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Cite This: ACS Appl. Mater. Interfaces XXXX, XXX, XXX−XXX

Multiscale Imaging of Brown Adipose Tissue in Living Mice/Rats with Fluorescent Polymer Dots Yixiao Guo,† Yao Li,† Yidian Yang,†,‡ Shiyi Tang,† Yufan Zhang,† and Liqin Xiong*,† †

Shanghai Med-X Engineering Center for Medical Equipment and Technology, School of Biomedical Engineering, Shanghai Jiao Tong University, Shanghai 200030, P. R. China ‡ The Key Laboratory of Resource Chemistry of Ministry of Education, Shanghai Key Laboratory of Rare Earth Functional Materials, and Shanghai Municipal Education Committee Key Laboratory of Molecular Imaging Probes and Sensors, Shanghai Normal University, Shanghai 200234, P. R. China S Supporting Information *

ABSTRACT: Brown adipose tissue (BAT) has been identified as a promising target for the treatment of obesity, diabetes, and relevant metabolism disorders because of the adaptive thermogenesis ability of this tissue. Visualizing BAT may provide an essential tool for pathology study, drug screening, and efficacy evaluation. Owing to limitations of current nuclear and magnetic resonance imaging approaches for BAT detection, fluorescence imaging has advantages in large-scale preclinical research on small animals. Here, fast BAT imaging in mice is conducted based on polymer dots as fluorescent probes. As early as 5 min after the intravenous injection of polymer dots, extensive fluorescence is detected in the interscapular BAT and axillar BAT. In addition, axillar and inguinal white adipose tissues (WAT) are recognized. The real-time in vivo behavior of polymer dots in rodents is monitored using the probe-based confocal laser endomicroscopy imaging, and the preferred accumulation in BAT over WAT is confirmed by histological assays. Moreover, the whole study is conducted without a low temperature or pharmaceutical stimulation. The imaging efficacy is verified at the cellular, histological, and whole-body levels, and the present results indicate that fluorescent polymer dots may be a promising tool for the visualization of BAT in living subjects. KEYWORDS: near-infrared fluorescence imaging, probe-based confocal laser endomicroscopy, conjugated polymers, brown adipose tissue, uncoupling protein-1, multi-scale, in vivo

1. INTRODUCTION Brown adipose tissue (BAT), as a special tissue that involves in adaptive thermogenesis, has attracted great attention in biomedical research.1−5 Medical studies have reported that BAT mass is significantly correlated with body mass index and other metabolic disorder parameters such as obesity and diabetes.8−15 BAT shows subcellular structures that are unique from white adipocyte [white adipose tissues (WAT]; it is rich in mitochondria equipped with ladder-shaped dense cristae, reflecting its high capacity for oxidative phosphorylation.1−3 It also contains abundant multilocular lipid droplets, which are located close to mitochondria, making the process of oxidative phosphorylation highly efficient. The thermogenesis process mainly depends on the high expression of uncoupling protein-1 (UCP-1). In addition, BAT is highly vascularized, which facilitates efficient heat dissipation to the blood and thus realizes body temperature maintenance. BAT is mainly distributed in specific body areas, including the interscapular region [interscapular BAT (iBAT)], axillar region, cervical region, mediastinum, and paravertebral region. The total BAT is up to approximately 5% of the human newborn or other small mammals, but in adults, the quantities are much smaller. BAT-related research has evolved from an underestimated to a rapid developing field in the past 3 decades.6−18 Dozens of or © XXXX American Chemical Society

even hundreds of small animals are often needed for screening drugs that modulate BAT metabolism and identifying genes or pathways that are associated with BAT development and regulation in preclinical research. It is of great importance to develop an imaging approach that realizes fast BAT identification to adapt the high throughput studies. However, fast imaging for whole body BAT in small animals still remains challenging, which hinders the clinical and preclinical studies.19−24 Currently, the most widely applied method for BAT localization is positron emission tomography (PET) imaging using 18F-FDG.25,26 It requires pretreatment such as cold stimulation or norepinephrine treatment to activate the metabolism of BAT for probe uptake. Magnetic resonance imaging (MRI) and computed tomography (CT) have also been used for BAT imaging27,28 in BAT-related studies. However, these modalities are suboptimal for high-throughput preclinical studies because of the expensive instrumentation, long scanning time, pretreatment limitations, and lowthroughput protocols. Received: April 15, 2018 Accepted: May 31, 2018

