Introduction

In the United States, prostate cancer is the second most common reason for cancer death in men (Jemal et al. 2010). Accurate detection methods are important for all aspects of the clinical management of prostate cancer, including diagnosis, risk assessment, staging, and prognosis. Such methods will result in individualized and efficacious treatments for patients at risk for prostate cancer or for its progression. Many of these tasks are currently managed by determination of the serum biomarker prostate specific antigen (PSA). For example, serum PSA levels are employed to evaluate prostate cancer risk and progression, and justify confirmatory biopsies to diagnose the presence of malignancy. However, biopsies have inherent risks such as bleeding and infection (Andrew et al. 2010), and cancer is not detected (false negative cases) in 30–50% of biopsies in patients with subsequently confirmed malignancy due to small and inconspicuous lesions (Rabbani et al. 1998). Another major issue is that ~25–40% of patients are over diagnosed using current detection methods leading to superfluous biopsies (Andrew et al. 2010). These findings indicate that the detection and staging of prostate cancer needs to be improved. Novel magnetic resonance molecular imaging methods promise to markedly increase the specificity of prostate tumor detection (Afnan and Tempany 2010).

Our goal is to develop targeted magnetic resonance imaging (MRI) agents for the specific detection of prostate cancer. A unifying theme in the development of novel imaging and therapeutic modalities in recent years has been specifically targeting these agents to cells of interest (Peng et al. 2008). The targeting motifs are often antibodies against antigens expressed on cancerous cells but not healthy cells. The most promising target antigen expressed specifically on prostate cancer cells is prostate specific membrane antigen (PSMA), which is most strongly expressed in the prostate, and expression has been found to increase as tumor grade and stage increases (Rajasekaran et al. 2005). In addition, many nonprostatic tumors have been found to express PSMA in the neovasculature, but expression in healthy vasculature has not been reported (Ghosh and Heston 2004; Chang et al. 1999), suggesting that PSMA may also be a general tumor antigen that could be used to detect numerous types of solid tumors. Thus, antibodies against PSMA are an appealing choice for use as targeting motifs for prostate cancer.

Superparamagnetic iron oxide nanoparticles (SPIONs) are the most common type of contrast agents used in agent-enhanced MRI (Zhang et al. 2010). Although SPIONs cause negative contrast in the MR images, the signal enhancement is still lower than other common imaging techniques (Lee et al. 2007). Therefore, novel nanoparticles with increased MRI relaxivities are needed to increase the signal enhancement in MRI and the detection of cancer, using lower doses of the contrast agents. Iron platinum (FePt) particles have been the focus of intense research in recent years due to their high magnetic anisotropy and high stability (Han et al. 2009; Kim et al. 2009; Antoniak et al. 2006; Sun 2006; Sun et al. 2000). Much of the interest in FePt has been placed on producing ferromagnetic face-centered tetragonal (fct) FePt particles, for use in magnetic storage devices, by annealing superparamagnetic face-centered cubic (fcc) nanoparticles at temperatures exceeding 500 °C (Basit et al. 2009; Varanda and Jafelicci 2006; Boyen et al. 2005; Sun et al. 2000). However, fcc superparamagnetic iron platinum particles (SIPPs) are of interest by themselves due to their possible use as contrast agents in MRI (Chen et al. 2010; Morales et al. 2009; Maenosono et al. 2008). For superparamagnetic MRI contrast agents, it is thought that a higher magnetic moment at a given magnetic field causes larger perturbations in the magnetic relaxation times of nearby water protons and, thus, higher moment particles should generate increased image contrast. SIPPs have previously been reported with volume magnetizations greater than 590 emu/cm3 (6 × 105 A m2), with some preparations approaching 1.140 emu/cm3 (1 × 106 A m2), the saturation magnetization of bulk FePt (Xu et al. 2009; Zhao et al. 2009; Barmak et al. 2004; Zeng et al. 2002). These reported high magnetic moments suggested that SIPPs would be superior MRI contrast agents.

