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Development and optimization of near-IR contrast agents for immune cell tracking

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Gold nanorods (NRs) are attractive for in vivo imaging due to their high optical cross-sections and tunable absorbance. However, the feasibility of using NRs for cell tracking has not been fully explored. Here, we synthesized dye doped silica-coated NRs as multimodal contrast agents for imaging of macrophages - immune cells which play an important role in cancer and cardiovascular diseases. We showed the importance of silica coating in imaging of NR-labeled cells. Photoacoustic (PA) imaging of NRs labeled macrophages showed high sensitivity. Therefore, these results provide foundation for applications of silica-coated NRs and PA imaging in tracking of immune cells.
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Development and optimization of near-IR
contrast agents for immune cell tracking
Pratixa P. Joshi,1 Soon Joon Yoon,1 Yun-Sheng Chen,1 Stanislav Emelianov,1,2 and
Konstantin V. Sokolov1,2,*
1 Department of Biomedical Engineering, University of Texas at Austin, 107 W Dean Keeton St, Austin, Texas 78712,
USA
2 Department of Imaging Physics, M.D. Anderson Cancer Center, Houston, Texas 77030, USA
* kostia@mail.utexas.edu
Abstract: Gold nanorods (NRs) are attractive for in vivo imaging due to
their high optical cross-sections and tunable absorbance. However, the
feasibility of using NRs for cell tracking has not been fully explored. Here,
we synthesized dye doped silica-coated NRs as multimodal contrast agents
for imaging of macrophages – immune cells which play an important role in
cancer and cardiovascular diseases. We showed the importance of silica
coating in imaging of NR-labeled cells. Photoacoustic (PA) imaging of NRs
labeled macrophages showed high sensitivity. Therefore, these results
provide foundation for applications of silica-coated NRs and PA imaging in
tracking of immune cells.
©2013 Optical Society of America
OCIS codes: (170.0170) Medical optics and biotechnology; (170.5120) Photoacoustic imaging;
(170.2655) Functional monitoring and imaging; (170.2520) Fluorescence microscopy;
(060.4230) Multiplexing; (160.4236) Nanomaterials
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Introduction
The tracking of immune cells is essential to understand the regulatory role of immune cells in
pathological conditions such as cancer and cardiovascular diseases [1] that can lead to
development of novel therapeutic approaches [2, 3]. Moreover, using immune cells as
transport vehicles for imaging and therapeutic agents can facilitate imaging and treatment of
cancer cells in vasculature inaccessible areas. Indeed, one of the major barriers in cancer
treatment is inaccessibility of the avascular necrotic core of a tumor by traditional drugs.
Cancer cells from these hypoxic regions can survive therapy and migrate to secondary sites
causing cancer metastasis [4, 5]. Immune cells can penetrate the tumor necrotic core by
following chemokines released by cancer cells that can be used as a strategy to deliver a
therapeutic payload [6, 7]. Optimization of these new approaches to treatment of cancer and
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1 November 2013 | Vol. 4, No. 11 | DOI:10.1364/BOE.4.002609 | BIOMEDICAL OPTICS EXPRESS 2611
other diseases requires development of sensitive imaging methods for in vivo tracking of
immune cells.
A direct approach for immune cell tracking is to load cells with contrast agents which are
suitable for in vivo imaging modalities. Recently, we have demonstrated that loading gold
nanoparticles into endosomal compartments of stem cells does not alter cellular functions
including cell viability and differentiation [8]. Furthermore, we have demonstrated that stem
cells labeled with gold nanoparticles can be tracked in vivo using combined photoacoustic
(PA) and ultrasound (US) imaging with penetration depth of several centimeters [9]. In PA
imaging, the tissue is irradiated with nanosecond pulses of low energy laser light. Then,
through the processes of optical absorption followed by thermal expansion, broadband
acoustic waves are generated within the irradiated volume. Using an ultrasound detector,
these waves can be detected and spatially resolved to provide an image. Furthermore, PA
imaging can be easily combined with US imaging because both imaging modalities can share
the same ultrasound sensor and associated receiver electronics [10–12]. In this combination,
PA signal from nanoparticle labeled immune cells can be put in context of surrounding tissue
anatomy using US imaging. Therefore, combined photoacoustic and ultrasound imaging can
provide complementary information.