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DOI: 10.1021/acsami.8b06094 ACS Appl. Mater. Interfaces XXXX, XXX, XXX−XXX

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ACS Applied Materials & Interfaces In comparison, fluorescence imaging is radical-free and available for high-throughput imaging for small animals because of its short scanning time and relatively low cost. Briefly, fluorescence imaging is based on the detection of signals from exogenous fluorochromes excited with lasers or lamps. The emitted photons are subsequently detected by a charge coupled device with high quantum efficiency.20 Therefore, developing fluorescence probes for BAT localization may provide a promising tool for preclinical and clinical research. However, few studies have examined whole-body BAT imaging of living subjects using fluorescence imaging.29−35 Bradley Smith et al. reported a micellar formulation of fluorescent dye SRFluor680 for iBAT imaging in living mice at 3 h postinjection.35 Kolonin et al. pioneered a peptide conjugated with a fluorescent tag for BAT-targeted imaging at 1 h postinjection.29 Yudasaka et al. reported a type of polymer-modified single-walled carbon nanotube for whole BAT imaging in living mice with the maximum photoluminescence (PL) signal reached at 1 h postinjection.31 These reports demonstrated promising imaging efficacy for BAT using various optical probes at least 1 h postinjection. Furthermore, the imaging capacity and efficiency could be improved. The aim of this study is to develop an effective approach to realize the fast imaging of whole body BAT in living subjects. In recent years, fluorescent polymer dots, especially nearinfrared (NIR) polymer dots, have emerged as attractive fluorescence imaging nanoprobes in living animals because of their excellent properties, including bright fluorescence intensity, excellent photostability, high emission rate, and low cytotoxicity.36−49 NIR fluorescence in the window of 650−800 nm has relatively deep tissue penetration and minimizes autofluorescence, which makes it promising for subcutaneous adipocyte identification. These findings suggested that the NIR polymer dots are suitable for biomedical imaging purposes. To date, no studies based on the polymer dots for BAT imaging have been reported to the best of our knowledge. Herein, for the first time, we report the results of the fast imaging of BAT in living mice/rats using a series of NIR polymer dots. We conducted the experiments and analysis in a multiscale and longitudinal manner. We tested the in vitro affinity between polymer dots and the mouse adipose tissue-derived stem cells (mADSCs) and carried out the comparative in vivo imaging study and ex vivo analysis. At microscopic resolution, we observed the real-time accumulation and retention of polymer dots in the BAT-related areas using probe-based confocal laser endomicroscopy (pCLE) imaging. The imaging results were also confirmed by a series of histological assays.

