparticles was examined using a FEI Tecnai F20 field emission, high resolution TEM (TEM/HRTEM) operating at an accelerating voltage of 200 kV and equipped with Eagle 4k CCD camera and an EDAX instruments EDS detector (point, line, and area profiling); Area Energy Dispersive X-Ray Spectra (EDS) were recorded on each specimen on at least four different few hundred nanometer areas using a thickness correction method for the elemental identification and Ag/Au ratio evaluation. Particle size distribution and Zeta-potential were measured by a Zetasizer NanoZS90 instrument (Malvern


10

Page 11 of 50

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


Instruments). Analysis was performed at a scattering angle of 90° and temperature of 25 °C. The SERS spectra of Raman tagged HNS were acquired with a portable Raman spectroscope (Raman Systems R3000 CN) equipped with a 785 nm diode laser with a power of 200 mW coupled to a 100 µm optical fiber.

2.4. Cell culture CD19 positive SKW 6.4 (ATCC® TIB-215™) cells (Epstein-Barr virus-transformed B lymphocytes, B-cell lymphoma) were cultured in RPMI 1640 medium, supplemented with 10% fetal bovine serum, and 50 U/mL penicillin and 50 µg/mL streptomycin. Cells were grown in a humidified atmosphere at 37 °C with 95% air and 5% carbon dioxide (CO2). 105 cells were incubated in 6-well plates with the antibody-conjugated, Raman-tagged nanoparticles, and a well was left un-treated, for use as a control sample. After 24 hours co-incubation in the presence of the particles the cells were centrifuged to remove the culture medium and washed twice with PBS. The cells were fixed with 4% paraformaldehyde, washed twice with PBS and kept at -20 °C prior imaging assays. CD19 negative, OCI-AML3 cell line (acute myeloid leukemia) was kindly gifted by Professor Gabriel Ghiaur, M.D., Ph.D. (Johns Hopkins Medicine, Baltimore, MD, USA) and the cells were cultured according to the indicated protocol.

2.5. Trasmission Electron Microscopy Cells incubated with nanoparticles were centrifuged, and the culture medium was replaced with the 2.7% glutaraldehyde (in 0.1 M phosphate buffer, pH 7.4) for a 2 h prefixation. Then, the cells were washed 4 times (3 x 1 h, and overnight) with the same buffer and then postfixed for 2 h with 1.5% OsO4 (in 0.15 M phosphate buffer, pH 7.4). Next, the cells were dehydrated in an


11


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

Page 12 of 50

acetone series and embedded in EMBed-812. Ultrathin sections obtained with a DiATOME diamond knife (DiATOME, USA) on a Bromma 8800 ULTRATOME III ultramicrotome (LKB, Sweden) were collected on 300 mesh copper grids and immediately contrasted with saturated alcoholic uranyl acetate for 5 min. Examination of the sections was performed on a JEOL JEM 1010 transmission electron microscope (JEOL Ltd., Japan) at 80 kV acceleration voltage, and different magnifications. Relevant images were captured using a Mega VIEW III camera (Olympus, Soft Imaging System, Germany).

2.6. Dark field microscopy Dark Field (DF) microscopy was performed on cells deposited onto Ibidi 30 µ-Dish (50 mm) using an inverted Zeiss Axio Observer Z1 microscope. A 100 W halogen lamp was used for DF illumination with constant intensity on each sample which was focused on the sample without using any thermal filter using a high numerical immersion DF condenser (NA=1.4). The scattered light was collected by a LD Plan-Neofluar 20× objective (NA = 0.4, Zeiss) or a PlanApochromat oil immersion 63× objective (NA = 0.7-1.4, Zeiss). Images were acquired using an AxioCam Icc Rev.4 CCD camera (1.4 megapixels, Zeiss) with the same integration time for each sample (300 s and 400 ms for images taken with objective 20× and 63×, respectively) and processed by the ZEN software.

2.7. SERS imaging For SERS imaging, paraformaldehyde fixed, nanoparticle loaded cells were deposited onto quartz microslides pretreated with poly-L-lysine. SERS mapping of cells incubated with nanoparticles was acquired using a confocal alpha300R Raman microspectrometer from WITec


12

Page 13 of 50

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


(Germany). The 785 nm diode laser and the 633 nm He-Ne laser were used for excitation and laser powers of 145 mW and 15 mW, respectively, were measured before entering the objective. The excitation beam and the backscattered light collected onto an air-cooled chargecoupled detector were delivered trough a W-plan Apochromat 63× (NA=1, WD=2.1 mm, Zeiss) water immersion objective. The system was precalibrated to the spectral line of silicon at 520,7 cm-1. An integration time of 0.75 s was used for collecting each SERS spectrum. For analyzing the light emerging from the 100 µm output optical fiber, an ultrahigh throughput spectrometer equipped with back-illuminated deep-depletion 1024 × 128 pixel CCD camera operating at -60 °C (DV401-BV, Andor) was used. An approximate 20 × 20 µm 2 area was chosen for scanning the fixed cells. The WITec Project Plus software was used for spectral analysis and image processing (cosmic ray removal, Savitzky-Golay smoothing, baseline correction, background subtraction). Univariate Raman maps of cells were obtained by integrating the intensity of the band at 1420-1480 cm-1 assigned to CH2 and CH3 deformations, specific to proteins and lipids, over the scanned area, and that of the 595 cm-1 band, specific to the phenoxazine ring stretching vibrations of the Raman reporter NB, for the visualization of the nanoparticle distribution inside the cells. The nanoparticle reference SERS spectra were obtained from the colloidal solutions using a 20× objective (NA=0.4) and an integration time of 10/20/30 s. Reflected-light brightfield optical images of cells were captured with a color video camera attached to the eyepiece output of the same microscope using for illumination a super-bright white LED source.

2.8. TPE FLIM imaging For TPE FLIM imaging assays by FLIM, the cells were deposited onto Ibidi 8 Well µ-Slides with glass bottom (no. 1.5H (170 µm +/- 5 µm); D 263 M Schott glass). Measurements were


13


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

Page 14 of 50

performed on a PicoQuant MicroTime 200 time-resolved confocal fluorescence microscope system based on an inverted microscope (IX 71, Olympus) equipped with a UPLSAPO 60×/NA 1.2 water immersion objective. The excitation beam was provided by a tunable Coherent Mira 900 Titanium: Sapphire pulsed laser operating at 800 nm, at a power of 40 mW (0.8 mW on sample) and 80 MHz. For FLIM image acquisition was employed a piezo x-y-scanning table and a PiFoc z-piezo actuator for microscope objective. The signal collected through the objective was spatially and spectrally filtered by a 50 µm pinhole and a FF01-750SP (Semrock, USA) emission filter, respectively, before being focused on a PDM Single Photon Avalanche Diode (SPAD) from MPD. The detector signals were processed by the PicoHarp 300 Time-Correlated Single Photon Counting (TCSPC) data acquisition unit, from PicoQuant. Data were recorded and analyzed using the SymPhoTime software from PicoQuant. TPL spectra from nanoparticles in solution and solid phase (dropcasted on microscope cover glass) were obtained with a SR-163 spectrograph equipped with a Newton 970EM CCD camera from Andor Technology coupled to an exit port ofthe main optical unit of MicroTime200 through a 50 µm optical fiber. The samples were excited at 800 nm and the signal was filtered by a FF01-750SP (Semrock, USA) emission filter. The integration time used for the acquisition of the fluorescence spectra was 60 s (solution phase) and 10 sec (solid phase).

