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Phasor–FLIM Analysis to Monitor Intercellular Drug Release from a pH-Sensitive Polymeric Nanocarrier Ting Zhou, Teng Luo, Jun Song, and Junle Qu Anal. Chem., Just Accepted Manuscript • DOI: 10.1021/acs.analchem.7b04511 • Publication Date (Web): 16 Jan 2018 Downloaded from http://pubs.acs.org on January 16, 2018
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Analytical Chemistry
Phasor-FLIM Analysis to Monitor Intercellular Drug Release from a pH-Sensitive Polymeric Nanocarrier Ting Zhou, Teng Luo, Jun Song* and Junle Qu* College of Optoelectronic Engineering, Shenzhen University, Shenzhen 518060, China ABSTRACT:The design of highly efficient drug carriers, and the development of appropriate techniques to monitor their mechanism of action and therapeutic effect, are both critical for improving chemotherapy. Herein, a polymeric nanoparticle, PAHCit/DOX, was synthesized and used as a nano-drug system for the efficient delivery and pH-responsive release of doxorubicin (DOX) into cancer cells. The PAH-Cit/DOX nanoparticles were stable at physiological pH, but effectively released DOX under weakly acidic conditions. The release efficiency was 90.6 % after 60 h dialysis in phosphate-buffered saline at pH 5.5. Confocal images showed the rapid movement of the drug from the cytoplasm to the nucleus, indicating the effective drug release MCF-7 cells. Notably, the combination of fluorescence lifetime imaging microscopy (FLIM) and phasor analysis (Phasor–FLIM) provides an approach to monitor the dynamic change of DOX fluorescence lifetime in intercellular environments. Phasor differentiated lifetime pixel intensity in FLIM images was quantified and used to evaluate the DOX release from nanocarriers, making it possible to detect the dynamics of intracellular release and transport of DOX.
An ideal drug carrier should both improve therapeutic effects and reduce side effects of drugs. The carrier thus needs to deliver therapeutic concentrations of the drug to the target disease site without harming normal tissues. Stimuli-responsive nanocarriers have recently attracted much attention1,2 since they can selectively release drugs at target sites in response to specific stimuli, such as pH3,4, light5, redox potential6,7 and enzymatic activity8,9. Since there are pH differences between healthy and diseased tissues, pH-responsive drug carriers have been especially widely investigated. At the tissue level, diseases such as tumors and inflammation usually result in an acidic microenvironment in the disease tissues10,11 and, at a cellular level, there are acidic intracellular compartments such as late endosomes and lysosomes12,13. Consequently, pHsensitive drug carriers can take advantage of these specific pH differences to release the transported drug into the target tissues or cells. Appropriate techniques for determining the intracellular dynamic distribution of drugs is essential for evaluating the release efficiency of drug carriers and understanding their mode of action14. Conventional intensity-based microscopic techniques, such as fluorescence imaging and spectral imaging, are of limited value since the intensity or spectral response does not differ significantly between the free drug and the drugpolymer carrier system, making their differentiation inside cells difficult15,16. In addition, the sensitivity of intensity-based techniques is also limited in the intracellular environment, which means that slight changes in drug distribution are not detectable. Some newer methods for intracellular monitoring of drug release also have shortcomings; for example, surface enhanced Raman scattering (SERS) is limited to metal-based nanocarriers.17,18 Fluorescence resonance energy transfer (FRET) needs the nanocarriers and drugs with specific fluorescence characteristics.19-20 Fluorescence lifetime imaging microscopy (FLIM) is a noninvasive imaging technique that can be used to monitor spatial variations of fluorescence lifetimes from fluorescent molecules. The fluorescence lifetime depends not on the concentration and fluorescence intensity but depends instead on the surrounding microenvironment of the fluorescent molecules.2123 FLIM can not only acquire information about the fluorescent molecules but also reflects changes in microenvironment.