2.2. Preparation of NIR Polymer Dots. The synthesis of polymer dots was through a coprecipitation method as in our previous reports. In a typical procedure, a solution of 2 mL tetrahydrofuran(THF) containing MEH-PPV or PFBT, PS-PEG-COOH or MMA-NH2, NIR775, and PE-FA was prepared as a stock solution. After 30 s of ultrasonication in a water bath, the mixture was quickly dispersed into 10 mL of Milli-Q water under vigorous sonication using an ultrasonic cell crusher for 1 min (10% power). Then, the extra THF was evaporated at 45 °C under the protection of nitrogen (Hannuo Instrument, China). Finally, the polymer dots were filtered with a polyvinylidene fluoride membrane (0.45 μm). Specific masses and ratios of different polymer dot synthesis methods are shown as Table S1. For further application, different concentrations of the NIR polymer dots were adjusted using a centrifugal filtration device (Amicon Ultra-15 Centrifugal Filter with a MW cutoff of 100 kDa). 2.3. Characterization of the NIR Polymer Dots. The morphologies of the polymer dots were determined using a 120 kV HITACHI JEM-2100F transmission electron microscopy (TEM) system. TEM samples were prepared by dripping the polymer dot solution onto carbon-supported copper grids and then drying at room temperature before observation. The hydrodynamic size and zeta potential of the polymer dots were measured using a dynamic light scattering (DLS) instrument (Malvern, UK). UV−vis absorption spectra were collected via a UV-2550 ultraviolet−vis spectrometer (Shimadu, China). Fluorescence emission spectra were collected on an ELISA molecular system (SpectraMax i3x). The fluorescence quantum yield (QY) of the NIR polymer dots was measured with a UV−NIR absolute PL QY spectrometer using an integrating sphere (Hamamatsu, Japan) with 509 nm excitation for MEH-PPV polymer dots and 467 nm excitation for PFBT polymer dots from a xenon lamp. In the physical stability test, four different polymer dots were dispersed in phosphate-buffered saline (PBS, 1×) and Dulbecco’s modified Eagle’s medium (DMEM) with 10% fetal bovine serum (FBS) cell culture media at 37 °C for 24 h. Then, the particle size variance was measured with the DLS, and the fluorescence spectra were collected to detect the fluorescence stability. In the blood circulation half-time assay, ∼50 μg of polymer dots was injected intravenously into the tail vain of ICR mice (∼20 g). At various time points postinjection, 5 μL of blood was collected from the tail into 100 μL of PBS. The fluorescence intensity of the blood samples at 775 nm was measured using the ELISA molecular system (SpectraMax i3x). 2.4. Cell Culture, Staining, and Confocal Laser Scanning Microscopy (CLSM) Imaging. mADSCs were isolated and cultured according to the standard methods in a previous study.69 NCI-H292 cells were purchased from the Cell Bank of Chinese Academy of Science (Shanghai China). The mADSCs were grown in a low-glucose DMEM, and NIC-H292 cells were cultured in PRMI 1640 medium. All the media were supplemented with 10% FBS and 1% penicillin/ streptomycin. Cells were maintained at 37 °C under a humidified atmosphere containing 5% CO 2 . In the log growth phase, approximately 106 mADSCs were seeded in a glass-bottom dish (NEST) and treated with ∼10 μg of MEH-PPV-COOH or PFBTCOOH without NIR775 doping. After 24 h incubation, cells were observed using a CLSM system (Leica, Germany). Channel settings were as follows: MEH-PPV (excitation: 488 nm, emission: 560−630 nm), PFBT (excitation: 458 nm, emission: 500−560 nm), 4′,6diamidino-2-phenylindole (DAPI; excitation: 405 nm, emission: 450− 520 nm). In the cell staining assay, MEH-PPV-COOH polymer dottreated cells were washed twice with PBS and incubated with LysoTracker Green FM for 20 min (50 nM, excitation: 458 nm, emission: 500−520 nm) and then stained with MitoTracker Deep Red for 10 min (100 nM, excitation: 633 nm, emission: 655−675 nm). After that, cells were fixed with 4% paraformaldehyde(PFA) for 10 min and stained with DAPI solution for 5 min. The MEH-PPV channel was excited with 561 nm to avoid cross color with LysoTracker Green FM. PFBT-COOH polymer dot-treated cells were incubated with LysoTracker Red for 20 min (50 nM, excitation: 561 nm, emission: 585−620 nm). Follow-up procedures were in accordance with the MEH-PPV-COOH group.

2. MATERIALS AND METHODS 2.1. Materials. Poly(phenylene vinylene) derivative poly[2methoxy-5-(2-ethylhexyloxy)-1,4-phenylenevinylene] [MEH-PPV; molecular weight (MW): 150 000−250 000 Da] was acquired from J&K, Inc. Poly(9,9-dioctylfluorene-alt-benzothiadiazole) (PFBT, average M n : 10k to 20k), silicon 2,3-naphthalocyanine bis(trihexylsilyloxide) (NIR775), and Evans Blue dye were purchased from Sigma Aldrich, Inc. Polystyrene (PS) graft ethylene oxide functionalized with PS and carboxylic end group (PS-PEG-COOH) and amino-terminated poly(methyl methacrylate) (MMA-NH2) were purchased from Polymer Source Inc. (Scheme S1). Folate Cap PE (PE-FA) was purchased from AVanti, Polar Lipids, Inc. MitoTracker Deep Red FM and LysoTracker Green FM were purchased from Thermo Fisher Scientific, Inc. (USA). LysoTracker Red was purchased from Beyotime, Inc. (China). All other chemicals were of analytical grade and used without purification. B


