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Logo of nihpaAbout Author manuscriptsSubmit a manuscriptHHS Public Access; Author Manuscript; Accepted for publication in peer reviewed journal;
Small. Author manuscript; available in PMC 2011 April 9.
Published in final edited form as:
PMCID: PMC3035053

Gold Nanocages as Photothermal Transducers for Cancer Treatment


Gold nanocages represent a new class of nanomaterials with compact size and tunable optical properties for biomedical applications. They exhibit strong light absorption in the near-infrared region in which light can penetrate deeply into soft tissue. After PEGylation, the Au nanocages can be passively delivered to tumors in animals. Analysis of tissue distribution for the PEGylated Au nanocages showed that the tumor uptake was 5.7 %ID/g at 96 h post injection. The Au nanocages were found not only on the surface, but also in the core of the tumor. By exposing tumors to a near-infrared diode laser (0.7 W/cm2, CW, λ=808 nm) for 10 min, the photothermal effect of the Au nanocages could selectively destroy tumor tissue with minimum damage to the surrounding healthy tissue. Data from functional [18F]fluorodexoyglucose positron emission tomography revealed a decrease in tumor metabolic activity upon the photothermal treatment. Histological examination identified extensive damage to the nuclei of tumor cells and tumor interstitium.

1. Introduction

Plasmonic nanomaterials have received considerable attention for cancer diagnosis and therapy.[1] Gold nanostructures with optical properties tunable in the near-infrared (NIR) region (650 to 900 nm) are particularly attractive for hyperthermia based on the photothermal effect.[26] In this optical window, the attenuation of light by blood and soft tissue is relatively low, allowing for deep penetration. The key component of this approach is a photothermal transducer capable of absorbing light with a large cross section and then converting the light into heat with high efficiency. When localized in the tumor, the photothermal transducers offer a highly selective method for cancer treatment with minimum side effects by controlling the intensity of light. Several types of Au nanostructures have been developed with localized surface plasmon resonance (LSPR) peaks tuned to the NIR region via wet chemical syntheses; notable examples include nanoshells,[7] nanorods,[8] and nanocages.[9] Recent studies have shown significantly improved local tumor hyperthermia and extended survival periods after the photothermal treatment.[1013]

Gold nanocages represent a novel class of nanomaterials which are particularly attractive as photothermal transducers for therapeutic applications.[4, 14] They can be routinely synthesized in large quantities using a simple galvanic replacement reaction between silver (Ag) nanocubes and chloroauric acid (HAuCl4) in water.[15] By controlling the titrated amount of HAuCl4 into the reaction, the LSPR peak position of Au nanocages can be precisely tuned to any wavelength of interest in the range of 600–1200 nm. The NIR absorption cross section of Au nanocages is five orders of magnitude greater than the conventional organic dyes such as indocyanine green (ICG) while maintaining a compact size of ~40 nm, which can facilitate in vivo delivery.[16] Additionally, the unique hollow and porous structures of Au nanocages make them well-suited for drug encapsulation and controlled release through the photothermal effect with NIR light.[17]

In the present study, we have investigated the in vivo photothermal efficacy of Au nanocages using a bilateral tumor model. We delivered the Au nanocages to the tumor via passive targeting through modification of the nanocage surface with a monolayer of poly(ethylene glycol) (PEG). Surface PEGylation allows the Au nanocages to maintain a long circulation time in the blood stream and to accumulate in the tumor through the enhanced permeability and retention (EPR) effect, whereby the leaky tumor vasculature contains wide inter-endothelial junctions and a malfunctioning lymphatic system.[18] We monitored the temperature increase during photothermal treatment using an infrared camera which can provide useful information for the in vivo treatment planning. The effects of photothermal therapy on tumor metabolism was evaluated noninvasively using [18F]fluorodeoxyglucose positron emission tomography (18F-FDG PET). Decrease in tumor metabolic activity, an indication of effective therapy, was only observed in tumors treated with a combination of Au nanocages and laser exposure. Irreversible damage to the tumor cells was readily found upon histological examination. Finally, biodistribution studies showed that the uptake of the PEGylated Au nanocages by tumors was efficient, and that the nanocages were distributed throughout the tumor with the concentration in the tumor periphery being slightly higher than that in the tumor core.