One obvious drawback to the synthesis of various types of nanoparticles is that toxic precursors are generally employed to produce the particles (Sun 2006; Sun et al. 2000; Inomata et al. 1988). Often, iron pentacarbonyl, Fe(CO)5, a very hazardous reagent (Bhalerao et al. 2009), is used as the iron precursor in FePt syntheses (Sun et al. 2000). A number of different FePt syntheses are described by Sun (2006). Recently, Zhao et al. (2009) described a method for producing ~11 nm SIPPs using the hydrophobic surface ligand 1-Octadecylamine (ODA) along with iron and platinum salts that are much less hazardous than Fe(CO)5. In this article, we report the synthesis of SIPP cores using modifications of this less hazardous method. In order to use hydrophobic core nanoparticles in vivo, the particles must first be made biocompatible. A plethora of methods for encapsulating drugs and hydrophobic imaging agents, to instill biocompatibility, have been reported and include encapsulation using hydrophilic or amphiphilic components such as monomers and phospholipids (Serda et al. 2010). In addition, polyethylene glycol (PEG) groups on nanoparticles have been shown to increase solubility and circulation times and decrease immunogenicity (Serda et al. 2010; Pasut and Veronese 2009). This decrease in immunogenicity imparts the nanoparticles with stealth capability in vivo. Furthermore, biocompatible contrast agents that also incorporate a fluorescent component offer the advantage of in vivo and ex vivo imaging using small animal fluorescence imagers and/or fluorescence microscopy for in vivo biodistribution studies and in vitro binding assays. In this article, we describe the physical and magnetic characterization of SIPP cores encapsulated with a mixture of 1,2-distearoyl-sn-glycero-3-phosphoethanolamine-N-[amino(polyethylene glycol)-2000] (DSPE-PEG), DSPE-PEG with biotin conjugated to the head group (biotin-DSPE-PEG), 1,2-dimyristoyl-sn-glycero-3-phosphoethanolamine-N-[lissamine rhodamine B sulfonyl] (Liss-Rhod), and subsequent conjugation to a monoclonal antibody (J591) against PSMA (J591-DSPE-SIPPs). In this study, we also demonstrate the specific binding of J591-DSPE-SIPPs to PSMA-positive prostate cancer cells using confocal microscopy and measure the MR relaxivities of the DSPE-SIPPs at 4.7 Tesla. Compared to the commercially available and clinically used SPIONs, the J591-DSPE-SIPPs are superior contrast agents in T 2-weighted MRI, specifically target PSMA-positive human prostate cancer cells, and can be detected with fluorescence microscopy. To our knowledge, this is the first report on the synthesis of multifunctional SIPP micelles and the first report of using SIPPs for the specific detection of prostate cancer cells.

Experimental details

Materials

Iron nitrate nonahydrate (Fe(NO3)3·9H2O), Platinum (II) acetylacetonate (Pt(Acac)2), and ODA were purchased from Fisher Scientific (Pittsburgh, PA). Temperature controller (model 210-J) was purchased from J-KEM Scientific, INC (St. Louis, MO). Heating mantle was purchased from Glas-Col, LLC (Terre Haute, IN), and glassware was purchased from Quark Glass (Vineland, NJ). The phospholipids 1,2-distearoyl-sn-glycero-3-phosphoethanolamine-N-[amino(polyethylene glycol)-2000] (DSPE-PEG), DSPE-PEG with biotin conjugated to the head group (biotin-DSPE-PEG), and 1,2-dimyristoyl-sn-glycero-3-phosphoethanolamine-N-[lissamine rhodamine B sulfonyl] (Liss-Rhod) were purchased from Avanti Polar Lipids (Alabaster, AL). SPIONs were purchased from Miltenyi Biotec (Carlsbad, CA) as their MACS® Streptavidin MicroBeads product. RPMI cell culture media, fetal bovine serum, and Penicillin–Streptomycin Solution were purchased from Sigma (St. Louis, MO). All other chemicals and supplies were purchased from common manufacturers.

Synthesis of SIPP cores

Nanoparticles were synthesized using a modification of a procedure Zhao et al. (2009). In brief, 1.0 mmol Fe(NO3)3·9H2O and 1.0 mmol Pt(Acac)2 were added to 12.5 mmol ODA in a 25 mL 3-neck round bottom flask fitted with a reflux condenser. The reaction was heated to 330 °C (200 °C/h) with 10 °C recirculated cooling in the reflux condenser. Refluxing was continued for 45 min at which point the reaction was removed from the heat and allowed to cool to room temperature. The resulting black particles were collected in hexane and subjected to repeated washing by collecting particles in conical tubes with an external magnet, removing the supernatant, and resuspending in hexane.