3. RESULTS AND DISCUSSION

3.1. Synthesis and characterization of hollow gold nanospheres The synthesis method used to prepare HNS is based on the templated galvanic replacement reaction, a common technique used to produce various nanomaterials with hollow structures with


14

Page 15 of 50

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


tunable porosity, optical and catalytic properties. It is an electrochemical redox process between two metals, which involves the oxidation of one metal, also referred as sacrificial template, by the ions of another metal with higher reduction potential, which will be reduced and plated onto the outer surface of the template 26. Herein, the high difference in the reduction potential between silver and gold was exploited to prepare hollow nanostructures with absorption in the NIR spectral domain. Spherical silver nanoparticles (AgNPs) were used as template for the galvanic replacement reaction to obtain nanospheres with hollow interior. First, AgNPs were synthesized by a modified method from Li et al., through the chemical reduction of AgNO3 by sodium citrate and ascorbic acid

24

. This synthesis yielded quasi-spherical silver nanoparticles with a mean

diameter of 51±7 nm as measured from TEM images (Supporting Information Figure S1.B), a hydrodynamic diameter of 58 nm as measured by Dynamic Light Scattering (DLS) and a SPR centered at 418 nm (Supporting Information, spectrum a from Figure S1.A). To enhance the reproducibility and stability of HNS, we introduced a stabilization step of the template AgNPs by using PVP polymer, method adapted from the protocols that are generally used for the synthesis of gold nanocages

27

or cobalt template based hollow gold nanospheres

28

. Compared to other

surfactants or organic molecules that are commonly used in the synthesis or stabilization of NIR absorbing gold or silver nanoparticles

29

, PVP has the advantage of good biocompability, being

approved by U.S. Food and Drug Administration (FDA) as food additive and also used as binder in pharmaceutical tablets and in cosmetics industry. The adsorption of PVP on the surface of AgNPs was first assessed by a 3 nm red-shift of the SPR peak. The increase of the hydrodynamic diameter from 55 to 69 nm measured by DLS technique also confirms the presence of polymer at the nanoparticles` surface, as well as the modification of the Zeta-potential values (from -48 to 40 mV).


15


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

Page 16 of 50

Five different samples of HNS (denoted as HNS1, HNS2, HNS3, HNS4, HNS5) having the SPR tuned over the red-NIR spectral domain were synthesized by varying the quantity of HAuCl4 added into the galvanic replacement reaction. The (normalized) extinction spectra of the obtained nanoparticles after the purification and separation are presented in Figure 1. Major spectral changes can be observed as function of the amount of HAuCl4 used: the extinction bands are shifted to longer wavelengths and are broadened as the Au concentration was increased. These spectral modifications were commonly observed by others in the synthesis of nanoparticles with hollow structures, being attributed to the deposition of thin gold layers on the surface of the silver nanoparticles templates which form a shell around the nanoparticles and as seen in the TEM images from Figure 1 30,31. On the other hand, the formation of pinholes in the nanoparticle morphology which lends a non-uniform and rough surface represents another factor that influences the aspect and the position of the plasmonic band. The PVP polymeric corona is also visible and appears as a faint shadow of about 3-4 nm that surrounds almost entirely the HNS. Specifically, in the case of HNS1, a clearly visible shoulder centered at around 420 nm is observed in the spectrum, which originates from the template AgNPs and indicates an unfinished replacement reaction

32

. This hypothesis is sustained by the TEM images which reveal the

existence of pinholes in a large number of nanoparticles and a non-compact shell. For the samples in which a higher HAuCl4 concentration was used (samples HNS3 to HNS5), the SPR band characteristic to solid spherical AgNPs disappears, proving the formation of hollow structures.


16

Page 17 of 50

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. UV-Vis-NIR extinction spectra (A) and corresponding TEM images (B-F) of HNS obtained by increasing the amount of HAuCl4. Insets show magnified images of a representative HNS from each sample and illustrate the formation of pinholes as function of HAuCl4 concentration. Scale bar in inset images equals 20 nm.

As the TEM images illustrate, starting from this point, the hollow architecture is confirmed by a clearly visible dark shell and a brighter interior core of the nanoparticles. Additionally, we observe that in most of the cases we deal with a non-uniform aspect of the shell, presenting several surface defects such as holes or cavities. Theoretically, the introduction of additional gold atoms in the reaction will be able to stopper these holes and smooth the shell. Contrary, we observed a partial defragmentation in case of HNS5, marked in the TEM image with arrows and illustrated in Figure S2 from the Supporting Information. This phenomenon was also reported by


17


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

Page 18 of 50

Wan et al in a study that shows that by further increasing the HAuCl4 quantity in the reaction mixture, the HNS start to collapse into small AuNPs fragments which spectroscopically manifests in the disappearance of the characteristic NIR band and the appearance of a new band in the visible plasmonic band which is characteristic to full AuNPs

33

. A similar behavior was

observed for the HNS5 sample for which the extinction spectrum presents a small shoulder in the SPR band, centered at 520 nm. The overall average diameters of all HNS samples measured based on TEM images and the hydrodynamic diameters measured by DLS (Table 1) are similar. This is an expected result since they are imposed by the size of the template AgNPs. Further on, no major changes in size are noticeable with the increase of the maximum absorption wavelength, except the case of HNS5 which, as previously said, started to collapse, as key structural changes that influence the spectral position occur in the interior and at the surface of nanoparticles. It has been demonstrated that galvanic replacement cannot yield completely caged nanostructure and usually partial Ag components still remain in the nanoparticles after the reaction

34,35

. Therefore, we performed

EDX measurements to determine the elemental ratio of gold and silver in the obtained nanostructures (Table 1). In the galvanic replacement reaction, the decreasing of the Ag/Au mass ratio by increasing the HAuCl4 amount is consistent with a growth model frequently met by other researchers in which gold was deposited on the surface of the silver template, followed by the partial dissolution of silver from the interior of nanoparticles, resulting in the formation of hollow structures 30,36.

Table 1. HAuCl4 quantity used in the galvanic replacement reaction, LSPR peak positions, average hydrodynamic diameters measured by DLS, Zeta-potential values and Ag/Au


18

Page 19 of 50

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


composition from EDX measurements of the obtained HNS and corresponding standard deviation (SD) values calculated for 3 measurements. HAuCl4 quantity (µl) Sample

λmax (nm)

(nm)

SD

PdI

SD

Ag/Au mass ratio

Dav

AgNP-PVP

0

417

69.3

0.6

0.59

0.005

-

HNS1

50

630

73

2

0.59

0.012

1.43

HNS2

62.5

652

73

2.4

0.32

0.005

1.13

HNS3

75

685

73.6

0.8

0.29

0.005

0.98

HNS4

100

720

74.2

1.1

0.25

0.003

0.83

HNS5

125

795

82.7

1.1

0.3

0.005

0.44

3.3. Evaluation of HNS SERS activity in solution To serve as efficient contrast agents for cellular imaging, nanoparticles need to possess a distinct, strong signal which can be unequivocally detected inside cells and allow their intracellular tracking. Due to the enhanced electromagnetic fields in their vicinity, plasmonic nanoparticles modified with Raman active molecules can generate a very intense, amplified signal of the reporter through the well-known phenomena SERS, and many schemes of SERSactive nanoparticles were developed lately by varying the type of the particle, the reporter molecule and the coating material 37,38. Herein, the SERS applicability of the nanoparticles was investigated using as Raman reporter Nile Blue (NB), a cationic dye molecule with affinities for the negative charged nanoparticle surface and strong Raman signal. A total volume of 100 µl NB (10-5 M) was added to 500 µl of HNS solutions in steps of 10 µl and SERS spectra were measured after each step using 785 nm


