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Multiple Functional Hyperbranched Poly(amido amine) Nanoparticles: Synthesis and Application in Cell Imaging Wen Yang,†,‡ Cai-Yuan Pan,*,† Xi-Qiu Liu,§ and Jun Wang§ †
Department of Polymer Science and Engineering, CAS Key Laboratory of Soft Matter Chemistry, University of Science and Technology of China, Hefei, Anhui, 230026, P. R. China ‡ Department of Polymer Science and Engineering, Anhui Key Laboratory of Controllable Chemistry Reaction & Materials Chemical Engineering, Hefei University of Technology, Hefei, Anhui, 230009, P. R. China § Hefei National Laboratory for Physical Sciences at Microscale and School of Life Sciences, University of Science and Technology of China, Hefei, Anhui 230027, P. R. China ABSTRACT: The hyperbranched poly(amido amine) nanoparticles (HPAMAM NPs) with multiple functions, such as biodegradability, autofluorescence, and specific affinity, were successfully prepared by Michael addition dispersion polymerization of CBA, AEPZ, and N-galactosamine hydrochloride (or N-glucosamine hydrochloride) in a mixture of methanol/water. The resultant NPs displayed strong photoluminescence, high photostability, broad absorption, and emission (from 430 to 620 nm) spectra. The fluorescence from HPAMAM NPs may be attributed to the tertiary amine chromophore. The incubation results of the liver cancer cells, HepG2, with the NPs showed that the NPs are nontoxic and can be recognized by asialoglycoprotein receptors on the surface of HepG2 and then can be internalized. Therefore, they have potential applications in bioimaging and drug or gene delivery.
’ INTRODUCTION Recently, polymeric nanoparticles with multiple functions have received a great deal of attention due to their potential applications in the biomedical field.1 Among these functionalities, fluorescence is of key importance since fluorescence-based methods have been widely applied in probing biomolecular interactions and the study of various biochemical processes.2 The fluorescent organic dyes, quantum dots (QDs), and fluorescent conjugated polymers are extensively used as fluorescent markers. However, the organic dyes suffered from many limitations, including narrow excitation/broad emission spectra, poor photostability, and substantial cytotoxicity,3 and the relatively low photobleaching thresholds of the organic dyes limited their effectiveness in long-term and three-dimensional imaging.4 QDs exhibit high photostability; broad exciting wavelength; and narrow, tunable emission spectra yet possess limitations, including low specificity, poor biocompatibility, and high toxicity.5 A strategy to resolve such issues is to conjugate the QDs on, or to encapsulate them in, biodegradable polymers;6 nevertheless, their potential carcinogenesis or toxicity, and other respective disadvantages, still remained.7 Thus, finding a new type of fluorescent materials is crucial for development of a watersoluble, biocompatible, and biodegradable fluorescent polymers with multiple functionalities. The poly(amido amine) (PAMAM) and poly(propylene imine) (PPI) dendrimers have been reported to emit fluorescence under suitable light excitation,810 but use of them as fluorescent probes in analytical and biological applications has not been studied, probably due to their weak fluorescence. Our previous results showed that when the aliphatic amines in the hyperbranched PAMAM (HPAMAM) were quaternized, the fluorescence was quenched almost completely. The linear r 2011 American Chemical Society
PAMAM possessing primary and secondary amines did not display fluorescence, but the HPAMAM with tertiary amines as branching units emitted fluorescence, though the emission was not strong enough for cell imaging.9 So, it is reasonable to presume that the tertiary amine unit in the HPAMAM is a chromophore, and this presumption was supported by previous studies.1115 Some tertiary amines, such as trimethylamine, triethylamine, and tri(n-propyl)amine, emitted fluorescence in the gas state,11 or in nonpolar solvent.12 But the primary and secondary amines do not emit fluorescence because a hydrogen atom is easily lost by predissociation.11 Since the tertiary amines are already known to act as chromophore in the peptides and other important biological systems, such as neurotransmitters,15 a study of fluorescent behavior and applications of the polymers containing tertiary amines should be interesting. Generally, the nanoparticles (NPs) displayed higher fluorescence and higher photostability than the small molecule labels;16 in addition, the cross-linking units composed of tertiary amine reduce their collision and resulted in high fluorescence. Thus, if the NPs containing tertiary amine as cross-linked units can be prepared, the fluorescent labels-doped step can be eliminated. However, the biological applications often need the multifunctional NPs, which are generally achieved through several steps.17 The aim of our investigation is to find a convenient and feasible strategy for the preparation of multifunctional NPs via one-pot polymerization. Since PAMAM is a well-known biocompatible polymer and has been applied in gene and drug delivery,18,19 and the surface Received: December 7, 2010 Revised: February 22, 2011 Published: March 16, 2011 1523
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Biomacromolecules modification of HPAMAM with D-mannose enhanced the fluorescence as high as 340 times that of its precursor,10 we selected the NPs with a structure similar to HPAMAM. To give the NPs a biodegradable property, the cross-linked NPs containing abundant SS bonds in their backbones were synthesized because disulfide linkage can be easily cleaved in the presence of biological or chemical stimuli.19 When the NPs are applied in drug delivery or cell imaging, they should have specific affinity to a target cell, such as the liver tumor cells. Herein, we reported the synthesis of multifunctional PAMAM NPs and their application in the cell imaging.
