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December 21, 2011

C 2011 American Chemical Society

Silver Nanoplate Contrast Agents forin VivoMolecular Photoacoustic ImagingKimberly A. Homan,† Michael Souza,† Ryan Truby,† Geoffrey P. Luke,† Christopher Green,† Erika Vreeland,‡

and Stanislav Emelianov†,*

†Biomedical Engineering Department, The University of Texas at Austin, 1 University Station C0800, Austin, Texas 78712, United States, and ‡Chemical EngineeringDepartment, University of New Mexico, 1 University of New Mexico, New Mexico 87131, United States

Molecular imaging of cancer is anarea of intense research. Many bio-medical imaging techniques are

traditionally anatomical in nature but canbe adapted, mostly through use of targetedcontrast agents, to detect cellular and mo-lecular behavior.1 One such technique isphotoacoustic (PA) imaging.2 PA imaging ishighly complementary to traditional ultra-sound (US) imaging. In US imaging, soundis used to interrogate thebody, and an imageis reconstructed from sound waves that arebackscattered from the tissue's mechanicalinhomogeneities. In PA imaging, pulsed laserlight is used to interrogate the body, and animage is reconstructed from broadbandsound waves generated by light absorptionevents in various tissue components. US andPA modalities can use the same electroniccomponents for receiving ultrasonic waves,allowing data to be collected from bothmodalities in a spatially coregistered way.As a result, complementary informationabout the tissue is obtained: US imagingcan visualize anatomy, while PA imagingcan visualize functional properties of thetissue based on optical absorption.3�5

Contrast in PA imaging arises from thenatural variation in the optical absorption oftissue components. Tissue components thatabsorb light strongly include hemoglobin,fat, andmelanin.6 Of these endogenous com-ponents, the use of contrast in PA imagingcoming from hemoglobin in blood has beenthe most widely researched and demon-strated.7�12 For example, PA imaging canbe used to determine the blood oxygensaturation in regions of particular interest,such as cancer, where hypoxia has beenshown to be strongly associated with malig-nancy.13 Combining US and PA imaging canreveal changes in blood vessel count or oxy-gen saturation inside tumors as comparedwith surrounding healthy tissue since anato-mical references, such as tumor location, size,

and boundaries, can be visualized and seg-mented using US imaging.The use of endogenous contrast in PA

imaging of cancer has made it a populartechnique in recent years.14 However, em-ploying exogenous contrast in PA imagingenables its molecular sensitivity and greatlyexpands the functionality of themodality.15�21

Exogenous contrast agents designed for PAimaging usually consist of highly absorbingdyes or nanoparticles that can be conjugatedto a targeting moiety of interest.22,23 Thetargeting moiety could be an antibody, apta-mer, or ligand that will facilitate localizationto cancer cells and highlight their presenceamonghealthy tissue. Design considerationsand requirements for a PA contrast agent

* Address correspondence toemelian@mail.utexas.edu.

Received for review October 24, 2011and accepted December 21, 2011.

Published online10.1021/nn204100n

ABSTRACT

Silver nanoplates are introduced as a new photoacoustic contrast agent that can be easily

functionalized for molecular photoacoustic imaging in vivo. Methods are described for

synthesis, functionalization, and stabilization of silver nanoplates using biocompatible

(“green”) reagents. Directional antibody conjugation to the nanoplate surface is presented

along with proof of molecular sensitivity in vitrowith pancreatic cancer cells. Cell viability tests

show the antibody-conjugated silver nanoplates to be nontoxic at concentrations up to

1 mg/mL. Furthermore, the silver nanoplates' potential for in vivo application as a molecularly

sensitive photoacoustic contrast agent is demonstrated using an orthotopic mouse model of

pancreatic cancer. Results of these studies suggest that the synthesized silver nanoplates are

well suited for a host of biomedical imaging and sensing applications.

KEYWORDS: contrast agents . photoacoustic imaging . optoacoustic imaging .silver . nanoplates . pancreatic cancer

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include (1) ease of synthesis with biocompatible re-agents, (2) an overall size on the nanoscale or smaller inorder to navigate the bloodstream and interact withbiological macromolecules, (3) a large absorption cross-section of light in the near-infrared (NIR) wavelengthregion where light penetration into tissue is maximized,(4) a surface chemistry that allows for simple bioconjuga-tion of targeting moieties, (5) low toxicity, (6) long shelflife, and (7) structural and molecular biostability in biolo-gical fluids.Several groups have shown that functionalized gold

nanoparticles fulfill most of these requirements andcan be successfully employed as PA imaging contrastagents.17,24�28 Here, we introduce the use of Ag nano-plates as a new PA contrast agent that meets all of therequirements. The steps of the inherently “green”synthesis methods (i.e., a synthesis using only biocom-patible reagents) are described along with the processof functionalizing the nanoplate surface to make itboth biostable (structurally stable in physiologicalmedia) and antibody-conjugated. Themolecular speci-ficity of the nanoplates is demonstrated in vitro usingpancreatic cancer cell lines and darkfield microscopy.The cytotoxicity of the nanoplates is assessed using

two pancreatic cancer cell lines and one noncancerouspancreatic cell line. Furthermore, the potential of usingsilver nanoplates for in vivo applications is demon-strated through combinedultrasoundandphotoacoustic(USPA) imaging of an orthotopic pancreatic tumor in amouse model following systemic injection of antibody-conjugated Ag nanoplates.

RESULTS AND DISCUSSION

“Green” Synthesis and Characterization of Ag Nanoplates.Researchers have designed numerous methods tosynthesize Ag nanoplates. Most are accomplishedthrough seed-mediated growthmechanisms29�40 thatcan include reducing agents such as hydrazine,41

increases in temperature,30 or even ingenious lightmediated methods.40,42 Interestingly, even bacteriahave been shown to produce silver nanoplates uponincubationwith Agþ in solution.43,44 Sincewe intend touse silver nanoplates in a biological context, weadopteda seed-mediated synthesis method that was particularly“green,” where only biocompatible chemicals wereemployed. For instance, ascorbic acid (vitamin C) wasused as the reducing agent, and sodium citrate (a well-known preservative) was used as a stabilizing and

Figure 1. Silver nanoplates directly after stepwise growth adding edge length from B1 to B4 (a�d) with their correspondingextinction spectra (e) and color change in solution (f). Scale bars are 100 nm.

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capping agent during synthesis. Using “green” reagentshelped to facilitate the formation of less toxic nano-plates and speed their transition to biocompatible con-trast agents for in vivo applications.

After exploring several different methods of pre-paring Ag seeds,38,45 we found that the seeds pro-posed by Xue et al.39 were most reproducible andprovided the twinned plane and stacking fault defectsthat help enable high-yield Ag nanoplate synthesis.46

To grow the Ag nanoplates from these seeds, wemodified procedures introduced by Zou et al.41 wherethe plates were synthesized in a stepwise process from4.2 nm seeds to large (>200 nm edge length) nano-plates. Figure 1 shows the progression of synthesisfrom seeds to large nanoplates in four steps. First, theyellow seedswere grown into 25 nmnanoplates (B1) asshown in Figure 1a. Subsequent growth steps in-creased the edge length of the nanoplates successively(Figure 1 panels b, c, and d, representing samples B2,B3, and B4, respectively). The colors of the resultingsolutions changed with the variation of their opticalextinction properties as depicted in Figure 1e,f. Oursynthesis method produced nanoplates with edgelengths and longitudinal surface plasmon resonances(LSPR) as shown in Table 1. The advantages of thissynthesis technique include the following: (1) onlybiocompatible reagents are used, (2) Ag nanoplatescan be tuned to absorb light throughout the entirevisible to near-infrared (NIR) light spectrum, and (3) theAg nanoplate surface is stabilized with sodium citrate, amolecule that can be easily displaced by amine or thiolfunctional groups for subsequent bioconjugation steps.

