Colloids and Surfaces B: Biointerfaces 116 (2014) 518–525
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Embedding fluorescent mesoporous silica nanoparticles into biocompatible nanogels for tumor cell imaging and thermo/pH-sensitive in vitro drug release Rijun Gui a,b,1 , Yanfeng Wang a,1 , Jie Sun a,∗ a b
Institute of Materia Medica, Shandong Academy of Medical Sciences, Jinan 250062, PR China Department of Chemistry, School of Chemistry and Chemical Engineering, Shanghai Jiao Tong University, Shanghai 200240, PR China
a r t i c l e
i n f o
Article history: Received 31 August 2013 Received in revised form 14 January 2014 Accepted 21 January 2014 Available online 6 February 2014 Keywords: N-Isopropylacrylamide Chitosan Nanospheres Cell imaging Drug release
a b s t r a c t Thermo/pH-sensitive/fluorescent/biocompatible nanospheres consisting of quantum dots-embedded mesoporous silica nanoparticles (Q-MS) as a core and poly(N-isopropylacrylamide (NIPAM))-graftchitosan (CS) nanogels as a shell (PNIPAM-g-CS) were prepared via temperature-regulated one-pot copolymerization of NIPAM monomer and CS in the presence of Q-MS. The prepared nanospheres exhibited remarkable fluorescence/thermo/pH-sensitivity. HepG2 cells treated with nanospheres displayed bright fluorescence imaging. Loading efficiency and capacity of Doxorubicin (Dox) into nanospheres were regularly increased with the increment of Dox concentration. At a high temperature and a low pH, cumulative in vitro release of Dox from Dox-loaded nanospheres was much great and fast. Released Dox still retained high anticancer activity, and blank nanosphere carriers produced neglectful toxicity to HepG2 cells. The multifunctional nanospheres could be further developed toward temperature/pH-regulated drug carriers for in vivo tumor therapy with a rapid drug release and fluorescence imaging in targeted tissues. © 2014 Elsevier B.V. All rights reserved.
1. Introduction Colloidal semiconductor quantum dots (QDs) have been extensively studied due to their unique properties such as size-tunable emission spectra, broad absorption profiles, high photochemical stability and quantum yields, which make QDs an attractive alternative to organic dyes in applications of light-emitting devices, lasers, multiplexed bio-labels and analysis, etc. [1–6]. Generally, a further modification to fabricate composite nanoparticles (NPs) is an important prerequisite for practical applications of NPs [7,8]. Thus, the formation of composite QDs, such as metal–QDs hybrids, biomolecule–QDs conjugates, or polymer–QDs nanospheres, is vital for applications of QDs-based nanomaterials [9–12]. Recently, polymer-embedded QDs nanospheres have been studied because the combination of QDs and polymers can not only improve mechanical and chemical stability of QDs, but also retains properties of polymers and QDs, thus showing promising applications in biomedical fields, such as fluorescence markers and drug delivery systems (DDS), biosensors, etc. [13–15]. To the best of our knowledge, three major methods have been adopted to develop
∗ Corresponding author. Tel.: +86 531 67816486; fax: +86 531 82919963. E-mail address: [email protected] (J. Sun). 1 These authors have the equivalent contribution to this work. 0927-7765/$ – see front matter © 2014 Elsevier B.V. All rights reserved. http://dx.doi.org/10.1016/j.colsurfb.2014.01.044
polymer-embedded QDs nanospheres, including in situ synthesis of QDs in polymer nanospheres [16], the entrapment of QDs within polymer matrix [17–19], and the monomer polymerization in the presence of QDs [20]. However, the in situ synthesized QDs in nanospheres are polydisperse and polycrystalline, displaying a broad size distribution and low quantum yield. The entrapment of QDs into polymers obtained via self-assembly, ligand exchange, covalent bonding, hydrogen bonding, etc., partly retains the properties of QDs and polymers. Once the external environment changes, the leakage of QDs probably occurs due to the weaker interaction between QDs and polymers. For encapsulating QDs into polymer matrix, monomer polymerization in the presence of QDs is simple, and resultant nanospheres demonstrate a high physicochemical stability. So far, the efficient encapsulation of QDs into polymers to achieve functionalized polymer–QDs nanospheres is still a challenging task. During the past decade, stimuli-sensitive polymers have generated longstanding interest due to their reversible changes of properties in response to external stimulus, such as pH, temperature, ionic strength, light, electrical and magnetic field [21–23]. In especial, thermosensitive poly(N-isopropylacrylamide) (PNIPAM) has attracted much attention as it has a lower critical solution temperature (LCST) of ∼32–34 ◦ C in water. Above the LCST, PNIPAM networks undergo an abrupt collapse to form hydrophobic nanospheres, showing a coil-globule volume transition [24,25].
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Scheme 1. Schematic illustration of the preparation of Q-MS/PNIPAM-g-CS core/shell hybrid nanospheres.
Since the first report by Pelton et al. in 1986 [26], PNIPAM has received considerable attention due to their outstanding thermo-sensitivity, used for the sorbent for proteins and enzymes, biomedical testing, pollution control, catalysis, controlled drug delivery, etc. [27–30]. When used for DDS, the transition temperature of PNIPAM can be modulated close to body temperature. To date, one key challenge for PNIPAM application is to modulate its LCST. Great efforts have been devoted to modulating the LCST of PNIPAM preferably up to 42–43 ◦ C, a temperature for hyperthermia treatment of cancer [31]. The copolymerization of PNIPAM and N,N -dimethylacrylamide has been reported, inducing a shift in LCST from 32 to 38 ◦ C. To improve biocompatibility of PNIPAM nanogels, rational surface modification is required, which facilitates their applications as biocompatible DDS [24,32]. Chitosan (CS) is a natural amphoteric polyelectrolyte and possesses excellent biocompatibility, which have made CS many biomedical applications, such as bacteriostatic agent, blood anticoagulant, wound dressing, artificial bone and skin [33,34]. CS owns a good pH-sensitivity in aqueous solution containing abundant OH and NH2 groups [35]. Hydrogels with a high substitution degree of CS will markedly swell in basic solution, and shrink dramatically in low-pH solution. These properties can be applied to DDS for controlled drug release [36]. To take the advantages of PNIPAM and CS, composite hydrogels (PNIPAM-g-CS) with thermo/pH-sensitivity have been prepared in past reports [37–39], and were further developed toward pH/temperature-sensitive release of coenzyme A. Nevertheless, QDs-embedded thermo/pH-responsive hydrogels have not been reported. These typical hydrogels (or nanospheres) combine thermo/pH-sensitivity, fluorescence and biocompatibility into one single entity, and their applied potential in the fields of biology and biomedicine would be expected. Herein, we described a facile method to prepare PNIPAMgraft-CS polymers coated QDs-embedded mesoporous silica NPs (Q-MS), i.e. Q-MS/PNIPAM-g-CS core/shell hybrid nanospheres. This method was divided into three steps. First, ligand exchange between 3-mercaptopropionic acid (MPA) and dodecyltrimethylammonium bromide (DTAB) occurred on the surface of QDs to prepare DTAB-stabilized QDs. Second, DTAB-QDs were incorporated into silica spheres using a modified Stöber method. Third, graft copolymerization of NIPAM monomer and CS was initiated
