Article pubs.acs.org/molecularpharmaceutics
Intracellular Degradation of Multilabeled Poly(Ethylene imine)− Mesoporous Silica−Silica Nanoparticles: Implications for Drug Release Lotta Bergman,†,‡ Pasi Kankaanpaä ,̈ § Silja Tiitta,§ Alain Duchanoy,† Ling Li,† Jyrki Heino,§ and Mika Lindén*,‡ †
Laboratory for Physical Chemistry, Åbo Akademi University, Porthansgatan 3-5, FI-20500 Turku, Finland Inorganic Chemistry II, University of Ulm, Albert-Einstein-Allee 11, D-89081 Ulm, Germany § Department of Biochemistry and Food Chemistry, Vatselankatu 2, Arcanum third floor, University of Turku, FI-20014 Turku, Finland ‡
S Supporting Information *
ABSTRACT: Mesoporous silica nanoparticles, MSNs, have emerged as an interesting carrier for drugs in vitro and in vivo. The particles are typically used in a surface functionalized form, where functional silanes or other covalently linked surface functions are used to provide anchoring sites for additional functionalities like targeting groups, imaging agents, and drugs. Here, we report results related to extra- and intracellular degradation of silica nanoparticles using multilabeled nonporous silica core−mesoporous silica shell−surface hyperbranched poly(ethylene imine) shell nanoparticles as model particles. Different fluorophores have been selectively covalently linked to different regions of the particles in order to study the particle degradation in detail under in vitro conditions in human SAOS-2 cells. A novel, quantitative method for nanoparticle degradation evaluation based on confocal fluorescence microscopy is applied. Our results suggest that the core−shell−shell MSNs degrade at a higher rate inside cells as compared to outside cells, which is of high importance for further application of this class of drug carriers. KEYWORDS: mesoporous silica nanoparticles, degradation, fluorescence microscopy, surface functionalization
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face functionalization is relatively straightforward13−15 which allows fine-tuning of the drug−support and particle−bioenvironment interactions. In most cases, active targeting of MSNs has been demonstrated under in vitro conditions but has also been shown to work in vivo.16−19 However, in order for a nanoparticulate carrier system to be effective, premature leakage of the drug before reaching the target cells has to be kept at a minimum. This is especially important for MSNs, as the release rate from carriers where the drug release is diffusioncontrolled is highest initially and also often connected with an initial burst release.20 The extent of drug leakage is naturally also dependent on the physicochemical properties of the drug, such as drug solubility (often pH-dependent), and the degradation rate of the MSNs, the circulation time before reaching the target cells. Much recent focus has been put on developing means for drug release that can be triggered by intracellular processes or by external stimuli. There are two
INTRODUCTION
Targeted delivery of drugs is one of the most promising approaches for the delivery of drugs associated with severe side effects, especially of importance for chemotherapy-based cancer treatment.1−3 Different nanoparticle-based drug delivery systems have been shown to accumulate in tumors either by passive or active targeting and to be taken up by target cells. While passive targeting is based on nanoparticle accumulation in tumors through the leaky nature of the tumor vasculature, active targeting is based on the attachment of cell-specific ligands onto the nanoparticle surface that are recognized by receptors overexpressed on the target cells.4,5 Nanoparticlebased drug delivery has several attractive features in addition to cellular targeting, the most important being the possibility to achieve high drug-loading levels and controlled release profiles. A promising new nanoparticle platform attracting wide current interest is based on amorphous mesoporous silica nanoparticles (MSNs).6−8 MSNs can be synthesized with controlled particle sizes and shapes,9 and the pore dimensions can be tuned within a range of some nanometers to tens of nanometers, allowing both small molecular drugs and proteins or genes to be accommodated inside the mesopores.10−12 Furthermore, sur© 2013 American Chemical Society
Received: Revised: Accepted: Published: 1795
October 13, 2012 February 21, 2013 March 19, 2013 March 19, 2013 dx.doi.org/10.1021/mp3005879 | Mol. Pharmaceutics 2013, 10, 1795−1803
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amount of the drug model has been detached from the particles mainly in the form of a PEI-mesoporous silica complex. The study represents a first step toward reaching a better understanding about the intracellular decomposition of functionalized MSNs.