2. Results and Discussion

The Au nanocages were prepared via a galvanic replacement reaction between Ag nanocubes and HAuCl4 in an aqueous solution using the procedure that has been optimized in our previous work.[15] The SPR peak of the Au nanocages was tuned to ~800 nm (Figure 1) to match the central wavelength of the diode laser (λ=808 nm). For the as-synthesized Au nanocages, the surface was covered by poly(vinyl pyrrolidone) (PVP, ~55,000 in molecular weight) and the size was 48±3.5 nm in edge length as measured by TEM (Figure 1 inset). The hydrodynamic diameter (intensity size, D) was measured as 99.8 nm with polydispersity index (PDI) of 0.16 by dynamic light scattering. The PVP layer was then replaced by heterofunctional poly(ethylene glycol) with one end terminated in the sulfhydryl group and the other end terminated in the methoxy group (HS-PEG5000-OMe, ~5,000 in molecular weight). After PEGylation, the surface of each nanocage was covered by HS-PEG5000-OMe molecules with an average total number on the order of 2×105 polymers per nanocage. This number is in agreement with the value estimated by assuming a footprint of 0.7 nm2 per thiol and an edge length of 48 nm for the nanocage. A previous study has measured the footprint of HS-PEG5000-OMe on 2.8 nm Au nanoparticle, and a value of 0.35 nm2 per thiol was obtained based on a full monolayer coverage.[19] The discrepancy can be attributed to two factors: i) the flat surface of a cubic nanostructure can increase the steric effects between polymer chains as compared to the curved surface of a spherical structure; and ii) the pores present in the nanocage can reduce the availability of the surface atoms for thiol binding. After PEGylation, the hydrodynamic diameter was reduced to 92.2 nm with PDI of 0.09. This difference in size is due to the conversion of surface coating from PVP to HS-PEG5000-OMe. Spectroscopic studies showed that the LSPR peak of the PEGylated Au nanocages was red shifted by 5 nm relative to the PVP-coated Au nanocages, while the peak profile remained the same, suggesting that the Au nanocages were well dispersed in phosphate buffered saline (PBS) before and after PEGylation. In addition, the PEGylated Au nanocages were stable in fetal bovine serum (FBS), showing no change to the LSPR peak for months until now (Figure 1). Using photoacoustic imaging, the absorption cross section of the Au nanocages was measured to be on the order of 6 × 10−15.[20]

Figure 1
UV-vis-NIR spectra showing the LSPR peaks of Au nanocages in different media: PVP-coated nanocages in PBS at pH 7.4 (solid line), PEGylated nanocages in PBS at pH 7.4 (dashed line), and PEGylated nanocages in fetal bovine serum (dotted line). The inset ...

To plan for in vivo photothermal treatment, we measured the temperature increase for a suspension of PEGylated Au nanocages in an aqueous solution under different conditions. For a given nanocage sample, the photothermal effect is determined by the particle concentration, as well as the power density and duration of laser irradiation.[14] We examined the temperature changes due to 10 min of irradiation by the diode laser at 1 W/cm2 (Figure 2A) and 0.5 W/cm2 (Figure 2B), respectively. The temperature profile was recorded by an infrared camera operating at a rate of 10 s per frame. For irradiation at a power density of 1 W/cm2, the temperature increased rapidly in the first two minutes and gradually reached a plateau after 5 min. When the same series of samples were exposed to the same laser at a power density of 0.5 W/cm2, the rate of temperature increase became markedly slower. Table 1 lists the temperature increase (ΔT) for aqueous suspensions of Au nanocages that were exposed to laser irradiation for 10 min. In the absence of nanocages, the temperature only increased by 2–3 °C, indicating that laser irradiation for 10 min at this power density poses minimal risk of adversely affecting cells or tissues. In the presence of 109 nanocage/mL (or ~1 ppm in terms of Au content), the temperature increased dramatically by 5–10 °C. Such a change could increase tissue temperature from 37 °C to >42 °C and cause an irreversible damage to the cells or tissues due to the denaturation of biomolecules.[21] Unlike the pulsed lasers with immense instantaneous power-per-pulse that can melt the nanocages to nanoparticles,[17] the continuous-wave (CW) diode laser caused no change to the optical properties of the Au nanocages (Figure 2C), indicating that the nanocages were stable under the irradiation conditions.