Gold nanoparticles, including nanospheres [13–20], nanoshells [21–24], nanocages [25–
28], and nanorods [29–37] are of increasing interest to biomedical engineers due to their
biocompatibility, facile surface modification and high optical cross sections which can
provide strong signal in imaging modalities based on optical contrast. The optical properties
of gold nanoparticles can be modulated by changing their size and shape. For example,
plasmon resonance of gold nanorods is easily tuned in the red to near infrared (NIR) spectral
region by changing their aspect ratio [38], facilitating simultaneous imaging of multiple
biomarkers [33, 39]. Strong NIR extinction cross-sections of nanorods have been used for
two-photon luminescence [29, 30] and photoacoustic [12,40–42] imaging of thick biological
samples as well as for photothermal destruction of cancer cells [43–46]. Thus, anisotropic
gold nanorods provide a convenient combination of properties for biomedical applications
including the possibility of cell tracking using PA imaging [47–49]. However, the optical
absorbance spectra of gold nanorods broaden when they interact with live cells [36]. This
effect is due to plasmon resonance coupling of closely spaced nanoparticles upon cellular
uptake and it diminishes the ability to simultaneously image multiple cell populations labeled
with different nanorods. In addition, recent reports showed that cells more readily uptake
spherical nanoparticles as compared to rod-shaped particles [50]. As spherical nanoparticles
do not have the desired near infrared absorption necessary for in vivo imaging, increasing
cellular uptake of nanorods is desirable.
Here we synthesized silica-coated gold nanorods to address the challenges associated with
applications of rod-shaped gold nanoparticles in cell tracking. The silica coating changes the
shape of nanorods from cylindrical to spheroidal. We demonstrated that the silica-shell
reduces plasmon resonance coupling between nanorods upon uptake by cells and, thus, can
facilitate multiplex imaging using nanorods. In addition, the silica shell was doped with
fluorescent dye to enable multimodal imaging with fluorescence and PA imaging modalities.
This combination enables in-depth in vivo imaging of labeled cells using photoacoustics
followed by a high-resolution ex vivo fluorescence imaging for validation and detailed
characterization of in vivo imaging data.
Experimental methods
Synthesis of PEG-coated gold nanorods (mPEG-NRs): Gold nanorods (NRs) were
synthesized by seed mediated growth mechanism using the previously published protocol
[51]. The as-prepared nanorods were washed twice in water by centrifugation at 18000g for
45 minutes. The final NRs pellet was dispersed in water and stored at room temperature at an
optical density (OD) of 15. Before silica-coating, the cetyl trimethyl ammonium bromide
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Received 14 Aug 2013; revised 30 Sep 2013; accepted 21 Oct 2013; published 24 Oct 2013
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1 November 2013 | Vol. 4, No. 11 | DOI:10.1364/BOE.4.002609 | BIOMEDICAL OPTICS EXPRESS 2612
(CTAB) layers on NRs were replaced with methoxy-PEG-thiol (mPEG-SH). One milliliter of
1 mg/mL mPEG-SH (5kDa) in water was mixed with 1 mL of CTAB-NRs (OD 15) and
sonicated using a water bath sonicator (Branson 1510R-MT) for 1 minute at room
temperature. After sonication, the NRs were stirred overnight in presence of mPEG-SH to
complete the replacement of CTAB. Free CTAB and mPEG-SH molecules were removed by
centrifugation through 100kDa centrifugal filters at 2500g for 15 minutes. Then, the NRs
were washed one more time in water to remove any residual mPEG and CTAB. mPEG-NRs
were dispersed in 1 mL water and stored at room temperature.
Synthesis of silica-coated gold nanorods: Fluorescent silica-coated gold nanorods were
synthesized by covalent attachment of a fluorescent dye (Rhodamine) to the silica matrix
(Fig. 1). Silica shells of controlled thickness were deposited on the PEG-coated nanorods
using a modified Stöber method [52]. One milliliter of mPEG-NRs (OD 15) was mixed with
1.5 mL of isopropanol. The solution was continuously stirred while adding 0.6 mL of 3 vol %
tetraethylorthosilicate (TEOS) and 0.625 mL of 3.84 vol % ammonium hydroxide (NH4OH).