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methanol/water mixture (85/15, v/v, 2 mL). The polymerization was carried out at 50 °C for 5 days in a sealed glass tube after three freezepumpthaw cycles. The solution was acidified with a 2N HCl aqueous solution to a pH of ∼2. Then, the solution was precipitated from 30 mL of acetone while being vigorously stirred. The polymer NPs were collected by filtration and purified by reprecipitation in acetone. The product was obtained in 45% yield after being dried in a vacuum for 1 day at room temperature. The Quantum Yield of the Poly(amido amine) Gels. The quantum yield of the NPs was calculated according to the following equation: ΦSA ¼ ΦST ðSSA =SST ÞðηSA =ηST Þ2
’ EXPERIMENTAL SECTION Materials. The 1-(2-aminoethyl)piperazine (AEPZ, 99%, Aldrich), cystamine dihydrochloride (g98%, Changzhou Furong Chemical Co.), D-glucosamine hydrochloride (>99%, Sigma), D-galactosamine hydrochloride (g99%, Fluka), and dithiothreitol (DTT, Sigma) were used as received without further purification. NaOH (g96%), dichloromethane (g99.5%), methanol (g99.5%), and acetone (g99.5%) were purchased from Sinopharm Chemical Reagent Co. Ltd. and were used as received. Acryloyl chloride was purified by distilling it under reduced pressure. Characterization. 1H and 13C NMR spectra were measured on a Bruker DMX-300 NMR nuclear magnetic resonance (NMR) instrument using tetramethylsilane (TMS) as an internal standard and CDCl3 or DMSO-d6 as the solvent. UVvis spectra were acquired on a Shimadzu UV-2401PC UVvisible scanning spectrophotometer at room temperature. The fluorescence spectra and photobleaching measurements were carried out at room temperature on a Shimadzu RF5301PC luminescence spectrophotometer. Laser confocal scanning microscope images were taken on a Carl Zeiss LSM 510. TEM images were obtained on a Hitachi model H-800 transmission electron microscope (TEM) with an accelerating voltage of 200 kV. The sample was prepared as follows. The galactosamine-modified HPAMAM NPs were dissolved in methanol (1 mg/mL), and then a drop of NPs solution was pipetted onto a copper grid and allowed to dry in the air before observation. The size and size distribution of NPs in aqueous solution measured by dynamic light scattering (DLS) were carried out on a Malvern Zetasizer Nano ZS90 with a HeNe laser (633 nm) and 90° collecting optics. All samples with a concentration of 0.2 mg/mL were prepared by dispersing the NPs in methanol, and then filtered through a 0.45 μm membrane filter (Millipore) prior to measurements. All measurements were carried out at 25 °C, and the data were analyzed by Malvern Dispersion Technology Software 4.20. Synthesis of N,N0 -Cystaminebisacrylamide (CBA). Cystamine dihydrochloride (5.8 g, 25 mmol) was added into 60 mL of an aqueous NaOH (4 g, 100 mmol) solution in a 500 mL three-necked round-bottom flask. A solution of acryloyl chloride (4.7 g, 50 mmol) in dichloromethane (5 mL) was added dropwise while stirring at 0 °C. After the addition was complete, the mixture was stirred at room temperature for 3 h. The solid product was obtained by filtration. The liquid phase was extracted with dichloromethane, and the organic extract was dried under anhydrous Na2SO4 overnight. The solvent was removed under reduced pressure. The final product was obtained as a white solid (4.2 g, 65%). 1H NMR (300 MHz, CDCl3, δ, TMS) 6.57 (2H, CONH); 6.36 and 6.22 (4H, CHHdCHCO); 5.69 (2H, CHHdCHCO); 3.67 (4H, CONHCH2CH2SS); 2.89 (4H, CONHCH2CH2S). 13C NMR (300 MHz, CDCl3, δ, TMS): 167 (CONHCH2); 131 (CH2dCH); 126 (CH2dCH); 39 (CONHCH2CH2SS); 38 (CONHCH2CH2SS). Preparation of HPAMAM Nanoparticles. The synthetic procedure is as follows. The CBA (0.26 g, 1 mmol), AEPZ (0.065 g, 0.5 mmol), and D-galactosamine hydrochloride (0.216 g, 1 mmol) or D-glucosamine hydrochloride (0.216 g, 1 mmol) were added into a