Consequently, changes in the fluorescence lifetime of drugs can indicate changes of the microenvironment around drugs, reflecting the intercellular drug release and transport.24-27 FLIM is thus an ideal tool for tracking the dynamics of intracellular drug distribution, especially for drugs with intrinsic fluorescence. However, quantitative analysis of drug release efficiency and distribution still cannot be achieved using FLIM alone. The phasor approach to lifetime has been used for the analysis of the FLIM images, due to its ability of separating pixels having different fluorescence lifetimes.28,29 Phasor analysis transfers the fluorescence lifetime data in a graphical form, negating the need for exponential fitting to the fluorescence decay. The approach can classify lifetime pixels in a FLIM image and separate areas of image with different lifetimes, has been used for separation of diseased or healthy tissues.30-33 Based on fluorescence lifetime difference exhibited by native doxorubicin (DOX) compared to conjugated DOX on nanocarriers, the combination of fluorescence lifetime imaging microscopy (FLIM) and phasor analysis (Phasor–FLIM) was successfully used to monitor the intercellular DOX release from nanocarriers and its accumulation in the cell nuclei. 34-35 In this study, we sought to obtain the relative quantitative analysis of drugs in different subcellular compartments with Phasor–FLIM, to further reflect the dynamic process of the intercellular drug release and movement. In this work, the anti-cancer drug, DOX was covalently loaded onto a pH-sensitive polymer (poly(allylamine)citraconic anhydride, PAH-Cit) and then self-assembled to form micellar nanoparticles. The nanoparticles are highly sensitive to pH and are rapidly hydrolyzed in weakly acidic environments. PAH-Cit is ideal for drug delivery since it has good biocompatibility and the DOX loaded nanoparticles are able to selectively release drugs to target sites, leading to high therapeutic effects with low side effects. Once PAH-Cit/DOX nanoparticles are internalized into cancer cells, the acidic pH in endosomes and lysosomes will trigger DOX releasing from the nanoparticles and the drug can then enter the nucleus to achieve its therapeutic effect (Scheme 1). Phasor–FLIM was used to monitor the release and subcellular distribution of DOX in cancer cells. The phasor analysis separated the subcellular compartments in FLIM images by different lifetimes. The phasor differentiated lifetime pixel intensity (PDLPI) in
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different subcellular compartments was quantified and first used to evaluate the release and dynamic movement of DOX. EXPERIMENATL SECTION Materials and characterization. Citraconic anhydride (Cit) and succinic anhydride (Sit) were purchased from Tokyo Chemical Industry Co.,Ltd. (Tokyo, Japan). Doxorubicin hydrochloride (DOX), poly(allyamine hydrochloride) (PAH, MW~15 kDa), Poly(D, L-lactide-co-glycolide) (PLGA, lactide:glycolide 50:50, MW~25 kDa), N-hydroxysuccinimide (NHS) and 1-ethyl-3-(3dimethylaminopropyl)carbodiimidehydrochloride (EDC) werepurchased from Sigma-Aldrich Co (St. Louis, MO, USA. CCK-8 cell counting kits were purchased from Dojindo Laboratories (Kumamoto, Japan). 4′, 6-Diamidino-2phenylindole (DAPI) was purchased from Thermo Fisher Scientific (Grand Island, NY, USA). All other chemicals were analytical grade and were used without further purification. High-purity deionized water (resistance >18 MΩ·cm) was used throughout. Visible absorption spectra were recorded using a Lambda35 UV-Vis spectrophotometer (Perkin-Elmer Inc., USA). Fluorescence spectra were recorded using an LS-55 fluorescence spectrophotometer (Perkin-Elmer Inc. USA), with an excitation wavelength of 490 nm. The morphologies of the nanoparticles were observed by transmission electron microscopy (TEM) using a JEM-2100 transmission electron microscope (JEOL, Tokyo, Japan), with an accelerating voltage of 100 kV. The average size and zeta potential of the nanoparticles were measured using a ZetaPlus particle sizer and zeta potential analyzer (Brookhaven Instruments Corporation, Holtsville, NY, USA) and a Malvern NanoSight NS300 Instrument (Malvern Instruments Ltd, Worcestershire, UK). Synthesis of the nanoparticles. The pH sensitive polymer PAH-Cit was synthesized as we have previously described36. 