Rhodamine (Rh) dye doped silica-coated nanorods (RhSilicaNRs) were synthesized by first
covalently attaching Rh isothiocyanate dye to [3-(2-
Aminoethylamino)propyl]trimethoxysilane (AEAPTMS) [53]. Briefly, 15 µL of [3-(2-
Aminoethylamino)propyl]trimethoxysilane was stirred with 10 mg Rhodamine B
isothiocyanate in 2.5 mL of anhydrous isopropanol (IPA) for 48 hours under nitrogen. This
solution was five times diluted in anhydrous IPA just prior to using. One milliliter of mPEG-
NRs (OD 15) was mixed with 1.5 mL of isopropanol. The solution was continuously stirred
while adding 0.6 mL of 3 vol % tetraethylorthosilicate (TEOS) and 0.625 mL of 3.84 vol %
ammonium hydroxide (NH4OH). After 30 minutes, four aliquots (12.5 µL each) of the diluted
Rhodamine-AEAPTMS solution were added to the reaction mixture (TEOS:Rh-AEAPTMS
molar ratio 1080:1). The aliquots were separated by 10 minutes time intervals. After 2 hours
of stirring at room temperature, the silica-coated nanorods were collected by centrifugation at
700g for 15 minutes using 100 kDa MWCO centrifugal filters. Both plain SilicaNRs and
RhSilicaNRs were washed three times with water prior to use.
Fig. 1. Schematic of approach used to synthesize fluorescent silica-coated nanorods.
Synthesis of polymer-coated nanorods: Polymer-coated nanorods were synthesized by
mixing 1 mL of CTAB-coated gold nanorods (OD 10) with 1 mL of polystyrene sulfonate
(PSS) solution (10 mg PSS per 1 mL of 1 mM NaCl) [30, 54]. The solution was kept on a
shaker for 30 minutes and then centrifuged at 18000g for 15 minutes. The nanorods
containing pellet was redispersed in 1 mL of 1 mM NaCl solution and was mixed with 1 mL
of PSS solution. The solution was placed on a shaker for 30 minutes and then centrifuged to
collect the nanorods coated with PSS. The PSS-nanorods were coated with Poly(allylamine
hydrochloride) (PAH) using the same procedure. PSS-nanorods pellet was dispersed in a 1
mL NaCl solution (1 mM) and mixed with 1 mL of PAH solution (10 mg PAH per mL of 1
mM NaCl). After 30 minutes of shaking, the nanorods pellet was collected using
centrifugation and dispersed in a salt solution. The PAH coating procedure was repeated once
more to obtain PAH-PSS-nanorods. The final nanorods pellet was dispersed in 1 mM NaCl
solution.
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Received 14 Aug 2013; revised 30 Sep 2013; accepted 21 Oct 2013; published 24 Oct 2013
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1 November 2013 | Vol. 4, No. 11 | DOI:10.1364/BOE.4.002609 | BIOMEDICAL OPTICS EXPRESS 2613
Cell viability study: The viability of monocyte/macrophage (RAW 264.7) cells was
performed after incubating the cells with RhSilicaNRs at a given concentration in phenol-free
DMEM cell culture media for 18 hours at 37°C. As the nanorods-loaded cells need to be
viable for a long period of time for successful tracking of cells, extended cell viability tests
were performed by culturing cells in regular cell culture media for an additional 24 to 48
hours after the incubation of cells with silica-nanorods. To perform cell viability study, cells
were washed once with PBS and a background absorbance was measured at 490 nm using
BioTek Synergy HT UV-Vis spectrophotometer. Then MTS reagent (mixture of MTS and
PMS prepared in cell culture media) was added to the cells. After 3 hours of incubation with
MTS solution, absorbance at 490 nm was measured again and the background absorbance
values were subtracted in order to determine the number of metabolically active live cells in a
sample.