where Φ = quantum yield, S = gradient of the curve obtained from the plot of intensity versus absorbance, η = refractive index of the solvent, SA = the sample, and ST = the standard. Anthracene (quantum yield = 0.27 in ethanol) was used as a standard. The NPs were dispersed in water, and anthracene was dissolved in ethanol. The slit width was kept the same for both the standard and the samples. Absorbance was measured on a Shimadzu UV-2401PC spectrophotometer. Cell Internalization. Human hepatocellular liver carcinoma HepG2 cells from the American Type Culture Collection (Manassas, VA) were maintained in Dulbecco’s Modified Eagle’s medium (DMEM, Invitrogen, Carlsbad, CA) supplemented with 10% (v/v) heat-inactivated fetal bovine serum (Hyclone), 100 mg/mL streptomycin, and 100 U/mL penicillin in a humidified atmosphere (relative humidity: 95%; 5% CO2) at 37 °C. For microscopic imaging, cells were seeded on sterile glass coverslips in 24-well cell culture plates to allow complete adhesion. The cells were then incubated with 2 mg/mL of gels in a DMEM medium without serum for 4 h. After being washed with phosphatebuffered saline (PBS, 1 mg/mL) three times, the cells were fixed with 4% formaldehyde. The slides were mounted and observed with a Zeiss LSM510 Laser Confocal Scanning Microscope imaging system. Viability Assay. The cytotoxicity of NPs was evaluated in vitro using an MTT assay, using PEI25k as the control. HepG2 cells were seeded in 96-well plates at 15 000 cells per well in 100 μL of complete DMEM medium and incubated at 37 °C in a 5% CO2 atmosphere for 24 h. The culture medium was replaced with 100 μL of a fresh medium containing NPs. Cells were further incubated for 24 h, followed by the addition of 25 μL of MTT stock solution (5 mg/mL in PBS) to achieve a final concentration of 1 mg/mL. After incubation for an additional 2 h, 100 μL of the extraction buffer (20% SDS in 50% DMF, pH 4.7, prepared at 37 °C) was added to the wells, and it was incubated overnight at 37 °C. The absorbance of the solution was then measured at 570 nm using a Bio-Rad 680 microplate reader. The cell viability was normalized to that of HepG2 cells cultured in the culture medium without nanoparticles.
’ RESULTS AND DISCUSSION Preparation of PAMAM NPs. Although various HPAMAMs have been successfully prepared through Michael addition polymerization of diacrylate and triamine monomers,20 we did not find the synthesis of HPAMAM NPs in the literature. To synthesize the stable and degradable polymer NPs with bright fluorescence, the selection of an appropriate reaction system is necessary. Since disulfide linkage can be easily cleaved in the presence of biological or chemical stimuli,19 N,N0 -cystaminebisacrylamide (CBA) with disulfide linkage was chosen as the diacrylate. Because galactose (Gal) has a strong affinity to liver tumor cells21 and can stabilize the NPs in water, its derivative, galactosamime (N-Gal), was used as a starting material. For synthesis of the stable polymer NPs, the selection of appropriate 1524
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Scheme 1. Synthesis of HPAMAM NPsa
a
(a) Starting materials. (b) HPAMAM with Gal (or Glu) surface groups. (c) Cross-linked HPAMAM NPs. (d) Degraded HPAMAM NPs.
solvents is crucial. Methanol is a good solvent for the starting materials and the resulting polymer, whereas water is a nonsolvent of CBA and the formed polymer. A mixture of methanol and H2O (85/15, v/v) was applied as a reaction medium in the dispersion polymerization after the effect of the CH3OH/H2O ratio on the stability of NPs was tested repeatedly. The architecture of PAMAM was determined by the feed molar ratio and reaction conditions. When a molar ratio of diacrylamide/1-(2aminoethyl)piperazine (AEPZ) was 2/1.5, the cross-linking reaction occurred.20 In order to obtain cross-linked polymer gels, the Michael addition dispersion polymerization with a feed molar ratio of CBA/AEPZ/N-Gal = 2/1/2 was carried out in the mixture of CH3OH and H2O (85/15, v/v) at 50 °C. As shown in Scheme 1, the initial polymerization proceeded homogeneously because all of the starting materials are soluble in the solvents (Scheme 1a). The continuous polymerization resulted in the hyperbranched chains with surface acrylate groups owing to polymerization of the AB2 intermediates with one amine and two acryloyl groups.20 At the same time, a Michael
addition reaction of the terminal acryloyl groups with N-Gal’s also took place. This resulted in HPAMAM chains with surface Gal groups that will stabilize the NPs formed later (Scheme 1b). Because the polymerization was carried out in a relatively dilute solution, the polymerization was slow, but it benefited control of the NPs’ formation. When the polymerization lasted 24 h, the HPAMAMs soluble in methanol were achieved. To confirm that the formed tertiary amine acted as a branched unit, 13C NMR spectra of the polymers formed at 24 h of polymerization were measured, and a typical spectrum is shown in Figure 1. Two signals at δ 50.1 and 50.3 ppm are ascribed to the three methylene carbons next to the nitrogen of the tertiary amine, which indicates the transformation of the secondary amine (formed) into a tertiary amine, forming a branched structure.20 The signal of methine carbon next to the ether oxygen of Gal at δ 73.8 ppm supports the Gal groups on the HPAMAM. When the polymerization continued further and the chain length of growing chains increased to a critical value, the polymer chains became insoluble in the reaction media, and the phase separation 1525