Encapsulation of SIPP cores

Phospholipid-encapsulated SIPP cores (DSPE-SIPPs) were prepared using a thin film method. 1.5 mL of SIPP cores (1.4% solids) in hexane was added to a 20.0 mL glass scintillation vial. A chloroform mixture of (56:1:1 mol ratio) DSPE-PEG, biotin-DSPE-PEG, and Liss-Rhod was then added to the SIPP cores. The mixture was further diluted in 1.5 mL of hexane and vortexed thoroughly. The vial was wrapped in aluminum foil and allowed to evaporate in the dark in a chemical fume hood overnight to produce a thin film. 5.0 mL of double-distilled water was heated to 67 °C and added to the thin film. Hydration of the thin film was then continued in a 67 °C water bath for 1.0 h with vortexing every 15 min to produce liposomes containing SIPP cores. The liposomes were then extruded at 67 °C through an 80-nm nuclepore track-etch membrane filter using a mini-extruder (Avanti Polar Lipids, Alabaster, AL) to produce ~45 nm DSPE-SIPP micelles. The DSPE-SIPPs were then purified from SIPP-free micelles and excess phospholipids by collecting the magnetic particles using an LS magnetic column placed in a VarioMACSTM magnetic separator (Miltenyi Biotec, Carlsbad, CA). After the non-magnetic material had passed through the column, 8.0 mL of double-distilled water was added to the top of the column to wash the particles. The washing was then repeated a second time. The column was removed from the magnet and placed in a tube rack with a 2.0 mL glass vial placed underneath the column. 2.0 mL of either double-distilled water or phosphate buffered saline (PBS), pH 7.4, was used to elute the purified DSPE-SIPPs from the column.

Physical characterization of SIPP cores and DSPE-SIPPs

Transmission electron microscopy (TEM) was utilized to determine the size and polydispersity of the particle populations. For SIPP cores, a drop of the hexane suspension was applied to a carbon-coated grid and dried. For DSPE-SIPPs, a drop of the aqueous suspension was applied to a carbon-coated grid, dried for 10 min, and the excess was absorbed using a kimwipe. Adding a drop of 2% Uranyl Acetate solution followed by a 2-min drying period negatively stained the grid. The excess was removed, and the grid was allowed to dry for at least 5 min. The samples were imaged on a Hitachi 7500 TEM with an acceleration voltage of 80 kV. Particle diameters were calculated using ImageJ Software (Rasband 1997–2009). At least 1000 particles were counted, and the mean Feret diameters and standard deviations were calculated. Diameters of the DSPE-SIPPs were additionally measured using Dynamic Light Scattering (DLS) with a Microtrac NanotracTM Ultra DLS (Microtrac, Largo, FL). The compositions of the SIPPs, phospholipids, and DSPE-SIPPs were investigated with thermogravimetric analysis (TGA). Aliquots of ODA, SIPP cores, phospholipids, or DSPE-SIPPs were placed in the TGA sample cup and evaporated at 30 °C under an argon stream for at least 90 min until all solvent had been removed and the mass of the sample stabilized. Weight loss profiles were then measured with a PyrisTM 1 thermogravimetric analyzer (PerkinElmer, Waltham, MA) under argon flow. The ODA, phospholipid, and SIPP content were determined by measuring the mass loss profile while the temperature was raised from 30 to 1000 °C at a 10 °C/min ramp rate. Inductively coupled plasma-optical emission spectroscopy (ICP-OES) was used to measure the metal content and iron to platinum ratio of each synthesis. Before analysis, aliquots of the particles were digested at 180 °C with nitric and hydrochloric acids in a PDS-6 Pressure Digestion System (Loftfields Analytical Solutions, Neu Eichenberg, Germany). After cooling, the samples were made up to a known volume, mixed, and centrifuged. Samples were then analyzed using a PerkinElmer Optima 5300DV ICP-OES. The recommended wavelengths for each of the analytes were used, and analysis was performed in an axial mode to improve detection limits. A blank and set of calibration standards were used to establish a three-point calibration curve. Calibration and instrument verification samples were analyzed before and after analyzing the samples, as well as periodically throughout the measurements. Analyte peaks were examined and peak identification and background points were adjusted for optimum recoveries.