Two mouse-macrophage cell lines, RAW 264.7 and P388D1, were used for cell labeling
and characterization throughout this study. While RAW 264.7 cells are highly phagocytic
macrophage cells, P388D1 cells have an inferior phagocytic activity [55, 56]. In order to
achieve camparable loading of nanorods in both cell lines, we used higher dosage of 1x107
NRs/cell for P388D1 cells as compared to 7x104 NRs/cell for RAW 264.7. The cells were
incubated with nanorods for 18 hours at 37°C in cell culture media. Labeled cells were
washed with cell culture media to remove any unbound nanorods and re-suspended in phenol
free media to measure absorbance. Absorbance spectra of unlabeled cells were subtracted
from the spectra of nanorod-loaded cells to obtain the contribution from nanorods. Before the
measurements, cells were counted using a standard hemocytometer in the presence of trypan
blue; trypan blue is a vital stain that selectively colors dead and lysed cells. More than 95% of
cells were vital during the absorption measurements.
Tissue mimicking phantom preparation: Gelatin based tissue-mimicking phantoms with
cell inclusions were fabricated for combined photoacoustic and ultrasound imaging. To
simulate tissue background, 8wt% gelatin solution with 0.2wt% of 15 µm silica particles was
used to prepare the base and top layers which encapsulated the cell inclusions. Suspensions of
P388D1 cells only and P388D1 cells loaded with RhSilicaNRs were mixed with the same
volume of 16wt% gelatin solution at 37°C. Specifically, five inclusions of P388D1 cells
loaded with RhSilicaNRs were prepared with cells concentrations 5 × 106, 2.5 × 106, 1.25 ×
106, 2.5 × 105, and 1.25 × 105 cells/mL. For the cells only inclusion, 5 × 106 cells/mL cell
concentration was used. For phantom preparation, suspension of silica particles in gelatin at
37°C was poured in a petri dish and was cooled down at 4°C. During the cool down step,
some silica particles settle down that results in a reduced ultrasound contrast in the top part of
the phantom. Then, suspensions of cells in gelatin were placed on the solidified base layer
and were allowed cool down. Finally, the top layer of gelatin/silica particles was added to
complete the phantom.
Photoacoustic imaging set-up: The photoacoustic signal from the tissue-mimicking
phatom was obtained using a Vevo® 2100 LAZR imaging system (VisualSonics, Inc.,
Toronto, Canada) and an array ultrasound transducer (LZ250, VisualSonics, Inc.) operating at
20 MHz center frequency. A laser beam with 5 ns pulses and 20 Hz repetition rate was
generated by a tunable OPO laser system pumped by Nd:YAG laser. The wavelength of the
light in this experiment was matched to the peak optical absorption wavelength of the silica-
nanorods, which is 780 nm. The laser beam was delivered through an optical fiber bundle
integrated with the ultrasound array transducer. A mechanical system was used to translate
the transducer to allow for collecting multiple cross-sectional US/PA images of the inclusions
in steps of 114 μm. In order to perform a quantitative photoacoustic signal comparision
between each inclusion, the fluence of the laser was maintained at 10 mJ/cm2, which is below
the damage threshold for silica-nanorods [52]. To compensate amplitude of PA signal for
laser fluctuations, laser fluence was recorded by the VEVO 2100 LAZR imaging system
during PA imaging and PA signals were normalized by corresponding laser fluences in post-
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(C) 2013 OSA
1 November 2013 | Vol. 4, No. 11 | DOI:10.1364/BOE.4.002609 | BIOMEDICAL OPTICS EXPRESS 2614
processing. PA signal intensity of each inclusion was measured by averaging PA signals from
11 different cross-sectional images of the inclusion.
Results and discussion
The RhSilicaNRs were characterized using TEM and fluorescence emission. Transmission
electron microscopy showed a uniform silica layer on the coated nanorods (Fig. 2(a)). The
fluorescence emission spectrum of the RhSilicaNRs indicated successful encapsulation of
fluorescent dye in the silica-coated nanorods (Fig. 2(b)).
Fig. 2. (a) Transmission Electron Microscopy of Rhodamine silica-coated nanorods
(RhSilicaNRs), scale bar: 100 nm; (b) fluorescence emission spectra of RhSilicaNRs (ex 540
nm) and plain silica-coated nanorods (SilicaNRs).
Cellular tracking requires long term viability of nanoparticle-loaded cells. Here we used a
monocyte macrophage cell line (RAW 264.7) to evaluate extended viability of cells loaded
with flourescent silica-coated and PEG-coated nanorods following 18 hours of incubation.