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Biomacromolecules occurred to form the spherical aggregates. Generally, the Michael addition polymerization of diacrylamide and AEPZ with a molar ratio of 2:1 produced the HPAMAMs with acryloyl groups on their surface.20 However, due to the reaction of N-Gal with the acryloyl group and its low reactivity, the acryloyl and the secondary amine groups on the surface of HPAMAM were retained, which was supported by the 13C NMR spectrum of the N-Gal NPs in Figure 1: the carbon signals of methylene groups next to nitrogen of the secondary amine at δ 43.6 and 43.7 ppm (n) and vinyl carbon signals at δ 125.8 and 131.6 ppm. Further polymerization must lead to intermolecular reaction in the aggregates, forming the cross-linked HPAMAM NPs (N-Gal NPs), as shown in Scheme 1c. TEM and DLS were used to identify whether the NPs were formed at 5 days of polymerization, and the results are shown in Figure 2. The TEM image in Figure 2a shows that the aggregates are spherical, and their average diameter (D) was approximately 137 nm (average value for 25 particles). The same sample was used in the DLS measurements, and the result is shown in Figure 2b. The D of the resulting N-Gal NPs was 146 nm. Compared to the results obtained from the aggregates formed by self-assembly,22 the difference between the sizes measured by TEM and DLS is relatively small, which might be due to cross-
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linking of the N-Gal NPs. In order to compare liver tumor affinity of the Gal with that of the glucose (Glu), the HPAMAM NPs with surface Glu groups (N-Glu NPs) were also prepared by the same method except that the N-Gal was replaced by the glucosamine (N-Glu). Both the N-Gal and N-Glu have similar solubility in water, so the size of N-Gal NPs (D = 146 nm) and N-Glu1 NPs (D = 149 nm), which were prepared at the same feed molar ratio and under the polymerization conditions, is similar (Table 1). These two NPs were used in the following study, except when specially mentioned. The NPs’ size can be adjusted by varying the ratio of methanol to water, and the amount of Gal or Glu added. When the polymerization with a feed molar ratio of CBA/AEPZ/D-Glu = 2/1/1 was carried out in the mixture of CH3OH/H2O (85/15, v/v) at 50 °C for 5 days, the size of the resulting NPs was measured by DLS, and the results are listed in Table 1. We can see that with a decrease of the N-Glu in the feed, the obtained N-Glu NPs increased from D = 149 nm (CBA/AEPZ/N-Glu = 2/1/2) to D = 280 nm. When the N-Glu hydrochloride dosage in the feed was reduced, the amount of N-Glu groups on the surface of the resulting HPAMAM chains was decreased, which lowered the stabilizing ability of the surface N-Glu groups, leading to the aggregation of more macromolecules, and the big spherical NPs were formed. The N-Gal and N-Glu groups on the surface of NPs are crucial for the affinity of the NPs with the liver tumor cells. Thus, 1H NMR spectra of the N-Gal and N-Glu NPs were measured and are shown in Figure 3a and b, respectively. The proton signals at δ 4.57 (c) and 4.91 ppm (c0 ) are ascribed to the methine group next to the ether and hydroxyl oxygen of the N-Gal and the N-Glu, respectively. This supports N-Gal or N-Glu being on the Table 1. Conditions and Results for Preparation of HPAMAM Nanoparticlesa feed molar ratio CBA/AEPZ/D-Gal (or
yieldb
diameterc
ζ potentialc
no
Glu)
(%)
(nm)
(mV)
N-Gald
2/1/2
55
146 ( 7.18 37.9 ( 3.26
N-Glu1e
2/1/2
60
149 ( 16.2 42.8 ( 2.82
N-Glu2e
2/1/1
58
280
Polymerization conditions: temperature, 50 °C; time, 5 days; solvents, CH3OH/H2O = 85/15 (v/v). b Measured by weight method. c Measured by DLS method d D-galactosamine hydrochloride was used. e D-glucosamine hydrochloride was used. a
Figure 1. 13C NMR spectrum of the hyperbranched PAMAM obtained from 1 day of polymerization at 50 °C. Feed molar ratio of CBA/AEPZ/ N-Gal hydrochloride = 2/1/2, CH3OH/H2O = 85/15 (v/v).