Figure 2
Plots of temperature increase for suspensions of Au nanocages at various concentrations as a function of irradiation time using the diode laser at different power densities: A) 1 W/cm2 and B) 0.5 W/cm2. C) UV-vis-NIR spectra of the Au nanocages in an ...
Table 1
Temperature increase (ΔT) for aqueous suspensions of Au nanocages upon irradiation by the diode laser for 10 min.

We further investigated the photothermal effect of the Au nanocages for selective destruction of the neoplastic tissue using a bilateral tumor model. Athymic mice were subcutaneously injected into the right and left rear flanks with U87wtEGFR cells. After the tumor volume had reached 200–400 mm3, the mice were randomly divided into Group 1 and 2 (n=5 per group). The mice in Group 1 were intravenously administrated with 100 μL of 10 mg/mL (15 nM or 9×1012 particle/mL) PEGylated Au nanocages in PBS. The mice in Group 2 served as control and were injected intravenously with 100 μL of saline. At 72 h post-injection, the tumor on the right rear flank of each mouse was subjected to photothermal treatment by exposure to the diode laser at a power density of 0.7 W/cm2 for 10 min. The spot size of the laser beam was adjusted to cover the entire tumor (Figure 3A). During the laser treatment, full-body thermographic images were captured using an infrared camera, as shown in Figure 3, B–I. The average temperature of the irradiated area was plotted as a function of the irradiation time (Figure 3J). For the nanocage-injected mice, the tumor surface temperature increased rapidly within one minute to reach 50 °C and began to plateau after 2 min at ~54 °C. In the case of saline-injected mouse, the surface temperature remained below 37 °C during the entire treatment.

Figure 3
A) Photograph of a tumor-bearing mouse under the photothermal treatment. 100 μL of PEGylated nanocages at a concentration of 9×1012 particles/mL or saline was administrated intravenously through the tail vein as indicated by an arrow. ...

Changes to tumor metabolism due to photothermal treatment were monitored using18F-FDG PET. Several human studies have shown that the use of 18F-FDG as a surrogate marker for tumor metabolism in patients undergoing therapy is superior to the Responsive Evaluation Criteria In Solid Tumors (RECIST), a method that simply evaluates the size of the tumor using an anatomical imaging technique such as computed tomography (CT).[2225] Measurement of tumor metabolism with 18F-FDG PET/CT imaging was performed before and after laser treatment for mice that had been intravenously injected with either saline or nanocages. Before laser irradiation, the 18F-FDG PET/CT imaging showed no significant difference between saline-injected mice (Figure 4A) and nanocage-injected mice (Figure 4B). At 24 h post-laser treatment, the metabolic activity in tumors of nanocage-injected mice (Figure 4D) was significantly reduced as compared to that of saline-injected mice (Figure 4C). We then normalized the PET signal of the laser-treated tumor to that of the untreated tumor to minimize the variation of 18F-FDG uptake at different time points (Figure 4E). The normalized value is ~0.3 after irradiation for the mice injected with Au nanocages as apposed to ~1 before irradiation, indicating a decrease of metabolic activity by 70%. For the saline-injected mice, the normalized value of 18F-FDG uptake was close to 1 before and after uptake, suggesting there is no benefit to laser treatment in the absence of Au nanocages.

Figure 4
18F-FDG PET/CT co-registered images of mice intravenously administrated with either saline or Au nanocages, followed by laser treatment: A) a saline-injected mouse prior to laser irradiation; B) a nanocage-injected mouse prior to laser irradiation; C) ...

Photothermal damage to tumor cells in mice injected with Au nanocages was confirmed by histological examination. Marked degenerative changes of coagulative necrosis, including abundant karyorrhectic debris and considerable regions of karyolysis, were found in laser-treated tumor tissue from mice injected with Au nanocages but were absent from tumors not exposed to laser irradiation or from mice injected with saline (Figure 5). In Figure 5D, a boundary between an area of karyorrhexis and an area of karyolysis is visible as vertical bands. A high magnification view of this boundary reveals extensive pyknosis, karyorrhexis, karyolysis, and interstitial edema (Figure 5H).