to form PNIPAM-g-CS nanogels (as an outer shell) on the surface of Q-MS, thus resulting in the formation of Q-MS/PNIPAMg-CS nanospheres (Scheme 1). A whole set of characteristics of nanospheres were investigated. Further researches on these nanospheres were conducted, including fluorescent imaging of tumor cells, cytotoxicity and thermo/pH-dual regulated in vitro drug release. 2. Experimental 2.1. Materials Chitosan (CS, ∼80 mesh, MW . 22.4 kDa, 95% of deacetylation degree) was purchased from Tokyo Kasei-Kogyo Corp. NIsopropylacrylamide (NIPAM, 99%), N,N -methylenebisacrylamide (MBA, 99%), ammonium persulphate (APS, 99%), 3mercaptopropionic acid (MPA, 98%), dodecyltrimethylammonium bromide (DTAB, 99%), sodium dodecyl sulfate (SDS, 97%), 3-(trimethoxylsilyl) propyl methacrylate (MPS, 95%) and tetraethyoxysilane (TEOS, 98%) were obtained from Sigma–Aldrich Corp. Doxorubicin (Dox) in the form of hydrochloride salts was obtained from Beijing Huafeng Corp. 3-[4,5-Dimethyl-thiazol-2-yl]-2,5diphenyl-tetrazolium bromide (MTT) and biological reagents were brought from Invitrogen Corp. Other chemicals with analytical grade were from Shanghai Chemical Reagent Corp. All chemicals can be used directly as received without any purification. Water used in experiments was of Milli-Q grade. Phosphate buffered salines (PBS, 1 mM) were prepared by modulating the molar ratio of Na2 HPO4 and NaH2 PO4 . 2.2. Preparation of Q-MS/PNIPAM-g-CS hybrid nanospheres MPA-capped CulnS2 /ZnS QDs were freshly prepared via hydrothermal route based on Ref. [40], and detailed preparation was available in Supplementary Data (Part S1). Typically, 5 mg of QDs dispersed in 1 mL chloroform were mixed with 9 mL of aqueous solution containing 0.1 g of DTAB under stirring. Chloroform was evaporated at 65 ◦ C to form DTAB-QDs composites, which were subsequently added into 90 mL of water. Then, 3 mL of NaOH solution (2 M), 0.5 mL of TEOS and 5 mL of ethyl acetate were
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successively added into the above mixture with stirring (100 rpm) for 1 min, and then kept for 6 h at 50 ◦ C. The products were centrifuged, repeatedly washed with water–ethanol. Finally, the products were transferred into 50 mL of ethanol solution containing 67 L of MPS, and the mixture was stirred for 12 h at 25 ◦ C. DTAB template was removed in an ethanol solution of NH4 NO3 (50 mL, 10 mg mL−1 ) via ion exchange [41], and the mixture was refluxed for 6 h. The final products were collected by centrifugation and repeatedly washed with ethanol. The resultant MPS-modified Q-MS was dispersed in water, and diluted to 1 wt% for subsequent experiments. Mixed solution (I) consisting of NIPAM (0.5 g), MBA (10 mg) and water (30 mL) was loaded in a three-necked flask equipped with a reflux condenser. The (I) was bubbled with N2 inlet for 30 min to remove dissolved oxygen. Then, MPS-modified Q-MS dispersed in water (1 g, 1 wt%) was added into the (I) under stirring to form mixed solution (II). Afterward, APS (3 mg) and SDS (5 mg) dissolved in 10 mL of water were rapidly added into the (II) under stirring to form mixed solution (III). After reaction for 2 h at 25 ◦ C, NIPAM monomers were initiated to form PNIPAM networks. Finally, 10 mL of aqueous solution containing 3 mg of APS, 0.1 g of CS and 0.1 mL of acetic acid was added into the (III), and then heated to 70 ◦ C. The polymerization reaction was allowed to proceed for 6 h under a continuous stirring of 300 rpm and a N2 flow of 200 mL min−1 . The products were centrifuged for 30 min at 12 000 rpm to remove unreactive monomers. Resultant products were dispersed in water, and dialyzed (Spectra/Por 7 dialysis membrane, MWCO 5 × 104 ) for 7 days against water frequently. Finally, purified products were prepared, named as Q-MS/PNIPAM-g-CS core/shell hybrid nanospheres. 2.3. Characterization Fourier transform infrared spectra (FTIR, Nicolet, USA) were recorded with a Nicolet 6700 FTIR spectrometer in the scanning region of 400–4000 cm−1 . N2 adsorption–desorption isotherms were given on a Micromeritics Tristar 3000 pore analyzer at 77 K, equipped with Brunauer–Emmett–Teller and Barrett–Joyner–Halenda for the analysis of surface area, pore size and volume. UV–vis spectra were recorded with a UV-2450 spectrophotometer (Shimadzu, Japan). PL spectra were measured by a FLSP 920 fluorescence spectrophotometer (Edinburgh Instruments, Britain). Transmission electron microscope (TEM, Jeol, Japan) images were acquired with a JEOL JEM-1400 TEM at 120 kV of acceleration voltage. Hydrodynamic radius (diameters) was obtained on dynamic light scattering apparatus (DLS, Malven Instruments, U.K.). Fluorescence images of tumor cells were obtained by a FV-300 IX71 confocal laser fluorescence microscope (CLFM, Olympus). 2.4. Cellular uptake of nanospheres and imaging Fluorescence imaging of HepG2 cells (bought from the cell bank of Shanghai Science Academe) was recorded by CLFM. The 6 × 104 cells/well was seeded on a 6-well plate for 24 h at 37 ◦ C. Then, the nanospheres dispersed in water (0.2 mg mL−1 ) were added into cell dishes. After incubation for 12 and 24 h, nanospheres-loaded cells were repeatedly washed with PBS (1 mM, pH 7.4) to remove free nanospheres absorbed or attached on the outer surface of cell membrane. Afterward, these treated cells were detected by CLFM, excited at 480 nm. 2.5. Drug loading and release of nanosphere carriers Nanospheres (20 mg) were ultrasonically dispersed in 25 mL of Dox solution (1 mg mL−1 , 1% of NaCl, pH 7.4 modulated by
0.1 M of NaOH) to form homogeneous solution, which was continuously stirred for 24 h at 25 ◦ C until Dox concentration in the solution stabilized. Then, the solution was centrifuged for 15 min at 5000 rpm, and washed with water twice to remove unbound or surface-absorbed Dox. The mass of Dox loaded in nanospheres was calculated by subtracting the Dox mass in the supernatant from the total Dox mass in the initial solution. Free Dox mass (Mfree-Dox ) in solution was determined by UV–vis spectrophotometer at 480 nm using Lambert–Beer Law. The Dox-loading efficiency (LE) and capacity (LC) were calculated according to the following equations: Dox-LE (%) = 100 ×
Mtotal-Dox − Mfree-Dox Mtotal-Dox
(1)
Dox-LC (%) = 100 ×
Mtotal−Dox − Mfree−Dox Mtotal−nanospheres
(2)
To investigate pH/temperature-dependent Dox release, 20 mg of Dox-loaded nanospheres in PBS (1 mM, pH 5.0, 6.5 and 7.4) were transferred into a dialysis bag (MWCO 5 × 104 ), which were then gently shaken in water bath at different temperatures (25, 37 and 42 ◦ C). At defined time intervals, 1 mL of sample in each mixture solution was withdrawn and analyzed by UV–vis at 480 nm. An equal volume of PBS was added after each sampling to retain a constant volume. Each experiment of drug release was performed in triplicate, and each result was expressed as the mean of three repeated measurements. 2.6. In vitro cytotoxicity of nanosphere carriers Cytotoxicity studies of the blank, Dox-loaded nanospheres and free Dox on HepG2 cells (as model tumor cells) were assessed by MTT assay. The 1 × 104 cells/well was incubated in Dulbecco’s modified Eagle medium containing calf serum (10 wt%) and 100 units mL−1 of penicillin in a fully humidified incubator with 5 vol% of CO2 at 37 ◦ C. When these cells reached 80% of confluence with normal morphology, the blank, Dox-loaded nanospheres with a concentration of 0.2–1.0 mg mL−1 , and free Dox were added into cell dishes, respectively. Then, these cell dishes were put into incubators for 48 h at 37 ◦ C, and no detectable damage to cells was observed. After incubation for a defined time, 20 L of MTT reagent (0.5 mg mL−1 , diluted in culture medium) was substituted for culture medium, followed by an additional 2 h incubation. Finally, MTT-medium was removed attentively, and 150 L of dimethyl sulfoxide was added into each well to dissolve formazan crystals. The absorbance (A) of respective wells (colored solutions) was recorded at 570 nm, using a Multiskan MK3 Enzyme-labeled Instrument (Thermo Scientific, USA). All cytotoxicity experiments were carried out in triplicate, and all results were averaged. The cell survival rate was calculated by the equation, as below: Cell survival rate (%) = 100 ×
Atest cells Acontrol cells
(3)
3. Results and discussion 3.1. Preparation and characterization of Q-MS/PNIPAM-g-CS nanospheres Q-MS composites were prepared using a base-catalyzed sol–gel approach in the presence of QDs [42,43]. Then, Q-MS were dispersed in anhydrous ethanol containing MPS for surface modification of C C bonds, which form covalent bonds between inorganic substrates and polymers [44]. Using MPS-modified Q-MS composites as seeds, Q-MS/PNIPAM-g-CS core/shell nanospheres were prepared via graft copolymerization of NIPAM monomer and CS. FTIR spectra of Q-MS and Q-MS/PNIPAM-g-CS show an evidence
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Fig. 1. (a) FTIR spectra, (b) N2 adsorption–desorption isotherms (inset: the pore diameter distribution), (c) UV–vis absorption and PL emission spectra, and TEM images of (d) QDs, (e) Q-MS and (f) Q-MS/PNIPAM-g-CS.