Figure 2. TEM image (a) and size and size distribution (b, concentration: 0.2 mg/mL) of the N-Gal NPs obtained from polymerization in the mixture of CH3OH/H2O (85/15, v/v) at 50 °C for 5 days with a feed molar ratio of CBA/AEPZ/N-Gal = 2/1/2. 1526
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Biomacromolecules surface of NPs because those groups inside the cross-linked particles cannot be detected. Fluorescence Properties of the NPs. Our previous study demonstrated that when the surface acryloyl group of HPAMAM was substituted by mannose, the fluorescence of the resulting HPAMAM increased significantly (approximately 340 times).10 To study the effect of polymer structure on the fluorescence, UVvis and fluorescence of the N-Gal NPs and N-Gal HPAMAM were measured. The UVvis spectrum of the NPs in Figure 4a shows a broad absorption band ranging from 250 to approximately 600 nm, but HPAMAM exhibits a relatively narrow absorption band ranging from 240 to approximately 350 nm. No fluorescent emission was observed when excited at 280 nm. But under excitation wavelengths of 400, 470, and 510 nm, the NPs emitted stronger fluorescence (b0 , b00 , and b000 ) than the N-Gal HPAMAM (a0 , a00 , and a000 ), as shown in
Figure 3. 1H NMR spectra (d6-DMSO) of N-Gal NPs (a) and N-Glu NPs (b) obtained from the polymerization with a feed molar ratio of CBA/AEPZ/N-Gal (or N-Glu) = 2/1/2 in CH3OH/H2O = 85/15 (v/ v) at 50 °C for 5 days.
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Figure 4b, this should be ascribed to contribution of the polymer network, which will be discussed later. In addition, the results in Figure 4b show that the fluorescence intensities of the N-Gal NPs relative to that of the HPAMAM increased progressively from 1.4 times to 4 and to 7 times with a change of the excitation wavelength from 400 to 470 nm and to 510 nm. Probably, the N-Gal NPs contain more excimers emitting the longer wavelength fluorescence than the HPAMAM. The exact reason needs to be further clarified, but this should be related to their structures and size of molecules because the NPs were produced from aggregation of the various molecular weight HPAMAM chains (Mw = 2.07 104 g/mol measured by light laser scattering method for HPAMAM, and D = 146 nm for N-Gal NPs). Further reactions in the aggregates formed after phase separation should change some structures of HPAMAM; thus, the NPs displayed UVvis spectra and fluorescent properties different from those of HPAMAM. To further ascertain the function of the cross-linked network in enhancing fluorescence, the dithiothreitol (DTT) was used to degrade the N-Gal NPs into fragments (Scheme 1d). The DLS results of the NPs before and after degradation are shown in Figure 5b; we can see a big decrease of the NPs’ size after degradation in addition to the degraded products in molecular size. The fluorescence of N-Gal NPs in methanol before and after DTT treatment was measured, and a comparison of the two fluorescent spectra in Figure 5a displayed a significant fluorescence decrease of the degraded products. The only change of the polymer structure after the degradation is the cleavage of SS to produce SH,19 leading to disassembly of the cross-linked NPs. Thus, the significant decrease of fluorescence should be attributed to disassembly of the cross-linked network; the reason is probably similar to that for the tertiary amines in gas phase: the network can reduce some degree of interaction between the nitrogen atoms in the tertiary amine arising at the orbital level or the state level.14 In contrast to the commonly used fluorescent marker, fluorescent proteins, the quantum yield of N-Gal NPs measured by Williams’ method23 was 7.9% at an excitation of 370 nm. This is nearly equal to the quantum yield of the green fluorescent protein (7.3%) and its blue derivative (7.9%). Although the quantum yield of the N-Gal NPs is lower than that of many organic dyes, these NPs with such a quantum yield are already satisfactory for biological applications. A study on the fluorescence of the NPs displayed some unique properties differing from those of the fluorescent organic dyes.
Figure 4. UVvis (a) and fluorescence spectra (b) of the N-Gal NPs and N-Gal functionalized HPAMAM in 0.2 wt % aqueous solution (solid, N-Gal NPs; dashed, N-Gal HPAMAM). 1527
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Figure 5. (a) Photoluminescence emission spectra of the N-Gal NPs solution in methanol (0.2 wt %) before (top) and after (bottom) being treated with DTT at room temperature (λex = 360 nm). (b) Size distribution of the N-Gal NPs obtained from polymerization with a feed molar ratio of CBA/ AEPZ/N-Gal = 2/1/1 before (b0 ) and after being treated with DTT (a0 ).
Figure 6. Fluorescent properties of the N-Gal NPs. (a) Absorption (Abs) and fluorescence spectra (with progressively longer excitation wavelengths from 320 nm on the left in 20 nm increments) of the NPs in aqueous solution (0.2 wt %) measured at room temperature. (b) Optical fluorescence microscope images of the letter “V” shaped film under excited wavelengths of 330385 (WU), 460490 (WB), and 510550 nm (WG) filters. (c) Photobleaching curves of the NPs and Rhodamine B in aqueous solution.