main approaches for achieving triggerable drug release; covalent linking of the drug to the support through cleavable bonds, or functionalization of the outer surface of the MSNs using sheddable coatings or coatings which change conformation upon environmental changes, typically pH, redox level, or temperature. Several in vitro studies have demonstrated that such strategies do indeed decrease the level of premature drug release and also allow for subsequent intracellular release of the cargo.21−24 Examples are bonds that can be cleaved at pH values lower than that in the plasma, as is the case in intracellular compartments and also within the interstitial space of solid tumors and within inflammatory tissues. Recently, we studied the therapeutic efficiency of MSNs surface functionalized by hyperbranched poly(ethylene imine), PEI, to which the cancer drug methotrexate (MTX) had been covalently linked.25 MTX is an antimetobolite which inhibits the enzymatic activity of dihydrofolate reductase (DHFR), thus blocking the biosynthetic pathway of nucleotides and proteins.26 Here, MTX served both as a targeting ligand and the drug, as MTX is structurally very similar to folic acid, an often employed targeting ligand, and both molecules are taken up by similar cellular routes. However, MTX has to reach the cytoplasm in order to be therapeutically active. Based on our findings we tentatively suggested that MTX predominantly remained covalently attached to the particles when outside the cells, and that MTX detachment occurred readily after particle endocytosis. Several possible explanations could account for this observation, including particle degradation through silica dissolution,27−29 detachment of the PEI layer to which MTX is attached from the MSNs after particle endocytosis by the cells, enzymatic degradation of the peptide bond30 linking MTX to the particles once the particles are inside endosomal vesicles, or a combination of these. In any case, the kinetics of detachment remains an open question. Furthermore, it was recently suggested that MSNs are exocytosed by cancer cells and that these are subsequently taken up by other cancer cells,31 and one important question is if there would still be drug molecules present in MSNs that can be released in such “second generation” cells or if all the cargo was already released in the cell of first entry. PEI-coated MSNs have also been suggested to preferentially lead to intracellular release of hydrophobic drugs or drug models physically adsorbed into the mesopores of the MSNs.32 Clearly, the answer to such questions is bound to be strongly system dependent, but a question is also how this could be studied experimentally. Here we present data where nonporous silica core−mesoporous silica shell particles have been synthesized with a PEI layer attached to the mesoporous shell of the particles, thus creating core−shell−shell MSNs. All three parts of the particle were covalently labeled with different fluorescent dyes so that the detachment of different parts of the particle could be studied and evaluated semiquantitatively. A three-layer model particle design was chosen to ensure that we would be able to track the main particle independently from that of the mesoporous portion of the MSNs, as nonporous silica is degrading at a slower rate than its mesoporous counterparts. Confocal fluorescence microscopy studies were carried out on human osteosarcoma SAOS-2 cells as a function of incubation time, and the images were analyzed using the BioimageXD software,33 allowing particles being located outside of the cells to be distinguished from particles internalized by the cells. The data suggests that, even after four hours, particles inside cells still do partly contain the PEIfluorophore layer representing the drug model, while a fair