19


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

Page 20 of 50

NIR excitation. The number of nanoparticles was kept constant for each type of HNS. Figure S3 from SI shows the concentration dependent SERS spectra of NB tagged HNS. The maximum peak intensity in function of NB concentration is presented in Figure 2, based on the peak centered at 591 cm-1 considered the marker Raman band of NB and attributed to the phenoxazine ring stretching vibrations. Other characteristic peaks can be clearly distinguished and are attributed to in-plane CCC and NCC deformations (497, 548 and 662 cm-1) and ring stretching modes (1357, 1495, 1641 cm-1). C-H bending modes can be also observed at higher NB concentration centered at 1132, 1193 and 1171 cm-1

39

. A higher amplification of the Raman

signal of the reporter molecule was observed in the case of HNS with absorption at higher wavelength (HNS3, HNS4 and HNS5), which can be attributed to the higher porosity of these type of nanoparticles surface which results in more hot-spots per nanoparticles. Based on the above described SERS results and its spectral response in the NIR domain correlated with the TEM analysis, we have identified HNS4 as the most suitable type of hollow nanoparticles to be further used as contrast agents for the in vitro assays. Consequently, the studies regarding functionalization with targeting moieties and cell imaging studies were conducted only on HNS4 sample, denoted thereafter as HNS4. Reproducibility of the SERS measurement is shown in Figure S4 from the Supporting Information.


20

Page 21 of 50

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 2. The maximum SERS intensities in function of NB concentration (a-e) based on the peak centered at 591 cm-1.

3.4. HNS functionalization with antibodies The most widely used scheme for specific delivery of nanoparticles at cancer cells and tumor locus rely on conjugating their surface with targeting molecules as are antibodies which are then selectively internalized by cancer cells that express the cell surface target for that antibody. Various strategies are used for antibody-nanoparticle conjugation starting from the relatively simple electrostatic interaction to the more complex cross-linking procedures

40

. For our assay

we adopted the later method as in this case, the bonding between the antibody and the particle is the most robust. First, a commonly used co-functionalization method was employed using a high molecular thiol terminated monofunctional PEG and a thiolated and carboxylated low molecular bifunctional PEG, creating a mixed monolayer/linker complex. The main advantage of using SHPEGs consist in the high affinity of thiol to the gold nanoparticle surface resulting in a chemical bonding which is one the most suitable surface modification method, especially for applications that require good stability of the samples

41

. However, it has been demonstrated that although


21


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

Page 22 of 50

high-molecular-weight PEGs are better colloidal stabilizers than low-molecular-weight PEGs, they are not very efficient in conjugation with biomacromolecules. Instead, low-molecularweight PEGs enable the conjugation of large amounts of biomacromolecules but provide only minimal colloidal stability which may induce the aggregation of nanoparticles 42. Hence, the use of mixed PEG layers offers an alternative to fulfill both the stability requirement and the functionalization with a sufficient number of macromolecules. In our case, the COOH active functional group from the low molecular polymer was exploited to covalently attach the antibody by its primary amine (NH2) groups to the outer ends of the PEG monolayer via carboxylic acid bonds through the EDC/NHS reaction

43

. The UV-Vis-NIR extinction spectra measured

systemically at each functionalization step are presented in Figure 3. A blue-shift of the SPR wavelength from 720 to 716 nm was observed after HNS PEGylation, which can be ascribed to the decrease of the refractive index (RI) around nanoparticles as a consequence of the PVP replacement (RI=1.52) by the PEG-SH (RI= 1.45) 38. DLS and zeta-potential measurements also confirm the surface modification of nanoparticles induced by PEGylation (Table 2). A slight increase of the hydrodynamic diameter was observed after replacement of PVP by PEG, without any significant modification of the polydispersity index. Considering the lower molecular weight of the PEG compared to that of PVP which would normally lead to a decrease of the hydrodynamic size, the observed increase can be attributed to a variation in the structure of the two polymers, and specifically to its conformational organization around nanoparticles, as well as to the number of the available functional head groups. While PEG is likely to attach in a tail form at the surface of the nanoparticles anchored by the thiol group to the NPs surface and the long carbon tail of the polymer exposed to the bulk medium, PVP chains are preferentially adsorbed in a looped and entrained conformations 44.


22

Page 23 of 50

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. UV-Vis-NIR extinction spectra of nanoparticles measured at each functionalization step: HNS-PEG (red spectrum), HNS-NB-PEG (blue spectrum) and HNS-NB-PEG-antiCD19 (green spectrum); UV-Vis absorption spectrum of NB (blue dotted spectrum) is also illustrated together with two axes (dotted lines) that indicate the laser excitation wavelength.

The zeta-potential was also modified upon conjugation with PEG from -39 mV for initial nanoparticles to -19.3 mV in the case of the PEGylated samples. Compared to HNS-PEG, NB loaded HNS-PEG (HNS-NB-PEG) present a LSPR red-shifted by 12 nm proving the interaction of HNS with NB without altering their stability, since neither spectral broadening nor increase of the hydrodynamic diameter of the SPR band was observed. No further LSPR shift occurred following conjugation of HNS-NB-PEG with antiCD19 antibody, which is not surprising as the LSPR sensitivity decreases while the distance from nanoparticle surface increases

46

. On the

other hand, the well recognizable broadening of the SPR band is ascribed to the increased scattering by the nanoparticles due to the presence of antiCD19 antibody onto the nanoparticles surface. DLS measurements also demonstrate the conjugation of nanoparticles with antiCD19


23


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

Page 24 of 50

antibody through an increase in diameter by approximately 10 nm, while the monodispersity of the nanoparticles was maintained. Notably, conjugation with antiCD19 antibody did not affect the SERS signal of the nanoparticles as shown in Figure S5 from Supporting Information. To assess the antibody concentration on the NP samples, the supernatant obtained in the last NP purification step was used as an indirect method, by measuring the fluorescence emission at 520 nm (excitation at 495 nm) of the FITC molecules stoichiometrically conjugated to the antibody, and fitting the value on a calibration curve (Supporting Information Figure S6). Then, this value was subtracted from the total incubated antibody concentration, to obtain the final amount of antiCD19 antibody attached to the nanoparticles used for further experiments. The calculations gave a final antiCD19 concentration of 5.7 mg/ml attached to HNS-NB-PEG.

Table 2. LSPR peak positions, average hydrodynamic diameters measured by DLS, polydispersity index (PdI) and Zeta-potential values. LSPR

Dav

PdI

Zeta-potential

(nm)

(nm)

HNS

720

74.2±1.1

0.25

-39

HNS-PEG

716

81.95±2.3

0.26

-19.3

HNS-NB-PEG

728

82.06± ±0.8

0.179

-16

HNS-NB-PEGantiCD19

728

91.13± ±1.52

0.238

-24.7

(mV)

3.5. Proving HNS-antiCD19 specificity for the targeted cells Although the protocol is time demanding and requires extensive training of the person that manipulates the samples which cannot be afterward recuperated and hence limiting its


24

Page 25 of 50

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


applicability only to in vitro experiments, transmission electron microscopy (TEM) is so far the method that provides the highest accuracy for imaging cells and cellular organelles. The literature is abundant with reports regarding nanoparticle internalization and intracellular localization, depending on the particle material, coating or morphology

47,48

. In our study, we

also employed TEM as a first-line method to prove the specificity of antiCD19 conjugated HNS for the investigated cells. For this, SKW6.4 cells were incubated in the presence of HNS-NBPEG-antiCD19 nanoparticles for 24 h after which were imaged by TEM (sample II). Four samples were used as control: cells without nanoparticles (sample IV), cells that were pretreated with the free antibody and then incubated with the HNS-NB-PEG-antiCD19 for competitive binding test (sample I), cells that were treated with nonconjugated HNS-NB-PEG nanoparticles (sample III) and a CD19 negative cell line of acute lymphoblastic leukemia (OCI-AML3)