The NPs can emit multicolor fluorescence when excited at different wavelengths. As shown in Figure 6a, the N-Gal NPs in aqueous solution emitted a broad fluorescent spectrum ranging from 430 to 620 nm, when excited with progressively increasing wavelengths from 320 nm in 20 nm increments. In order to directly observe the variation of color with a change of excitation wavelength, letter “V” films were prepared by writing “V” on the glass slide using the NPs solution and then drying them. As shown in Figure 6b, we can see the blue, green, and red letter “V”s when excited at 330385, 460490, and 510550 nm, respectively. This may be related to the high structural heterogeneity and the broad size distribution of the NPs Photostability is important for many fluorescence-based imaging applications, especially for long-term imaging and tracking experiments. Generally, the organic dyes suffered from low
photobleaching thresholds that limit their effectiveness in longterm imaging.3 The photobleaching kinetics of the N-Gal NPs in Figure 6c appeared to be remarkably photostable, where no obvious decrease in the fluorescent intensity was observed when photobleaching for 1 h. As a comparison, the photobleaching curve of Rhodamine B displayed relatively low photostability (Figure 6c). Differing from aggregation of the fluorescent dyes in a high concentration solution, the N-Gal NPs were stabilized by galactose groups and were homogeneously dispersed in water even in relatively high concentrations. Thus, the dependence of their fluorescence on concentration should be different from that of the fluorescent dyes. As shown in Figure 7b, we can see a linear increase of the fluorescent intensity with the increase of concentration from 0.05 wt % to 0.2 wt %, and the UVvis spectra in 1528
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Figure 7. Dependence of the UVvis spectra (a) and the fluorescence intensity (b) on the concentration of the N-Gal NPs in aqueous solution measured at room temperature (λex = 460 nm).
Figure 8. (A) Laser confocal scanning microscopic images of HepG2 cells after 4 h of incubation with N-Gal NPs. The excitation wavelengths were 543 (a), 488 (b), and 800 (dual-photo excitation) nm (c), respectively. (d) A merged picture of a, b, and c. (B) Specificity of the N-Gal and N-Glu NPs to HepG2 cells. The cells were incubated with the N-Glu NPs at 37 °C for 4 h, 20 (a) and 40 (c), and with the N-Gal NPs, 20 (b) and 40 (d). The concentrations of N-Gal and N-Glu NPs applied in the cell incubation experiments were the same (2 mg/mL).
Figure 7a show the absorbance increase of the N-Gal NPs with their concentration. This is understandable, when every NP is considered as an independent fluorescence or absorption unit, the fluorescence or absorption intensity should be proportional to the total amount of NPs in the solution. Cell Imaging and Selectivity of NPs. Since HepG2 cells can recognize galactose residues through an asialoglycoprotein receptor (ASGP-R) on the surface,24 we incubated HepG2 cells with N-Gal NPs (2 mg/mL) at 37 °C for 4 h, and then the cells were washed with phosphate-buffered saline for removal of the NPs that were not internalized. The cells obtained were observed under laser confocal scanning microscopy. As shown in Figure 8A, when excited at 543, 488, and 800 nm, colocalization of red, green, and blue fluorescence was clearly observed in the
HepG2 cells, indicating potential application of the N-Gal NPs in cell imaging. It is worthy to mention that we can see a bright nucleolus in Figure 8, and the N-Gal NPs did display the ability to effectively bind siRNA (the data were not shown). The investigation of this aspect is in progress. The N-Gal and N-Glu1 NPs have similar chemical structures except for surface groups. They also have similar autofluorescent properties and particle sizes, but Gal has a better specific binding ability to the HepG2 cells relative to Glu.21 To compare the specificity of N-Glu1 and N-Gal NPs to the HepG2, both NPs were separately incubated with HepG2 cells under the same conditions. The fluorescent images in Figure 8B show that the green fluorescence of HepG2 cells incubated with the N-Gal NPs (Figure 8Bb and Bd) is much brighter than that incubated with 1529
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’ ACKNOWLEDGMENT We thank the National Natural Science Foundation of China for financial support under contract Nos. 50673086 and 50633010. ’ REFERENCES
Figure 9. Cytotoxicity of the N-Gal and N-Glu NPs to HepG2 cells. Viability of HepG2 cells treated with PEI25k (2), N-Glu (9), and N-Gal (b) NPs.