500 mg PAH was dissolved in 10 mL of 1M NaOH. 2 mL citraconic anhydride was then added dropwise to the PAH solution, with addition of 6 M NaOH during the reaction to keep the pH above 8.0. After overnight reaction, the resultant mixture was dialyzed through a 5 kDa dialysis chamber to remove excess reagents. DOX (20 mg) was added to an ‘asprepared’ solution containing PAH-Cit (20 mg). EDC (1M, 500 µL) and NHS (1M, 500 µL) were then added successively to the resultant solution to catalyze attachment of DOX to PAH-Cit. 6 M NaOH was added to maintain the pH of the mixture above 7.4. After 24 h, unbound DOX was removed by dialysis through a 5 kDa dialysis chamber. The DOX loading efficiency (%) on PAH-Cit is defined as (weight of initially added DOX - weight of DOX in dialysis solution) / weight of initially added DOX × 100. The concentration of dialyzed DOX was determined by measuring fluorescence at 593 nm, using a calibration curve prepared under the same conditions. Dichloromethane and 3% PVA aqueous solution was then added to the PAH-Cit/DOX solution and the mixture was emulsified for 15 min using an ultrasonicator (Branson Instruments Co., Ltd., Stamford, CT, USA), operated at 450 W. Volatile solvent was evaporated by continuous magnetic stirring overnight. The resultant PAH-Cit/DOX nanoparticles were collected and washed by centrifugation at 13,000 rpm for 10 min and the precipitated PAH-Cit/DOX was redispersed in PBS.
PAH-Sit/DOX nanoparticles were prepared in the same way as PAH-Cit/DOX, using succinic anhydride instead of citraconic anhydride. PLGA/DOX nanoparticles were prepared using the emulsification-solvent evaporation technique37,38 with modification. 10 mg of DOX neutralized with triethylamine were dissolved in 10 mL of. The mixture was then added dropwise to 3% aqueous PVA (20 mL) containing 100 mg of PLGA. The mixture was emulsified for 10 min using an ultrasonicator operated at 450 dichloromethane W. Volatile solvent was removed by magnetic stirring overnight and the product was isolated by centrifugation at 13,000 rpm for 10 min. The resultant PLGA/DOX nanoparticles were redispersed in PBS. In vitro drug release. PAH-Cit/DOX and PAH-Sit/DOX solution (2 mg/mL) were placed separately into 5 kDa dialysis chambers and dialyzed in 50 mL buffers with different pH values (pH 5.5, 6.8 or 7.4). The dialysis was carried out by stirring inside an incubator shaker at 37 °C and the solutions were protected from light. Drug release was assumed to start as soon as the dialysis chambers were placed into the buffer reservoirs. At 10 min, 20 min, 30 min, 45 min, 1 h, 2 h, 4 h, 6 h, 12 h, 18 h, 24 h, 36 h, 48 h and 60 h, aliquots (100 µL) of the solutions in the release reservoirs were taken out for characterization. The concentration of released DOX was determined by measuring fluorescence at 593 nm, using a calibration curve prepared under the same conditions. Cell culture. MCF-7 human breast cancer cells were cultured in Dulbecco’s modified Eagle’s medium (DMEM), supplemented with 10% fetal bovine serum and 1% penicillin– streptomycin, at 37 °C in a humidified incubator under an atmosphere containing 5% CO2. Determination of cytotoxicity using CCK-8 assay. MCF7 cells were seeded in 96-well plates at 1×104 cells per well and incubated with various concentrations of PAH-Cit, and free DOX, PAH-Cit/DOX, PAH-Sit/DOX or PLGA/DOX at a DOX concentration of 5 µg/mL. After incubation for 48 h, CCK-8 was added to each well and the cells were incubated at 37 °C for 1 h under an atmosphere containing 5% CO2. The absorbance of each well at 450 nm was then measured using a Varioskan Flashplate reader (Thermo Scientific). Results are expressed as the mean percentage of cell viability relative to untreated cells. IC50 values for free DOX and PAH-Cit/DOX in MCF-7 cells were measured by incubating free DOX and PAHCit/DOX (at DOX-equivalent concentrations of 0, 0.065, 0.25, 1, 4, 16 µg/mL) with the cells for 48 h. Confocal laser scanning microscopy (CLSM). MCF-7 cells seeded in dishes were treated with free DOX, PAHCit/DOX or PAH-Sit/DOX at concentrations equivalent to a DOX concentration of 5 µg/mL. At 0.5, 3, 6 and 20 h, the cells were washed with PBS and fixed with 4% paraformaldehyde. DAPI was then used to stain the nuclei and images were captured using a TCS SP2 confocal laser scanning microscope (Leica Microsystems, Wezlar, Germany), equipped with an objective (63× PL APO CS, NA = 1.4, Leica Microsystems). A mode-locked titanium (Ti): sapphire laser (Mira 900, Coherent Inc., Santa Clara, CA, USA), operating at 76 MHz and 120 fs, was tuned to a wavelength of 780 nm for two-photon excitation of DAPI (emission λ=420-480 nm). DOX fluorescence was excited using a 488 nm argon laser and the emission was collected from 550 nm to 620 nm.