The nanorod-loaded macrophages were grown in cell culture media for 24 and 48 hours
before testing cell viability using MTS assay that measures metabolic activity of live cell and
also reflects the number of viable cells. Most of macrophages loaded with RhSilicaNRs
showed no change or a slight increase in metabolic activity possibly due to macrophage
activation by the nanoparticle uptake. A slight decrease in viability of cells loaded with
mPEG-NRs after 24 hours culture can be attributed to a small amount of residual CTAB
molecules which are present after ligand exchange with PEG [36].
Fig. 3. Viability of mouse monocyte macrophage cell line (RAW 264.7) loaded with mPEG-
NRs and RhSilicaNRs immediately after 18 hrs of incubation with nanoparticles and 24 and 48
hours of cell culture after the incubation. RhSilicaNRs show no toxicity during extended cell
culture of labeled cells. Statistically different results based on student t-test (p-value < 0.01)
are identified by (*).
Silica-coated nanorods exhibit negligible changes in the longitudinal peak position and the
full-width at half-maximum (FWHM) values after uptake by two macrophage cells lines -
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(C) 2013 OSA
1 November 2013 | Vol. 4, No. 11 | DOI:10.1364/BOE.4.002609 | BIOMEDICAL OPTICS EXPRESS 2615
RAW 264.7 and P388D1 (Fig. 4(a), 4(b) and Table 1). In contrast, nanorods coated using
layer-by-layer deposition of polystyrene sulfonate (PSS) and poly(allylamine hydrochloride)
(PAH) polymers [57] show significant change in the optical absorption upon cellular uptake
(Fig. 4(c), Table 1). Note that RhSilicaNRs, SilicaNRs and PAH-PSS-NRs have different
longitudinal peak maxima. These results demonstrate that silica coating of ca. 30 nm provides
sufficient spacing to prevent plasmon resonance coupling of nanorods after they are uptaken
by live cells. Since silica-coated gold nanorods do not change optical spectra inside cells, we
used extinction coefficients of the nanorods in suspension to determine nanorod loading of
cells. RAW 264.7 and P388D1 cells accumulated ca. 104 and 2x104 silica-coated nanorods
per cell, respectively. Cells loaded with RhSilicaNRs can be readily viasualized using
fluorescence imaging (Fig. 5(e) and 5(f)). Negligible nanorod-specific contrast was observed
in the dark-field images of RhSilicaNRs-loaded cells due to low sensitivity of our optical
microscope to NIR logitudinal resonances of the nanorods used in this study (Fig. 5(b) and
5(c)). Thus, fluorescence imaging provides a convenient way of identifying cells labeled with
dye-dopped silica-coated gold nanorods.
Previous literature reports have shown a good contrast in dark-field imaging of gold
nanorods with a longitudinal peak in the visible range [58, 59]. It has been also reported by us
[36] and other groups [43, 60, 61] that cells loaded or labeled with gold nanorods coated by a
thin layer of organic ligands and targeting biomolecules produce detectable contrast in dark-
field images as compared to unlabeled cells. We attribute this contrast to aggregation of gold
nanorods upon cellular uptake that results in an overall increase in scattering in the visible
region due to plasmon resonance coupling between closely spaced nanoparticles. In the
present study, silica coating around nanorods significantly diminishes the effect of plasmon
resonance coupling as described above and, therefore, silica-coated nanorods do not provide
significant contrast in the presence of endogenous scattering background from cells.
Fig. 4. Extinction spectra of: (a) RhSilicaNRs in suspension and P388D1 cells loaded with
RhSilicaNRs; (b) SilicaNRs in suspension and RAW 264.7 cells loaded with SilicaNRs; (c)
polymer-coated nanorods (PAH-PSS-NRs) in suspension and RAW 264.7 cells loaded with
PAH-PSS-NRs. Longitudinal peak position and extinction spectra of silica-coated nanorods do
not undergo significant changes after cell uptake in contrast to polymer-coated NRs.
Table 1. Characterization of absorbance spectra of nanorods and cells-loaded with
nanorods; FMHM – full width at half maximum.