N-Glu NPs (Figure 8Ba and Bc), indicating the enhanced cellular uptake of the N-Gal NPs due to strong interaction of the galactosyl moieties with the ASGP-R of HepG2 cells.24 Cytotoxicity of N-Gal and N-Glu NPs. Cytotoxicity is very important for biological applications. We tested the cytotoxicity using an MTT assay. Branched PEI with a molecular weight of 25k (PEI25k) was used as the control. In Figure 9, the PEI25k exhibited high cytotoxicity to HepG2 cells with 50% cell viability at 0.015 mg/mL. In contrast, the cells could tolerate the culture with both NPs at a dose of up to 0.250 mg/mL. These preliminary results demonstrate that the as-prepared PAMAM NP is a suitable candidate for cell imaging, biosensing, and drug delivery.
’ CONCLUSION We devised a simple and feasible strategy for the preparation of fluorescent polymer NPs with multifunctions including biodegradability, fluorescence, and specificity, and the fluorescence was attained due to formation of the tertiary amine in the polymerization because the tertiary amine acting as a fundamental chromophore was well studied in the previous reports. The resulting N-Gal NPs displayed strong photoluminescence, high photostability, broad absorption, and emission (from 430 to 620 nm) spectra. The photoluminescence behaviors including quantum yields can be adjusted by altering the structures of polymer chains and the architectures of polymer materials, which should be further studied in order to clarify the relationship of fluorescence with structures of the polymers. The results from incubation of the liver cancer cells, HepG2, with the NPs showed that they have low cytotoxicity and can be recognized by an asialoglycoprotein receptor on the surface of HepG2 and then can be internalized. Therefore, they have potential applications in bioimaging and drug or gene delivery. The molecular design could be expanded to different functional biopolymers for various applications. Thus, the design of the tertiary amine based polymer materials may represent a new direction in developing fluorescent biomaterials. ’ AUTHOR INFORMATION Corresponding Author
*To whom correspondence should be addressed. E-mail:
[email protected].
(1) (a) Goto, Y.; Matsuno, R.; Konno, T.; Takai, M.; Ishihara, K. Biomacromolecules 2008, 9, 3252–3257. (b) Yu, J.; Wu, C.; Sahu, S. P.; Fernando, L. P.; Szymanski, C.; McNeill, J. J. Am. Chem. Soc. 2009, 131, 18410–18414. (c) Stich, M. I. J.; Schaeferling, M.; Wolfbeis, O. S. Adv. Mater. 2009, 21, 2216–2220. (2) (a) Iwasaki, Y.; Maie, H.; Akiyoshi, K. Biomacromolecules 2007, 8, 3162–3168. (b) Gao, D.; Xu, H.; Philbert, M. A.; Kopelman, R. Angew. Chem., Int. Ed. 2007, 46, 2224–2227. (c) Xia, T.; Kovochich, M.; Liong, M.; Meng, H.; Kabehie, S.; George, S.; Zink, J. I.; Nel, A. E. ACS Nano 2009, 3, 3273–3286. (d) Yang, Z.; Zheng, S.; Harrison, W. J.; Harder, J.; Wen, X.; Gelovani, J. G.; Qiao, A.; Li, C. Biomacromolecules 2007, 8, 3422–3428. (3) (a) Gao, X.; Yang, L.; Petros, J. A.; Marshall, F. F.; Simons, J. W.; Nie, S. Curr. Opin. Biotech. 2005, 16, 63–72. (b) Weng, K. C.; Noble, C. O.; Papahadjopoulos-Sternberg, B.; Chen, F. F.; Drummond, D. C. D.; Kirpotin, B.; Wang, D.; Hom, Y. K.; Hann, B.; Park, J. W. Nano Lett. 2008, 8, 2851–2857. (c) Pecher, J.; Huber, J.; Winterhalder, M.; Zumbusch, A.; Mecking, S. Biomacromolecules 2010, 11, 2776–2780. (4) Lichtman, J. W.; Conchello, J. A. Nat. Methods 2005, 2, 910–919. (5) (a) Bruchez, M., Jr.; Moronne, M.; Gin, P.; Weiss, S.; Alivisatos, A. P. Science 1998, 281, 2013–2016. (b) Smith, A. M.; Nie, S. J. Am. Chem. Soc. 2008, 130, 11278–11279. (c) Gopalakrishnan, G.; Danelon, C.; Izewska, P.; Prummer, M.; Bolinger, P.; Geissbhler, I.; Demurtas, D.; Dubochet, J.; Vogel, H. Angew. Chem. 2006, 118, 5604–5609. (d) Mahendra, S.; Zhu, H.; Colvin, V. L.; Alvarez Environ. Sci. Technol. 