for the preparation of nanospheres (in Fig. 1a). For Q-MS, a strong signal for water bending vibration at 1655 cm−1 is observed, and is attributed to hydrophilic silanol surface after template extraction [45]. In addition, C O stretching at 1718 cm−1 and CH2 stretching at 2978 cm−1 indicate successful modification of Q-MS by MPS. For Q-MS/PNIPAM-g-CS, characteristic absorption bands at 2973 and 1390 cm−1 testify that NIPAM chains exist in nanospheres. The peak at 1560 cm−1 is assigned to NH2 bending vibration of CS. For both samples, characteristic peaks at 460, 805 and 1035 cm−1 denote that the silica framework of Q-MS maintains unchanged in nanospheres upon the coating of polymers. These results suggest the polymerization of NIPAM and CS, and the integration of silica spheres and PNIPAM-g-CS polymers. Fig. 1b displays the N2 adsorption–desorption isotherms and pore size distribution (insert) of both samples. The surface area, pore volume and pore diameter of Q-MS are 857 m2 g−1 , 1.1 cm3 g−1 and 2.8 nm, respectively. For Q-MSN/PNIPAM-g-CS, corresponding parameters are 698 m2 g−1 , 0.9 cm3 g−1 and 2.7 nm, respectively. The pore diameter has a slight decrease from 2.8 to 2.7 nm, implying that PNIPAM-g-CS is predominantly distributed on the exterior surface of Q-MS [46]. The nanospheres with nearly identical mesoporosity as Q-MS exhibit a promising application in stimulus-responsive drug release platforms. Photoluminescence (PL) properties of nanospheres were characterized by UV–vis and PL spectrometry. As elaborated in Fig. 1c, no remarkable change in PL spectra of QDs appears after encapsulated into PNIPAM-g-CS [40], indicating that the silica on outer layers indeed protects QDs.
Especially, these nanospheres possess well-stabilized dispersion with coupling shells, and intense PL can retain for several weeks (in Fig. S1). Morphology and structure of QDs, Q-MS and Q-MS/PNIPAM-gCS nanospheres were tested by wide-field TEM images (in Fig. 1d–f). The QDs (Fig. 1d) show a uniform, discrete spherical shape with an average diameter of 4.8 ± 0.5 nm. In Fig. 1e, multiple dark spots are observed in each particle with a size of 50 ± 5 nm. The average diameter of black dots coated by the silica shell is ∼20 nm, denoting that the QDs were embedded within mesoporous silica matrix. Fig. 1f displays core/shelled, spherical and polymer-encapsulated (bright ring) spheres, and the average size is 95 ± 6 nm. These nanospheres consist of black dots (QDs) encapsulated in the gray silica interlayer (SiO2 ) and a dark shell of PNIPAM-g-CS polymers. In addition, average hydrodynamic size of nanospheres measured by DLS is 125 nm at 25 ◦ C, and there is a low diameter polydispersity index (0.075) [47]. The hydrodynamic size from DLS is larger than the size from TEM due to hydrate layers in aqueous environment [48], further implying the existence of copolymer shell layers in nanospheres. TEM images of QDs, Q-MS and Q-MS/PNIPAM-gCS (in Fig. 1d–f) are consistent with corresponding absorption and emission spectra (in Fig. S4). Dramatic red-shift in spectra from the initial QDs to Q-MS (with mesoporous silica shells around QDs) and to Q-MS/PNIPAM-g-CS (with polymer shells around Q-MS) were observed, implying a gradual increment in size [15]. In addition, 1 H-NMR spectra of PNIPAM-g-CS nanogels (in Part S2) testified the graft copolymers (in Fig. S2a). An acute proton peak
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Fig. 2. (a) Diameter as a function of temperature of PNIPAM-g-CS nanogels and Q-MS/PNIPAM-g-CS nanospheres in PBS (1 mM, pH 7.4). (b) Diameter as a function of pH of the two samples in PBS at 25 ◦ C. (c) DLS data of nanospheres stored in PBS at 37 ◦ C containing different NaCl concentrations.
( CH CH2 ) at 2.1 ppm, a peak ( NH CH 10.0. By contrast, the diameter of nanospheres is nearly unchanged at pH > 8.0. These results reveal that two samples are pH-sensitive, and Q-MS in nanospheres hardly affect this sensitivity, in an agreement with previous reports [49]. At a lower pH, OH from PNIPAM-g-CS is protonated. Increasing the pH, the degree of protonation reduces, and OH gradually translates to O− [35]. As a result, electrostatic repulsion and hydrophilia of O− cause an increase in the swelling degree of nanospheres. When the pH exceeds a certain value, hydrogen-bonding between water molecules and polymer chains gradually increases, thus resulting in the diameter of nanospheres trending to stability [44]. In addition, the effect extent of pH on the diameter of nanogels is larger than nanospheres under the same conditions (pH 2–12), which may be ascribed to the existence of Q-MS that partly restrains the swelling degree of nanospheres [44,49].
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Fig. 3. CLFM images of HepG2 cells after (a) 12 h and (b) 24 h treatment with 0.2 mg mL−1 of nanospheres in PBS (1 mM, pH 7.4). The excitation wavelength is 480 nm, and the scale bars are 20 m.