49

(sample V). The results are illustrated in Figure 4 and show a selective and preferential uptake of nanoparticles by SKW6.4 cells incubated with HNS-NB-PEG-antiCD19 nanoparticles. In these cells, the nanoparticles were found in important amounts either packed in endocytosis vesicles located in the proximity of plasma membrane (Figure 4A,B), or dispersed within the cytosol (Figure 4B). Some of the studied cells contained both nanoparticles still surrounded by membranes that indicate a recent endocytosis process, and nanoparticles in direct contact with the cytosol, predominantly in the peripheral region of the cytoplasm (Figure 4B). However, the plasma membrane of these cells was “decorated” with nanoparticles tightly attached at the cell surface, especially on the cell extensions (Figure 4A). No particles were found in the cells pretreated with the free antibody prior to incubation with the HNS-NB-PEG-antiCD19 (Figure 4C,D), but were present in the proximity of the cells. Is worth mentioning that very rare nanoparticles were found attached to the plasma membrane of these cells. Most of the SKW6.4


25


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

Page 26 of 50

cells incubated with nonconjugated HNS-NB-PEG nanoparticles do not contain any particles and when present, they are in very low quantity and located in the cytosol Figure 4E, F, including in more profound regions (Figure 4E, and inset of Figure 4F), close to nucleus. For negative control OCI-AML3 cells, the nanoparticles were intracellularly present, possible internalized by nonspecific endocytosis, but unlike in the case of CD19+ cells, a semnificative lower percent of HNS was observed and the particles were localized solely in endolysosomal compartments and mostly arranged in a cluster-like structure. The SKW6.4 cells incubated without nanoparticles displayed a normal ultrastructure, which was also kept for the cells incubated with nanoparticles.


26

Page 27 of 50


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 ACS Paragon Plus Environment

27


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

Page 28 of 50

Figure 4. TEM images showing a differentiated uptake of nanoparticles by SKW6.4 cells. In the SKW6.4 cells incubated for 24 h with HNS-NB-PEG-antiCD19 nanoparticles, important amounts of nanoparticles were found in endocytosis vesicles (ev) (A, B), and dispersed within the cytosol (B). The cells pretreated with CD19 and incubated with the HNS-NB-PEG-antiCD19, did not contain nanoparticles (C, D). After incubation with HNS-NB-PEG nanoparticles, only rare cells contained one or a few nanoparticles (arrows) located in cytosol (E, F). SKW6.4 cells without nanoparticles (G, H). OCI-AML3 cells with rare clusters of HNS-NB-PEG-antiCD19 nanoparticles (I, J). pm, plasma membrane; n, nucleus; nu, nucleolus; m, mitochondria, er, endoplasmic reticulum; ly, lysosomes; a, autophagosome.

3.6. Cell imaging by dark field microscopy Due to their enhanced light scattering properties, gold nanoparticles are widely used as highly efficient contrast agents in live cell imaging using dark field (DF) microscopy

50

. It is well-

known that the scattering abilities of gold nanoparticles are largely dependent on their size and shape and both aggregation and deviation from spherical to anisotropic shape increase the scattering cross section of the particles

51

. Considering these aspects, the proposed HNS

represent excellent candidates for DF imaging and hence, we used the method as an alternative, less invasive tool to evaluate the uptake of HNS by the malignant lymph cells. To evaluate the optimal incubation time required for a maximum uptake, the cells were incubated with HNS-NBPEG-antiCD19 for different periods of time (30 min, 2 h, 6 h, 14 h and 24 h). Cells without nanoparticles were used as control. As seen from Figure 5, the nanoparticles are visible as shiny red dots while the white dots seen in the samples of untreated cells are due to the characteristic scattering of the intrinsic cellular organelles. A time dependent trend is clearly observed as


28

Page 29 of 50

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


starting from the 30 min and up 2 h incubation and the nanoparticles can be seen mostly at the surface of cells, bound to the cell membrane. At 6 h incubation, a large number of nanoparticles are internalized into the cytoplasmic region of the cells while dark field image recorded after 24 h incubation shows an efficient nanoparticles uptake and internalization without a considerable amount of nanoparticles bound to the cell membrane. Moreover, by focusing at different Z levels, is possible to detect the presence of the particles at multiple depths inside the cells.

Figure 5. Dark field microscopy images of SKW6.4 cells without nanoparticles (A) and incubated with HNS-NB-PEG-antiCD19 for 30 min (B), 2 h (C), 6 h (D), 14 h (E) and 24 h (F).


29


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

Page 30 of 50

Larger, more intense bright spots indicate the agglomeration or aggregation of nanoparticles, a common phenomenon observed for gold nanoparticles after internalization in cellular organelles such as lysosomes or endosomes. Such aggregation of the nanoparticles may be induced by the salts both in the growth medium and in the cytoplasm of the cells 52. In the majority of the cells, the nuclear region can be clearly delimited as a dark zone, without evident intrinsic cellular scattering or signal from nanoparticles. Observation of the nanoparticles using DF technique was facilitated by their presence in high number in cytoplasm (and sometimes aggregated as said above), or in small endocytosis vesicles, results that correlate with those obtained by TEM.

3.7. Cell imaging by SERS Besides DF microscopy, SE(R)RS spectroscopy also proved as being an efficient technique for evaluating the uptake and intracellular distribution of plasmonic nanoparticles 53,54. By providing molecular information through the measurement of characteristic vibrational frequencies present within a sample, the method has demonstrated its versatility in characterizing the composition of complex biochemical mixtures, such as those found in cells

55

. SERS mapping of cellular

populations containing both labeled and label-free nanoparticles by SERS imaging can provide a quantitative method by which the number of intracellular nanoparticles could be monitored in time and upon multiple rounds of cell division

56

. Moreover, it was demonstrated that by

selecting the type of the colloidal substrate, and specifically the wavelength domain of the characteristic nanoparticle plasmonic response one can simultaneously image nanoparticle localization and cellular components distribution which makes SERS imaging as a veritable noninvasive and nondestructive alternative method to TEM 57.


30

Page 31 of 50

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


We have used SERS mapping for imaging uptake and distribution of antiCD19 conjugated NB labeled HNS inside SKW6.4 cells using two laser excitation lines (785 nm and 633 nm). As already proven in the dark field images above, the particles are strong light scatterers; further evidence of nanoparticle uptake was provided also by the bright field images of the cells where some luminous spots can be distinguished from the rest of the cell organelles which are poorly discernible (Figure 6C and G). To confirm that the observed spots are related to nanoparticles, we corroborated the images with the SERS maps on both nanoparticle treated cells and on cells without nanoparticles, as control. Figure 6 presents the confocal Raman maps of selected cells (indicated by rectangles), obtained by plotting the intensity of the 1450 cm-1 CH2, CH3 deformation band characteristic to cellular material recorded at 785 nm (B, D) and 633 nm (F, H) excitation. Other bands characteristic to cellular components such as the 1003 cm-1 phenylalanine ring breathing and the ~1660 cm-1 amide I band

58

are evidenced from the extracted spectra (2, 3, 6, 7 in Figure 6 J).