2. EXPERIMENTAL SECTION 2.1. Synthesis of Three-Layer Fluorescent Silica Nanoparticles. The solid silica particle core was prepared based on the procedure described by Stöber et al.34 In a typical synthesis, 250 μg (1 mg/mL) of fluorescein isothiocyanate isomer I, FITC (minimum 90% HPLC, Sigma-Aldrich), was mixed with 3-aminopropyltriethoxysilane, APTS (SigmaAldrich), under inert atmosphere and added to an alkaline (ammonium hydroxide solution, max 33% NH3, puriss., SigmaAldrich) solution together with tetraehoxyothosilicate TEOS (purum ≥98% GC, Fluka). The resulting synthesis mixture had molar ratios of 0.1 FITC:242 APTS:4630 TEOS:1892 NH4OH:129684 H2O:266744 EtOH. The solution was stirred overnight 350 rpm at RT. The mesoporous surface layer was then introduced based on the method described by Kim et al.35 The nonporous silica nanoparticles were separated, washed carefully, and dispersed into basic reaction solution. The structure-directing agent CTAB (Sigma-Aldrich) and finally TEOS together with tetramethylrhodamine isothiocyanate TRITC (Sigma-Aldrich) fluorophore conjugated with APTS were continuously added to the synthesis, here proceeding step by step according to the reference. The resulting solution had molar ratios of TRITC 0.01:APTS 0.025:CTAB 158:TEOS 440:NH4OH 2882:EtOH 152574:H2O 615220. Synthesis was stirred overnight at 500 rpm, and particles were separated, washed, and dried in vacuo at 298 K. To remove surfactant, particles were extracted under sonication; 30 min in acidic ethanol (1:8-mixture of HCl and absolute ethanol). This treatment was repeated three times to ensure complete surfactant removal. Particles were further separated by centrifugation and carefully vacuum-dried overnight at 298 K. The so-synthesized core−shell particles were further surface-modified by hyperbranching polymerization of polyethyleneimine (PEI), using aziridine as a precursor. Aziridine was synthesized from aminoethylsulfuric acid (Sigma-Aldrich, Miss, USA) according to the procedure described by Allen et al.36 The surface polymerization of PEI was performed in one step under argon as the protective gas. Particles were dispersed in toluene, and catalytic amounts of acetic acid were added, after which aziridine was added. In a typical conjugation, 35 μL of aziridine for 100 mg particles was used. The suspension was refluxed under stirring overnight at 348 K, filtered, washed, and vacuum-dried at 298 K. The third fluorophor, Alexa633, was linked to the particle described above by attaching it to the surface amino groups. In a typical procedure, 100 mg of particles were dispersed in toluene. Surface primary amino groups were activated with DIPEA (100 μL, 1 μL/mL in DMF) and further conjugated with Alexa633 fluorophore (200 μL, 1 μL/mL in DMF) under shaking for 2 h at RT (298 K). Finally the particles were separated, washed with ethanol, and vacuum-dried at 298 K. 2.2. Particle Characterization. The structure of the nanoparticles was confirmed by low angle, main reflection at 2.2° 2Θ, powder-XRD using a Kratky compact small-angle system (Hecus Braun, Austria), Seifert ID-300 X-ray generator with maximum intensity of 50 kV and 40 mA, and sample-todetector distance of 267 mm. 1796
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ships of the different colors inside the cells, and all pixels in the images were analyzed, unlike in segmentation-based analyses. Colocalization analyses were done by using automatic thresholding and then recording all six possible Manders’ colocalization coefficients, and observing changes in these between the three time points; 5 min, 1 h, and 4 h. All image analysis was done with the BioImageXD software.33 The results were statistically analyzed with a t test for unequal variances and sample sizes, and significance was marked onto graphs as follows: ns = p > 0.05, * = p < 0.05, ** = p < 0.01, *** = p < 0.001.