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Scheme 1. Schematic Diagram of This Study; (a) Chemical Structures of MEH-PPV, PFBT, PS-PEG-COOH, NIR775, and PFFA; (b) Representative Diagram Depicting the Preparation of NIR-Emitting PFBT Polymer Dots via the Coprecipitation Method; (c) Physiological Process Illustration of the BAT Imaging; after Intravenous Injection, the Prepared Polymer Dots Show a Fast-Accumulating Retention in the iBAT, Axillar Adipocytes, and Inguinal Adipocytes through Blood Transport

2.5. Flow Cytometry Assay. mADSCs were seeded into a six-well plate (1 × 106 cells per well). Then, 20 μg of MEH-PPV-COOH polymer dots with or without PE-FA was added for 24 h. After washing with PBS three times and centrifuging (1000 rpm, 4 min), cells were resuspended in PBS and analyzed with a flow cytometry Accuri C6 (BD, USA). The fluorescence intensity plot was acquired and processed using the Accuri C6 software. 2.6. In Vivo and ex Vivo Fluorescence Imaging and Region of Interest (ROI) Analysis of BALB/c Mice. Animal procedures were reviewed and approved by the Institutional Animal Care Use Committee of Shanghai Jiao Tong University. BALB/c nude mice were divided into four groups (n = 5), and each mouse was anesthetized with 200 μL of PBS solution of pentobarbital sodium by intraperitoneal injection (0.1 mg/kg). After that, each mouse was intravenously treated with 20 μg of polymer dots (∼100 μL) and immediately imaged using an IVIS Lumina K imaging system (PerkinElmer, Inc.). Excitation and emission settings are as in Table S2. The imaging was recorded at 5 and 30 min followed by dosing with the polymer dots [Binning: 8, field-of-view (FOV): 10 cm × 10 cm]. Mice were euthanized by cervical dislocation, and the dorsal skin was removed prior to imaging to expose the interscapular adipose tissue. Subsequently, the fluorescence emitting interscapular adipose tissue was isolated from the mouse and imaged. Finally, the interscapular, axilla, and inguinal adipose depots and major organs (heart, liver, spleen, lung, and kidney) were isolated from the corresponding mouse and imaged with the same settings. ROI analysis was carried out on the Living Image software (PerkinElmer, Inc.). Signal-to-skin analysis of interscapular adipose depots circled two regions. ROI-1 represented the interscapular adipose depots, and ROI-2 represented the skin of the hind limb as a baseline control. Then, the corresponding ROI value (radiant efficiency) was measured based on the average intensity mode. The signal-to-skin ratio represents the radiant efficiency ratio of ROI-1/ROI-2. 2.7. pCLE Imaging of Adipocytes in Living BALB/c Nude Mice and Sprague Dawley (SD) Rats. Experiments were conducted using a Cellvizio Dual Band system (Mauna Kea Technologies, Paris,