One can also observe signals specific to the genetic material as are the 785 cm-1 pyrimidine ring breathing and phosphodiester-stretch, and the 1094 cm-1 and 1126 cm-1 phosphate symmetric and antisymmetric stretches, respectively

59,60

, which were more prominent under NIR laser

excitation. The localization of the particles was assigned by plotting the intensity of the 595 cm1

band, which is characteristic to the NB-labeled HNS as corroborated with the spectra of the

colloidal HNS-NB-PEG-antiCD19 (spectra 1 and 5 in Figure 6) and the spectra specific of the HNS-rich areas (spectra 4, 8 in figure 6). Although colloidal nanoparticles present quite a good signal of the reporter molecules under 785 nm excitation, one can observe that SERS mapping following the NB signal from the particles internalized by cells was obtained mostly under 633 nm laser excitation (Figure 6I). A possible explanation for this is that at 633 nm excitation, the


31


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

Page 32 of 50

laser line overlaps the absorption of the NB molecules (Figure 3) and hence their resonant excitation leads to the supplementary enhancement characteristic to Surface Enhanced Resonant Raman Scattering (SERRS) 61. An interesting observation is that the two maps (Figure 6H and I), although plotted over bands that correspond to different materials, both identify the regions specific to the localization of the nanoparticles within the cell.


32

Page 33 of 50

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 6. Bright field reflectivity images of SKW6.4 cells without (A, E) and with (C, G) HNSNB-PEG-antiCD19. Raman/SERS maps of correspondent cells (indicated by quadrants), obtained by plotting the intensity of the 1450 cm-1 CH2, CH3 deformation band recorded at 785


33


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

Page 34 of 50

nm (B, D) and 633 nm (F, H) excitation. SERS map obtained by plotting the intensity of the 595 cm-1 NB band (I). (J) Background subtracted spectra extracted from the marked regions in images B, D, F, H, representing non amplified Raman cell signal (2, 3, 6, 7) and nanoparticlerich areas specific signal (4, 8). Spectra 1 and 5 are representative SERS spectra of the colloidal HNS-NB-PEG-antiCD19.

This result is strongly related to the shape of the SERS spectra extracted from the cells with internalized nanoparticles which, beyond the characteristic signal from the reporter molecules, also present an intensity enhancement of the so called SERS background

62

(data not shown).

This enhancement of the SERS background occurs over the entire measured interval, engulfs most of the relevant Raman and even SERS peaks, and can be observed only in localized regions that correspond to accumulated nanoparticles. Compared to the brightest spots, zones closely adjacent have spectral features combining the aforementioned cases, an overall rise of the spectral intensity over the whole spectral range and the presence of some visible cell-specific peaks. This was observed for both laser excitations and hence it is not excessive to account such feature for mapping the internalization and intracellular localization of HNS-NB-PEG-antiCD19 not only by the signal of the label molecule but also by the intrinsic signal provided by the SERS background.

3.8. Cell imaging by TPE-FLIM As previously reported, gold nanoparticles can emit photoluminescence upon two photon excitation due to the sequential recombination of the photoexcited electrons in the sp conduction band with holes in the d-band of metal surface 63. It is noteworthy that this process is different


34

Page 35 of 50

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


than two-photon absorption in case of fluorophores, which requires near simultaneous absorption of two coherent photons

64

. It was shown that two photon luminescence (TPL) is more

pronounced for noble metal nanoparticles which feature anisotropic shapes with enhanced electromagnetic fields, especially when excited in resonance with the LSPR peak

65,66

.

Additionally, time-resolved fluorescence/luminescence technique under TPE allows the measurement of the lifetime of specimens which permits discrimination between signals having overlapping emissions but with different fluorescence decay times

67

. Therefore, the very short

lifetime of gold nanoparticles can be more clearly delimitated from the anywise weak cellular background at NIR excitation. Herein, TPE-FLIM was used for the cell imaging using specifically targeted antiCD19 conjugated HNS as contrast agents. First, the TPL response of the nanoparticles was measured in solution and the obtained spectrum is presented in Figure S7A from Supporting Information. Figure 7 presents TPE-FLIM of nanoparticle-incubated and control cells at 800 nm excitation wavelength. The images of HNS-NB-PEG-antiCD19 incubated cells show successful internalization and cellular distribution of HNS-NB-PEG-antiCD19 having a very short lifetime, meanwhile only a very slight signal was observed from control cells (with a lifetime of ~ 4 ns) free of nanoparticles. Similar to previous study 22, the fluorescence lifetime measured by TCSPC technique was found to be shorter than 100 ps, comparable with the instrument response function of the TCSPC system (see decay in Figure S7B from Supporting Information). Depth sectioning images of the HNS-NB-PEG-antiCD19 incubated cells (Supporting Information Figure S8) demonstrate intracellular localization of the nanoparticles, which can be visualized in the entire cytoplasmic region at the entire depth of the cells. As the signal intensity can be correlated with the extent of the electromagnetic field generated around nanoparticles, agglomeration of HNS-


35


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

Page 36 of 50

NB-PEG-antiCD19 in intravesicle compartments can facilitate their detection inside cells. It has been shown that plasmonic coupling can induce TPL enhancement in case of spherical noble metal nanoparticles 68,69 and gold nanocubes 70. The authors stated that the plasmon coupling can cause a dynamical charge redistribution with concentrated charges at the gap region which could further enhance the local-field intensity, especially for the resonators with sharp-tip or edgedirected coupling orientation. On the other hand, plasmon resonance may also increase the quantum yield of photoluminescence, which may also contribute to the observed TPL enhancement.

Figure 7. Transmission and corresponding TPE-FLIM images of SKW6.4 control cells (A, B) and incubated with HNS-NB-PEG-antiCD19 nanoparticles (C, D).


36

Page 37 of 50

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


4. CONCLUSIONS

This work successfully demonstrates the applicability of antibody conjugated, Raman reporter labeled hollow gold nanospheres for the in vitro specific detection and multimodal imaging of CD19 positive lymphoma cells via combined microspectroscopic techniques: Dark Field (DF), Surface Enhanced Raman Spectroscopy (SERS), and Two Photon Excited Fluorescence Lifetime Imaging Microscopy (TPE-FLIM) in NIR. The particles were evaluated for SERS efficiency and proved a robust and stable signal. The specificity of the antibody-nanocomplex for the targeted cells was demonstrated by TEM which showed a preferential and abundant internalization of the antibody conjugated particles by the SKW6.4 cancer lymphoblast cells in comparison with control samples of antibody-pretreated cells or nonconjugated particles. Imaging results of the nanoparticle loaded cells by DF microscopy, SERS mapping and TPE-FLIM are in good agreement with those obtained by TEM and prove that HNS-NB-PEG-anti-CD19 are promising agents for specific targeting and noninvasive microspectroscopic imaging of CD19(+) lymphoma cells. Using of more accessible, non invasive, and rapid and microspectroscopic techniques such as those presented here might represent a step forward for the early and accurate diagnostic of lymphoblastic cancers.

ASSOCIATED CONTENT Supporting Information is available free of charge and contains: UV-Vis extinction spectra and TEM image of AgNP; TEM images of HNS5 showing defragmented nanoparticles; SERS spectra of HNS at increasing NB concentrations; Reproducibility of the SERS measurement; SERS spectra of HNS before and after functionalization with the antibody; AntiCD19 antibody


37


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

Page 38 of 50

calibration curve; TPL spectra of HNS and fluorescence lifetime decay of HNS measured inside SKW6.4 cells and transmission and depth sectioning TPE-FLIM images of SKW6.4 cells incubated with HNS-NB-PEG-antiCD19.

Corresponding Author * [email protected] Author Contributions The manuscript was written through contributions of all authors. All authors have given approval to the final version of the manuscript. Notes The authors declare no competing financial interest.