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Analytical Chemistry Fluorescence lifetime imaging microscopy (FLIM). MCF-7 cells seeded in dishes were treated with free DOX, PAH-Cit/DOX or PAH-Sit/DOX, with DOX concentrations of 5 µg/mL. At 0.5, 3, 6 and 20 h, the cells were washed with PBS and images were collected using a time-resolved fluorescence measurement system, which incorporated the confocal microscope and a time-correlated single photon counting (TCSPC) module. For the time-resolved experiment, a cutting short-pass filter (700 nm, Chroma Technology Corp., Bellows Falls, VT, USA) was used to block the reflected laser. The fluorescence signal was passed through a band-pass filter (550±30 nm, Chroma Technology Corp.) and the fluorescence lifetime was determined using a micro-channel plate photomultiplier tube (MCP-PMT, Hamamatsu) mounted on the optional port of the confocal microscope and connected to a single photon counting module (SPC150, Becker & Hickl GmbH, Berlin, Germany). The average fluorescence lifetime (τm) at each pixel of a 256×256 image was calculated for all stained tissue sections, with bi-exponential components fitting expressed as: I(t)/I(0) = a1 exp( -t/τ1 ) +a2 exp( -t/τ2 ) (1) where τ1, τ2 and α1, α2 denote the lifetime and amplitude, respectively, of two different components. A distribution of the average lifetime, weighted to pixel intensity, was determined for each image. The average lifetime was derived according to the following equation: τm= (a1 τ1 +a2 τ2 )/(a1 +a2 ) (2) Pseudo-colored lifetime images were generated by assigning a color to the value of τm at each pixel. The minimum time channel width of the TCSPC module was 813 fs and the response time of the whole system was less than 30 ps. Lifetime calculations and fitting were performed using SPCImage software (Becker & Hickl GmbH). In fluorescence mode, epifluorescence illumination was focused via a 63×objective and fluorescence passing through a long-pass filter (425 nm, Leica Microsystems) was collected by the same digital camera (DFC310 FX CCD, Leica Microsystems) used for bright field imaging. Phasor approach to FLIM data analysis. The phasor-FLIM analysis was done using SPCImage 6.4. The fluorescence collected from each pixel of the image was transformed to the Fourier space and a phasor plot, i.e., a graphical representation of intensity decays for a FLIM image, was constructed. Points in the two-dimensional phasor plot are defined by the values of sine (S) and cosine (G) transforms derived using the following equations:
plot).All possible single exponential lifetimes lie on the “universal circle”, defined as the semicircle going from point (0, 0) to point (1, 0), with radius 1/2. Point (1, 0) corresponds to τ=0, and point (0, 0) to τ=∞. In the phasor coordinates, the single lifetime components add directly because the phasor follows vector algebra. A mixture of two distinct single lifetime components, each of which lies separately on the single lifetime semicircle, does not lie on the semicircle. Clusters of pixel values are detected in specific regions of the phasor plot. The cluster assignment is performed not only by taking into account similar fluorescence properties in the phasor plot but also by exploiting the spatial distribution and localization in cellular substructures. RESULTS AND DISCUSSION Synthesis and Characterization of PAH-Cit/DOX nanoparticles. PAH is a cationic polymer that has been widely used in bionanotechnology because it can be used for electrostatic layer-by-layer assemblies39,40 and has low toxicity and good biocompatibility41. PAH reacts with a number of citraconic anhydride molecules to form a hydrophilic polymer,
Scheme 1. Schematic illustration of formation of drug-loaded nanostructure and intracellular drug release triggered by acidic pH environments. Phasor-FLIM analysis was used to monitor the drug release process.