Using Ag nanoplates as a NIR beacon for any type ofbiological sensor or imaging contrast agent applicationrequires that they are stable in saline solutions. WhileAg has excellent optical properties for use as a sensor,its lack of structural stability in salt solutions has largelylimited its use in biomedical applications. As shownpreviously,41 when Ag nanoplates are built, the face ofthe nanoplate grows along the (111) plane, the lowestpossible energy surface. A high resolution transmissionelectron micrograph (TEM) of a silver nanoplate face(fromB4) is shown in Supporting Information, Figure S1a.The electron diffraction pattern from that same nano-plate face is shown in Supporting Information, Figure S1bwhere the first ring of intense spots can be attributedto the (220) Bragg reflections with a lattice spacing

of 1.44 Å. This nanoplate surface is most unstableat the tips, where its electromagnetic field enhance-ments are greatest.47 Usually it is the loss of thesetips that causes the nanoplates' LSPR peak48 to blueshift, even during idle suspension in water. There-fore, the synthesis technique we chose for silvernanoplates does not yield nanoplates with perfecttips, but instead allows for rounded, more stable tipsas shown in Figure 1 and Supporting Information,Figure S1a to be produced.

Allowing for rounded nanoplate tips was only thefirst step to solving the stability issue. Even Ag nano-plates with rounded tips will degrade and dissolve insalt solutions without additional passivation. To makenanoplates suitable for a contrast agent application,they must be biostable (i.e., structurally stable in aharsh salt or cell media environment) for aminimum ofthe time period required to circulate in the bloodstreamand interact with diseased cells (for cancer applicationsthis time period could be from several hours to a fewdays). Therefore, we tested the biostability of the Agnanoplates in Dulbecco's phosphate buffered saline(DPBS) after conjugation of various amounts of thiol-terminated methoxy-poly(ethylene glycol) (mPEG-SH,5 kDa) to the nanoplate surface (note that PEG waschosen for its well-established potential to passivatenanostructures and limit opsonization49). The initialconjugation of mPEG-SH to the nanoplates caused ared shift in the LSPR peak of up to 60 nm in the caseswhere enough mPEG-SH was used to passivate theentire surface. For the example shown in Figure 2,a 30-fold excess of mPEG-SH was reacted with Agnanoplates. The nanoplates were washed with watervia centrifugation and subsequently resuspended inDPBS at day zero. As shown in this case, with correctpassivation (an optimal surface area coverage of PEG),the Ag nanoplates were biostable in salt solutions. Theirdegradation in DPBS was very slow after passivationsuch that, even after 62 days, the shift in LSPR wasless than 40 nm, there was no evident change in thewidth of the LSPR peak over time, and only a 6% decreasein extinction was observed. As a control, we created a

TABLE 1. Size Chart of Ag Nanoplate Edge Length and

Thickness for Each Growth Step alongwith the LSPR Peak

for Each Batch

nanoplate edge length (nm) nanoplate thickness (nm) LSPR peak (nm)

B1 25.3 ( 5.5 10.4 ( 1.6 550B2 60.9 ( 10.1 12.5 ( 1.9 720B3 128.0 ( 25.9 18.0 ( 2.7 900B4 218.6 ( 35.6 25.6 ( 6.6 1080

Figure 2. Demonstration of the biostability of PEGylatedAgnanoplates suspended in saline solution over a period of 62days.

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suboptimal coverage of mPEG-SH on Ag nanoplates(surface area coverage of only 50%), and upon suspendingin DPBS at day zero, a blue shift of over 250 nm wasimmediate (see Supporting Information, Figure S2). Notethat with no passivation or surface modifications, as-pre-pared Ag nanoplates will completely dissolve when sus-pended in DPBS. Therefore, functionalizing the Ag nano-plate surface with mPEG-SH passivated the Ag nanoplates,making them suitable for contrast agent applications.

Molecular Specificity and Cytotoxicity of Ag Nanoplates.Functionalizing the surface of noble metals with mol-ecules containing thiols creates strong, covalent thi-olate bonds.50,51 These thiolate bonds are exploitedhere to not only create biostabilized nanoplates, butalso to bind antibodies to the nanoplate surface via

directional methods. The epidermal growth factorreceptor (EGFR) is commonly overexpressed on thesurface of cancer cells and promotes tumor growth andproliferation.52�54 A monoclonal antibody to EGFR(a-EGFR) was used here as a model to target Ag nano-plates to pancreatic cancer cells. The directional con-jugation method we used for binding a-EGFR to nano-plates was previously described by Kumar et al.55 Thisdirectional method employs linker molecules that bindthe Fc portion of the antibody directly to the Agsurface, leaving the entire variable region free tointeract with cells. To confirm that the antibody-conjugated Ag nanoplates would target pancreaticcancer cells, while PEGylated Ag nanoplates wouldhave limited interaction, wemixed both types of nano-plates with pancreatic cancer cells in vitro for 4 h. Theresulting cells imaged using darkfield microscopy areshown in Figure 3. The control cells with no nanoplateincubation are seen as blue in darkfield microscopy(Figure 3a). The cells that were incubated with PEGy-latedAgnanoplates haveno significant accumulation ofsilver (Figure 3b), but the cells that were incubated witha-EGFR-conjugated nanoplates show significant inter-action (Figure 3c). The accumulation of a-EGFR conju-gated nanoplates in cells is due to receptor-mediatedendocytosis of the nanoplates, as has been shownextensively with a-EGFR conjugated gold nanoparticlesand quantum dots.52,56�58 These in vitro results demon-strate the ability of a-EGFR conjugated nanoplates tomolecularly target pancreatic cancer cells.

Since the a-EGFR conjugated nanoplateswere prov-en to target cancer cells, we decided next to study ifthese interactions induce any cytotoxicity. Therefore,various amounts of a-EGFR conjugated nanoplateswere mixed with three different cell lines in vitro:MPanc96 and L3.6pl (pancreatic cancer cell lines thatoverexpress EGFR), and HPNE (a noncancerous pan-creatic cell line). The viability of the cells was testedafter incubation with the a-EGFR conjugated nano-plates for 24 husing anMTS assay. As shown in Figure 4,even up to concentrations of 1 mg/mL Ag nanoplates,no statistically significant cytotoxicity was found with

any of the three cells lines. These results were surpris-ing since the cytotoxicity of 5�20 nm Ag nanostruc-tures has been characterized at much lower concen-trations (less than 0.4 mg/mL).59,60 Our results suggestthat there is possibly a size- or coating-dependentcytotoxicity of silver that deserves further study asrecommended in a recent review.61

In Vivo Photoacoustic and Ultrasound Imaging with AgNanoplates. All in vivo imaging studies were conductedon Nu/Nu transgenic mice with xenograft humanpancreatic cancer grown orthotopically. Before injec-tion of nanoplates, volumetric PA and US images of thetumor, consistingofmultiple2-D transverse sectionsof thetumor, were acquired by translating the mouse through

Figure 3. Darkfield microscopy of pancreatic cancer cellsafter incubation with no nanoplates (a), PEGylated nano-plates (b), and a-EGFR conjugated nanoplates (c). All scalebars are 20 μm.