ACKNOWLEDGMENT This work was supported by a grant of the Romanian National Authority for Scientific Research and Innovation, CNCS - UEFISCDI, project number PN-II-RU-TE-2014-4-2426. AnaMaria Craciun acknowledges the financial support of the Romanian National Authority for Scientific Research, CNCS – UEFISCDI, project number PN-II-RU-TE-2014-4-1991.

REFERENCES


38

Page 39 of 50

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)

Wang, K.; Chi, C.; Hu, Z.; Liu, M.; Hui, H.; Shang, W.; Peng, D.; Zhang, S.; Ye, J.; Liu,

H.; Tian, J. Optical Molecular Imaging Frontiers in Oncology: The Pursuit of Accuracy and Sensitivity. Engineering 2015, 1 (3), 309–323. (2)

Kneipp, J.; Kneipp, H.; Wittig, B.; Kneipp, K. Novel Optical Nanosensors for Probing

and Imaging Live Cells. Nanomedicine (N. Y., NY, U. S.). 2010, 6 (2), 214–226. (3)

Dykman, L. A.; Khlebtsov, N. G. Multifunctional Gold-Based Nanocomposites for

Theranostics. Biomaterials 2016, 108, 13–34. (4)

Gao, J.; Xu, B. Applications of Nanomaterials inside Cells. Nano Today 2009, 4 (1), 37–

51. (5)

Lane, L. A.; Qian, X.; Nie, S. SERS Nanoparticles in Medicine: From Label-Free

Detection to Spectroscopic Tagging. Chem. Rev. 2015, 115 (19), 10489–10529. (6)

Boca, S.; Rugina, D.; Pintea, A.; Barbu-Tudoran, L.; Astilean, S. Flower-Shaped Gold

Nanoparticles: Synthesis, Characterization and Their Application as SERS-Active Tags inside Living Cells. Nanotechnology 2011, 22 (5), 055702. (7)

Boca-Farcau, S.; Potara, M.; Simon, T.; Juhem, A.; Baldeck, P.; Astilean, S. Folic Acid-

Conjugated, SERS-Labeled Silver Nanotriangles for Multimodal Detection and Targeted Photothermal Treatment on Human Ovarian Cancer Cells. Mol. Pharmaceutics 2014, 11 (2), 391–399. (8)

Aldea, M.; Florian, I. A.; Kacso, G.; Craciun, L.; Boca, S.; Soritau, O.; Florian, I. S.

Nanoparticles for Targeting Intratumoral Hypoxia: Exploiting a Potential Weakness of Glioblastoma. Pharm. Res. 2016, 33 (9), 2059–2077.


39


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

(9)

Page 40 of 50

Ren, Q.-Q.; Bai, L.-Y.; Zhang, X.-S.; Ma, Z.-Y.; Liu, B.; Zhao, Y.-D.; Cao, Y.-C.

Preparation, Modification, and Application of Hollow Gold Nanospheres. J. Nanomater. 2015, 2015, 1–7. (10) Lu, W.; Huang, Q.; Ku, G.; Wen, X.; Zhou, M.; Guzatov, D.; Brecht, P.; Su, R.; Oraevsky, A.; Wang, L. V.; Li, C. Photoacoustic Imaging of Living Mouse Brain Vasculature Using Hollow Gold Nanospheres. Biomaterials 2010, 31 (9), 2617–2626. (11) Park, J.; Park, J.; Ju, E. J.; Park, S. S.; Choi, J.; Lee, J. H.; Lee, K. J.; Shin, S. H.; Ko, E. J.; Park, I.; Kim, C.; Hwang, J. J.; Lee, J. S.; Song, S. Y.; Jeong, S.-Y.; Choi, E. K. Multifunctional Hollow Gold Nanoparticles Designed for Triple Combination Therapy and CT Imaging. J. Controlled Release 2015, 207, 77–85. (12) Gong, X.; Tang, J.; Ji, Y.; Wu, B.; Wu, H.; Liu, A. Adjustable Plasmonic Optical Properties of Hollow Gold Nanospheres Monolayers and LSPR-Dependent Surface-Enhanced Raman Scattering of Hollow Gold Nanosphere/Graphene Oxide Hybrids. RSC Adv. 2015, 5 (53), 42653–42662. (13) Mahmoud, M. A.; Snyder, B.; El-Sayed, M. A. Surface Plasmon Fields and Coupling in the Hollow Gold Nanoparticles and Surface-Enhanced Raman Spectroscopy. Theory and Experiment †. J. Phys. Chem. C 2010, 114 (16), 7436–7443. (14) Lee, S.; Chon, H.; Lee, M.; Choo, J.; Shin, S. Y.; Lee, Y. H.; Rhyu, I. J.; Son, S. W.; Oh, C. H. Surface-Enhanced Raman Scattering Imaging of HER2 Cancer Markers Overexpressed in Single MCF7 Cells Using Antibody Conjugated Hollow Gold Nanospheres. Biosens. Bioelectron. 2009, 24 (7), 2260–2263.


40

Page 41 of 50

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


(15) Lee, S.; Chon, H.; Lee, J.; Ko, J.; Chung, B. H.; Lim, D. W.; Choo, J. Rapid and Sensitive Phenotypic Marker Detection on Breast Cancer Cells Using Surface-Enhanced Raman Scattering (SERS) Imaging. Biosens. Bioelectron. 2014, 51, 238–243. (16) Zhang, J. Z. Biomedical Applications of Shape-Controlled Plasmonic Nanostructures: A Case Study of Hollow Gold Nanospheres for Photothermal Ablation Therapy of Cancer. J. Phys. Chem. Lett. 2010, 1 (4), 686–695. (17) Hoover, E. E.; Squier, J. A. Advances in Multiphoton Microscopy Technology. Nat. Photonics 2013, 7 (2), 93–101. (18) Durr, N. J.; Larson, T.; Smith, D. K.; Korgel, B. A.; Sokolov, K.; Ben-Yakar, A. TwoPhoton Luminescence Imaging of Cancer Cells Using Molecularly Targeted Gold Nanorods. Nano Lett. 2007, 7 (4), 941–945. (19) Zhao, T.; Shen, X.; Li, L.; Guan, Z.; Gao, N.; Yuan, P.; Yao, S. Q.; Xu, Q.-H.; Xu, G. Q. Gold Nanorods as Dual Photo-Sensitizing and Imaging Agents for Two-Photon Photodynamic Therapy. Nanoscale 2012, 4 (24), 7712–7719. (20) Wu, X.; Wang, J.; Chen, J.-Y. The Effect of Aspect Ratio of Gold Nanorods on Cell Imaging with Two-Photon Excitation. Plasmonics 2013, 8 (2), 685–691. (21) Sironi, L.; Freddi, S.; Caccia, M.; Pozzi, P.; Rossetti, L.; Pallavicini, P.; Donà, A.; Cabrini, E.; Gualtieri, M.; Rivolta, I.; Panariti, A.; D’Alfonso, L.; Collini, M.; Chirico, G. Gold Branched Nanoparticles for Cellular Treatments. J. Phys. Chem. C 2012, 116 (34), 18407– 18418.