Magnetic characterization of SIPP cores and DSPE-SIPPs

Superconducting quantum interference device (SQUID) magnetometry was employed to measure the blocking temperatures of the SIPP cores, DSPE-SIPPs, and MACS® SPIONs and saturation magnetizations of the SIPPs and MACS® SPIONs. An aliquot (100 μL) of the samples were applied to the end of cotton Qtips® (Unilever, Englewood Cliffs, NJ). Magnetic measurements were then made on a Quantum Design MPMS-7 SQUID magnetometer. Temperature sweeps between 0 and 310 K were performed by zero-field cooling the sample and then measuring the magnetic moment as a function of temperature under the influence of a weak magnetic field (1 mT) during warming and subsequent cooling. This procedure yields both a zero-field-cooled (ZFC) and field-cooled (FC) curve, respectively. Values of the blocking temperature (T B ) were recorded by determining the peak location in each ZFC curve. Saturation magnetizations were measured at 310 K (37 °C) by varying the applied field from −5 to 5 Tesla. Mass magnetizations were calculated from the known iron concentrations determined by ICP-OES.

Magnetic resonance relaxometry

Increasing concentrations of SPIONs (0.08–0.48 mM iron) or DSPE-SIPPs (0.04–0.20 mM iron) were added to 1% agarose in 2.0 mL self-standing micro-centrifuge tubes (Corning, Corning, NY). Samples were imaged on a 4.7 Tesla Bruker Biospin (Billerica, MA) MRI system with Paravision 4.0 software. Samples were imaged with a 256 × 256 matrix, a variable TE, and TR = 10 s. T 1 measurements were acquired by inversion-recovery with 15 interpulse delays. Spin- and gradient- echo sequences were employed to measure T 2, and T 2*, respectively. The MRI samples were then digested as above, and the iron concentration was determined with ICP-OES. The relaxation rates, \( R_{n} = \frac{1}{{T_{n} }} \), were calculated and plotted versus the ICP-OES-determined iron concentration of each sample. Linear regression was used to fit the data, and the relaxivity (r n ) of each sample is given as the slope of the resulting line in units of s−1 mM−1 of iron.

Antibody conjugation, cell culture, and confocal binding assay

Humanized monoclonal antibody against PSMA (J591) (purchased from Neal Bander, Cornell College of Medicine) and polyclonal goat anti-rabbit IgG (Sigma, St. Louis, MO) were conjugated to streptavidin in an overnight reaction using a Lightning-LinkTM Streptavidin Conjugation Kit (Innova Biosciences, Cambridge, UK) according to the manufacturers’ instructions. Concentrations of streptavidin, antibodies, and streptavidin-antibody conjugates were quantitated using a NanoDropTM 2000 Spectrophotometer (Wilmington, DE). Streptavidin-conjugated antibodies (~30 μg) were then incubated with DSPE-SIPPs (100 μg iron) overnight at 4 °C to conjugate the antibodies to the DSPE-SIPPs through the biotin groups of the biotin-DSPE-PEG. A Micro BCATM Protein Assay (Thermo Scientific, Rockford, IL) was employed to quantitate the antibody concentrations and the amount of antibody conjugated to the DSPE-SIPP surface using a BioSpec-mini Spectrophotometer (Shimadzu, Columbia, MD) at a wavelength of 562 nm. 20,000 C4-2 or PC-3 human prostate cancer cells in RPMI media containing 10% fetal bovine serum and 100 U/mL Penicillin–Streptomycin solution were seeded onto polylysine-coated cover slips in 6-well polystyrene plates (Corning, Corning, NY) and incubated at 37 °C, 5% CO2 for 24 h. The media was then exchanged with media containing J591-DSPE-SIPPs (20 μg iron), IgG-DSPE-SIPPs (20 μg iron), or PBS (20 μL). The cells were incubated with the particles for 10 min at 37 °C, 5% CO2, and the media was then aspirated, and 5.0 mL PBS was added to wash unbound particles away from the cells. Washing was repeated three times. Cover slips were mounted on slides containing a drop of ProLong® Gold Antifade Reagent with DAPI (Invitrogen, Eugene, OR). Confocal Images were acquired using a 60X oil objective with an Olympus IX-81 inverted spinning disk confocal microscope. Cells were also imaged by light microscopy, using a Zeiss Axiovert 25 CA inverted light microscope with a 63X phase-contrast objective.