Nanospheres’ pH-sensitivity could be further verified by the effect of NaCl concentration on the size and stability of nanospheres (in Fig. 2c). For the system consisting of nanospheres (0.2 mg mL−1 ), PBS (1 mM, pH 5.0/7.4) and NaCl (0.5–0.8 M), the average diameter of nanospheres shows relatively slight fluctuation (105–110 nm), which is consistent with the results from TEM, implying monodisperse NPs. It is mainly ascribed to PNIPAM-g-CS shells that bring more ions in a dilute salt solution and thus lead to electrostatic repulsion and low viscosity. As a result, the aggregation of NPs is restrained. In 0.8–1.2 M of NaCl, nanospheres have a larger diameter by DLS compared to TEM, revealing a possible aggregation. It is probably due to the competition of excess salts with PNIPAM-g-CS nanogels for water molecules in solution, and the competition results in high viscosity and the following aggregation of nanospheres. Under the same concentration, the diameter of nanospheres at pH 5.0 is less than at pH 7.4, which can be explained by electrostatic shielding of polymer shells and excessive charges appeared in acid environment. The electrostatic shielding causes the shrink of polymers, and the decline of intermolecular entangled degree. So, the nanospheres have a higher stability at pH 5.0, and the diameter is desirable in acidic and low-salt solutions.
affected by Dox, and increase with the increment of Dox concentration ([Dox]). Increasing [Dox] from 0.05 to 1.0 mg mL−1 , LC increases from 3.1% to 45.2%, while LE increases from 11.9% to 61.1%. These denoted that Dox-loading into nanospheres was [Dox]-dependent. Dox molecule consists of acidic phenolichydroxyl groups and alkaline amino groups that can interact
3.3. In vitro tumor cell imaging of nanospheres The encapsulation of QDs into nanospheres gives a possibility of PL imaging. In this regard, CLFM was adopted to evaluate the nanospheres for tumor cell imaging. The qualitative cell uptake of nanospheres was investigated, as demonstrated in Fig. 3. CLFM images of HepG2 cells treated with nanospheres denoted that the nanospheres were endocytosed into HepG2 cells. When irradiated at 480 nm, the QDs immobilized in nanospheres produced bright green fluorescence. Fluorescence intensity increases (from Fig. 3a and b), suggesting that the internalized extent of nanospheres by cells enhances with the increase of incubation time. It also means that the amounts of nanospheres uptaken by HepG2 cells are dependent on incubation time. In addition, no remarkable autofluorescence is observed under the same conditions. The areas where nanospheres did not permeate are distinct, and there is no dark region appearing in cells, implying a potential of the nanospheres as effective PL imaging in tumor cells. 3.4. Drug loading and thermo/pH-regulated release in vitro Dox was used for a typical anticancer drug to evaluate potential application of nanospheres. In Fig. 4a, Dox with different concentrations was loaded in nanospheres. Both LC and LE are markedly
Fig. 4. (a) Effects of Dox concentration on Dox-loading capacity (LC) and efficiency (LE) in PBS (1 mM, pH 7.4) at 37 ◦ C. (b) Effects of pH on Dox-loading content and embed efficiency for nanosphere carriers in PBS at 25 ◦ C.
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Fig. 5. Cumulative Dox release profiles from nanosphere carriers at selected temperatures (25, 37 and 42 ◦ C) in PBS (1 mM, pH 5.0, 6.5 and 7.4). The data points are averages of six measurements.
with NH2 and OH of PNIPAM-g-CS polymers (as a shell in nanospheres). This helps to form intermolecular complexes via hydrogen bonding [50], inducing a high LE. Moreover, Doxloading into nanospheres was efficient because nanospheres had no adsorption interaction with Dox, otherwise distinct decrease in LE would be observed upon the increase of [Dox] [51]. Loading content (LC) and embed efficiency (EE) of Dox into nanosphere carriers were investigated, as shown in Fig. 4b. Both two were markedly influenced by pH. Upon the pH increase from 4.9 to 9.2, LC increased from 0.02 to 0.45 mmol g−1 , while EE increased from 4.2 to 87.1 wt%. The EE has a sharp increase at pH 8.3 (compared to pH 7.4), which could be explained by Dox-loading process based on physical adsorption in mesoporous channels. Isoelectric point of Dox was 8.25 [44], and net charge of Dox was around zero. On this account, the electrostatic interactions between polymer shells and Dox molecules were minimal. Dox molecules would be easily absorbed into mesoporous channels across the larger mesh size of polymer shells at an elevated pH. Dox molecule is water-soluble at a lower pH, but becomes insoluble in alkaline environment. In aqueous solution of Dox with a higher pH, electrostatic repulsion among adsorbed Dox molecules markedly decreased, which caused the adsorption of Dox molecules into nanospheres. To estimate Dox release of nanosphere carriers (in Fig. S3), related experiments were conducted. As displayed in Fig. 5, Dox release responses of nanosphere carriers in PBS (pH 5.0, 6.5 and 7.4) were studied at 25 ◦ C (LCST). Both temperature and pH have a remarked influence on Dox release. At 25 ◦ C, Dox release was less than 15.1% at pH 5.0, and greater than at a higher pH, which was due to acid-labile physical adsorption. Dox release at 37 ◦ C was apparently similar to that at 42 ◦ C, but the release extent of Dox was intermediate between 25 and 42 ◦ C. In other words, Dox release at 42 ◦ C was greater and faster than at a lower temperature. At each investigated pH, a rapid release occurred during the initial stage (0–2 h) due to the changes of chemical environment in carriers, followed by a slower release. At a longer duration (2–24 h), the extent of Dox release continued to increase before reaching a sustained release, which was similar to the cases at different temperatures. Internal structure of PNIPAM was loose while drug molecules diffused from the interior of nanogels in an easy way [52]. In Dox-loaded nanospheres, Dox diffusion across nanogel shells may have a distance-limiting
Fig. 6. In vitro cytotoxicity of the blank, Dox-loaded nanospheres and free Dox at different concentrations against HepG2 cells after 48 h incubation at 37 ◦ C.
mechanism. At pH 7.4 or 25 ◦ C, nanogel shells were in the swelling state, and hydrodynamic size of nanospheres was large, thus causing a slow release. Enhanced electrostatic attraction between positively charged drug molecules and negatively charged nanogels was unfavorable for the mass-transport of Dox molecules. By contrast, nanogel shells were collapsed at a higher temperature (>37 ◦ C). At a lower pH (LCST) and mild acidic medium were favorable for Dox release, controlled via small changes in the vicinity of LCST (37 ◦ C) and a lower pH (mild acidic medium), Doxloaded nanospheres exhibited a faster cumulative in vitro Dox release. Released Dox still maintained high anticancer activity, and blank nanosphere carriers produced neglectful toxicity to HepG2 cells. Thus, the nanospheres could be developed as a targeting probe for tumor cells, and exhibit promising applications for diagnosis, therapy, controllable drug release, and real-time detection of drug treatment in the intracellular process. Acknowledgments This work was financially supported by Grants from the Shandong Provincial Doctoral Foundation of China (No. 2011BSE27195), the Postdoctoral Science Foundation of China (No. 2013M531164) and the Postdoctoral Science Foundation of Shanghai (No. 13R21413800). Appendix A. Supplementary data Supplementary data associated with this article can be found, in the online version, at http://dx.doi.org/10.1016/j.colsurfb. 2014.01.044. References [1] M. Bruchez, M. Moronne, P. Gin, S. Weiss, A.P. Alivisatos, Science 281 (1998) 2013–2016. [2] W.C.W. Chan, S. Nie, Science 281 (1998) 2016–2018. [3] N. Tessler, V. Medvedev, M. Kazes, S.H. Kan, U. Banin, Science 295 (2002) 1506–1508. [4] C. Burda, X. Chen, R. Narayanan, M.A. El-Sayed, Chem. Rev. 105 (2005) 1025–1102. [5] R.C. Somers, M.G. Bawendi, D.G. Nocera, Chem. Soc. Rev. 36 (2007) 579–591. [6] A.L. Rogach, N. Gaponik, J.M. Lupton, C. Bertoni, D.E. Gallardo, S. Dunn, Angew. Chem. Int. Ed. 120 (2008) 6638–6650. [7] Y. Dai, P. Ma, Z. Cheng, X. Kang, X. Zhang, Z. Hou, C. Li, D. Yang, X. Zhai, J. Lin, ACS Nano 6 (2012) 3327–3338. [8] X. Zhang, P. Yang, Y. Dai, P. Ma, X. Li, Z. Cheng, Z. Hou, X. Kang, C. Li, J. Lin, Adv. Funct. Mater. 23 (2013) 4067–4078.