∞
si,j ω=
0 It sinnωt dt ∞
0 Itdt
3
∞
gi,j ω=
0 It cosnωt dt ∞
0 Itdt
(4)
Where the indices i and j represent pixels of the image and si,j(ω) and gi,j(ω) are the y and x coordinates, respectively, of the phasor plot. ω = 2πf, where f is the laser repetition frequency (i.e., 76 MHz in our experiments) and n is the harmonic frequency. The fluorescence collected from each pixel of an image was thus transformed to a point in the phasor plot. Analysis of the phasor distribution was performed by cluster identification. There is a direct relationship between a phasor location and lifetime. Every possible lifetime can be mapped onto this universal representation of the decay (phasor
Figure 1. Characterization of PAH-Cit/DOX nanoparticles. Morphology and size of PAH-Cit/DOX nanoparticles measured by (A) TEM and (B) Dynamic Scattering Light analysis. (C) Absorbance spectra and (D) fluorescence spectra of PAHCit/DOX. Scale bar for TEM is 200 nm.
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PAH-Cit, containing pH-sensitive cis-aconityl groups. As shown in Scheme 1, after DOX being covalently loaded onto PAH-Cit (with a loading efficiency of 92.6%), the hydrophilic polymer then becomes amphiphilic and self-assembles into nanostructures in the form of spherical micelles. The morphology of the PAH-Cit/DOX nanoparticles, determined by TEM, is shown in Figure 1A. The average particle size was around 120 nm (Figure 1B, Figure S1). PAH-Cit/DOX had similar absorbance and fluorescence characteristics to those of free DOX (Figure 1C and D). It is thus difficult to differentiate PAH-Cit/DOX from free DOX by fluorescence imaging or spectral imaging. pH-dependent DOX releasing. The nanostructures were stable at neutral pH (Figure 2A) and, at pH 7.4, less than 10% of DOX was released from the nanostructures after 60 h of dialysis with stirring. Under acidic conditions, DOX was released from the nanostructures much faster. At pH 6.8, 37.8% of DOX was released after 60 h of dialysis. At pH 5.5, 90.6% of DOX was released, demonstrating that the delivery and controlled release of DOX from the nanostructures is highly pH dependent. This is because the cis-aconityl linker connecting PAH-Cit with DOX is very acid-labile (Figure S2) and hydrolysis can take place even under very weakly acidic conditions.
7 cells, even at a high concentration of 0.5 mg/mL (Figure 3A), demonstrating that the polymer is biocompatible and suitable for drug delivery. PAH-Cit/DOX, however, significantly decreased the viability of MCF-7 cells. PAH-Cit/DOX showed markedly higher cytotoxicity than the other polymeric nanodrugs, PAH-Sit/DOX and PLGA/DOX (Figure 3B). PAH-Sit/DOX has a similar structure to PAH-Cit/DOX, but it cannot effectively release DOX in a weakly acidic environment. PLGA nanoparticles are widely used for drug delivery since they are biocompatible and biodegradable through natural pathways.44,45 However, PLGA cannot selectively release drugs at target sites and the release rate for DOX is relatively low compared with PAH-Cit/DOX.46-48 When PAH-Cit/DOX is internalized into cancer cells, it can rapidly release DOX, which then translocates to the nucleus and causes cell death. Viability of MCF-7 cells was reduced to a similar extent by PAH-Cit/DOX and free DOX at the DOX-equivalent concentrations (IC50 values of 1.54 and 1.14 µg/mL, respectively, Figure 3C).