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the imaging plane. Then a-EGFR, PEG-passivatedAgnano-plates (∼1012 nanoplates in 200 μL of PBS with LSPR peakat∼800 nm) were injected into the tail vein. Imaging wasperformed every hour after injection up to 6 h postinjec-tion.At 4hpostinjection, peakaccumulationofnanoplatesin the tumor was observed as determined by monitoringthe photoacoustic signal intensity with time postinjection.USandcombinedUSPA images are shown in Figure 5. Thetumor is outlined in the ultrasound image as the hypoe-choic region inside the dotted white line (Figure 5a) andappears under the spleen and adjacent to the kidney. PAsignals captured ina rangeofwavelengths (740 to940nm)were used to spectroscopically resolve the signal fromblood and nanoplates in Figure 5b. Translating themousethrough the 2-D cross-sectional USPA imaging planeallowed for reconstructionof a3-D imageof theorthotopictumor (Figure 6). To see further spatial distribution of the

contrast agent and blood within the tumor space, a videoshowing a rotating 3-D view of the imaged tissue wascreated (Supporting Information, Figure S3). As has beenshown with other cancer models, a heterogeneous dis-tribution of nanoparticle accumulation in the tumor wasobserved.62,63 Furthermore, histological slices of the tumorstained with silver stain and nuclear fast red (Figure 7)confirmed the heterogeneous nanoplate distribution seenin USPA imaging. These in vivo imaging results indicate

Figure 4. Results of the MTS assay for cell viability on three cell lines after incubation for 24 h with a-EGFR conjugated Agnanoplates at various concentrations.

Figure 5. 2-D cross sections of an orthotopic pancreatictumor in a nude mouse model. US image (a) shows theanatomical features, while the USPA image (b) shows mo-lecular accumulation of a-EGFR conjugated nanoplates(yellow), oxygenated blood (red), and deoxygenated blood(blue). Image dimensions are 14.5 mm by 11.8 mm.

Figure 6. Reconstructed 3-D USPA image of an orthotopicpancreatic tumor in a mouse model. The PA visualization ofaccumulated a-EGFR-conjugated nanoplates (yellow), oxy-genated blood (red), and deoxygenated blood (blue) isshown overlaid on the US image of the mouse anatomy.Image dimensions are 8 mm by 17.5 mm by 11.8 mm.

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that silver nanoplates have the ability to accumulate inthe tumor space and provide enhanced contrast of thetumor region using PA signals that are easily differen-tiated from the endogenous tissue contrast (e.g., blood).Employing molecularly targeted nanoplates in thisway can be useful for studying the molecular profileof tumors and monitoring molecular therapeuticapproaches in future applications.

CONCLUSIONS

Ag nanoplates were synthesized using only biocom-patible, “green” methods that resulted in a clean, non-toxic surface ready for thiol-mediated, directional con-jugation of targeting and passivating molecules, such asantibodies and PEG. Once conjugated to a-EGFR, func-tionalized nanoplates underwent receptor-mediated

endocytosis in pancreatic cancer cells that overexpressEGFR, demonstrating their potential formolecular speci-ficity in vitro. Furthermore, once properly passivatedwith PEG, Ag nanoplates remained biostable in saltsolutions and cell media for over two months. For theconcentrations tested up to 1 mg/mL, Ag nanoplatesdid not inhibit proliferation of three different pancreaticcell lines, indicating they had low cytotoxicity. Further-more, spectroscopic PA imaging was able to visualizea-EGFR conjugated nanoplates in a mouse model oforthotopic pancreatic cancer and differentiate the Agnanoplate PA signal from that of the endogenousblood.Overall, the results of these studies suggest that Agnanoplates are well suited as PA contrast agents andhave the potential for use in a host of biomedicalimaging and sensing applications.

METHODSChemicals. All chemicals were used without further purifica-

tion and were at a minimum of ACS grade. Silver nitrate (>99%),trisodium citrate (TSC), sodium periodate, the antibody toEpidermal Growth Factor Receptor (EGFR) clone C225, poly-(ethylene glycol) bisphenol A epichlorohydrinMn = 14 kDa, andbis(p-sulfonatophenyl) phenylphosphine dihydrate dipotas-sium salt (BSPP) were purchased from Sigma-Aldrich (St. Louis,MO, USA). ACS grade L(þ) ascorbic acid was a product of Acros(Morris Plains, NJ, USA). Dulbecco's phosphate buffered saline(DPBS) came from Mediatech, Inc. (Herndon, VA, USA). Themethoxy-poly(ethylene glycol)-thiol (mPEG-SH of 5 kDa) wasfrom Laysan Bio (Arab, AL, USA). All cell culture products, in-cluding the MTS assay, were purchased from Invitrogen(Carlsbad, CA, USA) unless otherwise specified. Slide mountingmedia, Vectashield with DAPI stain, was from Vector Labora-tories (Burlingame, CA, USA). The linker hydrazide-polyethyleneglycol-dithiol (SPT-014B) was a product of Sensopath (Bozeman,MT, USA).

Ag Nanoplate Synthesis. Ag nanoplates were formed via aseed-mediated growth process. Ag seeds were synthesizedvia procedures previously described by Xue et al.40 Specifically,a 250 mL flask was placed in an ice bath and filled with 95mL ofdeionized and ultrafiltered (18.2 MΩ-cm) (DIUF) water. In an Arenvironment, 1 mL of 30 mM trisodium citrate (TSC) and 0.5 mL

of 20 mM silver nitrate (AgNO3) were added. The mixture wasstirred at 500 rpm for 20 min. To reduce the silver ions fromAgNO3, 1 mL of ice cold 50 mM sodium borohydride (NaBH4)was added quickly, and the solution immediately turned brightyellow. Onemoremilliliter of the 50mMNaBH4 was then addeddropwise over the following 15min. In a separate ice cold vial, asolution of 1 mL of 50 mM NaBH4 was added to 1 mL of 5 mMBSPP. That 2mLmixture of NaBH4 and BSPPwas slowly droppedinto the vigorously stirring Ag seed solution over a period of5 min. The Ag seeds were then stirred at 350 rpm and kept at0 �C for 5 h. Finally, the flask containing the seeds was trans-ferred to 4 �C and allowed to age undisturbed overnight. Notethat an argon environment was maintained throughout theentire procedure and the flask was wrapped in foil to minimizelight exposure.