41


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

Page 42 of 50

(22) Zhang, Y.; Yu, J.; Birch, D. J. S.; Chen, Y. Gold Nanorods for Fluorescence Lifetime Imaging in Biology. J. Biomed. Opt. 2010, 15 (2), 020504-020504-3. (23) Simon, T.; Potara, M.; Gabudean, A.-M.; Licarete, E.; Banciu, M.; Astilean, S. Designing Theranostic Agents Based on Pluronic Stabilized Gold Nanoaggregates Loaded with Methylene Blue for Multimodal Cell Imaging and Enhanced Photodynamic Therapy. ACS Appl. Mater. Interfaces 2015, 7 (30), 16191–16201. (24) Li, H.; Xia, H.; Wang, D.; Tao, X. Simple Synthesis of Monodisperse, Quasi-Spherical, Citrate-Stabilized Silver Nanocrystals in Water. Langmuir 2013, 29 (16), 5074–5079. (25) Qian, X.; Peng, X.-H.; Ansari, D. O.; Yin-Goen, Q.; Chen, G. Z.; Shin, D. M.; Yang, L.; Young, A. N.; Wang, M. D.; Nie, S. In Vivo Tumor Targeting and Spectroscopic Detection with Surface-Enhanced Raman Nanoparticle Tags. Nat. Biotechnol. 2008, 26 (1), 83–90. (26) Xia, X.; Wang, Y.; Ruditskiy, A.; Xia, Y. 25th Anniversary Article: Galvanic Replacement: A Simple and Versatile Route to Hollow Nanostructures with Tunable and WellControlled Properties. Adv. Mater. 2013, 25 (44), 6313–6333. (27) Chen, J.; Wiley, B.; Li, Z.-Y.; Campbell, D.; Saeki, F.; Cang, H.; Au, L.; Lee, J.; Li, X.; Xia, Y. Gold Nanocages: Engineering Their Structure for Biomedical Applications. Adv. Mater. 2005, 17 (18), 2255–2261. (28) Preciado-Flores, S.; Wang, D.; Wheeler, D. A.; Newhouse, R.; Hensel, J. K.; Schwartzberg, A.; Wang, L.; Zhu, J.; Barboza-Flores, M.; Zhang, J. Z. Highly Reproducible Synthesis of Hollow Gold Nanospheres with near Infrared Surface Plasmon Absorption Using PVP as Stabilizing Agent. J. Mater. Chem. 2011, 21 (7), 2344–2350.


42

Page 43 of 50

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


(29) Zhang, Y.; Newton, B.; Lewis, E.; Fu, P. P.; Kafoury, R.; Ray, P. C.; Yu, H. Cytotoxicity of Organic Surface Coating Agents Used for Nanoparticles Synthesis and Stability. Toxicol. Vitro Int. J. Publ. Assoc. BIBRA 2015, 29 (4), 762–768. (30) Vongsavat, V.; Vittur, B. M.; Bryan, W. W.; Kim, J.-H.; Lee, T. R. Ultrasmall Hollow Gold–Silver Nanoshells with Extinctions Strongly Red-Shifted to the Near-Infrared. ACS Appl. Mater. Interfaces 2011, 3 (9), 3616–3624. (31) Sun, Y.; Mayers, B. T.; Xia, Y. Template-Engaged Replacement Reaction: A One-Step Approach to the Large-Scale Synthesis of Metal Nanostructures with Hollow Interiors. Nano Lett. 2002, 2 (5), 481–485. (32) Prevo, B. G.; Esakoff, S. A.; Mikhailovsky, A.; Zasadzinski, J. A. Scalable Routes to Gold Nanoshells with Tunable Sizes and Response to Near-Infrared Pulsed-Laser Irradiation. Small 2008, 4 (8), 1183–1195. (33) Wan, D.; Chen, H.-L.; Lin, Y.-S.; Chuang, S.-Y.; Shieh, J.; Chen, S.-H. Using Spectroscopic Ellipsometry to Characterize and Apply the Optical Constants of Hollow Gold Nanoparticles. ACS Nano 2009, 3 (4), 960–970. (34) Polavarapu, L.; Zanaga, D.; Altantzis, T.; Rodal-Cedeira, S.; Pastoriza-Santos, I.; PérezJuste, J.; Bals, S.; Liz-Marzán, L. M. Galvanic Replacement Coupled to Seeded Growth as a Route for Shape-Controlled Synthesis of Plasmonic Nanorattles. J. Am. Chem. Soc. 2016, 138 (36), 11453–11456.


43


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

Page 44 of 50

(35) Goris, B.; Polavarapu, L.; Bals, S.; Van Tendeloo, G.; Liz-Marzán, L. M. Monitoring Galvanic Replacement Through Three-Dimensional Morphological and Chemical Mapping. Nano Lett. 2014, 14 (6), 3220–3226. (36) Li, C.-H.; Jamison, A. C.; Rittikulsittichai, S.; Lee, T.-C.; Lee, T. R. In Situ Growth of Hollow Gold–Silver Nanoshells within Porous Silica Offers Tunable Plasmonic Extinctions and Enhanced Colloidal Stability. ACS Appl. Mater. Interfaces 2014, 6 (22), 19943–19950. (37) Merican, Z.; Schiller, T. L.; Hawker, C. J.; Fredericks, P. M.; Blakey, I. Self-Assembly and Encoding of Polymer-Stabilized Gold Nanoparticles with Surface-Enhanced Raman Reporter Molecules. Langmuir 2007, 23 (21), 10539–10545. (38) Boca, S. C.; Astilean, S. Detoxification of Gold Nanorods by Conjugation with Thiolated Poly(ethylene Glycol) and Their Assessment as SERS-Active Carriers of Raman Tags. Nanotechnology 2010, 21 (23), 235601. (39) Gabudean, A. M.; Biro, D.; Astilean, S. Hybrid Plasmonic Platforms Based on SilicaEncapsulated Gold Nanorods as Effective Spectroscopic Enhancers for Raman and Fluorescence Spectroscopy. Nanotechnology 2012, 23 (48), 485706. (40) Jazayeri,

M.

H.;

Amani,

H.;

Pourfatollah,

A.

A.;

Pazoki-Toroudi,

H.;

Sedighimoghaddam, B. Various Methods of Gold Nanoparticles (GNPs) Conjugation to Antibodies. Sens. Bio-Sens. Res. 2016, 9, 17–22. (41) Eghtedari, M.; Liopo, A. V.; Copland, J. A.; Oraevsky, A. A.; Motamedi, M. Engineering of Hetero-Functional Gold Nanorods for the in Vivo Molecular Targeting of Breast Cancer Cells. Nano Lett. 2009, 9 (1), 287–291.


44

Page 45 of 50

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


(42) Liu, T.; Thierry, B. A Solution to the PEG Dilemma: Efficient Bioconjugation of Large Gold Nanoparticles for Biodiagnostic Applications Using Mixed Layers. Langmuir 2012, 28 (44), 15634–15642. (43) Marega, R.; Karmani, L.; Flamant, L.; Nageswaran, P. G.; Valembois, V.; Masereel, B.; Feron, O.; Borght, T. V.; Lucas, S.; Michiels, C.; Gallez, B.; Bonifazi, D. AntibodyFunctionalized Polymer-Coated Gold Nanoparticles Targeting Cancer Cells: An in Vitro and in Vivo Study. J. Mater. Chem. 2012, 22 (39), 21305. (44) Tejamaya, M.; Römer, I.; Merrifield, R. C.; Lead, J. R. Stability of Citrate, PVP, and PEG Coated Silver Nanoparticles in Ecotoxicology Media. Environ. Sci. Technol. 2012, 46 (13), 7011–7017. (45) Smith, J. N.; Meadows, J.; Williams, P. A. Adsorption of Polyvinylpyrrolidone onto Polystyrene Latices and the Effect on Colloid Stability. Langmuir 1996, 12 (16), 3773–3778. (46) Willets, K. A.; Duyne, R. P. V. Localized Surface Plasmon Resonance Spectroscopy and Sensing. Annu. Rev. Phys. Chem. 2007, 58 (1), 267–297. (47) Murphy, C. J.; Gole, A. M.; Stone, J. W.; Sisco, P. N.; Alkilany, A. M.; Goldsmith, E. C.; Baxter, S. C. Gold Nanoparticles in Biology: Beyond Toxicity to Cellular Imaging. Acc. Chem. Res. 2008, 41 (12), 1721–1730. (48) Nativo, P.; Prior, I. A.; Brust, M. Uptake and Intracellular Fate of Surface-Modified Gold Nanoparticles. ACS Nano 2008, 2 (8), 1639–1644. (49) Quentmeier, H.; Martelli, M. P.; Dirks, W. G.; Bolli, N.; Liso, A.; MacLeod, R. A. F.; Nicoletti, I.; Mannucci, R.; Pucciarini, A.; Bigerna, B.; Martelli, M. F.; Mecucci, C.; Drexler, H.