France). In this study, a ProFlex S1500 scanning probe was used (diameter = 1.5 mm, spatial resolution = 3.3 μm, λexcitation = 488 and/ or 660 nm), and dual-wavelength channels are equipped with two detection units for the spectral range λdetection = 505−700 and 680−900 nm, respectively. Each nude mouse (two groups, n = 3) was anesthetized with 200 μL of PBS solution of pentobarbital sodium (1%) by intraperitoneal injection and then intravenously treated with 80 μg of MEH-PPV-COOH-30 or PFBT-COOH-30 polymer dots without NIR775. After 25 min, 40 μL of Evans Blue solution (0.5%) was injected into the mice via the tail vain. Approximately 5 min later, the laser scanning probe was inserted into the incised interscapular, axillar, and inguinal adipocytes. Real time video sequences were recorded (12 frames per second), while having direct contact between probe and the tissues. Polymer dots were excited at 488 nm, and the emission fluorescence was collected from 505 to 700 nm. Evans Blue was excited at 660 nm, and the emission fluorescence was collected from 680 to 900 nm. pCLE video sequences and images were processed offline using the matching software (IC viewer, Mauna Kea Technologies, Paris, France). The protocol for SD rats (approximately 150 g each) was in accordance with that of the nude mice, except that all the doses increased 7.5-fold according to the body weight ratio. 2.8. Histology Analysis of Adipose Tissues. Adipose tissues were fixed with PBS buffer containing 4% PFA. Haemotoxylin and eosin (H&E) staining was carried out on paraffin-embedded sections. UCP-1 immunohistochemical staining was performed after activating the tissue antigen of paraffin sections with ethylenediaminetetraacetic acid solution (diluted with water) in a steamer for 1 h. Bovine serum albumin (BSA) (10%) was added to the slices after washing with PBS. Then, samples were incubated with the primary antibody (1:4000) for 18 h at 4 °C (Abcam Plc., Catalogue no. ab10983). A horseradish peroxidase-conjugated goat anti-rabbit IgG polyclonal (1:50) antibody was used as the secondary antibody (Abcam Plc., Catalogue no. ab97051). The observation was carried out on a microscope (Olympus IX71). For immunofluorescence staining, 10 μm-thick frozen sections were obtained using a freezing microtome (Leica CM1950). Then, frozen sections were fixed with 4% PFA for 10 min, followed by C


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Figure 1. Characterization of the fluorescent polymer dots. Color labels of different polymer dots are as follows: black for MEH-PPV-COOH-30, blue for MEH-PPV-COOH-50, red for MEH-PPV-NH2-30, and green for PFBT-COOH-30. (a) Sample images of MEH-PPV-COOH and PFBTCOOH polymer dots under room light (left) and UV light (center) before NIR775 doping. The fluorescence images (right) show NIR fluorescence emission of NIR775-doped MEH-PPV-COOH and PFBT-COOH polymer dots. (b) Hydrodynamic diameter of four different polymer dots. (c) Zeta potential of four different polymer dots. (d−g) TEM images of the four polymer dots: MEH-PPV-COOH-30 (d), MEH-PPV-COOH-50 (e), MEH-PPV-NH2-30 (f), and PFBT-COOH-30 (g). Scale bar: 20 nm. (h) UV−vis spectra of four different polymer dots. (i) Fluorescence spectra of four different polymer dots. (j) Particle size variance of four different polymer dots after treatment with PBS or DMEM containing 10% FBS for 24 h at 37 °C. Solid lines represent the PBS dispersant and dashed lines represent the 10% FBS dispersant. (k) Fluorescence stability of four polymer dots in PBS or DMEM containing 10% FBS for 24 h at 37 °C. Solid lines represent the PBS dispersant and dashed lines represent the 10% FBS dispersant. (l) The fluorescence signals of blood samples of mice injected with polymer dots (approximately 50 μg) from 2 min to 6 h. Data represent mean ± standard deviation (n = 3). washing with 0.3% Triton X-100 (10 min) and BSA (1 h). The primary antibody was applied upon dilution with PBS (1:300) for 12 h at 4 °C. Primary antibodies used were rabbit anti-UCP1 (Abcam Plc., Catalogue no. ab23841) and goat anti-m/r CD31 (RD, Inc., AF3268). Then, the secondary antibody was applied to the slides (1:500) for 2 h at room temperature. Secondary antibodies used were donkey antirabbit IgG H&L (Abcam Plc., Cambridge, UK, Catalogue no. ab150067) and donkey anti-goat IgG H&L (Invitrogen, Alexa 594). Nuclei were stained with DAPI for 5 min. After washing with PBS, slides were mounted and then observed using confocal microscopy. Channel settings were as follows: DAPI (excitation: 405 nm, emission: 460−520 nm), MEH-PPV (excitation: 488 nm, emission: 560−630 nm), PFBT (ion: 458 nm, emission: 480−560 nm), UCP-1 (excitation: 633 nm, emission: 653−700 nm), and CD31 (excitation: 594 nm, emission: 604−635 nm).