The Ag seeds were used to grow Ag nanoplates up to14 days after synthesis, with the peak efficacy (largest transla-tion to plates) occurring on the day after synthesis. The typicalpeak resonance wavelength of the seeds as prepared was395 nm at an optical density (OD) of 1 using a 1 cm path lengthcuvette. Growing nanoplates of sizes up to 250 nm in diameterinvolved four separate growth steps where the product of onestep was used as the starting material for the subsequent step,asmodified from that described by Zou et al.41 First, a 20mL vialwas filled with 8.0mL of DIUF water, 50 μL of 40mM TSC, 150 μLof 40 mM Lþascorbic acid, and 1.8 mL of the as prepared

Figure 7. Histological cross-section of the orthotopic pancreatic tumor imaged using USPA imaging. The section is stainedwith nuclear fast red (pink areas) and silver stain (dark areas in the slice that indicate the presence of Ag nanoplates). Thedistribution of the nanoplates is heterogeneous and correlates with the distribution of nanoplates displayed in USPAimaging.

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Ag seeds at an OD of 0.65. The mixture was vigorously stirred atroom temperature while 5.05mL of 584 μMAgNO3was drippedin via syringe pump over 10 min. After the 10 min, the solutionturned color from bright yellow to orange, red, and maroon.After the AgNO3 addition was complete, the solution wasallowed to stir for an additional 10 min and the light extinctionspectrum was taken using a DU series 600 spectrophotometer(Beckman Coulter). This first solution was called B1. Second, forthe next Ag nanoplate growth step, another 20mL vial was filledwith 4.8 mL of DIUF water, 50 μL of 40mM TSC, 150 μL of 40mML(þ) ascorbic acid, and 5 mL of B1. Again, the mixture wasvigorously stirred at room temperature while 5.05mL of 584 μMAgNO3was dripped in via syringe pump over 10min. During the10 min, the solution turned color from red to purple to blue toteal-green. After the AgNO3 additionwas complete, the solutionwas allowed to stir for an additional 10 min and the lightextinction spectrum was taken. This second solution was calledB2. Third, for the next Ag nanoplate growth step, another 20mLvial was filled with 4.8 mL of DIUF water, 50 μL of 40 mM TSC,150 μL of 40 mM L(þ) ascorbic acid, and 5 mL of B2. Again, themixture was vigorously stirred at room temperature while5.05 mL of 584 μM AgNO3 was dripped in via syringe pumpover 10min. During the 10min, the solution turned color from tealto forest green to light forest green with a purple hue. After theAgNO3 addition was complete, the solution was allowed to stirfor an additional 10 min, and the light extinction spectrum wastaken. This third solution was called B3. For the fourth and finalgrowth step, another 20 mL vial was filled with 4.8 mL of DIUFwater, 50μL of 40mMTSC, 150μL of 40mMLþascorbic acid, and5 mL of B3. Again, the mixture was vigorously stirred at roomtemperature while 5.05 mL of 584 μM AgNO3 was dripped in viasyringepumpover 10min.During the10min, the solution turnedcolor from purple to pastel blue. After the AgNO3 addition wascomplete, the solution was allowed to stir for an additional10 min and the light extinction spectrum was taken. This fourthand final solution was called B4. The ImageJ application64 wasused to manually size representative populations of the nano-plate and seed batches. Note that Ag nanoplate batches didcontain a subpopulation of silver spheres (∼15% by particlenumber). All nanoplate solutions were analyzed using electronmicroscopy with both a LEO S5500 scanning transmission elec-tron microscope (STEM) and a JEOL 2010F transmission electronmicroscope (TEM).

Ag Nanoplate Antibody and Poly(ethylene glycol) Conjugation. A direc-tional conjugation of the C225 antiepidermal growth factorreceptor antibody (a-EGFR) from Sigma-Aldrich (product numberE2156) to nanoplateswas performed. Directional procedureswereadapted fromthosepreviously described for antibody conjugationto gold nanoparticles by Kumar et al.55 A linker molecule wasemployed to achieve directionality, meaning that only the Fcregion, or nonbinding part of the antibody, attached to the silver.The particular linker we used, a hydrazide�polyethylene gly-col�dithiol (Sensopath SPT-014B), covalently bound to the glyco-sylated portion of the antibody (found in the Fc region) through itshydrazide group, while the dithiol groups bound to silver.

Before directional conjugation of a-EGFR to Ag nanoplatescould be achieved, first the linker molecule was attached to theantibody. As previously described,55 the antibody was preppedby changing the buffer to a mono/dibasic sodium phosphatebuffer (pH = 7.5) using two wash steps in a 50 kDa molecularweight cut off (MWCO) centrifugal filter from Millipore(centrifuge at 3600 rpm for 15 min at 4 �C). The antibody wasreconstituted in 1.3 mL of the sodium phosphate buffer andplaced in a sterile 2mL tube. To create an aldehyde group on theglycosylated portion of the antibody, 80 μL of fresh 100 mMNaIO4 was added to the antibody and allowed to shake in thedark for 30 min at room temperature. The reaction wasquenched by the addition of 500 μL of 1� phosphate bufferedsaline. Then, 6 μL of 427.4 mM linker in anhydrous ethanol (keptat �80 �C prior to use) was added to the antibody solution andallowed to react for 1 h while shaking at room temperature.Finally, the antibody�linker (AB�linker) conjugate was cleanedusing HEPES buffer in the same 50 kDa MWCO Millipore filtersdescribed above and then reconstituted in 300 μL HEPES buffer(pH = 8.0) for storage. The AB�linker conjugate can be stored at

4 �C and will remain viable for the life of the antibody. A 75% to85% yield of the AB�linker conjugate can be expected usingthis technique if strict and clean pipetting techniques areemployed throughout the process.

Once the AB�linker conjugatewas built, further conjugation tothe Ag nanoplates was facile. Briefly, a B4 batch of nanoplates wasplaced in a 50 kDa MWCO Millipore centrifugal filter directly aftersynthesis and spun at 1500 rcf for 5 min at 20 �C. The nanoplateswere then resuspended in 1mL of DIUFwater and placed in a 2mLtube. Then, enoughof theAB�linker solutionwasadded toaccountfor 0.05 mg of antibody�linker (the exact volume of the AB�linkersolution varied based on the final concentration of the AB�linker inHEPES as determined by a Thermo Scientific Nanodrop read at awavelength of 280 nm with an assumed extinction coefficient of210,000M�1cm�1). The AB�linker and nanoplateswere allowed toshake at room temperature in the dark for 20 min. No shift in peakresonance of the nanoplates was observed after conjugation of theAB�linker to the nanoplates.

To enhance biocompatibility and stability, the surface areaof the silver not covered by antibody was passivated by attach-ment of poly(ethylene glycol) (PEG). Specifically, after allowingthe AB�linker and Ag nanoplate solution to shake for 20 min,50 μL of a 2 wt % PEG (Mn = 14 kDa from Sigma-Aldrich) wasadded to prevent aggregation and 40 μL of 50 mM mPEG-SH(5 kDa from Laysan Bio) was also added to react with the Agnanoplates. After shaking in the dark for 20 min, the nanoplateswere cleaned using 50 kDa MWCO Millipore centrifugal filtersspun at 1500 rcf for 5 min at 4 �C. The nanoplates were thenresuspended in 500 μL of PBS and sent through a 0.22 μm filterinto a sterile 2 mL tube for future use. The antibody-conjugatedAg nanoplates can be stored at 4 �C for the life of the antibody.A 40 to 60 nm red shift in the peak resonance of the nanoplateswas observed directly after the mPEG-SH passivating step.