45


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

Page 46 of 50

G.; Falini, B. Cell Line OCI/AML3 Bears Exon-12 NPM Gene Mutation-A and Cytoplasmic Expression of Nucleophosmin. Leukemia 2005, 19 (10), 1760–1767. (50) Wax, A.; Sokolov, K. Molecular Imaging and Darkfield Microspectroscopy of Live Cells Using Gold Plasmonic Nanoparticles. Laser Photonics Rev. 2009, 3 (1–2), 146–158. (51) Jain, P. K.; Lee, K. S.; El-Sayed, I. H.; El-Sayed, M. A. Calculated Absorption and Scattering Properties of Gold Nanoparticles of Different Size, Shape, and Composition: Applications in Biological Imaging and Biomedicine. J. Phys. Chem. B 2006, 110 (14), 7238– 7248. (52) El-Sayed, I. H.; Huang, X.; El-Sayed, M. A. Surface Plasmon Resonance Scattering and Absorption of Anti-EGFR Antibody Conjugated Gold Nanoparticles in Cancer Diagnostics: Applications in Oral Cancer. Nano Lett. 2005, 5 (5), 829–834. (53) von Maltzahn, G.; Centrone, A.; Park, J.-H.; Ramanathan, R.; Sailor, M. J.; Hatton, T. A.; Bhatia, S. N. SERS-Coded Gold Nanorods as a Multifunctional Platform for Densely Multiplexed Near-Infrared Imaging and Photothermal Heating. Adv. Mater. 2009, 21 (31), 3175– 3180. (54) Kneipp, K.; Haka, A. S.; Kneipp, H.; Badizadegan, K.; Yoshizawa, N.; Boone, C.; Shafer-Peltier, K. E.; Motz, J. T.; Dasari, R. R.; Feld, M. S. Surface-Enhanced Raman Spectroscopy in Single Living Cells Using Gold Nanoparticles. Appl. Spectrosc. 2002, 56 (2), 150–154. (55) De Gelder, J.; De Gussem, K.; Vandenabeele, P.; Moens, L. Reference Database of Raman Spectra of Biological Molecules. J. Raman Spectrosc. 2007, 38 (9), 1133–1147.


46

Page 47 of 50

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


(56) Sirimuthu, N. M. S.; Syme, C. D.; Cooper, J. M. Monitoring the Uptake and Redistribution of Metal Nanoparticles during Cell Culture Using Surface-Enhanced Raman Scattering Spectroscopy. Anal. Chem. 2010, 82 (17), 7369–7373. (57) Potara, M.; Boca, S.; Licarete, E.; Damert, A.; Alupei, M.-C.; Chiriac, M. T.; Popescu, O.; Schmidt, U.; Astilean, S. Chitosan-Coated Triangular Silver Nanoparticles as a Novel Class of Biocompatible, Highly Sensitive Plasmonic Platforms for Intracellular SERS Sensing and Imaging. Nanoscale 2013, 5 (13), 6013–6022. (58) Rygula, A.; Majzner, K.; Marzec, K. M.; Kaczor, A.; Pilarczyk, M.; Baranska, M. Raman Spectroscopy of Proteins: A Review: Raman Spectroscopy of Proteins. J. Raman Spectrosc. 2013, 44 (8), 1061–1076. (59) Camp Jr, C. H.; Lee, Y. J.; Heddleston, J. M.; Hartshorn, C. M.; Walker, A. R. H.; Rich, J. N.; Lathia, J. D.; Cicerone, M. T. High-Speed Coherent Raman Fingerprint Imaging of Biological Tissues. Nat. Photonics 2014, 8 (8), 627–634. (60) Deng, H.; Bloomfield, V. A.; Benevides, J. M.; Thomas, G. J. Dependence of the Raman Signature of genomicB-DNA on Nucleotide Base Sequence. Biopolymers 1999, 50 (6), 656–666. (61) McNay, G.; Eustace, D.; Smith, W. E.; Faulds, K.; Graham, D. Surface-Enhanced Raman Scattering (SERS) and Surface-Enhanced Resonance Raman Scattering (SERRS): A Review of Applications. Appl. Spectrosc. 2011, 65 (8), 825–837. (62) Farcau, C.; Astilean, S. Evidence of a Surface Plasmon-Mediated Mechanism in the Generation of the SERS Background. Chem. Commun. 2011, 47 (13), 3861.


47


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

Page 48 of 50

(63) Dulkeith, E.; Niedereichholz, T.; Klar, T. A.; Feldmann, J.; von Plessen, G.; Gittins, D. I.; Mayya, K. S.; Caruso, F. Plasmon Emission in Photoexcited Gold Nanoparticles. Phys. Rev. B 2004, 70 (20), 205424. (64) So, P. T. C.; Dong, C. Y.; Masters, B. R.; Berland, K. M. Two-Photon Excitation Fluorescence Microscopy. Annu. Rev. Biomed. Eng. 2000, 2 (1), 399–429. (65) Mohamed, M. B.; Volkov, V.; Link, S.; El-Sayed, M. A. The `lightning’ Gold Nanorods: Fluorescence Enhancement of over a Million Compared to the Gold Metal. Chem. Phys. Lett. 2000, 317 (6), 517–523. (66) Siddiquee, A. M.; Taylor, A. B.; Syed, S.; Lim, G.-H.; Lim, B.; Chon, J. W. M. Measurement of Plasmon-Mediated Two-Photon Luminescence Action Cross Sections of Single Gold Bipyramids, Dumbbells, and Hemispherically Capped Cylindrical Nanorods. J. Phys. Chem. C 2015, 119 (51), 28536–28543. (67) Marcu, L. Fluorescence Lifetime Techniques in Medical Applications. Ann. Biomed. Eng. 2012, 40 (2), 304–331. (68) Guan, Z.; Polavarapu, L.; Xu, Q.-H. Enhanced Two-Photon Emission in Coupled Metal Nanoparticles Induced by Conjugated Polymers. Langmuir 2010, 26 (23), 18020–18023. (69) Han, F.; Guan, Z.; Tan, T. S.; Xu, Q.-H. Size-Dependent Two-Photon Excitation Photoluminescence Enhancement in Coupled Noble-Metal Nanoparticles. ACS Appl. Mater. Interfaces 2012, 4 (9), 4746–4751. (70) Guan, Z.; Li, S.; Cheng, P. B. S.; Zhou, N.; Gao, N.; Xu, Q.-H. Band-Selective CouplingInduced Enhancement of Two-Photon Photoluminescence in Gold Nanocubes and Its


48

Page 49 of 50

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


Application as Turn-on Fluorescent Probes for Cysteine and Glutathione. ACS Appl. Mater. Interfaces 2012, 4 (10), 5711–5716.


49


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

Page 50 of 50

Table of Contents Graphic


50

Antibody Conjugated, Raman Tagged Hollow GoldâSilver

Recommend Documents

Antibody Conjugated, Raman Tagged Hollow GoldâSilver