Thermogravimetric analysis was performed in air with a Netzsch STA 449C cell setup with a heating rate of 10 K/min. Dynamic light-scattering (DLS) and zeta-potential measurements were performed using a Nano ZS (Malvern, Worcestershire, UK) setup in a HEPES buffer. Measurements were performed at 298 K, using a monochromatic laser, with a working wavelength of 632.8 nm and using non-invasive backscatter (NIBS), with the detector positioned at 173° relative to the laser beam. Scanning electron microscopy (Jeol JSM-6335F, Jeol Ltd., Japan) was performed using an acceleration voltage of 10 kV and working distance of 9.6 mm. Transmission electron microscopy (TEM) micrographs were measured using a Zeiss Libra 120 TEM setup operated at 80 kV. Nitrogen adsorption−desorption experiments (ASAP 2010 sorptometer, Micromeritics) were carried out at 77 K. All samples were degassed for 8 h at 323 K before measurements. The specific surface area was determined by the BET method, and the pore dimensions were determined using the BJH method (desorption branch). 2.3. Confocal Imaging and Image Analysis/Quantification. Particles were dispersed (1 mg/mL) in water one hour before their delivery to cells. As silica particles do not hold intact in aqueous solvents and a relatively fast process of dissolution is reported for contact times below 1 h, the delay has to be taken into account when interpreting the imaging results. In this study, we concentrate on what happens to the particles in in vitro conditions at the exposion time point and 1 and 4 h thereafter. Human SAOS-2 cells were incubated in Dulbecco’s modified Eagle’s medium with the particles at a concentration of 125 μg/ mL. The cells were fixed with 4% paraformaldehyde (20 min RT) after 5 min, 1 h, or 4 h, and embedded in Mowiol. Thickness-selected cover glasses (0.17 ± 0.01 mm/Assistent) were used to minimize fluorescence intensity fluctuations. 2D confocal images were acquired with a Zeiss AxioObserver Z1 (objective 63x, 1.4) equipped with LSM 510, both near the glass surface and from the upper half of the cell. All three wavelengths (488, 543, and 633 nm) were excited and recorded separately (multitracking) line-by-line, with pinhole-calibrated dichroics and filters optimized for minimum bleed-through, and pinholes were adjusted for an optical slice thickness of 700 nm and pixel size of 94 nm × 94 nm for every wavelength. Acquisition settings and conditions were kept as constant as possible to enable the comparison of fluorescence intensities. Importantly, images were always acquired with the same pixel density, roughly following the Nyquist theorem, and the detector sensitivity and background offset were kept constant at such values that the whole intensity range of the samples was recorded without saturation. Regions of interest (ROI) were drawn onto the images so that particles outside cells were analyzed separately from particles inside cells. In total approximately 40 images from inside and 40 from outside the cells were analyzed for each time point. Segmentation-based analyses were done by first filtering the images with hybrid median 2D and Gaussian smoothing (dimensionality 3:4, 4, 2), then thresholding them for maximum object number (minimum object size 10) and finally running object separation (level 1, image spacing used). The segmented objects were then analyzed for the intensities of each of the three wavelengths as well as average object size. Colocalization studies were performed to confirm the relation-
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RESULTS AND DISCUSSION An scanning electron microscopy (SEM) image of the core− shell−shell MSNs is shown in Figure 1a. The particles have a diameter of about 240 nm and have a very narrow size distribution. A TEM image of a native silica core−shell MSN is shown in Figure 1b. The core−shell structure of the MSNs is clearly seen, and the mesoporous shell is uniform. From the TEM image a mesoporous silica layer thickness of about 12 nm can be estimated, which corresponds to about 30% of the total volume of the MSN. A nitrogen sorption isotherm measured for the native core−shell MSNs is shown in Figure 2. A pronounced uptake at a relative pressure close to 0.3p/po is characteristic for MSNs synthesized using C16TAB as a structure-directing agent and corresponds to filling of mesopores with a diameter of about 4 nm with a narrow pore size distribution. The BET specific surface area was 344 m2/g, and the specific pore volume of the primary mesopores was 0.34 cm3/g, which is slightly higher than that expected from the relative contributions of the mesoporous surface layer and the solid core, suggesting that the thickness of the surface layer based on TEM might be underestimated. The pronounced uptake at higher p/po is due to filling