Ag Nanoplate Biostability. Silver in nanoform is known to be amesostable compound. Passivation of the reactive Ag surfacewith polymers such as poly(ethylene glycol) (PEGMW= 5 kDa) isrequired to maintain the shape and optical properties of Agnanoplates. For instance, if Ag nanoplates are left in growthsolution overnight, their peak resonance will blue shift by up to120 nm by the next day. If Ag nanoplates are cleaned andresuspended in DIUF water, they will blue shift by∼80 nm overthe next 20 days. This blue shift in peak resonance is generallydue to loss of sharp triangular plate tips since they are theregions of highest surface energy on the plate. Furthermore,without surface passivation, when Ag nanoplates are placed inPBS, they dissolve. For biological applications, it is required thatAg nanoplates remain stable for days in PBS and cell media.Therefore, we tested the stability of Ag nanoplates over timeafter passivating the surface with mPEG-SH (5 kDa, Laysan Bio).

To test the passivating effect of mPEG-SH on Ag nanoplates,the nanoplates were reacted with various amounts ofmPEG-SH,washed, and then resuspended in DPBS. The extinction spec-trum of the nanoplates was then monitored over time. Briefly,the growth solution was removed from a B3 after synthesisusing a 50 kDaMWCOMillipore centrifugal filter spun at 1500 rcffor 5 min at 20 �C. The B3 nanoplates were then resuspended in3mL of DIUF water. To one batch of nanoplates, 445 μL of 5mMmPEG-SH was added, and to the other batch, 445 μL of0.083 mM mPEG-SH was added (representing a 30-fold excessof mPEG-SH surface area coverage on the first nanoplate sample,and only 50% surface area coverage on the second sample). Eachsample was allowed to react at room temperature, in the dark,while shaking for 15min. ExcessmPEG-SHwas then removed fromeach sample using a 50 kDaMWCOMillipore centrifugal filter spunat 1500 rcf for 5min at 20 �C. Each sample was resuspended to anoptical density of ∼1 (using a 1 cm path length) in Dulbecco'sphosphate buffered saline (DPBS). The extinction spectrum of thenanoplates was captured at various times over the next 60 days.Each sample was stored in a sealed plastic cuvette at 4 �C for theduration of the experiment.

In vitro Cell Labeling with Antibody and Poly(ethylene glycol) Con-jugated Ag Nanoplates. The human pancreatic cancer cell line,MPanc96, was mixed in media with two types of Ag nanoplates:nanoplates conjugated to a-EGFR and PEG, and nanoplatesconjugated only to PEG. Briefly, approximately 300000 MPanc96

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cells were suspended in 500 μL of media in two separate sterile2 mL tubes. To each respective tube, 50 μL of the as prepareda-EGFR and PEG conjugated nanoplates, and the PEG conjugatednanoplates weremixed. The tubes were shaken every 30min overa 4 h period while in the incubator. After 4 h, the cells were spundown in their tubes at 125 rcf for 5 min at 4 �C. They wereresuspended in 2 wt % formalin in DPBS, allowed to sit for 20 min,and then washed 2� with the 2 wt % formalin in DPBS solution.The cells were fixed on a microscope slide using Vectashield withDapi mounting solution and imaged in fluorescence and darkfieldmodes of a Leica DMI3000 B invertedmicroscopewith a Leica DFC290 kit and 3 mega pixel color camera.

In vitro Cytotoxicity Studies of Ag Nanoplates. Two human pan-creatic cancer cell lines, L3.6pl and MPanc96, and the noncan-cerous pancreatic line, HPNE, were incubated with Agnanoplates and tested for cell viability. Both cancer cell lineswere cultured in vitro using Dulbecco's modified eagle medium(DMEM) (with 4500 mg glucose/L, L-glutamine, NaHCO3, andpyridoxine HCl) supplemented with 10% fetal bovine serum(FBS) and 1% penicillin�streptomycin (Pen/Strep) while beingmaintained at 37 �C and 5% CO2 in a humidified incubator. Forthe pancreatic cancer cell lines, the DMEM and the FBS werepurchased from Sigma, while the Pen/Strep was purchasedfrom Invitrogen. The noncancerous pancreatic cell line, HPNE,was cultured in vitro using Medium D that contained onevolume of medium M3, three volumes of glucose-free DMEM(with L-glutamine, phenol red, and pyridoxine HCl from Invitrogen),5% FBS, 5.5mMglucose, 10 ng/mL EGF (Invitrogen), and 50μg/mLgentamycin (Sigma). Medium M3 is a proprietary formulationoptimized for the growth of neuroendocrine cells (InCell Corp.,San Antonio, TX, USA). For cell viability studies, cells were seededin a 96 well plate (each well had 5000 cells per 100 μL of media).The cells were allowed to attach and grow in the 96well plate for48 h, after which the cell media was removed and replaced with100 μL suspensions of a-EGFR- and PEG-conjugated Ag nano-plates at various concentrations representing 1, 0.5, 0.25, 0.125,0.0625, and 0mg/mLof silver (at least fivewells were seededwitheach concentration). Note that the highest silver concentration at1mg/mL represented∼1.2� 1012 nanoplates permilliliter, whilethe lowest silver concentration at 0.0625 mg/mL represented∼7.3� 1010 nanoplates permilliliter. After 24 h of incubation, themedia was removed and replaced with fresh media containingno nanoplates. The absorbance of each well in the plate wasmeasured at 490 nm using a Synergy HT Multimode MicroplateReader from BioTek. To perform the assay, 20 μL of a tetrazoliumcompound [3-(4,5-dimethylthiazol-2-yl)-5-(3-carboxymethoxy-phenyl)-2-(4-sulfophenyl)-2H-tetrazolium (MTS) and an electroncoupling reagent (phenazine methosulfate) PMS from theCellTiter 96 AQueous Non-Radioactive Cell Proliferation Assay(a Promega product) was added to each well. Over a period of1 h in the incubator, the MTS was bioreduced by cells into aformazan product that had an absorbance peak at 490 nm.Dehydrogenase enzymes found inmetabolically active cellswereresponsible for the conversion of MTS into the soluble formazanproduct. Therefore, the absorbance of each well at 490 nm wasdirectly proportional to the number of viable cells. The absor-bance of each well at 490 nm taken before adding MTS wassubtracted from the after MTS incubation value. Cell viability wasdetermined by comparing the resulting absorbance of wellscontaining no nanoplates to wells containing nanoplates usingan F test for a one-way ANOVA.

Animal Models. Xenograft tumors were grown orthotopicallyin Nu/Nu mice to test the in vivo targeting and imagingcapabilities of the a-EGFR conjugated nanoplates (the nano-plates were also passivated with mPEG-SH as described above).Specifically, after administering anesthesia to the mouse(avertin), surgery was performed to expose the pancreas andinject 50 μL containing 200 000 L3.6pl cells suspended in sterileDPBS. After approximately two weeks of growth, the tumor wasmature and the combined USPA imaging experiment wasconducted with the mouse situated on a heated electrocardio-gram pad where temperature and heart rate could be mon-itored while the animal was under anesthesia (isoflurane). Aftercollecting US and PA data on the tumor area in what we term“before” images, the mouse was injected (via the tail vein) with

∼1012 a-EGFR conjugated, PEG passivated Ag nanoplates in200 μL of PBS (LSPR peak of 800 nm). US and PA data was thencollected every hour for the first 6 h after injection. All proce-dures were conducted in accordance with the protocols ap-proved by the Institutional Animal Care and Use Committee atThe University of Texas at Austin.