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Colloids and Surfaces B: Biointerfaces journal homepage: www.elsevier.com/locate/colsurfb
Embedding fluorescent mesoporous silica nanoparticles into biocompatible nanogels for tumor cell imaging and thermo/pH-sensitive in vitro drug release Rijun Gui a,b,1 , Yanfeng Wang a,1 , Jie Sun a,∗ a b
Institute of Materia Medica, Shandong Academy of Medical Sciences, Jinan 250062, PR China Department of Chemistry, School of Chemistry and Chemical Engineering, Shanghai Jiao Tong University, Shanghai 200240, PR China
a r t i c l e
i n f o
Article history: Received 31 August 2013 Received in revised form 14 January 2014 Accepted 21 January 2014 Available online 6 February 2014 Keywords: N-Isopropylacrylamide Chitosan Nanospheres Cell imaging Drug release
a b s t r a c t Thermo/pH-sensitive/fluorescent/biocompatible nanospheres consisting of quantum dots-embedded mesoporous silica nanoparticles (Q-MS) as a core and poly(N-isopropylacrylamide (NIPAM))-graftchitosan (CS) nanogels as a shell (PNIPAM-g-CS) were prepared via temperature-regulated one-pot copolymerization of NIPAM monomer and CS in the presence of Q-MS. The prepared nanospheres exhibited remarkable fluorescence/thermo/pH-sensitivity. HepG2 cells treated with nanospheres displayed bright fluorescence imaging. Loading efficiency and capacity of Doxorubicin (Dox) into nanospheres were regularly increased with the increment of Dox concentration. At a high temperature and a low pH, cumulative in vitro release of Dox from Dox-loaded nanospheres was much great and fast. Released Dox still retained high anticancer activity, and blank nanosphere carriers produced neglectful toxicity to HepG2 cells. The multifunctional nanospheres could be further developed toward temperature/pH-regulated drug carriers for in vivo tumor therapy with a rapid drug release and fluorescence imaging in targeted tissues. © 2014 Elsevier B.V. All rights reserved.
1. Introduction Colloidal semiconductor quantum dots (QDs) have been extensively studied due to their unique properties such as size-tunable emission spectra, broad absorption profiles, high photochemical stability and quantum yields, which make QDs an attractive alternative to organic dyes in applications of light-emitting devices, lasers, multiplexed bio-labels and analysis, etc. [1–6]. Generally, a further modification to fabricate composite nanoparticles (NPs) is an important prerequisite for practical applications of NPs [7,8]. Thus, the formation of composite QDs, such as metal–QDs hybrids, biomolecule–QDs conjugates, or polymer–QDs nanospheres, is vital for applications of QDs-based nanomaterials [9–12]. Recently, polymer-embedded QDs nanospheres have been studied because the combination of QDs and polymers can not only improve mechanical and chemical stability of QDs, but also retains properties of polymers and QDs, thus showing promising applications in biomedical fields, such as fluorescence markers and drug delivery systems (DDS), biosensors, etc. [13–15]. To the best of our knowledge, three major methods have been adopted to develop
∗ Corresponding author. Tel.: +86 531 67816486; fax: +86 531 82919963. E-mail address: [email protected] (J. Sun). 1 These authors have the equivalent contribution to this work. 0927-7765/$ – see front matter © 2014 Elsevier B.V. All rights reserved. http://dx.doi.org/10.1016/j.colsurfb.2014.01.044
polymer-embedded QDs nanospheres, including in situ synthesis of QDs in polymer nanospheres [16], the entrapment of QDs within polymer matrix [17–19], and the monomer polymerization in the presence of QDs [20]. However, the in situ synthesized QDs in nanospheres are polydisperse and polycrystalline, displaying a broad size distribution and low quantum yield. The entrapment of QDs into polymers obtained via self-assembly, ligand exchange, covalent bonding, hydrogen bonding, etc., partly retains the properties of QDs and polymers. Once the external environment changes, the leakage of QDs probably occurs due to the weaker interaction between QDs and polymers. For encapsulating QDs into polymer matrix, monomer polymerization in the presence of QDs is simple, and resultant nanospheres demonstrate a high physicochemical stability. So far, the efficient encapsulation of QDs into polymers to achieve functionalized polymer–QDs nanospheres is still a challenging task. During the past decade, stimuli-sensitive polymers have generated longstanding interest due to their reversible changes of properties in response to external stimulus, such as pH, temperature, ionic strength, light, electrical and magnetic field [21–23]. In especial, thermosensitive poly(N-isopropylacrylamide) (PNIPAM) has attracted much attention as it has a lower critical solution temperature (LCST) of ∼32–34 ◦ C in water. Above the LCST, PNIPAM networks undergo an abrupt collapse to form hydrophobic nanospheres, showing a coil-globule volume transition [24,25].
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Scheme 1. Schematic illustration of the preparation of Q-MS/PNIPAM-g-CS core/shell hybrid nanospheres.
Since the first report by Pelton et al. in 1986 [26], PNIPAM has received considerable attention due to their outstanding thermo-sensitivity, used for the sorbent for proteins and enzymes, biomedical testing, pollution control, catalysis, controlled drug delivery, etc. [27–30]. When used for DDS, the transition temperature of PNIPAM can be modulated close to body temperature. To date, one key challenge for PNIPAM application is to modulate its LCST. Great efforts have been devoted to modulating the LCST of PNIPAM preferably up to 42–43 ◦ C, a temperature for hyperthermia treatment of cancer [31]. The copolymerization of PNIPAM and N,N -dimethylacrylamide has been reported, inducing a shift in LCST from 32 to 38 ◦ C. To improve biocompatibility of PNIPAM nanogels, rational surface modification is required, which facilitates their applications as biocompatible DDS [24,32]. Chitosan (CS) is a natural amphoteric polyelectrolyte and possesses excellent biocompatibility, which have made CS many biomedical applications, such as bacteriostatic agent, blood anticoagulant, wound dressing, artificial bone and skin [33,34]. CS owns a good pH-sensitivity in aqueous solution containing abundant OH and NH2 groups [35]. Hydrogels with a high substitution degree of CS will markedly swell in basic solution, and shrink dramatically in low-pH solution. These properties can be applied to DDS for controlled drug release [36]. To take the advantages of PNIPAM and CS, composite hydrogels (PNIPAM-g-CS) with thermo/pH-sensitivity have been prepared in past reports [37–39], and were further developed toward pH/temperature-sensitive release of coenzyme A. Nevertheless, QDs-embedded thermo/pH-responsive hydrogels have not been reported. These typical hydrogels (or nanospheres) combine thermo/pH-sensitivity, fluorescence and biocompatibility into one single entity, and their applied potential in the fields of biology and biomedicine would be expected. Herein, we described a facile method to prepare PNIPAMgraft-CS polymers coated QDs-embedded mesoporous silica NPs (Q-MS), i.e. Q-MS/PNIPAM-g-CS core/shell hybrid nanospheres. This method was divided into three steps. First, ligand exchange between 3-mercaptopropionic acid (MPA) and dodecyltrimethylammonium bromide (DTAB) occurred on the surface of QDs to prepare DTAB-stabilized QDs. Second, DTAB-QDs were incorporated into silica spheres using a modified Stöber method. Third, graft copolymerization of NIPAM monomer and CS was initiated