of interparticulate porosity. The low-angle XRD pattern (Figure 3c) exhibited a main reflection at 2.2° 2Θ and additional intensity in the regions expected for (11) and (20) reflections of a 2D hexagonal mesophase, suggesting that the mesoporous surface layer is not disordered but has a structural motif similar to that observed for corresponding MSNs void of the solid core. Assuming a 2D hexagonal arrangement of pores, the lattice spacing derived from the (10) reflection is 4.83 nm. Before PEI functionalization, the core−shell MSNs have a zetapotential of −27 mV in 25 mM HEPES pH 7.2, which increases to +52 mV after PEI functionalization, reflecting successful attachment of the highly cationic PEI layer. Thermogravimetric analysis shows a mass loss of 7.1 wt % in comparison to reference particles lacking PEI. Upon attachment of the Alexa 633 dye to PEI, the zeta-potential increased slightly to 55 mV. The particles could be fully dispersed in HEPES, as evidenced by dynamic light-scattering (Figure S1). The fluorescence emission colors of the different parts of the core−shell MSNs are schematically shown in Figure 1a. The total amounts of fluorophores covalently attached to the PEI-MSNs were analyzed by dissolving the particles in 2 M NaOH-solution 1 mg/mL and analyzing the dye content in the supernatant by fluorescence spectrometry. The PEI-MSNs contained Alexa 633 1.09 μg/mg particles (attached to the PEI layer), TRITC 0.12 μg/mg particles (attached to the mesoporous silica layer), and FITC 0.04 μg/mg particles (attached to the nonporous silica core). Fluorophore leakage was studied under in sink conditions (particle concentration 75 μg/mL) in HEPES buffer (pH 7.2) at 37 °C, and no leakage of FITC or Alexa 633 was observed, 1797
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Figure 2. (a) SAXS graph shows the ordered porosity of the mesoporous surface layer. (b) Nitrogen physisorption isotherm for prepared core−shell particles. (c) Nitrogen physisorption BJH analysis shows narrow pore size distribution in the mesoporous layer.
any detectable fluorescence signals could positively be attributed to fluorescence originating from the particles. While Alexa633 and TRITC show pH-independent fluorescence intensities in the pH range of interest in our study (pH about 5−7.2), the fluorescence of FITC is known to be strongly pH dependent due to the pKa of FITC of 6.4. Tests using free FITC in buffers showed a decrease in the fluorescence intensity of about 50% at pH 6.5 and of about 85% at pH 5.5 as compared to the intensities measured for the same FITC concentration at pH 7.5, in good agreement with literature values.37 Performing similar experiments using the core−shell particles void of PEI resulted in a FITC fluorescence intensity decrease of 60% at pH 5.5 as compared to pH 7.5; that is, the decrease in emission intensity was lower than that of the free fluorophore. The smaller pH dependency of the FITC fluorescence emission intensity when incorporated into Stöbertype silica is tentatively attributed to different local pH values experienced by the probe inside the Stöber particles (silica
Figure 1. Three color core−shell particles. (a) Schematic representation of the particle structure. Fluorescent dyes are covalently linked inside the silica network and likewise covalently linked to the surface polyethylene imine amino groups. (b) Scanning electron microscope image (SEM), scale bar corresponds to 1 μm. (c) Transmission electron microscope image (TEM), scale bar corresponds to 20 nm.
but about 10% of the TRITC fluorophore was detected in the supernatant after the 4 h incubation time. It was also ensured that the free fluorophore, defined by the maximum concentration of FITC used in the study, could not be detected by confocal fluorescence microscopy under the applied imaging conditions. In the presence of cells no clumps or clusters of free fluorophore could be detected, and therefore 1798
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Figure 3. Typical examples of the confocal fluorescence microscopy images analyzed, here from the green channel at the 1 h time point. Left: original images, middle: original images with regions of interests (ROIs) drawn, right: segmentation results within the ROIs. Regions marked with A: segmented objects analyzed as intracellular. Regions marked with B: segmented objects analyzed as extracellular. Image intensities have been enhanced by linear intensity transfer functions to improve the visualization. Upper row: Image taken close to the upper part of the cells. Lower row: Image taken close to the glass substrate. The scale bar corresponds to 10 μm. Pseudocolored versions of all of the three fluorescence channels observed is available as Supporting Information for the image in the lower row.