Combined Ultrasound and Photoacoustic (USPA) Imaging System. Toperform in vivo imaging experiments, a custom designedcombined USPA imaging system was used as previouslydescribed.6,65�67 The setup for the combined USPA imagingsystem consisted of pulsed light in the range of wavelengthsbetween 740 and 940 nm generated by an optical parametricoscillator (OPO) Spectraphysics laser system operating at 10 Hzwith a 5�7 ns pulse duration. The pulsed light illuminated theside of the mouse, encompassing the tumor region, at fluencesranging from 10 to 15 mJ/cm2. An array transducer (9 MHzcenter frequency, 14 mm wide, 128 element linear array) col-lected the ultrasound data from above. To collect radiofre-quency (RF) data, the transducer was interfaced with a Cortexultrasound imaging engine (Winprobe Corp) capable of RF dataacquisition for spatially coregistered US and PA RF signals. Toform images, the RF data was beam-formed using a delay-and-sum approach. The PA spectrum measured at each pixelwas compared to known endogenous absorption spectra in theliterature68 and the extinction spectrum of Ag nanoplates. Eachcomponent was classified using a least squared errors estima-tion method67,69 to form a spectroscopic PA image. The USimages (displayed using dB scale) and fluence-compensatedspectroscopic PA images (displayed using linear scale) wereplotted. Three-dimensional renderings were obtained by stack-ing 21 two-dimensional slices spaced 400 μm apart.

Acknowledgment. The authors thank Dr. Craig Logsdon andDr. Thiru Arumugan at The University of Texas M.D. AndersonCancer Center for graciously providing all pancreatic cell linesand assisting with the generation of orthotopic, xenograftpancreatic cancer models in mice; Dr. Srivalleesha Mallidi atHarvard University for her expertise in animal imaging and dataprocessing; Aaron Chockla at The University of Texas at Austinfor taking high resolution TEM images; the Animal ResourceCenter for their animal handling training; and the WinprobeCorp for providing the Cortex imaging engine. We thank theNational Science Foundation (Grant No. 0821312) for fundingthe Hitachi S-5500 SEM/STEM used in this work. Partial supportfrom the National Institutes of Health under Grants CA 141203,HL 096981 and EB 008101 and from the National ScienceFoundation under the Grant OISE-1103935 are acknowledged.

Supporting Information Available: Additional figures andsupporting video. This material is available free of charge viathe Internet at http://pubs.acs.org.

REFERENCES AND NOTES1. Weissleder, R. Molecular Imaging in Cancer. Science 2006,

312, 1168–1171.2. Xu, M.; Wang, L. V. Photoacoustic Imaging in Biomedicine.

Rev. Sci. Instrum. 2006, 77, 041101.3. Emelianov, S. Y.; Aglyamov, S. R.; Karpiouk, A. B.; Mallidi, S.;

Park, S.; Sethuraman, S.; Shah, S.; Smalling, R. W.; Rubin,J. M.; Scott, W. G. Synergy and Applications of Ultrasound,Elasticity, and Photoacoustic Imaging. Proc. IEEE Ultrason.Symp. 2006, 405–415.

4. Kolkman, R. G.; Brands, P. J.; Steenbergen, W.; van Leeuwen,T. G. Real-Time in Vivo Photoacoustic and Ultrasound Ima-ging. J. Biomed. Opt. 2008, 13, 050510.

5. Kruger, R. A.; Liu, P.; Fang, Y. R.; Appledorn, C. R. Photo-acoustic Ultrasound (PAUS) Reconstruction Tomography.Med. Phys. 1995, 22, 1605–1609.

6. Homan, K.; Kim, S.; Chen, Y. S.; Wang, B.;Mallidi, S.; Emelianov,S. Prospects of Molecular Photoacoustic Imaging at 1064 nmWavelength. Opt. Lett. 2010, 35, 2663–2665.

7. Jose, J.; Manohar, S.; Kolkman, R. G.; Steenbergen, W.;van Leeuwen, T. G. Imaging of Tumor Vasculature UsingTwente Photoacoustic Systems. J Biophoton. 2009, 2, 701–717.

ARTIC

LE

HOMAN ET AL. VOL. 6 ’ NO. 1 ’ 641–650 ’ 2012

www.acsnano.org

649

8. Kolkman, R. G.; Mulder, M. J.; Glade, C. P.; Steenbergen, W.;van Leeuwen, T. G. Photoacoustic Imaging of Port-WineStains. Lasers Surg. Med. 2008, 40, 178–182.

9. Ku, G.; Wang, X.; Xie, X.; Stoica, G.; Wang, L. V. Imaging ofTumor Angiogenesis in Rat Brains in Vivo by PhotoacousticTomography. Appl. Opt. 2005, 44, 770–775.

10. Staley, J.; Grogan, P.; Samadi, A. K.; Cui, H.; Cohen, M. S.;Yang, X. Growth of Melanoma Brain Tumors Monitored byPhotoacoustic Microscopy. J. Biomed. Opt. 2010, 15,040510.

11. Zhang, H. F.; Maslov, K.; Stoica, G.; Wang, L. V. FunctionalPhotoacoustic Microscopy for High-Resolution and Nonin-vasive in Vivo Imaging. Nat. Biotechnol. 2006, 24, 848–851.

12. Yao, J.; Wang, L. V. Photoacoustic Tomography: Funda-mentals, Advances and Prospects. Contrast Media Mol.Imaging 2011, 6, 332–345.

13. Hockel, M.; Vaupel, P. Tumor Hypoxia: Definitions andCurrent Clinical, Biologic, and Molecular Aspects. J. Natl.Cancer Inst. 2001, 93, 266–276.

14. Mallidi, S.; Luke, G. P.; Emelianov, S. Photoacoustic Imagingin Cancer Detection, Diagnosis, and Treatment Guidance.Trends Biotechnol. 2011, 29, 213–221.

15. Homan, K.; Gomez, S.; Gensler, H.; Shah, J.; Brannon-Peppas, L.; Emelianov, S. Design and Development ofMultifunctional Contrast Agents for Photoacoustic Ima-ging. Proc. SPIE 2009, 7190, 71900I.

16. Joshi, P.; Chen, Y.-S.; Kim, S.; Shah, J.; Emelianov, S. Molec-ular Therapeutic Agents for Noninvasive PhotoacousticImage-Guided Photothermal Therapy. Proc. 31st Ann. Int.IEEE, EMBC 2009, 1158.

17. Kim, C.; Cho, E. C.; Chen, J.; Song, K. H.; Au, L.; Favazza, C.;Zhang, Q.; Cobley, C. M.; Gao, F.; Xia, Y.; et al. In VivoMolecular Photoacoustic Tomography of Melanomas Tar-geted by Bioconjugated Gold Nanocages. ACS Nano 2010,4, 4559–4564.