Figure 5
Representative histology images of tumor tissues from the two mice intravenously administrated with saline and Au nanocages, respectively, followed by different treatments: A) tumor from saline-injected mouse with no irradiation; B) tumor from saline-injected ...

To quantitatively assess the biodistribution and passive targeting of PEGylated Au nanocages, organs of interest were collected after the photothermal treatment to determine Au content. Inductively coupled plasma mass spectrometry (ICP-MS) was used to analyze the accumulation of exogenously given Au in tissues. Figure 6A plots the distribution of PEGylated-Au nanocages in various organs at 96 h post-injection. The nanocages were completely cleared from the blood (0.04±0.03 %ID/g), and relatively little Au remained in normal tissues (e.g., 0.80±0.12 %ID/g in muscles). Since the size of the nanocage is above the renal filtration limit (<8 nm),[26] the PEGylated Au nanocages were found to clear via reticuloendothelial system (RES) uptake with splenic clearance dominating, a pattern similar to that observed previously for PEGylated spherical Au nanoparticles with a diameter of 10 nm.[27] Most importantly, passive accumulation of the PEGylated Au nanocages within tumors was found to be highly efficient, reaching a particle concentration of 5.1×1010 particles/g (or 5.7% ID/g) at 96 h post-injection. The entire surface of the dissected tumor appeared in dark blue color (inset of Figure 6B), owing to the presence of Au nanocages. To examine the spatial distribution of the nanocages in a tumor, we cut a rectangular core through each tumor and sectioned them into five pieces. Each small piece was weighed and digested for ICP-MS analysis of Au content. Figure 6B displays the distribution of Au in different portions of the tumor. The edges were found to contain more nanocages than the center portion of the tumor because blood vessels are typically most abundant at the host interface where they may form a prominent, circumferential mantle enveloping tumor and leaving internal portions of tumors less well vascularized.[28] This result implies that over time the PEGylated Au nanocages penetrated through the leaky blood vessels of the tumor and diffused into the interstitium region of the tumor, allowing for uniform heat generation within the tumor.

Figure 6
A) Tissue distribution of the PEGylated Au nanocages intravenously administrated (100 μL, with a concentration of 9×1012 particles/mL) into tumor-bearing mice. The amount of Au in the tissue sample was analyzed by ICP-MS at 96 h post injection. ...

3. Conclusions

We have studied the use of PEGylated Au nanocages for in vivo photothermal treatment. The compact Au nanocages (~45 nm in edge length) have strong optical absorption in the NIR region with an LSPR peak precisely tunable for matching the center wavelength of the laser. Surface modification with PEG allowed the nanocages to accumulate in tumors with a relatively high efficiency. The intratumoral distribution revealed that the outer surface of the tumor contained slightly more nanocages than the inner core of the tumor. Non-invasive 18F-FDG PET/CT imaging technique serves as a useful tool for monitoring the treatment response, and allows for guiding and repeating of the therapeutic procedure. The short-term result suggested that the Au nanocages can serve as effective transducers for photothermal treatment of cancer. Long-term observation of treatment response will need to be evaluated by examining the change to the size and pathological characteristics of tumors in the future studies.

4. Experimental Section

Synthesis and PEGylation of Au nanocages

Gold nanocages were prepared using a galvanic replacement reaction between Ag nanocubes and HAuCl4.[15] The LSPR peak of the nanocages was tuned to ~800 nm as monitored using a UV-vis-NIR spectrometer (Cary 50). The surface of the nanocages was then derivatized with methoxy-terminated poly(ethylene glycol) thiol (SH-PEG-OMe). Briefly, 10 mg of SH-PEG5000-OMe (M.W.≈5,000, Laysan Bio) was dissolved in 18 mL of water in a 100-mL round-bottom flask, followed by adding 2 mL of Au nanocages at a final concentration of 2 nM. The reaction solution was stirred in dark for 12 h. The excess SH-PEG-OMe was removed by centrifugation at 10,000 rpm for 15 min and decanting. The PEGylated Au nanocages were then washed with water twice and re-suspended in PBS for in vivo studies at a concentration of 10 mg/mL (15 nM or 9×1012 particles/mL). The nanocages had an edge length of 48 ± 3.5 nm as determined by TEM analysis. The hydrodynamic diameter (intensity size) was 92 nm with a PDI of 0.09 as measured by dynamic light scattering (Malvern, Nano-ZS). To quantify the number of SH-PEG5000-OMe per particle, the nanocages were coated with amine-terminated PEG thiol (SH-PEG5000-NH2) using the same conjugation procedure. The average number of PEG5000 per Au nanocage was derived from the number of amine groups obtained from a Kaiser test kit (Sigma-Aldrich).[29]