Results and discussion

SIPP cores and DSPE-SIPPs were prepared as described in the “Experimental details” section. Figure 1 shows TEM images of the SIPP cores and DSPE-SIPPs. The TEM images indicate that both the SIPP cores and the DSPE-SIPPs are spherical in shape. Using ImageJ software to analyze TEM images of the SIPP cores and DSPE-SIPPs, we measured average diameters of 8.5 ± 1.6 nm and 44.2 ± 13.1 nm, respectively. DLS was also employed to measure the size of the DSPE-SIPPs and revealed diameters of 42.9 ± 8.2 nm, showing good agreement with the diameters measured with the TEM images. In this study, we also used ImageJ to analyze the number of SIPP cores per DSPE-SIPP. We analyzed 175 DSPE-SIPPs and found that there were 7.2 ± 6.8 SIPP cores per DSPE-SIPP. The TEM images suggest that the number of SIPP cores per DSPE-SIPP is quite variable even though the overall encapsulated population is quite monodisperse, as is evident from the DLS and TEM size distribution data. Considering that the SIPP cores were found to be 8.5 nm in diameter, 5 SIPP cores would encompass a diameter of 42.5 nm, which is approximately the diameter of the DSPE-SIPPs (42.9 nm) and, on average, the greatest number of SIPP cores we observed spanning the diameter of the DSPE-SIPPs in the TEM images. This suggests that the DSPE-SIPPs do not contain an inner aqueous layer characteristic of a liposome, but rather have a purely hydrophobic inner layer that contains the hydrophobic SIPP cores. DSPE-PEG phospholipid bilayers are reported to be ~5.0 nm in thickness (Johnsson and Edwards 2003). It is extremely unlikely that an 8.5 nm hydrophobic SIPP core would fit into a 5-nm bilayer. Moreover, the TEM images in Fig. 1 show that there is space in between the hydrophobic SIPP cores in the inner layer of some of the DSPE-SIPPs. It is very unlikely that water would reside at this boundary between hydrophobic phospholipid tail and hydrophobic ODA on the SIPP core surface. Therefore, we suggest that the DSPE-SIPPs are not liposomes, but rather micellar contrast agents. Johnsson and Edwards (2003) analyzed particles prepared with increasing concentrations of DSPE and DSPE-PEG and found that concentrations >33 mol% DSPE-PEG resulted in micelle formation rather than liposomes. Our DSPE-SIPPs are prepared with ~98 mol% DSPE-PEG, and although liposomes and micelles reported by Johnsson and Edwards (2003) did not contain an additional hydrophobic superparamagnetic nanoparticle at the core, their results support the idea of micelle formation in our system. To investigate the stability of the DSPE-SIPP micelles, we also used TEM to image the DSPE-SIPPs up to 21 days post-synthesis. The particles were stored in PBS, pH 7.4 at 4 °C. The TEM images revealed no physical changes up to 12 days post-synthesis, at which point the DSPE-SIPPs began to merge and aggregate into larger particles (data not shown).

Fig. 1
figure 1

TEM and DLS of SIPP Cores and DSPE-SIPPs. TEM images of a SIPP cores and b, c DSPE-SIPPs. Scale bars are 20, 50 and 50 nm, respectively. Arrows denote internal areas of the DSPE-SIPPs where space can be seen between the hydrophobic SIPP cores. d DLS of DSPE-SIPPs in PBS

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To investigate the composition of the SIPPs, we used ICP-OES and measured an iron to platinum ratio (Fe:Pt) of 1.24:1 for the SIPP cores. The encapsulation process did not appear to significantly affect the Fe:Pt stoichiometry. To further investigate the composition of the SIPPs and DSPE-SIPPs, TGA was used to thermally decompose the particles and determine the weight percents of ODA, phospholipid, SIPP core, and naked FePt. The thermograms of ODA, SIPPs, phospholipids, and DSPE-SIPPs are shown in Fig. 2. ODA has a boiling point around 314 °C, and both ODA and the SIPP core samples show pronounced weight loss from ~180 to 375 °C due to the removal of ODA from the SIPP surface. The hump in the middle of the curve in Fig. 2a suggests the SIPP decomposition is a two-step process. It is possible that a portion of the ODA is not bound, but rather entrapped and being removed from the particles before the bound fraction. The TGA results suggest that the organic ODA layer comprised approximately 72% of the SIPP core mass and indicated that the SIPPs were 28% naked FePt by mass. The phospholipid and DSPE-SIPP samples showed similar weight loss profiles and continued to lose mass up to ~400 °C. The DSPE-SIPP thermogram revealed that the phospholipids comprised ~55% of the DSPE-SIPP mass, while SIPP cores made up the remaining ~45% of the DSPE-SIPP mass. The mass reduction seen in the thermogram of phospholipids (prepared in chloroform) at ~65 °C is likely due to release of residual chloroform which has a boiling point of 61.2 °C.