Figure 2. DOX releasing efficiency of (A) PAH-Cit/DOX and (B) PAH-Sit/DOX in PBS at pH 7.5 and pH 6.8 and in acetate buffer at pH 5.5. The inset images show buffer after dialysis of PAH-Cit/DOX and PAH-Sit/DOX for 60 h. To confirm that the pH sensitivity of PAH-Cit/DOX is conferred by the cis-aconityl group, PAH-Sit/DOX nanoparticles were synthesized for comparison. PAH-Sit/DOX, which has a very similar chemical structure to that of PAH-Cit/DOX, except that it does not contain cis-aconityl groups, is stable and does not release DOX under weakly acidic conditions (Figure S3). Less than 10% of DOX was released from PAH-Sit/DOX after 60 h dialysis in buffer at pH 7.4, 6.8 or 5.5 (Figure 2B). The release of DOX from PAH-Sit/DOX also showed no pHdependency under these conditions since the efficiency of DOX release at pH 6.8 was higher than that at either pH 7.4 or pH 5.5.These results confirm that the ability of PAH-Cit/DOX to rapidly release DOX in a weakly acidic environment is attributable to the high pH sensitivity of the cis-aconityl groups42,43. Cytotoxicity studies of PAH-Cit/DOX naoparticles. The side effects of anti-tumor drugs are a major problem in chemotherapy and an ideal way to reduce side effects is to deliver the drugs for release at the target tumor site using drug carriers. To avoid bringing side effects, the drug carriers themselves should have good biocompatibility and low cytotoxicity. To evaluate the biocompatibility of PAH-Cit, MCF-7 cells were incubated with various concentrations of this pH-sensitive polymer. After 48 h, a CCK-8 kit was used to measure cell viability. PAH-Cit did not show obvious cytotoxicity to MCF-
Figure 3. (A) Viability of MCF-7 cells after incubation for 48 h with PAH-Cit (0, 5, 10, 50, 100 and 500 µg/mL). (B) Viability of MCF-7 cells after incubation for 48 h with PAHCit/DOX, free DOX, PAH-Sit/DOX and PLGA/DOX at a DOX-equivalent concentration of 5 µg/mL. ***P < 0.001. (C) Viability of MCF-7 cells after incubation for 48 h with PAHCit/DOX and free DOX (at DOX-equivalent concentrations of 0, 0.065, 0.25, 1, 4, 16 µg/mL). IC50 values were 1.54 and 1.14 µg/mL. Error bars represent the relative standard deviations calculated from 10 wells. In vitro studies of controlled drug release from PAHCit/DOX nanoparticles with fluorescence imaging. DOX is used clinically as an anti-cancer drug and works by entering the nucleus where it intercalates with DNA and prevents DNA replication.49,50 The ability to kill cancer cells thus depends on
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Analytical Chemistry how much DOX can be released from the drug carrier, which determines how much DOX is available to be transported from the cytoplasm to the nucleus. To verify that PAH-Cit/DOX nanoparticles can effectively release DOX after internalization into the cytoplasm, the nanoparticles were incubated with MCF-7 cells and DOX distribution was measured at various time points. Free DOX and PAH-Sit/DOX were used as controls. Confocal images (Figure S6) showed that, after 24 h treatment, PAH-Sit/DOX was internalized into MCF-7 cells but DOX was mainly located in the cytoplasm. In contrast, DOX carried by PAH-Cit/DOX was mainly located in the nucleus. This is consistent with free DOX, which can rapidly move into the nucleus (Figure S7), and demonstrates that DOX can enter the nucleus of cancer cells after delivery by the pH-sensitive drug carrier, PAH-Cit/DOX nanoparticles .The controlled release of DOX from PAH-Cit/DOX nanoparticles in cancer cells is illustrated in Figure 4. DOX was internalized
times can be selected in the phasor plot and the FLIM image is separated and painted accordingly. As shown in the FLIM images (Figure 5B), after incubating PAH-Cit/DOX nanoparticles with MCF-7 cells for 6 h, some of the DOX had already moved into the nucleus, where its lifetime is shorter than that in the cytoplasm. After the Phasor-FLIM analysis, the fluorescence lifetimes of DOX were separated into four segments in the phasor plot. (Figure 5D). Each segment was indicated by a colored, rectangular cursor, and the corresponding FLIM images (1, 2, 3 and 4) were colored red, yellow, green and blue according to the chosen cursors (Figure 5C). It was shown that the four parts of FLIM images mainly located in cell membrane, cytoplasm, nucleus membrane and nucleus respectively. And the average lifetimes in these four parts were 4.46, 3.16, 2.34 and 1.52 ns (Figure 5E). The results meant that DOX had a similar fluorescence lifetime in one of these cellular compartments. When coming to another compartment, its lifetime changed. The lifetime difference in these compartments might be attributed to the change of physicochemical environments and the drug release from the nanocarrier. Free DOX has a fluorescence lifetime about 1 ns. When DOX is encapsulated in the polymeric nanocarriers, its lifetime becomes longer.35,51 After PAH-Cit/DOX being internalized into MCF-7 cells, DOX was gradually released from the nanocarrier and the average lifetime decreased. When DOX got to the nucleus, it has the shortest fluorescence lifetime.