Cell line and animal model

The U87MGwtEGFR human glioblastoma cell line was cultured in Dulbecco’s Modified Eagle Medium with high glucose (DMEM, HG, Gibco) supplemented with 10% Fetal Bovine Serum (FBS, Gibco) and 500 mg/L of Geneticin (G418, Gibco) at 37 °C with 5% CO2. Athymic nu/nu mice, age of 3–4 weeks, were obtained from Charles River Laboratory. The U87MGwtEGFR tumor model was generated by subcutaneous injection of 5×106 cells in 100 μL PBS into the right and left rear flanks. Animals used in photothermal studies had a tumor volume of 200–400 mm3 (typically 2–3 weeks after inoculation). All animal experiments were conducted in compliance with the guidelines for the care and use of research animals established by the Washington University Animal Studies Committee.

In vivo photothermal treatment

Ten tumor-bearing mice were randomly divided into two groups (n=5 per group). The mice in Group 1 and 2 were injected intravenously with 100 μL of 10 mg/mL (15 nM or 9×1012 particles/mL) PEGylated Au nanocages and saline, respectively. At 72 h post injection, animals were anesthetized with 2% isofurane (ISoFlo, Abott Laboratories) in 100% oxygen and placed in a prone position on a heated tray. The right tumor of each animal in Group 1 and 2 were exposed to a diode laser coupled to a 100-μm core fiber (Power Technology Inc.) at a power density of 0.7 W/cm2 for 10 min. The spot size of the laser beam was adjusted to cover the entire region of the tumor. During irradiation, the thermographs and temperature were recorded by an infrared camera (ICI7320, Infrared Camera Inc.). A circular region of interest (ROI) encompassing the irradiated tumor was drawn on thermographs and analyzed using IRFlash thermal imaging analysis software (Infrared Cameras Inc.).

18F-FDG PET/CT imaging studies

The effect of photothermal treatment on tumor metabolism was evaluated using 18F-FDG PET/CT imaging. Mice were imaged with 18F-FDG immediately prior to injection of Au nanocages and again at 24 h after photothermal therapy. Eight to twelve hours before each imaging session, the mice were fasted while drinking water was given ad libitum. The mice were anesthetized by inhalation of isoflurane (2% in 100% oxygen) and administered by tail vein injection of 300–400 μCi (11.1–14.8 MBq) of 18F-FDG in 100 μL saline. Sixty minutes after 18F-FDG injection, the mice were anesthetized by inhalation of isoflurane (2% in 100% oxygen) and secured in a supine position inside an acrylic imaging cradle. MicroPET/CT imaging was performed using an Inveon preclinical PET/CT scanner (Siemens Medical Solutions). The maximum a posteriori (MAP) reconstruction algorithm[30] was used on PET images. Image reconstruction was performed with registered CT-derived attenuation correction and scatter correction. Absolute quantitation was obtained with cross-calibration to a dose calibrator (CRC-15, Capintec, Inc.) using a known amount of activity in a mouse-sized cylindrical phantom. Tumor uptake of 18F-FDG was calculated in terms of the standardized uptake value (SUV) in 3-dimensional (3D) regions of interest (ROIs). In general, SUV is defined as the tissue concentration of radiotracer divided by the activity injected per body weight[31] and is calculated according to the following equation:


Upon decay of 18F-FDG, the mice were anesthetized and sacrificed by cervical dislocation. Tissues of interest were removed, blotted dry, weighed, and placed in pre-weighed vials of neutral buffered formalin and stored at 4 °C.

Histology study

The excised tumors were embedded in paraffin blocks, sectioned into 7 μm slices, and stained with hematoxylin and eosin (Sigma-Aldrich) to assess tissue and cellular morphology. The sections were viewed using 2.5× and 20× Plan Neofluar objectives and images were acquired using a Zeiss AxioCam-HR camera.