to form PNIPAM-g-CS nanogels (as an outer shell) on the surface of Q-MS, thus resulting in the formation of Q-MS/PNIPAMg-CS nanospheres (Scheme 1). A whole set of characteristics of nanospheres were investigated. Further researches on these nanospheres were conducted, including fluorescent imaging of tumor cells, cytotoxicity and thermo/pH-dual regulated in vitro drug release. 2. Experimental 2.1. Materials Chitosan (CS, ∼80 mesh, MW . 22.4 kDa, 95% of deacetylation degree) was purchased from Tokyo Kasei-Kogyo Corp. NIsopropylacrylamide (NIPAM, 99%), N,N -methylenebisacrylamide (MBA, 99%), ammonium persulphate (APS, 99%), 3mercaptopropionic acid (MPA, 98%), dodecyltrimethylammonium bromide (DTAB, 99%), sodium dodecyl sulfate (SDS, 97%), 3-(trimethoxylsilyl) propyl methacrylate (MPS, 95%) and tetraethyoxysilane (TEOS, 98%) were obtained from Sigma–Aldrich Corp. Doxorubicin (Dox) in the form of hydrochloride salts was obtained from Beijing Huafeng Corp. 3-[4,5-Dimethyl-thiazol-2-yl]-2,5diphenyl-tetrazolium bromide (MTT) and biological reagents were brought from Invitrogen Corp. Other chemicals with analytical grade were from Shanghai Chemical Reagent Corp. All chemicals can be used directly as received without any purification. Water used in experiments was of Milli-Q grade. Phosphate buffered salines (PBS, 1 mM) were prepared by modulating the molar ratio of Na2 HPO4 and NaH2 PO4 . 2.2. Preparation of Q-MS/PNIPAM-g-CS hybrid nanospheres MPA-capped CulnS2 /ZnS QDs were freshly prepared via hydrothermal route based on Ref. [40], and detailed preparation was available in Supplementary Data (Part S1). Typically, 5 mg of QDs dispersed in 1 mL chloroform were mixed with 9 mL of aqueous solution containing 0.1 g of DTAB under stirring. Chloroform was evaporated at 65 ◦ C to form DTAB-QDs composites, which were subsequently added into 90 mL of water. Then, 3 mL of NaOH solution (2 M), 0.5 mL of TEOS and 5 mL of ethyl acetate were
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successively added into the above mixture with stirring (100 rpm) for 1 min, and then kept for 6 h at 50 ◦ C. The products were centrifuged, repeatedly washed with water–ethanol. Finally, the products were transferred into 50 mL of ethanol solution containing 67 L of MPS, and the mixture was stirred for 12 h at 25 ◦ C. DTAB template was removed in an ethanol solution of NH4 NO3 (50 mL, 10 mg mL−1 ) via ion exchange [41], and the mixture was refluxed for 6 h. The final products were collected by centrifugation and repeatedly washed with ethanol. The resultant MPS-modified Q-MS was dispersed in water, and diluted to 1 wt% for subsequent experiments. Mixed solution (I) consisting of NIPAM (0.5 g), MBA (10 mg) and water (30 mL) was loaded in a three-necked flask equipped with a reflux condenser. The (I) was bubbled with N2 inlet for 30 min to remove dissolved oxygen. Then, MPS-modified Q-MS dispersed in water (1 g, 1 wt%) was added into the (I) under stirring to form mixed solution (II). Afterward, APS (3 mg) and SDS (5 mg) dissolved in 10 mL of water were rapidly added into the (II) under stirring to form mixed solution (III). After reaction for 2 h at 25 ◦ C, NIPAM monomers were initiated to form PNIPAM networks. Finally, 10 mL of aqueous solution containing 3 mg of APS, 0.1 g of CS and 0.1 mL of acetic acid was added into the (III), and then heated to 70 ◦ C. The polymerization reaction was allowed to proceed for 6 h under a continuous stirring of 300 rpm and a N2 flow of 200 mL min−1 . The products were centrifuged for 30 min at 12 000 rpm to remove unreactive monomers. Resultant products were dispersed in water, and dialyzed (Spectra/Por 7 dialysis membrane, MWCO 5 × 104 ) for 7 days against water frequently. Finally, purified products were prepared, named as Q-MS/PNIPAM-g-CS core/shell hybrid nanospheres. 2.3. Characterization Fourier transform infrared spectra (FTIR, Nicolet, USA) were recorded with a Nicolet 6700 FTIR spectrometer in the scanning region of 400–4000 cm−1 . N2 adsorption–desorption isotherms were given on a Micromeritics Tristar 3000 pore analyzer at 77 K, equipped with Brunauer–Emmett–Teller and Barrett–Joyner–Halenda for the analysis of surface area, pore size and volume. UV–vis spectra were recorded with a UV-2450 spectrophotometer (Shimadzu, Japan). PL spectra were measured by a FLSP 920 fluorescence spectrophotometer (Edinburgh Instruments, Britain). Transmission electron microscope (TEM, Jeol, Japan) images were acquired with a JEOL JEM-1400 TEM at 120 kV of acceleration voltage. Hydrodynamic radius (diameters) was obtained on dynamic light scattering apparatus (DLS, Malven Instruments, U.K.). Fluorescence images of tumor cells were obtained by a FV-300 IX71 confocal laser fluorescence microscope (CLFM, Olympus). 2.4. Cellular uptake of nanospheres and imaging Fluorescence imaging of HepG2 cells (bought from the cell bank of Shanghai Science Academe) was recorded by CLFM. The 6 × 104 cells/well was seeded on a 6-well plate for 24 h at 37 ◦ C. Then, the nanospheres dispersed in water (0.2 mg mL−1 ) were added into cell dishes. After incubation for 12 and 24 h, nanospheres-loaded cells were repeatedly washed with PBS (1 mM, pH 7.4) to remove free nanospheres absorbed or attached on the outer surface of cell membrane. Afterward, these treated cells were detected by CLFM, excited at 480 nm. 2.5. Drug loading and release of nanosphere carriers Nanospheres (20 mg) were ultrasonically dispersed in 25 mL of Dox solution (1 mg mL−1 , 1% of NaCl, pH 7.4 modulated by
0.1 M of NaOH) to form homogeneous solution, which was continuously stirred for 24 h at 25 ◦ C until Dox concentration in the solution stabilized. Then, the solution was centrifuged for 15 min at 5000 rpm, and washed with water twice to remove unbound or surface-absorbed Dox. The mass of Dox loaded in nanospheres was calculated by subtracting the Dox mass in the supernatant from the total Dox mass in the initial solution. Free Dox mass (Mfree-Dox ) in solution was determined by UV–vis spectrophotometer at 480 nm using Lambert–Beer Law. The Dox-loading efficiency (LE) and capacity (LC) were calculated according to the following equations: Dox-LE (%) = 100 ×
Mtotal-Dox − Mfree-Dox Mtotal-Dox
(1)
Dox-LC (%) = 100 ×
Mtotal−Dox − Mfree−Dox Mtotal−nanospheres
(2)
To investigate pH/temperature-dependent Dox release, 20 mg of Dox-loaded nanospheres in PBS (1 mM, pH 5.0, 6.5 and 7.4) were transferred into a dialysis bag (MWCO 5 × 104 ), which were then gently shaken in water bath at different temperatures (25, 37 and 42 ◦ C). At defined time intervals, 1 mL of sample in each mixture solution was withdrawn and analyzed by UV–vis at 480 nm. An equal volume of PBS was added after each sampling to retain a constant volume. Each experiment of drug release was performed in triplicate, and each result was expressed as the mean of three repeated measurements. 2.6. In vitro cytotoxicity of nanosphere carriers Cytotoxicity studies of the blank, Dox-loaded nanospheres and free Dox on HepG2 cells (as model tumor cells) were assessed by MTT assay. The 1 × 104 cells/well was incubated in Dulbecco’s modified Eagle medium containing calf serum (10 wt%) and 100 units mL−1 of penicillin in a fully humidified incubator with 5 vol% of CO2 at 37 ◦ C. When these cells reached 80% of confluence with normal morphology, the blank, Dox-loaded nanospheres with a concentration of 0.2–1.0 mg mL−1 , and free Dox were added into cell dishes, respectively. Then, these cell dishes were put into incubators for 48 h at 37 ◦ C, and no detectable damage to cells was observed. After incubation for a defined time, 20 L of MTT reagent (0.5 mg mL−1 , diluted in culture medium) was substituted for culture medium, followed by an additional 2 h incubation. Finally, MTT-medium was removed attentively, and 150 L of dimethyl sulfoxide was added into each well to dissolve formazan crystals. The absorbance (A) of respective wells (colored solutions) was recorded at 570 nm, using a Multiskan MK3 Enzyme-labeled Instrument (Thermo Scientific, USA). All cytotoxicity experiments were carried out in triplicate, and all results were averaged. The cell survival rate was calculated by the equation, as below: Cell survival rate (%) = 100 ×
Atest cells Acontrol cells
(3)
3. Results and discussion 3.1. Preparation and characterization of Q-MS/PNIPAM-g-CS nanospheres Q-MS composites were prepared using a base-catalyzed sol–gel approach in the presence of QDs [42,43]. Then, Q-MS were dispersed in anhydrous ethanol containing MPS for surface modification of C C bonds, which form covalent bonds between inorganic substrates and polymers [44]. Using MPS-modified Q-MS composites as seeds, Q-MS/PNIPAM-g-CS core/shell nanospheres were prepared via graft copolymerization of NIPAM monomer and CS. FTIR spectra of Q-MS and Q-MS/PNIPAM-g-CS show an evidence
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Fig. 1. (a) FTIR spectra, (b) N2 adsorption–desorption isotherms (inset: the pore diameter distribution), (c) UV–vis absorption and PL emission spectra, and TEM images of (d) QDs, (e) Q-MS and (f) Q-MS/PNIPAM-g-CS.