18. Kim, J. W.; Galanzha, E. I.; Shashkov, E. V.; Moon, H. M.;Zharov, V. P. Golden Carbon Nanotubes as MultimodalPhotoacoustic and Photothermal High-ContrastMolecularAgents. Nat. Nanotechnol. 2009, 4, 688–694.

19. Li, P. C.; Wei, C. W.; Liao, C. K.; Chen, C. D.; Pao, K. C.; Wang,C. R.; Wu, Y. N.; Shieh, D. B. Photoacoustic Imaging ofMultiple Targets Using Gold Nanorods. IEEE Trans. Ultra-sonic Ferroelectr. Freq. Control 2007, 54, 1642–1647.

20. Wilson, K.; Homan, K.; Emelianov, S. Synthesis of a DualContrast Agent for Ultrasound and Photoacoustic Ima-ging. Proc. SPIE 2010, 7576, 75760M.

21. Luke, G. P.; Yeager, D.; Emelianov, S. Y. Biomedical Applicationsof Photoacoustic Imaging with Exogenous Contrast Agents.Ann. Biomed. Eng. 2011, published online 3 Nov 2011.

22. Homan, K.; Mallidi, S.; Cooley, E.; Emelianov, S. CombinedPhotoacoustic and Ultrasound Imaging of Metal Nanopar-ticles in Vivo. In Nanoimaging; Goins, B., Phillips, W., Eds.;Pan Stanford Publishing: Singapore, 2011; Vol. 3.

23. Oraevsky, A. A. Gold and Silver Nanoparticles as ContrastAgents for Optoacoustic Imaging. In Photoacoustic Ima-ging and Spectroscopy; Wang, L. V., Ed.; Taylor and FrancisGroup: New York, 2009.

24. Agarwal, A.; Huang, S. W.; O'Donnell, M.; Day, K. C.; Day, M.;Kotov, N.; Ashkenazi, S. Targeted Gold Nanorod ContrastAgent for Prostate Cancer Detection by PhotoacousticImaging. J. Appl. Phys. 2007, 102, 064701.

25. Chen, Y. S.; Frey, W.; Kim, S.; Kruizinga, P.; Homan, K.;Emelianov, S. Silica-Coated Gold Nanorods as Photoacous-tic Signal Nanoamplifiers. Nano Lett. 2011, 11, 348–354.

26. Pan, D.; Pramanik, M.; Senpan, A.; Ghosh, S.; Wickline, S. A.;Wang, L. V.; Lanza, G. M. Near Infrared PhotoacousticDetection of Sentinel Lymph Nodes with Gold Nanobea-cons. Biomaterials 2010, 31, 4088–4093.

27. Skrabalak, S. E.; Chen, J.; Sun, Y.; Lu, X.; Au, L.; Cobley, C. M.;Xia, Y. Gold Nanocages: Synthesis, Properties, and Appli-cations. Acc. Chem. Res. 2008, 41, 1587–1595.

28. Pan, D.; Pramanik, M.; Wickline, S. A.; Wang, L. V.; Lanza,G. M. Recent Advances in Colloidal Gold Nanobeacons forMolecular Photoacoustic Imaging. Contrast Media Mol.Imaging 2011, 6, 378–388.

29. Biju, V.; Itoh, T.; Anas, A.; Sujith, A.; Ishikawa, M. Semicon-ductor Quantum Dots and Metal Nanoparticles: Synthe-ses, Optical Properties, and Biological Applications. Anal.Bioanal. Chem. 2008, 391, 2469–2495.

30. Hao, E.; Schatz, G. C.; Hupp, J. T. Synthesis and OpticalProperties of Anisotropic Metal Nanoparticles. J. Fluoresc.2004, 14, 331–341.

31. Jiang, X. C.; Chen, C. Y.; Chen, W. M.; Yu, A. B. Role of CitricAcid in the Formation of Silver Nanoplates through aSynergistic Reduction Approach. Langmuir 2010, 26,4400–4408.

32. Martínez-Casta~n�on, G.; Ni~no-Martínez, N.; Loyola-Rodrí-guez, J.; Pati~no-Marín, N.; Martínez-Mendoza, J.; Ruiz, F.Synthesis of Silver Particles with Different Sizes and Mor-phologies. Mater. Lett. 2009, 63, 1266–1268.

33. Metraux, C.; Mirkin, C. Rapid Thermal Synthesis of SilverNanoprisms with Chemically Tailorable Thickness. Adv.Mater. 2005, 17, 412–415.

34. Murphy, C. J.; Sau, T. K.; Gole, A. M.; Orendorff, C. J.; Gao, J.;Gou, L.; Hunyadi, S. E.; Li, T. Anisotropic Metal Nanoparti-cles: Synthesis, Assembly, andOptical Applications. J. Phys.Chem. B 2005, 109, 13857–13870.

35. Sharma, V. K.; Yngard, R. A.; Lin, Y. Silver Nanoparticles:Green Synthesis and Their Antimicrobial Activities. Adv.Colloid Interface Sci. 2009, 145, 83–96.

36. Suber, L.; Sondi, I.; Matijevic, E.; Goia, D. V. Preparation andthe Mechanisms of Formation of Silver Particles of Differ-ent Morphologies in Homogeneous Solutions. J. ColloidInterface Sci. 2005, 288, 489–495.

37. Sun, Y.; Xia, Y. Shape-Controlled Synthesis of Gold andSilver Nanoparticles. Science 2002, 298, 2176–2179.

38. Wiley, B.; Sun, Y.; Xia, Y. Synthesis of Silver Nanostructureswith Controlled Shapes and Properties. Acc. Chem. Res.2007, 40, 1067–1076.

39. Xue, C.; Metraux, G. S.; Millstone, J. E.; Mirkin, C. A. Mechanis-tic Study of Photomediated Triangular Silver NanoprismGrowth. J. Am. Chem. Soc. 2008, 130, 8337–8344.

40. Xue, C.; Mirkin, C. A. pH-Switchable Silver NanoprismGrowth Pathways. Angew. Chem., Int. Ed. 2007, 46, 2036–2038.

41. Zou, X.; Ying, E.; Chen, H.; Dong, S. An Approach forSynthesizing Nanometer-to Micrometer-Sized Silver Nano-plates. Colloids Surf. Physicochem. Eng. Aspects 2007, 303,226–234.

42. Sakamoto, M.; Fujistuka, M.; Majima, T. Light as a Construc-tion Tool of Metal Nanoparticles: Synthesis and Mecha-nism. J. Photochem. Photobiol., C 2009, 10, 33–56.

43. Ramanathan, R.; O'Mullane, A. P.; Parikh, R. Y.; Smooker,P. M.; Bhargava, S. K.; Bansal, V. Bacterial Kinetics-Controlled Shape-Directed Biosynthesis of Silver Nano-plates Using Morganella Psychrotolerans. Langmuir 2011,27, 714–719.

44. Xie, J.; Lee, J. Y.; Wang, D. I.; Ting, Y. P. Silver Nanoplates:From Biological to Biomimetic Synthesis. ACS Nano 2007, 1,429–439.