ICP-MS analysis

Each tissue sample was completely digested by 8 mL of aqua regia in a 100-mL beaker at boiling temperature. The solution was evaporated to 5 mL and subsequently diluted to 12 mL using 0.5% HCl and 2% HNO3. Samples were passed through a 0.45-μm filter to remove any undigested debris prior to ICP-MS measurement. The analysis of Au content was performed on ICP-MS (7500 CE, Agilent). Quantification was carried out by external five-point-calibration with internal standard correction and the percentage of injected Au dose per gram of tissue (%ID/g) was calculated.


This work was supported in part by a Director’s Pioneer Award from the NIH (DP1 OD000798) and startup funds from Washington University in St. Louis (to Y.X.); as well as a Research Development Award from the Alvin J. Siteman Cancer Center at Barnes-Jewish Hospital and Washington University School of Medicine (to J.C.). Siteman is supported by Grant P30 CA91842 from the NIH. The Inveon small animal PET/CT system, a component of the Siteman Small Animal Cancer Imaging Core, was acquired using an NIH-NCRR shared instrumentation grant (S10 RR025097, to R. L.). Part of the work was performed at the Nano Research Facility (NRF), a member of the National Nanotechnology Infrastructure Network (NNIN), which is supported by the NSF under award no. ECS-0335765.

Contributor Information

Jingyi Chen, Department of Biomedical Engineering, Washington University in St. Louis, St. Louis, Missouri 63130 (USA)

Dr. Charles Glaus, Department of Radiology, Washington University in St. Louis, St. Louis, Missouri 63110 (USA)

Prof. Richard Laforest, Department of Radiology, Washington University in St. Louis, St. Louis, Missouri 63110 (USA)

Qiang Zhang, Department of Biomedical Engineering, Washington University in St. Louis, St. Louis, Missouri 63130 (USA)

Miaoxian Yang, Department of Biomedical Engineering, Washington University in St. Louis, St. Louis, Missouri 63130 (USA)

Michael Gidding, Department of Chemical Engineering, Washington University in St. Louis, St. Louis, Missouri 63130 (USA)

Prof. Michael J. Welch, Department of Radiology, Washington University in St. Louis, St. Louis, Missouri 63110 (USA)

Prof. Younan Xia, Department of Biomedical Engineering, Washington University in St. Louis, St. Louis, Missouri 63130 (USA)