Figure 4. Confocal images taken from the yellow channel from the top part of the cell.
contains Bröstedt acidic silanol groups),38 but the results also highlight that the Stö ber particles are not completely nonporous, as external pH changes can indeed be felt by the dye. Importantly, no pH-dependent changes in the fluorescence intensity of FITC were observed for PEI-coated core−shell particles, suggesting that the presence of PEI buffered the pH inside the core−shell particles. These results will be discussed below as a means for investigating the detachment kinetics of PEI from the MSNs, in addition to the analysis of the locus of the different fluorescent dyes. For clarity, the different fluorescence emission colors will be indicated together with their locus in the particles as follows; green (C) for FITC-core, yellow (MP) for TRITC-mesoporous shell, and red (PEI) for Alexa 633-PEI. 3.3. Confocal Image Analysis. 3.3.1. Segmentations. Representative segmentations of the SAOS-2 cells as a function of particle incubation times ranging from 5 min to 4 h are shown in Figure 3. The images were acquired in such a way that it could be distinguished whether fluorescence was originating from particles located outside or inside the cells. When close to glass surface (but still slightly above it) and inside the cell perimeter, one can specify from the 3D imaging characteristics of the confocal microscope the objects located inside and outside the cell. The same is true higher up, close to the summits of the cells, when images were taken slightly below the summit and also inside the cell perimeter. The cell surface is clearly identifiable in the microscopy images shown in Figure 3, and regions of interest (ROI) were drawn to allow independent analyses of intracellular and extracellular regions. As can be seen in Figure 4 (5′), MSNs are internalized already at the 5 min time point, and the number of internalized MSNs increases
with time. The MSNs are compartmentalized inside the cells, in agreement with the well-established endosomal uptake of this type of particles, while particles located outside the cells appear to be well-dispersed. 2D image analysis of particle cluster size inside and outside of the cells (Figure 5) appreciates a decrease of cluster size inside the cells with time. Here, cluster sizes defined by confocal microscopy are to be evaluated for relative differences only, not as absolute values, because of light scattering as defined by the point spread function. Fast initial (5′) internalization leads to
Figure 5. Segmented object size (pixels) from the yellow channel, inside cells (■) and outside cells (▲) as a function of incubation time. The analysis is based on the yellow channel (TRITC-MP), as it showed highest fluorescence intensity, but qualitatively similar results were obtained also when performing the analysis based on the green or the red channel. 1799
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Figure 6. Intensity of the segmented objects over time, measured both inside and outside the cells. (a) Separate segmentations outside the cells. (b) All colors segmented in relation to yellow channel outside the cells. (c) Separate segmentations inside the cells. (d) All colors segmented in relation to yellow channel inside the cells. [ns] indicates nonsignificant change, and stars (*), up to three, describe the degree of a significant change. (yellow ■, red ●, green ▲).