Figure 4. Confocal images of MCF-7 cells after incubation with PAH-Cit/DOX for 30 min, 3, 6 and 20 h. Scale bar is 20 µm. by PAH-Cit/DOX into MCF-7 cells after 30 min and the fluorescence of DOX then gradually moved from the cytoplasm to the nucleus. After 3 h, there was more DOX fluorescence in the cytoplasm than in the nucleus. After 6 h, DOX fluorescence in the nucleus had increased and, after 20 h, DOX fluorescence was mainly concentrated in the nucleus. These results indicate that PAH-Cit/DOX nanoparticles can deliver DOX into the cytoplasm of cancer cells and then effectively release the drug, which translocates to the nucleus and kills the cells. Monitor of intercellular drug release and distribution with Phasor-FLIM. In order to further quantitatively analyze the releasing efficiency and transport of DOX, Phasor-FLIM was used to study the PAH-Cit/DOX treated MCF-7 cells. The fluorescence lifetime of a fluorophore (DOX in this case) is sensitive to physicochemical factors such as pH changes, protein binding and interactions with the drug carrier. After internalization of DOX into cells, changes in its fluorescence lifetime can, therefore, indicate changes in the subcellular microenvironment of DOX, reflecting release and transport of the drug. In the Phasor-FLIM approach, the analysis of the FLIM images is performed by detecting clusters of pixel values in specific regions of the phasor plot. Pixels having similar life-
Figure 5. Phasor-FLIM analysis of MCF-7 cells after incubation with PAH-Cit/DOX for 6 h. (A) Fluorescence image, (B) FLIM image and (C) Phasor separated and pseudo-colored FLIM images of MCF-7 cells. Scale bar is 20 µm. (D) Phasor plot of FLIM images. With phasor analysis, MCF-7 cells were separated into cell membrane, cytoplasm, nucleus membrane and nucleus based on the different fluorescence lifetime of DOX in these four areas. (E) The average lifetime of DOX in these four areas were 4.46, 3.16, 2.34 and 1.52 ns. Error bars indicate means ± standard deviation. (n=5) Phasor-FLIM analysis was further used to image and analyze the cells treated with PAH-Cit/DOX nanoparticles at various time points. In the FLIM images, it was difficult to tell how the DOX lifetime changed with time (Figure 6A2). After phasor analysis, the cell membrane, cytoplasm, nucleus membrane and nucleus were separated for their distinctively different lifetimes, and colored as in Figure 5 (Figure 6A3). Then the according phasor differentiated lifetime pixel intensity (PDLPI) which was attributed by the DOX with a similar fluorescence lifetime, was quantified in each compartment. PDLPI
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was analyzed as the average pixel intensity in 5 random areas in each kind of compartment from different cells. The lifetime distribution histogram (Figure 6B) showed that DOX had two main fluorescence lifetimes in MCF-7 cells. One was in cytoplasm about 3.2 nm and another was in nucleus around 1.7 nm. From 30 min to 20 h, the pixel intensity in the cytoplasm (longer fluorescence lifetime) decreased and that in the nucleus (shorter fluorescence lifetime) increased. After 20 h, most of DOX had the shorter lifetime, indicating that most drugs had already been released and moved from the nanocarriers and had moved into the nucleus. PDLPI in cytoplasm and nucleus is in consistent with the results (Figure 6C). At 30 min, PDLPI in cytoplasm was about 3 times of PDLPI in nucleus.