1. For reviews: a) Hu M, Chen J, Li ZY, Au L, Hartland GV, Li X, Marquez M, Xia Y. Chem Soc Rev. 2006;35:1084. [PubMed] b) Sperling RA, Gil PR, Zhang F, Zanella M, Parak WJ. Chem Soc Rev. 2008;37:1896. [PubMed] c) Biosselier E, Astrue D. Chem Soc Rev. 2009;38:1759. [PubMed]
2. Hirsch LR, Stafford RJ, Bankson JA, Sershen SR, Rivera B, Price RE, Hazle JD, Halas NJ, West JL. Proc Natl Acad Sci USA. 2003;100:13549. [PubMed]
3. Huang X, El-Sayed IH, Qian W, El-Sayed MA. J Am Chem Soc. 2006;128:2115. [PubMed]
4. Chen J, Wang D, Xi J, Au L, Siekkinen A, Warsen A, Li ZY, Zhang H, Xia Y, Li X. Nano Lett. 2007;7:1318. [PMC free article] [PubMed]
5. Melancon MP, Lu W, Yang Z, Zhang R, Cheng Z, Elliot AM, Stafford J, Olson T, Zhang JZ, Li C. Mol Cancer Ther. 2008;7:1730. [PMC free article] [PubMed]
6. Hasan W, Stender CL, Lee MH, Nehl CL, Lee J, Odom TW. Nano Lett. 2009;9:1555. [PMC free article] [PubMed]
7. Oldenburg SJ, Averitt RD, West JL, Halas NJ. Chem Phys Lett. 1998;288:243.
8. Gou L, Murphy CJ. Chem Mater. 2005;17:3668.
9. a) Sun Y, Xia Y. J Am Chem Soc. 2004;126:3892. [PubMed] b) Chen J, McLellan JM, Siekkinen A, Xiong Y, Li ZY, Xia Y. J Am Chem Soc. 2006;128:14776. [PubMed] c) Skrabalak SE, Chen J, Sun Y, Lu X, Au L, Cobley CM, Xia Y. Acc Chem Res. 2008;41:1587. [PubMed]
10. O’Neal DP, Hirsch LR, Halas NJ, Payne JD, West JL. Cancer Lett. 2004;209:171. [PubMed]
11. Lu W, Xiong C, Zhang G, Huang Q, Zhang R, Zhang JZ, Li C. Clin Cancer Res. 2009;15:876. [PMC free article] [PubMed]
12. Dickerson EB, Dreaden EC, Huang X, El-Sayed IH, Chu H, Pushpanketh S, McDonald JF, El-Sayed MA. Cancer Lett. 2008;269:57. [PMC free article] [PubMed]
13. von Maltzahn G, Park JH, Agrawal A, Bandaru NK, Das SK, Sailor MJ, Bhatia SN. Cancer Res. 2009;69:3892. [PMC free article] [PubMed]
14. Au L, Zheng D, Zhou F, Li ZY, Li X, Xia Y. ACS Nano. 2008;2:1645. [PMC free article] [PubMed]
15. Skrabalak SE, Au L, Li X, Xia Y. Nat Protl. 2007;2:2182. [PubMed]
16. Chen J, Saeki F, Wiley BJ, Cang H, Cobb MJ, Li ZY, Au L, Zhang H, Kimmey MB, Li X, Xia Y. Nano Lett. 2005;5:473. [PubMed]
17. Yavuz MS, Cheng Y, Chen J, Cobley CM, Zhang Q, Rycenga M, Xie J, Kim C, Song KH, Schwartz AG, Wang LV, Xia Y. Nat Mater. 2009;8:935. [PMC free article] [PubMed]
18. Matsumura YMH. Cancer Res. 1986;46:6387. [PubMed]
19. Wuelfing WP, Gross SM, Miles DT, Murray RW. J Am Chem Soc. 1998;120:12696.
20. Cho EC, Kim C, Zhou F, Cobley CM, Song KH, Chen J, Li ZY, Wang LV, Xia Y. J Phys Chem C. 2009;113:9023. [PMC free article] [PubMed]
21. Roti Roti JL. Int J Hyperthermia. 2008;24:3. [PubMed]
22. Schuetze SM, Eary JF, Griffith KA, Rubin BP, Hawkins DS, Vernon CB, Mann GN, Conrad EU. J Clin Oncology. 2005;23(Suppl):9005.
23. Sunagaa N, Oriuchib N, Kairaa K, Yanagitania N, Tomizawaa Y, Hisadaa T, Ishizukaa T, Endob K, Moria M. Lung Cancer. 2008;59:203. [PubMed]
24. Pottgen C, Levegrun S, Theegarten D, Marnitz S, Grehl S, Pink R, Eberhardt W, Stamatis G, Gauler T, Antoch G, Bockisch A, Stuschke M. Clin Cancer Res. 2006;12:97. [PubMed]
25. Hellwig D, Graeter TP, Ukena D, Georg T, Kirsch CM, Schäfers DIHJ. J Thor Cardiov Surg. 2004;128:892. [PubMed]
26. Choi HS, Liu W, Misra P, Tanaka E, Zimmer JP, Ipe BI, Bawendi MG, Frangioni JV. Nat Biotech. 2007;25:1165. [PMC free article] [PubMed]
27. Cai QY, Kim SH, Choi KS, Kim SY, Byun SJ, Kim KW, Park SH, Juhng SK, Yoon KH. Invest Radiol. 2007;42:797. [PubMed]
28. Jain RK. Ann Rev Biomed Engin. 1999;1:241. [PubMed]
29. Sarin VK, Kent SBH, Tam JP, Merrifield RB. Anal Biochem. 1981;117:147. [PubMed]
30. Ruangma A, Bai B, Lewis JS, Sun X, Welch MJ, Leahy R, Laforest R. Nucl Med Biol. 2006;33:217. [PubMed]
31. Zasadny KR, Wahl RL. Radiology. 1993;189:847. [PubMed]