for the preparation of nanospheres (in Fig. 1a). For Q-MS, a strong signal for water bending vibration at 1655 cm−1 is observed, and is attributed to hydrophilic silanol surface after template extraction [45]. In addition, C O stretching at 1718 cm−1 and CH2 stretching at 2978 cm−1 indicate successful modification of Q-MS by MPS. For Q-MS/PNIPAM-g-CS, characteristic absorption bands at 2973 and 1390 cm−1 testify that NIPAM chains exist in nanospheres. The peak at 1560 cm−1 is assigned to NH2 bending vibration of CS. For both samples, characteristic peaks at 460, 805 and 1035 cm−1 denote that the silica framework of Q-MS maintains unchanged in nanospheres upon the coating of polymers. These results suggest the polymerization of NIPAM and CS, and the integration of silica spheres and PNIPAM-g-CS polymers. Fig. 1b displays the N2 adsorption–desorption isotherms and pore size distribution (insert) of both samples. The surface area, pore volume and pore diameter of Q-MS are 857 m2 g−1 , 1.1 cm3 g−1 and 2.8 nm, respectively. For Q-MSN/PNIPAM-g-CS, corresponding parameters are 698 m2 g−1 , 0.9 cm3 g−1 and 2.7 nm, respectively. The pore diameter has a slight decrease from 2.8 to 2.7 nm, implying that PNIPAM-g-CS is predominantly distributed on the exterior surface of Q-MS [46]. The nanospheres with nearly identical mesoporosity as Q-MS exhibit a promising application in stimulus-responsive drug release platforms. Photoluminescence (PL) properties of nanospheres were characterized by UV–vis and PL spectrometry. As elaborated in Fig. 1c, no remarkable change in PL spectra of QDs appears after encapsulated into PNIPAM-g-CS [40], indicating that the silica on outer layers indeed protects QDs.
Especially, these nanospheres possess well-stabilized dispersion with coupling shells, and intense PL can retain for several weeks (in Fig. S1). Morphology and structure of QDs, Q-MS and Q-MS/PNIPAM-gCS nanospheres were tested by wide-field TEM images (in Fig. 1d–f). The QDs (Fig. 1d) show a uniform, discrete spherical shape with an average diameter of 4.8 ± 0.5 nm. In Fig. 1e, multiple dark spots are observed in each particle with a size of 50 ± 5 nm. The average diameter of black dots coated by the silica shell is ∼20 nm, denoting that the QDs were embedded within mesoporous silica matrix. Fig. 1f displays core/shelled, spherical and polymer-encapsulated (bright ring) spheres, and the average size is 95 ± 6 nm. These nanospheres consist of black dots (QDs) encapsulated in the gray silica interlayer (SiO2 ) and a dark shell of PNIPAM-g-CS polymers. In addition, average hydrodynamic size of nanospheres measured by DLS is 125 nm at 25 ◦ C, and there is a low diameter polydispersity index (0.075) [47]. The hydrodynamic size from DLS is larger than the size from TEM due to hydrate layers in aqueous environment [48], further implying the existence of copolymer shell layers in nanospheres. TEM images of QDs, Q-MS and Q-MS/PNIPAM-gCS (in Fig. 1d–f) are consistent with corresponding absorption and emission spectra (in Fig. S4). Dramatic red-shift in spectra from the initial QDs to Q-MS (with mesoporous silica shells around QDs) and to Q-MS/PNIPAM-g-CS (with polymer shells around Q-MS) were observed, implying a gradual increment in size [15]. In addition, 1 H-NMR spectra of PNIPAM-g-CS nanogels (in Part S2) testified the graft copolymers (in Fig. S2a). An acute proton peak
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Fig. 2. (a) Diameter as a function of temperature of PNIPAM-g-CS nanogels and Q-MS/PNIPAM-g-CS nanospheres in PBS (1 mM, pH 7.4). (b) Diameter as a function of pH of the two samples in PBS at 25 ◦ C. (c) DLS data of nanospheres stored in PBS at 37 ◦ C containing different NaCl concentrations.
( CH CH2 ) at 2.1 ppm, a peak ( NH CH 10.0. By contrast, the diameter of nanospheres is nearly unchanged at pH > 8.0. These results reveal that two samples are pH-sensitive, and Q-MS in nanospheres hardly affect this sensitivity, in an agreement with previous reports [49]. At a lower pH, OH from PNIPAM-g-CS is protonated. Increasing the pH, the degree of protonation reduces, and OH gradually translates to O− [35]. As a result, electrostatic repulsion and hydrophilia of O− cause an increase in the swelling degree of nanospheres. When the pH exceeds a certain value, hydrogen-bonding between water molecules and polymer chains gradually increases, thus resulting in the diameter of nanospheres trending to stability [44]. In addition, the effect extent of pH on the diameter of nanogels is larger than nanospheres under the same conditions (pH 2–12), which may be ascribed to the existence of Q-MS that partly restrains the swelling degree of nanospheres [44,49].
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Fig. 3. CLFM images of HepG2 cells after (a) 12 h and (b) 24 h treatment with 0.2 mg mL−1 of nanospheres in PBS (1 mM, pH 7.4). The excitation wavelength is 480 nm, and the scale bars are 20 m.