45. Skrabalak, S. E.; Au, L.; Li, X.; Xia, Y. Facile Synthesis of AgNanocubes and AuNanocages.Nat. Protoc. 2007, 2, 2182–2190.

46. Xia, Y.; Xiong, Y.; Lim, B.; Skrabalak, S. E. Shape-ControlledSynthesis of Metal Nanocrystals: Simple ChemistryMeets Complex Physics? Angew Chem., Int. Ed. 2009, 48,60–103.

47. Sherry, L. J.; Jin, R.; Mirkin, C. A.; Schatz, G. C.; Van Duyne,R. P. Localized Surface Plasmon Resonance Spectroscopyof Single Silver Triangular Nanoprisms. Nano Lett. 2006, 6,2060–2065.

48. Kelly, K.; Coronado, E.; Zhao, L.; Schatz, G. The OpticalProperties of Metal Nanoparticles: The Influence of Size,Shape, and Dielectric Environment. J. Phys. Chem. B 2003,107, 668–677.

49. Owens, D. E., 3rd; Peppas, N. A. Opsonization, Biodistribu-tion, and Pharmacokinetics of Polymeric Nanoparticles.Int. J. Pharm. 2006, 307, 93–102.

50. Bergen, J. M.; von Recum, H. A.; Goodman, T. T.; Massey,A. P.; Pun, S. H. Gold Nanoparticles as a Versatile Platform

ARTIC

LE

HOMAN ET AL. VOL. 6 ’ NO. 1 ’ 641–650 ’ 2012

www.acsnano.org

650

for Optimizing Physicochemical Parameters for TargetedDrug Delivery. Macromol. Biosci. 2006, 6, 506–516.

51. Levin, C. S.; Bishnoi, S.W.; Grady,N. K.;Halas, N. J. Determiningthe Conformation of Thiolated Poly(Ethylene Glycol) on AuNanoshells by Surface-Enhanced Raman Scattering Spectro-scopic Assay. Anal. Chem. 2006, 78, 3277–3281.

52. Diagaradjane, P.; Orenstein-Cardona, J. M.; Colon-Casasnovas,N. E.; Deorukhkar, A.; Shentu, S.; Kuno, N.; Schwartz,D. L.; Gelovani, J. G.; Krishnan, S. Imaging EpidermalGrowth Factor Receptor Expression in Vivo: Pharmaco-kinetic and Biodistribution Characterization of a Bio-conjugated Quantum Dot Nanoprobe. Clin. Cancer Res.2008, 14, 731–741.

53. Rosenthal, E. L.; Kulbersh, B. D.; King, T.; Chaudhuri, T. R.;Zinn, K. R. Use of Fluorescent Labeled Anti-epidermalGrowth Factor Receptor Antibody to Image Head andNeck Squamous Cell Carcinoma Xenografts. Mol. CancerTher. 2007, 6, 1230–1238.

54. Sokolov, K.; Follen, M.; Aaron, J.; Pavlova, I.; Malpica, A.;Lotan, R.; Richards-Kortum, R. Real-Time Vital OpticalImaging of Precancer Using Anti-epidermal Growth FactorReceptor Antibodies Conjugated to Gold Nanoparticles.Cancer Res. 2003, 63, 1999–2004.

55. Kumar, S.; Aaron, J.; Sokolov, K. Directional Conjugation ofAntibodies to Nanoparticles for Synthesis of MultiplexedOptical Contrast Agents with Both Delivery and TargetingMoieties. Nat. Protoc. 2008, 3, 314–320.

56. Aaron, J.; Travis, K.; Harrison, N.; Sokolov, K. DynamicImaging of Molecular Assemblies in Live Cells Based onNanoparticle Plasmon Resonance Coupling. Nano Lett.2009, 9, 3612–3618.

57. Bhattacharyya, S.; Bhattacharya, R.; Curley, S.; McNiven,M. A.; Mukherjee, P. Nanoconjugation Modulates theTrafficking and Mechanism of Antibody Induced ReceptorEndocytosis. Proc. Natl. Acad. Sci. U.S.A. 2010, 107, 14541–14546.

58. El-Sayed, I. H.; Huang, X.; El-Sayed, M. A. Surface PlasmonResonanceScattering andAbsorptionofAnti-EGFRAntibodyConjugated Gold Nanoparticles in Cancer Diagnostics: Ap-plications in Oral Cancer. Nano Lett 2005, 5, 829–834.

59. AshaRani, P. V.; LowKahMun, G.;Hande,M. P.; Valiyaveettil, S.Cytotoxicity and Genotoxicity of Silver Nanoparticles inHuman Cells. ACS Nano 2009, 3, 279–290.

60. Asharani, P. V.; Hande, M. P.; Valiyaveettil, S. Anti-proliferative Activity of Silver Nanoparticles. BMC Cell Biol.2009, 10, 65.

61. Johnston, H. J.; Hutchison, G.; Christensen, F. M.; Peters, S.;Hankin, S.; Stone, V. A Review of the in Vivo and in VitroToxicity of Silver and Gold Particulates: Particle Attributesand Biological Mechanisms Responsible for the ObservedToxicity. Crit. Rev. Toxicol. 2010, 40, 328–346.

62. Li, M. L.; Wang, J. C.; Schwartz, J. A.; Gill-Sharp, K. L.; Stoica, G.;Wang, L. V. In-Vivo Photoacoustic Microscopy of NanoshellExtravasation from Solid Tumor Vasculature. J Biomed Opt2009, 14, 010507.

63. Brigger, I.; Dubernet, C.; Couvreur, P. Nanoparticles inCancer Therapy and Diagnosis. Adv. Drug Delivery Rev.2002, 54, 631–651.

64. Rasband, W. S. ImageJ. http://imagej.nih.gov/ij/. Accessedon 3/24/2010.

65. Homan, K.; Shah, J.; Gomez, S.; Gensler, H.; Karpiouk, A.;Brannon-Peppas, L.; Emelianov, S. In Combined Ultra-sound and Photoacoustic Imaging of Pancreatic CancerUsing Nanocage Contrast Agents. SPIE Photonics West,San Jose, CA, USA, 2009; Proc. SPIE 2009, 71771M.

66. Homan, K.; Shah, J.; Gomez, S.; Gensler, H.; Karpiouk, A.;Brannon-Peppas, L.; Emelianov, S. Silver Nanosystems forPhotoacoustic Imagingand Image-GuidedTherapy. J. Biomed.Opt. 2010, 15, 021316.

67. Kim, S.; Chen, Y. S.; Luke, G. P.; Emelianov, S. Y. In VivoThree-Dimensional Spectroscopic Photoacoustic Imagingfor Monitoring Nanoparticle Delivery. Biomed. Opt. Express2011, 2, 2540–2550.

68. Prahl, S. A.Optical Property Spectra Compiled by Scott Prahl.http://omlc.ogi.edu/spectra/. Accessed on 8/25/2010.

69. Li, M. L.; Oh, J. T.; Xie, X.; Ku, G.; Wang, W.; Li, C.; Lungu, G.;Stoica, G.; Wang, L. Simultaneous Molecular and HypoxiaImaging of Brain Tumors in Vivo Using SpectroscopicPhotoacoustic Tomography. Proc. IEEE 2008, 96, 481–489.

ARTIC

LE