Fig. 2
figure 2

SIPP Core and DSPE-SIPP TGA. TGA thermograms of a SIPP cores (solid curve) and ODA (dashed curve) and b DSPE-SIPPs (solid curve) and phospholipids (dashed curve). Vertical dashed line denotes the temperature, reported to the left of the dashed line, at which the a SIPP cores and b DSPE-SIPPs stopped loosing significant mass. % values are the percent mass of each sample remaining at the temperature denoted by the vertical dashed line

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To characterize the magnetic properties of the SIPP cores and DSPE-SIPPs, we used SQUID magnetometry. Figure 3 shows the mass magnetization as a function of the applied magnetic field for the SIPP cores. Blank Qtips® were also scanned as controls but did not have any measureable effect in the SQUID (data not shown). The mass magnetization of the SIPP cores was 120 A m2/kg Fe. As a comparison, we also measured commercially available SPIONs (~50 nm MACS® MicroBeads, Miltenyi Biotec) that we have previously used as an MRI contrast agent (Serda et al. 2007). The SPIONs had a mass magnetization of 82 A m2/kg Fe, which is 1.5-fold lower than the SIPPs. SQUID magnetometry was also employed to measure the blocking temperatures of the SIPPs (Fig. 3), DSPE-SIPPs, and SPIONs, which were 210, 180–210, and 155 K (−63, −93 to −63, and −118 °C), respectively. All of these blocking temperatures are below body temperature, and no hysteresis is seen in the magnetization curves confirming the SIPPs, and DSPE-SIPPs, and SPIONs are superparamagnetic for biological applications. The broad blocking transition observed upon lipid encapsulation of the SIPPs is likely caused by the varying environments of the nanoparticles within the micelle, which alters the effective anisotropy energy of the particles. It has been shown that packing multiple magnetic cores into a single particle alters the measured anisotropy of the particles (Schaller et al. 2010). Next, we calculated the effective anisotropy of the SIPPs and SPIONs based on the blocking temperatures. The relationship between the effective anisotropy energy and the blocking temperature is \( K = \frac{{25kT_{B} }}{V}, \) where k is Boltzmann’s constant, T B is the blocking temperature, and V is the volume of the magnetic core in units of m3. The constant 25 is calculated using a relaxation time of 1 × 10−9 s and a measurement time of 100 s. Table 1 summarizes the physical and magnetic characteristics of the SIPP cores, DSPE-SIPPs, and SPIONs and shows that the SIPPs effective anisotropy energy is ~2-fold greater than for the SPIONs. The effective anisotropy constants for the SIPPs and SPIONs are in excellent agreement with anisotropy constants for SIPP cores (Maenosono et al. 2008; Salgueirino-Maceira et al. 2004) and SPIONs (Demortiere et al. 2010; Sohn et al. 1998) previously reported.

Fig. 3
figure 3

Magnetization of SIPPs. Saturation magnetization curves for the mass magnetization of SIPP cores versus the applied magnetic field from −5 to 5 Tesla. Inset shows the zero-field-cooled (ZFC) and field-cooled (FC) curves. Values of the blocking temperature (T B ) were recorded by determining the peak location in the ZFC curve