Nanospheres’ pH-sensitivity could be further verified by the effect of NaCl concentration on the size and stability of nanospheres (in Fig. 2c). For the system consisting of nanospheres (0.2 mg mL−1 ), PBS (1 mM, pH 5.0/7.4) and NaCl (0.5–0.8 M), the average diameter of nanospheres shows relatively slight fluctuation (105–110 nm), which is consistent with the results from TEM, implying monodisperse NPs. It is mainly ascribed to PNIPAM-g-CS shells that bring more ions in a dilute salt solution and thus lead to electrostatic repulsion and low viscosity. As a result, the aggregation of NPs is restrained. In 0.8–1.2 M of NaCl, nanospheres have a larger diameter by DLS compared to TEM, revealing a possible aggregation. It is probably due to the competition of excess salts with PNIPAM-g-CS nanogels for water molecules in solution, and the competition results in high viscosity and the following aggregation of nanospheres. Under the same concentration, the diameter of nanospheres at pH 5.0 is less than at pH 7.4, which can be explained by electrostatic shielding of polymer shells and excessive charges appeared in acid environment. The electrostatic shielding causes the shrink of polymers, and the decline of intermolecular entangled degree. So, the nanospheres have a higher stability at pH 5.0, and the diameter is desirable in acidic and low-salt solutions.
affected by Dox, and increase with the increment of Dox concentration ([Dox]). Increasing [Dox] from 0.05 to 1.0 mg mL−1 , LC increases from 3.1% to 45.2%, while LE increases from 11.9% to 61.1%. These denoted that Dox-loading into nanospheres was [Dox]-dependent. Dox molecule consists of acidic phenolichydroxyl groups and alkaline amino groups that can interact
3.3. In vitro tumor cell imaging of nanospheres The encapsulation of QDs into nanospheres gives a possibility of PL imaging. In this regard, CLFM was adopted to evaluate the nanospheres for tumor cell imaging. The qualitative cell uptake of nanospheres was investigated, as demonstrated in Fig. 3. CLFM images of HepG2 cells treated with nanospheres denoted that the nanospheres were endocytosed into HepG2 cells. When irradiated at 480 nm, the QDs immobilized in nanospheres produced bright green fluorescence. Fluorescence intensity increases (from Fig. 3a and b), suggesting that the internalized extent of nanospheres by cells enhances with the increase of incubation time. It also means that the amounts of nanospheres uptaken by HepG2 cells are dependent on incubation time. In addition, no remarkable autofluorescence is observed under the same conditions. The areas where nanospheres did not permeate are distinct, and there is no dark region appearing in cells, implying a potential of the nanospheres as effective PL imaging in tumor cells. 3.4. Drug loading and thermo/pH-regulated release in vitro Dox was used for a typical anticancer drug to evaluate potential application of nanospheres. In Fig. 4a, Dox with different concentrations was loaded in nanospheres. Both LC and LE are markedly
Fig. 4. (a) Effects of Dox concentration on Dox-loading capacity (LC) and efficiency (LE) in PBS (1 mM, pH 7.4) at 37 ◦ C. (b) Effects of pH on Dox-loading content and embed efficiency for nanosphere carriers in PBS at 25 ◦ C.
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Fig. 5. Cumulative Dox release profiles from nanosphere carriers at selected temperatures (25, 37 and 42 ◦ C) in PBS (1 mM, pH 5.0, 6.5 and 7.4). The data points are averages of six measurements.
with NH2 and OH of PNIPAM-g-CS polymers (as a shell in nanospheres). This helps to form intermolecular complexes via hydrogen bonding [50], inducing a high LE. Moreover, Doxloading into nanospheres was efficient because nanospheres had no adsorption interaction with Dox, otherwise distinct decrease in LE would be observed upon the increase of [Dox] [51]. Loading content (LC) and embed efficiency (EE) of Dox into nanosphere carriers were investigated, as shown in Fig. 4b. Both two were markedly influenced by pH. Upon the pH increase from 4.9 to 9.2, LC increased from 0.02 to 0.45 mmol g−1 , while EE increased from 4.2 to 87.1 wt%. The EE has a sharp increase at pH 8.3 (compared to pH 7.4), which could be explained by Dox-loading process based on physical adsorption in mesoporous channels. Isoelectric point of Dox was 8.25 [44], and net charge of Dox was around zero. On this account, the electrostatic interactions between polymer shells and Dox molecules were minimal. Dox molecules would be easily absorbed into mesoporous channels across the larger mesh size of polymer shells at an elevated pH. Dox molecule is water-soluble at a lower pH, but becomes insoluble in alkaline environment. In aqueous solution of Dox with a higher pH, electrostatic repulsion among adsorbed Dox molecules markedly decreased, which caused the adsorption of Dox molecules into nanospheres. To estimate Dox release of nanosphere carriers (in Fig. S3), related experiments were conducted. As displayed in Fig. 5, Dox release responses of nanosphere carriers in PBS (pH 5.0, 6.5 and 7.4) were studied at 25 ◦ C (LCST). Both temperature and pH have a remarked influence on Dox release. At 25 ◦ C, Dox release was less than 15.1% at pH 5.0, and greater than at a higher pH, which was due to acid-labile physical adsorption. Dox release at 37 ◦ C was apparently similar to that at 42 ◦ C, but the release extent of Dox was intermediate between 25 and 42 ◦ C. In other words, Dox release at 42 ◦ C was greater and faster than at a lower temperature. At each investigated pH, a rapid release occurred during the initial stage (0–2 h) due to the changes of chemical environment in carriers, followed by a slower release. At a longer duration (2–24 h), the extent of Dox release continued to increase before reaching a sustained release, which was similar to the cases at different temperatures. Internal structure of PNIPAM was loose while drug molecules diffused from the interior of nanogels in an easy way [52]. In Dox-loaded nanospheres, Dox diffusion across nanogel shells may have a distance-limiting
Fig. 6. In vitro cytotoxicity of the blank, Dox-loaded nanospheres and free Dox at different concentrations against HepG2 cells after 48 h incubation at 37 ◦ C.
mechanism. At pH 7.4 or 25 ◦ C, nanogel shells were in the swelling state, and hydrodynamic size of nanospheres was large, thus causing a slow release. Enhanced electrostatic attraction between positively charged drug molecules and negatively charged nanogels was unfavorable for the mass-transport of Dox molecules. By contrast, nanogel shells were collapsed at a higher temperature (>37 ◦ C). At a lower pH (LCST) and mild acidic medium were favorable for Dox release, controlled via small changes in the vicinity of LCST (37 ◦ C) and a lower pH (mild acidic medium), Doxloaded nanospheres exhibited a faster cumulative in vitro Dox release. Released Dox still maintained high anticancer activity, and blank nanosphere carriers produced neglectful toxicity to HepG2 cells. Thus, the nanospheres could be developed as a targeting probe for tumor cells, and exhibit promising applications for diagnosis, therapy, controllable drug release, and real-time detection of drug treatment in the intracellular process. Acknowledgments This work was financially supported by Grants from the Shandong Provincial Doctoral Foundation of China (No. 2011BSE27195), the Postdoctoral Science Foundation of China (No. 2013M531164) and the Postdoctoral Science Foundation of Shanghai (No. 13R21413800). Appendix A. Supplementary data Supplementary data associated with this article can be found, in the online version, at http://dx.doi.org/10.1016/j.colsurfb. 2014.01.044. References [1] M. Bruchez, M. Moronne, P. Gin, S. Weiss, A.P. Alivisatos, Science 281 (1998) 2013–2016. [2] W.C.W. Chan, S. Nie, Science 281 (1998) 2016–2018. [3] N. Tessler, V. Medvedev, M. Kazes, S.H. Kan, U. Banin, Science 295 (2002) 1506–1508. [4] C. Burda, X. Chen, R. Narayanan, M.A. El-Sayed, Chem. Rev. 105 (2005) 1025–1102. [5] R.C. Somers, M.G. Bawendi, D.G. Nocera, Chem. Soc. Rev. 36 (2007) 579–591. [6] A.L. Rogach, N. Gaponik, J.M. Lupton, C. Bertoni, D.E. Gallardo, S. Dunn, Angew. Chem. Int. Ed. 120 (2008) 6638–6650. [7] Y. Dai, P. Ma, Z. Cheng, X. Kang, X. Zhang, Z. Hou, C. Li, D. Yang, X. Zhai, J. Lin, ACS Nano 6 (2012) 3327–3338. [8] X. Zhang, P. Yang, Y. Dai, P. Ma, X. Li, Z. Cheng, Z. Hou, X. Kang, C. Li, J. Lin, Adv. Funct. Mater. 23 (2013) 4067–4078.
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