FWHM, nm % Increase
in FWHM
Longitudinal Peak
Wavelength, nm
RhSilicaNRs 120 782
P388D1 cells with RhSilicaNRs 138 15.00% 778
SilicaNRs 83 704
RAW 264.7 cells with SilicaNRs 87 4.82% 700
PAH-PSS-NRs 139 725
RAW 264.7 cells with PAH-PSS-NRs 222 59.71% 800
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(C) 2013 OSA
1 November 2013 | Vol. 4, No. 11 | DOI:10.1364/BOE.4.002609 | BIOMEDICAL OPTICS EXPRESS 2616
Fig. 5. Dark-field (a, b, c) and fluorescence (d, e, f) images of RAW 264.7 cells alone (a, d),
RAW 264.7 cells loaded with RhSilicaNRs (b, e) and P388D1 cells loaded with RhSilicaNRs
(c, f). Scale bar 50 µm. Images were acquired with Leica DM600 upright microscope using
20x 0.5 NA objective. Fluorescence imaging was performed using Cy3 filter cube, ex/em
555/590 nm.
Having successfully loaded cells with silica-coated nanorods, the next step was to
evaluate the feasibility of sensitive detection of the nanorod-loaded cells using PA imaging.
To this end, inclusions containing different concentrations of P388D1 cells loaded with
RhSilicaNRs were incorporated in the tissue-mimicking phantoms and multiple cross-
sectional PA images of each inclusion were collected at 780 nm wavelength, which
corresponds to the longitudinal plasmon resonance peak of the nanorods (Fig. 6). Although
the same matrix was used to prepare the top and bottom layers of the phantom, settlement of
silica particles in the bottom layer of the phantom results in a decreased US contrast at the
boundary between the top and the bottom layers (approximately in the middle of the images
on Fig. 6(a). The PA signal intensity showed linear behavior as a function of concentration of
labeled cells (the R2 value for the linear regression fit is 0.9753). From the linear regression
fit in Fig. 6(b), we determined that PA imaging can detect ca. 1.25 x 106 cells evenly
dispersed in a 1 cm3 volume. After taking into account the imaging kernel size of the
photoacoustic system of approximately 160 μm x 110 μm x 220 μm, we calculated that PA
imaging could detect as few as five nanorod-loaded cells per imaging kernel.
Fig. 6. Photoacoustic images of tissue-mimicking phantoms prepared with different
concentrations of P388D1 cells loaded with Rh-Silica-NRs (a). Each image covers a 6.3 x 8.8
mm field of view. Dependence of PA signal amplitudes on concentration of nanorod-loaded
cells at 780 nm excitation wavelength (b); the top horizontal axis shows a number of cells per
imaging kernel size of the photoacoustic system of approximately 160 μm x 110 μm x 220 μm.
The solid blue line represents the linear regression fit of the data and the black line shows the
noise level in PA imaging (b).
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(C) 2013 OSA
1 November 2013 | Vol. 4, No. 11 | DOI:10.1364/BOE.4.002609 | BIOMEDICAL OPTICS EXPRESS 2617
Conclusions
In summary, we systematically addressed challenges associated with applications of gold
nanorods in cell imaging. First, we demonstrated long-term biocompatibility of fluorescent
silica-coated nanorods for cell tracking experiments. Then, we showed that silica-coated
nanorods do not alter optical properties upon cellular uptake that can facilitate multiplex
imaging of various cell populations. Furthermore, our previous studies demonstrated that
silica coating increases stability [52] and signal strength [62] of gold nanorods in PA imaging.
The combination of these properties that are afforded by silica coating results in the detection
limit of just few labeled cells in tissue mimicking phantoms. These promising results provide
the foundation for applications of silica-coated gold nanorods and PA imaging in tracking of
immune cells in studies ranging from the mechanistic understanding of immune responses in
cancer and cardiovascular diseases to the cell mediated delivery of therapy.
Acknowledgments
This research was supported by National Institute of Health grants EB008101, HL096981 and
CA149740. We also would like to thank Ms. Maria Jimenez for proofreading the manuscript.
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Received 14 Aug 2013; revised 30 Sep 2013; accepted 21 Oct 2013; published 24 Oct 2013
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