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Table 1 Physical and magnetic characterizations of SIPPs, DSPE-SIPPs, and MACS® MicroBeads
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The DSPE-SIPPs are prepared from a 56:1:1 mol ratio of DSPE-PEG, biotin-DSPE-PEG, and Liss-Rhod, respectively. The biotin-labeled phospholipid allowed us to conjugate streptavidin-labeled J591 to the DSPE-SIPPs. We measured ~2 streptavidin per J591 antibody and after conjugation, we calculated ~6 J591 antibodies per J591-DSPE-SIPP. DSPE-SIPPs were also conjugated to rabbit IgG antibodies as a non-targeted control (IgG-DSPE-SIPPs). We also measured ~2 streptavidin per IgG antibody, but ~12 IgG antibodies were measured per DSPE-SIPP. To determine if our J591-DSPE-SIPPs could specifically target PSMA-expressing human prostate cancer cell lines, we incubated the J591-DSPE-SIPPs and IgG-DSPE-SIPPs with C4-2 and PC-3 human prostate cancer cells grown on polylysine-coated cover slips. C4-2 cells were used as our PSMA-positive cell line. C4-2 is an androgen-deprivation therapy resistant cell line that over-expresses PSMA (Wolf et al. 2010; Sobel and Sadar 2005). PC-3 cells were used as a PSMA-negative cell line. PC-3 cells originate from a bone metastasis, are androgen-deprivation therapy resistant, and do not express or only minimally express PSMA (Kuroda et al. 2010; Sobel and Sadar 2005). Figure 4 shows the confocal microscopy images of C4-2 and PC-3 cells incubated with PBS (mock), J591-DSPE-SIPPs, and IgG-DSPE-SIPPs. Since the stealth immunomicelles are made with 1 mol% Liss-Rhod, the particles are fluorescent red in the confocal images. Both C4-2 and PC-3 cells lacked red fluorescence in the mock samples. Also, both cell lines only show minimal non-specific or IgG-specific uptake of the IgG-DSPE-SIPPs (non targeting control). The PSMA-negative cell line, PC-3, also only showed minimal non-specific or IgG-specific uptake of the J591-DSPE-SIPPs. The amount of non-specific J591-DSPE-SIPP uptake in the PC-3 cells appears to be comparable to the non-specific uptake of the IgG-DSPE-SIPPs. In stark contrast to the other images, significant uptake of the J591-DSPE-SIPPs can be seen in the C4-2, PSMA-positive cell line. Clearly, J591-DSPE-SIPPs are internalized only by the C4-2 cell line with only minimal non-specific pick up by the PC-3 cell line and only minimal non-specific pick up when IgG-DSPE-SIPPs were used. The confocal data demonstrates the successful targeting of the multifunctional DSPE-SIPPs and detection of PSMA-expressing human prostate cancer cells in vitro with no, or minimal, non-specific binding to cell lines that do not express PSMA.

Fig. 4
figure 4

Specific detection of PSMA-expressing Prostate Cancer Cells using J591-DSPE-SIPPs. C4-2 (top row), PSMA-positive, and PC-3 (bottom row), PSMA-negative, cell lines were imaged using phase-contrast light microscopy with a 63X objective (a, e). Cells were incubated for 10 min with either PBS (Mock) (b, f), J591-DSPE-SIPPs (c, g), or IgG-DSPE-SIPPs (d, h) and imaged using confocal microscopy with a 60X oil objective. Blue DAPI nuclear stain and Red Liss-Rhod incorporated in the DSPE-SIPPs

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Finally, to test whether the DSPE-SIPPs could be beneficial as MRI contrast agents, we measured the longitudinal (T 1), transverse (T 2), and T 2-star (T 2*) relaxation rates of the DSPE-SIPPs and commercially available SPIONs. Table 2 shows the relaxivities measured at 4.7 Tesla, while Fig. 5 shows the T 2-weighted MR image of the DSPE-SIPP agarose samples, as well as the transverse relaxation rates of the DSPE-SIPPs and SPIONs as a function of iron concentration. It is apparent that the DSPE-SIPPs have a ~13-fold higher r 2 than the SPIONs, a measure of the particles ability to create negative contrast in the MR images, and a 1.5-fold increase in the r 2/r 1 ratio. As expected, the SIPPs had increased magnetizations compared with the SPIONs and far superior transverse relaxivities. Since the commercially available SPIONs had such low transverse relaxivities, we also compared relaxivities of the DSPE-SIPPs with relaxivities of the clinically used SPION contrast agents Feridex® and Resovist® that are reported in the literature (Patel et al. 2010; Yang et al. 2010; Figuerola et al. 2008; Maenosono et al. 2008; Hinds et al. 2003). Table 3 shows the comparison of the relaxivities at 4.7 Tesla for the DSPE-SIPPs, Feridex®, and Resovist®. Compared to Feridex® and Resovist®, the DSPE-SIPPs produce superior negative contrast enhancement in MRI, as is evident from the 1.6- to 3-fold higher r 2. Overall, our data suggest that J591-DSPE-SIPPs are stable, superparamagnetic, specifically target PSMA-positive human prostate cancer cells, useful for fluorescence detection for in vitro binding applications, and superior contrast agents in T 2-weighted imaging when compared with both commercially available and clinically used SPIONs in vitro.