penetration for in vivo imaging. The amphiphilic polymer encapsulation agent, PS-PEG-COOH or MMA-NH2, was introduced to coat the NIR polymer dots with a biocompatible shell and to prevent possible leakage of the NIR775 from the polymer dots. As shown in Figure 1a, both MEH-PPV polymer dots and PFBT polymer dots emit bright fluorescence under UV light without NIR775 doping, and the radiant efficiency in the NIR region at 780 nm determined by imaging system can reach up to approximately 109 to 1010

(

p / sec /cm 2 / sr μ W / cm 2

) with

NIR775 doping (∼20 μg/mL). To examine, compare, and optimize the imaging capacity of BAT, we synthesized four types of NIR polymer dots. The hydrodynamic diameter of the polymer dots was determined by DLS as follows (Figure 1b,c): (1) MEH-PPV-COOH-30: MEH-PPV polymer dots coated with PS-PEG-COOH had a hydrodynamic diameter of 30 ± 3 nm mean polydispersity index (PDI) of 0.147 and a zeta potential of −30 ± 2 mV. (2) MEH-PPV-COOH-50: MEH-PPV polymer dots coated with PS-PEG-COOH had a hydrodynamic diameter of 50 ± 2 nm, a PDI of 0.134, and a zeta potential of −34 ± 4 mV. (3) MEHPPV-NH2-30: MEH-PPV polymer dots coated with MMA-NH2

3. RESULTS 3.1. Preparation and Characterization of the Polymer Dots. A coprecipitation method was applied to synthesize the NIR polymer dots (Scheme 1a,b). Fluorescent conjugated polymer, MEH-PPV or PFBT, as the fluorescence resonance energy transfer (FRET) donor was encapsulated into the nanoparticle core. An NIR dye, NIR775, was doped into the dots as the FRET acceptor to give the advantage of high tissue D


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ACS Applied Materials & Interfaces Table 1. Fluorescence QY of the Fluorescent Polymer Dots with or without NIR775 Doping

a

polymer dots

MEH-PPV/PFBT emissiona (NIR775 undoped)

MEH-PPV/PFBT emissiona (NIR775 doped)

NIR775 emissiona (NIR775 doped)

MEH-PPV-COOH-30 MEH-PPV-COOH-50 MEH-NH2-COOH-30 PFBT-COOH-30

0.085 0.076 0.075 0.166

0.045 0.033 0.024 0.050

0.074 0.064 0.079 0.060

All the MEH-PPV polymer dots were excited at 509 nm and PFBT polymer dots were excited at 467 nm.