against the total yellow fluorescence intensity. Also in these measurements no significant changes in the dye intensities outside the cells over time were detected. This suggests that particles located extracellularly remained stable or dissolved in a homogeneous manner. The time-dependency of the fluorescence intensity measured for the three different fluorophores originating from particles located inside the cells followed a distinctively different pattern, as shown in Figure 6c and d. While the total fluorescence emission intensity of the yellow (MP) remained virtually constant over time, a clear decrease in the corresponding total intensities of the red (PEI) and green (C) was observed significantly with increasing incubation time. The decrease in the red (PEI) intensity, both in the individual segmentations and when normalized against the intensity of the yellow fluorescence, is consistent with a partial detachment of PEI from the particles with time together with dilution of the detached PEI, possibly due to endosomal escape.40,41 This leads to a decrease of the red fluorescence observed in these pixels to values below the set detection limits. The decrease in the green (C) fluorescence intensities with time supports this suggestion, due to the observed decrease in the FITC fluorescence emission intensities with decreasing pH in the absence of pH-buffering PEI on the particles. 3.3.2. Colocalization Analyses. Colocalization analyses of the three different fluorophores were carried out in order to get a more detailed picture of the particle degradation process. The
high particle concentration in intracellular compartments. Our earlier internalization studies with SAOS-2 cells show an increase in integrin cluster size up to 45 min, whereafter the cluster size starts to decline.39 Similarly, in the present study, the point measurement at 1 h shows a decrease of intracellular cluster size. Confocal images from the yellow channel at all three time points (Figure 4) support the numerical data: initially (5′) particles are seen to form larger clumps. At the 1 h time point, the clusters are smaller and fainter, and they are perhaps closer to the nucleus as internalization has progressed. This is a trend that continues further at the 4 h time point. A more detailed discussion about the potential locus of MSNs as a function of time is given below, in connection with a more detailed analysis of the time-dependent fluorescence measurements of the different colors. All three fluorescent colors were segmented separately, and the absolute intensity of each fluorescent color was plotted over time. Interestingly, outside the cells no significant changes in the fluorescence intensities of single fluorophores from individual channel-spesific segmentations were observed (Figure 6a). Additionally, the fluorescence intensities of all colors in relation to the yellow channel were calculated (Figure 6b). Intensities of all colors were then measured based on the yellow channel segmentations; that is, pixels having yellow fluorescence were used as the basis of the total intensity of the green and red fluorescence, and the intensities were normalized 1800
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results are expressed as Manders’ coefficients, always for two colors at a time. First, the number of pixels that contain both colors are determined, and the total intensity of a given color within these pixels is divided by the total intensity of one or the other color within that segment. Thus, the Manders’ coefficients of, for example, red toward yellow and yellow toward red, are not necessarily the same. For example, red could be completely colocalized with yellow, which would correspond to a Manders’ coefficient of 1, while there could be yellow pixels which would not coincide with pixels showing both red and yellow, thus resulting in a Manders’ coefficient smaller than 1. It should also be noted that the determined Manders’ coefficients are more sensitive to the stable particles as compared to potentially detached fluorophores, as detached fluorophores are diluted and may fall below the set threshold values. This also implies that intracellular analyses are more sensitive to colors remaining inside cellular compartments rather than being located in the cytoplasm for the same reasons. The analyses of the Manders’ coefficients were carried out separately for particles located inside or outside of the cells. Interestingly, the changes in the Manders’ coefficients for particles located outside the cells was within experimental error for the time-period studied (data not shown), indicating that the particles contained all fluorophores up to the time-point of 4 h. Importantly, this does not mean that there was no particle dissolution, but it indicates that the particles did at least not completely disintegrate nor did any of the layers completely detach from the main particle during the time of observation. Inside the cells, the situation was quite different. The Manders’ coefficients of all possible color-combinations inside the cells are shown in Figure 7a−c. Depending on the combinations, the Manders’ coefficients, that is, the colocalization of different colors, are remarkably different when followed as a function of incubation time. The Manders’ coefficient for green (C)−yellow (MP) remain high over the time of observation, while the corresponding values for yellow (MP)−green (C) decrease significantly over time. This suggests that the mesoporous silica layer is detaching from the main particles, leaving some of the mesoporous layer behind. Also the Manders’ coefficient for red (PEI)/yellow (MP) remain high throughout the experiment, while the Manders’ coefficient for yellow (MP)/red (PEI) decreases significantly over time. This result is consistent with a dissolution process where the mesoporous layer is dissolved in a way that some of the mesoporous silica layer is detaching together with the PEI layer from the particles, while a portion of the mesoporous silica layer is still associated with the main particle. This is also seen in the high Manders’ coefficient for green (C)/yellow (MP) throughout the experiments. Interestingly, the corresponding Manders’ coefficients for red (PEI)/ green (C) and green (C)/red (PEI) cannot be explained using this simple dissolution model. The Manders’ coefficients indicate that the colocalization of green (C)/yellow (MP) remain high throughout the experiment, as expected based on the values of the discussed Manders’ coefficients, while significantly decreasing colocalization is seen in the decreasing values of the Manders’ coefficient for yellow (MP)/green (C) with time. This suggests that the core is separating from the PEI layer, while the PEI layer remains on the Stöber particle core. This finding can be explained by a coexistence of particles where some fraction of the particles have lost the PEI layer, while some particles still have a PEI layer on them. As discussed above, the detachment of PEI, also supported by the time-
Figure 7. (a) Manders’ colocalization coefficients as a function of incubation time (red/green ■ and green/red ⧫), (b) Manders’ colocalization coefficients as a function of incubation time (green/ yellow ■, yellow/green ⧫), and (c) Manders’ colocalization coefficients as a function of incubation time (red/yellow ■, yellow/ red ⧫).