Table 2 MR relaxivities of DSPE-SIPPs and MACS® MicroBeads at 4.7 Tesla
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Fig. 5
figure 5

Magnetic resonance relaxometry of DSPE-SIPPs and MACS® MicroBeads at 4.7 Tesla. a T 2-weighted MRI of 1% agarose samples containing increasing concentrations of DSPE-SIPPs. Top left sample is agarose that did not contain DSPE-SIPPs. The other samples have increasing concentrations of DSPE-SIPPs going from left to right in the top row images and continuing from left to right in the lower row images. b Transverse relaxation rates (1/T 2) versus iron concentration (mM) for the DSPE-SIPPs (squares) and MACS® MicroBeads (triangles). Linear regression was used to fit the data (solid lines) and the transverse relaxivities (r n ) of the DSPE-SIPPs and MACS® MicroBead SPIONs, given as the slope of the resulting line, were 300.8 ± 8.5 and 23.8 ± 6.9 s−1 mM−1, respectively

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Table 3 Contrast agent relaxivity comparison at 4.7 Tesla
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Summary and conclusions

In 2009, more than 200,000 men were newly diagnosed with and over ~30,000 men died due to prostate cancer in the United States, making carcinoma of the prostate the second most lethal cancer in men in the United States (Jemal et al. 2010). New detection methods are critically needed to achieve earlier diagnosis and better staging of the disease. SPION contrast agents have been used to enhance the contrast of tumors in MRI, but novel contrast agents with increased relaxivities could be useful in detecting smaller tumors earlier and with lower doses of the contrast agents. In addition, the specific targeting of contrast agents and therapeutics to cells of interest is now widely accepted as a cornerstone to the development of individualized diagnosis and treatment. In this article, we report the synthesis of SIPP core particles from simple salts and their subsequent encapsulation in a mixture of phospholipids and conjugation to a monoclonal antibody against PSMA to produce stable, water soluble, multifunctional contrast agents with targeting, fluorescent, and MRI capabilities for the specific detection of prostate cancer cells. To our knowledge, this is the first report of the synthesis of multifunctional SIPP micelles and the first report of using SIPPs for the specific detection of prostate cancer.

The SIPP cores have a large effective anisotropy energy of 2.5 × 105 J/m3 and magnetic moment of 120 A m2/kg Fe. In this study, we expected that the particles with higher mass magnetizations would be better contrast agents for MRI. We found this to be true when comparing the different composition particles. Compared to the SPIONs, the SIPPs have a higher magnetization accompanied by a ~13-fold higher transverse relaxivity at 4.7 Tesla. TGA suggests that the particles are ~45% SIPP core and ~55% phospholipid. The TEM images show that the SIPP cores and DSPE-SIPPs have diameters of 8.5 ± 1.6 nm and 42.9 ± 8.2 nm, respectively. The DSPE-SIPPs are spherical and contain 7.2 ± 6.8 SIPP cores per DSPE-SIPP. These structural characterizations suggest that the DSPE-SIPPs are micellar contrast agents. Using fluorescence confocal microscopy, we determined that the J591-DSPE-SIPPs specifically bound to C4-2 human prostate cancer cells that over-express PSMA and did not bind to PC-3 cells that do not express PSMA. In addition, IgG-DSPE-SIPPs did not accumulate in either cell line. This shows the specific detection of PSMA-expressing human prostate cancer cells using the fluorescent capabilities of the SIPP immunomicelles. Finally, we show that the DSPE-SIPPs were 13-fold better than commercially available SPIONs and 1.6- to 3-fold better than Feridex® and Resovist® at producing negative contrast in MRI, at 4.7 Tesla. Taken together, our data suggest that the multifunctional SIPP immunomicelles are superior contrast agents for T 2-weighted MRI, specifically target PSMA-expressing human prostate cancer cells, can be used to specifically detect human prostate cancer cells in vitro using fluorescence microscopy, and should be beneficial as MRI contrast agents. Future studies will include using MRI to specifically detect human prostate cancer cells in vivo using the SIPP immunomicelles. It is important to note that the DSPE-SIPPs could be conjugated to any antibody or peptide for selective targeting and non-invasive detection of other types of tumors, using MRI. An additional benefit to this multimodal platform is that the in vivo biodistribution of the nanoparticles could be measured by examining the tissues and tumors in vivo and/or ex vivo, using small animal fluorescent imagers and fluorescence microscopy. Overall, our data suggest that J591-DSPE-SIPPs specifically target human prostate cancer cells in vitro, can be easily detected using fluorescence microscopy, and are superior contrast agents in T 2-weighted MRI.