2008, 42, 9424–9430. (e) Jamieson, T.; Bakhshi, R.; Petrova, D.; Pocock, R.; Imani, R.; Seifalian, A. M. Biomaterials 2007, 28, 4717–4732. (6) (a) Zheng, Y.; Gao, S.; Ying, J. Y. Adv. Mater. 2007, 19, 376–380. (b) Zhu, L.; Wu, W.; Zhu, M.-Q.; Han, J. J.; Hurst, J. K.; Li, A. D. Q. J. Am. Chem. Soc. 2007, 129, 3524–3526. (c) Huang, C.; Lo, C.; Chen, H.; Hsiue, G. Adv. Funct. Mater. 2007, 17, 2291–2297. (d) Chen, Y.; Thakar, R.; Snee, P. T. J. Am. Chem. Soc. 2008, 130, 3744–3745. (e) Sanghvi, A. B.; Miller, K. P-H.; Belcher, A. M.; Schmidt, C. Nat. Mater. 2005, 4, 496–502. (7) Yang, J.; Zhang, Y.; Gautama, S.; Liu, L.; Dey, J.; Chen, W.; Mason, R. P.; Serrano, C. A.; Schuge, K. A.; Tang, L. Proc. Natl. Acad. Sci. U.S.A. 2009, 106, 10086–10091. (8) (a) Wu, D. C.; Liu, Y.; He, C. B.; Goh, S. H. Macromolecules 2005, 38, 9906–9909. (b) Wang, D.; Imae, T. J. Am. Chem. Soc. 2004, 126, 13204–13205. (c) Wang, D.; Imae, T.; Miki, M. J. Colloid Interface Sci. 2007, 306, 222–227. (d) You, Y. Z.; Yu, Z. Q.; Cui, M. M.; Hong, C. Y. Angew. Chem., Int. Ed. 2010, 49, 1099–1102. (9) Yang, W.; Pan, C. Y. Macromol. Rapid Commun. 2009, 30, 2096–2101. (10) Yang, W.; Pan, C. Y. Biomacromolecules 2010, 11, 1840–1846. (11) Freeman, C. G.; Mcewan, M. J.; Claridge, R. F. C.; Phillips, L. F. Chem. Phys. Lett. 1971, 8, 77–78. (12) Beecroft, R. A.; Davidson, R. S. J. Chem. Soc., Perkin Trans. 2 1985, 1069–1072. (13) (a) Halpern, A.; Wryzykowska, K. J. Photochem. 1981, l5, 147–157. (b) Muto, Y.; Nakato, Y.; Tsubomura, H. Chem. Phys. Lett. 1971, 8, 597–599. (14) Halpern, A. M.; Gartman, T. J. Am. Chem. Soc. 1974, 96, 1393–1398. (15) Cardoza, J. D.; Rudakov, F. M.; Weber, P. M. J. Phys. Chem. A 2008, 112, 10736–10743. (16) Fernando, L. P.; Kandel, P. K.; Yu, J.; McNeill, J.; Ackroyd, P. C.; Christensen, K. A. Biomacromolecules 2010, 11, 2675–2682. 1530
dx.doi.org/10.1021/bm1014816 |Biomacromolecules 2011, 12, 1523–1531
Biomacromolecules
ARTICLE
(17) Kumar, R.; Roy, I.; Ohulchanskyy, T.; Goswami, L.; Bonoiu, A.; Bergey, E.; Tramposch, K.; Maitra, A.; Prasad, P. ACS Nano 2008, 2, 449–456. (18) Wu, D.; Liu, Y.; Jiang, X.; He, C.; Goh, S. H.; Leong, K. W. Biomacromolecules 2006, 7, 1879–1883. (19) (a) Lin, C.; Zhong, Z.; Lok, M. C.; Jiang, X.; Hennink, W. E.; Feijen, J.; Engbersen, J. F. J. Bioconjugate Chem. 2007, 18, 138–145. (b) Hong, C.-Y.; You, Y.-Z.; Wu, D.-C.; Liu, Y.; Pan, C.-Y. J. Am. Chem. Soc. 2007, 129, 5354–5355. (20) (a) Wang, D.; Liu, Y.; Hong, C. Y.; Pan, C. Y. J. Polym. Sci., Part A: Polym. Chem. 2005, 43, 5127–5137. (b) Wang, D.; Liu, Y.; Hu, Z.; Hong, C. Y.; Pan, C. Y. Polymer 2005, 46, 3507–3514. (c) Wang, D.; Zheng, Z. J.; Hong, C. Y.; Liu, Y.; Pan, C. Y. J. Polym. Sci., Part A: Polym. Chem. 2006, 44, 6226–6242. (d) Wang, D.; Liu, Y.; Hong, C. Y.; Pan, C.Y. Polymer 2006, 47, 3799–3806. (21) Brannon-Peppas, L.; Blanchette, J. O. Adv. Drug Delivery Rev. 2004, 56, 1649–1659. (22) Wan, W. M.; Sun, X. L.; Pan, C. Y. Macromolecules 2009, 42, 4950–4952. (23) Williams, A. T. R.; Winfield, S. A.; Miller, J. N. Analyst 1983, 108, 1067–1071. (24) Seymour, L. W. Adv. Drug Delivery Rev. 1994, 14, 89–111.
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