dependent total fluorescence intensity analysis, together with the escape of PEI into the cytoplasm, will lead to an overestimation of the amount of PEI present in intracellular compartments. Here PEI may be attached to particles or to fragments of mesoporous silica still being present within intracellular compartments. However, the results clearly suggest that the PEI layer is detaching from the particles, most probably both in the form of free PEI and PEI attached to fragments of mesoporous silica, and that this process preferentially occurs 1801
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Schrott-Fischer on Department of Otolaryngology, Medical University of Innsbruck, Innsbruck, Austria for recording the TEM image. This work was supported by Academy of Finland project Biotarget (Contract 118196).
inside the cells. A schematic representation of the suggested intracellular degradation process of the studied particles is given in Figure 8. Our results do not allow for a more detailed
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Figure 8. Schematic representation of the suggested particle degradation process inside the cells. Within 4 h incubation time parts of the PEI surface layer and mesoporous silica shell are seen to separate from the original particle.
analysis of the nature of the intracellular compartments (different types of endosomes and lysosomes), but the observed preferential intracellular detachment of PEI from the particles do indeed give clear support for our intuitive claims based on drug release studies, that PEI must detach from the particles inside cells, providing a means for intracellularily triggered drug release. This mechanism seems to be operational irrespective of whether the drug is covalently linked to the PEI layer as in the case of MTX, or if the drug is physically adsorbed into the mesopores of the MSNs.
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CONCLUSIONS We have studied the extracellular and intracellular degradation of amino functionalized mesoporous silica core−shell−shell nanoparticles. A preferential intracellular degradation of the particles was observed, mainly driven by the dissolution of the mesoporous silica shell. These results explain previous observations using corresponding fully mesoporous silica nanoparticles and suggest that covalent attachment of drugs, targeting groups, or other functionalities and their subsequent intracellular detachment is a promising route toward intrinsically triggered intracellular release. Our results also demonstrate new imaging-based methodology that can be used to effectively analyze the degradation of nanoparticles.
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ASSOCIATED CONTENT
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AUTHOR INFORMATION
REFERENCES
S Supporting Information *
Pseudocolored images of all the three fluorescence channels observed, for the image on the lower row of Figure 3. This material is available free of charge via the Internet at http:// pubs.acs.org. Corresponding Author
*E-mail:
[email protected]. Tel.: +49-731-5022730. Fax: +49-731-5022733. Notes
The authors declare no competing financial interest.
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ACKNOWLEDGMENTS We would like to thank Dr. Cecilia Sahlgren, Turku Centre for Biotechnology, University of Turku and Åbo Akademi University, and Department of Biology, Åbo Akademi University, Turku, Finland, for performing initial confocal fluorescence microscopy studies and for useful discussions, Anna-Brita Puranen for valuable assistance, and Dr. Anneliese 1802
dx.doi.org/10.1021/mp3005879 | Mol. Pharmaceutics 2013, 10, 1795−1803
Molecular Pharmaceutics
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