PAH-Cit/DOX less. Thus its lifetime decreased and became closer to that in nucleus membrane with time. CONCLUTION In conclusion, a pH-sensitive polymer was synthesized and covalently loaded with DOX. The loaded polymer selfassembled to form nanoparticles, PAH-Cit/DOX, which acted as nanocarriers for efficient delivery and controlled release of DOX to cancer cells. The polymer itself had good biocompatibility and very low cytotoxicity but, after loading with DOX, it markedly reduced cell viability. The PAH-Cit/DOX nanoparticles efficiently released DOX in a weakly acidic environment and, after internalization into MCF-7 cells, the nanoparticles rapidly released DOX, which was then transported from the cytoplasm to the nucleus. Phasor-FLIM analysis was successfully separated subcellular compartments with DOX under a different releasing rates, based on the difference of DOX fluorescence lifetime. PDLPI quantification in different subcellular compartments was used to evaluate the releasing and intercellular distribution of DOX. Combining FLIM and phasor plot analysis allowed the dynamic analysis of DOX releasing and transport at different time points, confirmed that PAHCit/DOX nanoparticles were able to release DOX in a highly efficient manner in cancer cells.
ASSOCIATED CONTENT Supporting Information
Figure 6. Phasor-FLIM analysis of MCF-7 cells after incubation with PAH-Cit/DOX for 30 min, 3 h, 6 h and 20 h. (A1) Fluorescence images (A2) FLIM images and (A3) Phasor separated and pseudo-colored FLIM images of MCF-7 cells. Scale bar is 20 µm. (B) Fluorescence lifetime histogram of DOX at different time points. (C) The change of PDLPI in cytoplasm and nucleus with time. Error bars indicate means ± standard deviation (n=5). *P < 0.05, ****P < 0.0001; ordinary one-way ANOVA. (D) The change of fluorescence lifetimes in Phasor-FLIM separated four areas. Error bars indicate means ± standard deviation (n=8). *P < 0.05; NS, not significant. PDLPI in cytoplasm increased from 30 min to 3 h, and then started to decrease. While PDPI in nucleus kept increasing with time. At 20 h, PDLPI in cytoplasm was only about half of PDLPI in nucleus. These results indicated that DOX was delivered into MCF-7 cells with PAH-Cit/DOX nanoparticles and then gradually accumulated into cytoplasm. When it got to the cytoplasm, the acidic environments made the DOX began to release from the polymeric nanocarriers and then free DOX moved to nucleus. Therefore, free DOX kept accumulating in the nucleus. In cytoplasm, DOX accumulated first and then decreased. In addition, in these four compartments, that their average lifetime changed with time also reflects the dynamic release of DOX from the nanocarriers (Figure 6D). The form of DOX was steady in cell membrane, nucleus membrane and nucleus, so the lifetime of DOX did not change regularly with time. When DOX came to the cell membrane, it was not released and manly loaded on nanocarriers. And it got to nucleus only as free DOX. While in cytoplasm, DOX was gradually released from nanocarriers. Free DOX became more and
Nanoparticle tracking analysis of PAH-Cit/DOX nanoparticles. Chemical structural formula of PAH-Cit/DOX and PAH-Sit/DOX and their stability under weakly acidic conditions. Zeta potentials of PAH-Cit and PAH-Cit/DOX. 1H NMR (nuclear magnetic resonance) spectra of PAH-Cit. Confocal images of MCF-7 cells after incubation with PAH-Cit/DOX and PAH-Sit/DOX. Confocal images of MCF-7 cells after incubation with free DOX. PhasorFLIM analysis of MCF-7 cells after incubation with free DOX.
AUTHOR INFORMATION Corresponding Author
* E-mail:
[email protected],
[email protected] Notes
The authors declare no competing financial interest.
ACKNOWLEDGMENT The authors gratefully acknowledge financial support by the National Basic Research Program of China (2015CB352005); the National Natural Science Foundation of China (61605124, 61620106016, 61525503, 61378091, 61405123); the Guangdong Natural Science Foundation Innovation Team (2014A030312008); Hong Kong, Macao, and Taiwan cooperation innovation platform & major projects of international cooperation in Colleges and Universities in Guangdong Province (2015KGJHZ002); Shenzhen Basic Research Project (JCYJ20170412110212234, JCYJ20150324141711561, JCYJ20150930104948169, JCYJ20160328144746940, JCYJ20160308093035903); and the China Postdoctoral Fund (Grant NO. 2016M602512).
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