had a hydrodynamic diameter of 30 ± 2 nm, a PDI of 0.172, and a zeta potential of −26 ± 2 mV. (4) MEH-PPV-COOH30: PFBT polymer dots coated with PS-PEG-COOH had a hydrodynamic diameter of 30 ± 5 nm, PDI of 0.140, and zeta potential = −36 ± 4 mV. DLS results showed the good dispersity and colloidal stability of polymer dots in water. The size distribution and morphologies were further determined by TEM. TEM measurements showed that the NIR polymer dots possessed a spherical morphology with ultrasmall size (Figure 1d−g). MEH-PPV-COOH-30, MEH-PPV-COOH-50, and MEH-PPV-NH2-30 had a mean diameter of approximately 2.6, 2.8, and 1.6 nm, respectively, and the mean diameter of PFBT-COOH-30 was approximately 3.0 nm. There exists an approximately 20 nm increase in the hydrodynamic size for MEH-PPV-COOH-30 and MEH-PPV-COOH-50. It is worth noting that all the TEM average diameters of the four types of polymer dots are less than 5 nm. Compared with TEM data, the increase in size determined by DLS may be attributed to the hydration corona formed by the PS-PEG-COOH or MMANH2 polymer chains around NIR polymer dots. Similar results have also been reported in a study of superparamagnetic iron oxide nanoparticles for T1-weighted MRI.50 MEH-PPV polymer dots exhibited a broad UV−vis absorbance spectrum and with a maximum at 510 nm and a weak NIR peak at 772 nm (Figure 1h). PFBT-COOH-30 polymer dots exhibited a maximum absorbance peak at 468 nm and a weak peak at 772 nm. The intensity increase of the MEHPPV-COOH-50 was ascribed to the original amount of MEHPPV during the synthesis (Table S1). Under excitation at 500 nm, MEH-PPV polymer dots exhibited its distinctive characteristic emission peak at 592 nm and a narrow-band NIR peak at 778 nm, which belongs to NIR775 (Figure 1i). Under excitation at 450 nm, PFBT showed its distinctive characteristic emission peak at 538 nm and an NIR peak at 778 nm. Different intensities at 778 nm in the normalized spectra indicate different FRET efficiencies between the donor (MEH-PPV or PFBT) and the acceptor, NIR775. As shown in Table 1, a fluorescence QY test clearly reflected the light-harvesting efficiency and energy transfer offered by polymer dots. After NIR775 doping, QY values of the MEH-PPV (FRET donor) emission decreased 47−68% and PFBT emission decreased 70%. Comparatively, the QY of the doped NIR775 emission under excitation at 509 or 467 nm maintained at 0.064−0.079, which was fairly close to the 0.07 value of free NIR775 excited at 763 nm and determined in THF.45 The results indicated the efficient energy transfer from the donor (MEH-PPV or PFBT) to the accepter (NIR775), and the doped NIR775 was free of aggregation. If the aggregation occurred, a greatly reduced QY would have been noted. We further tested the physical stability of these polymer dots by dispersing them in PBS and DMEM with 10% FBS cell culture media at 37 °C, respectively. Over 24 h, these polymer dots showed no distinct size variance (Figure 1j). As shown in Figure 1k, the NIR fluorescence intensities of polymer dots

treated with PBS maintained approximately 90% of their original intensities, and in the DMEM with 10% FBS group, the polymer dots all maintained over 80% of their original intensities, suggesting that these polymer dots have good physical stability. We further tested the blood half-life time of the polymer dots in mice (Figure 1l). The circulation time was measured to be 29.0 min for MEH-PPV-COOH-30, 30.6 min for MEH-PPV-COOH-50, 28.5 for MEH-PPV-NH2-30, and 31.3 min for PFBT-COOH-30. The results implied that the PSPEG-COOH coating prolonged the blood circulation time of polymer dots to avoid quick filtration by the reticuloendothelial system (RES). 3.2. mADSC Staining, Imaging, and Flow Cytometry Assay. To test the cellular uptake of the polymer dots, mADSCs were treated with MEH-PPV-COOH and PFBTCOOH polymer dots and incubated for 24 h (Figure 2a). Under confocal microscopy, strong fluorescence was obtained from both MEH-PPV- and PFBT-treated cells, indicating that these polymer dots have intrinsic affinity to mADSCs and can be internalized into the cytoplasm. The cell staining assay showed that the polymer dots in the cells were more inclined to localize in the lysosome than in the mitochondria (Figure 2b). In comparison with adipose stem cells, we also stained lung tumor cells (NCI-H292) to examine and compare their subcellular distribution tendency. The results revealed that the fluorescence of polymer dots was more similar to that of LysoTracker than MitoTracker, indicating that the polymer dots may be internalized into lysosomes via the endocytosis pathway in both stem cells and tumor cells. We also found that PE-FA modification greatly increased the cell labeling efficiency of the polymer dots. As shown in the flow cytometry plot (Figure 2c,d), approximately 99.3% of the cells were labeled by PE-FA functionalized MEH-PPC-COOH polymer dots, while only approximately 16.9% of the cells were labeled by MEHPPV-COOH polymer dots without PE-FA. 3.3. In Vivo Fluorescence Imaging of brown Adipocytes in Mice. We subsequently tested the imaging capacity of these polymer dots for whole body BAT localization (Figure 3). Four groups of nude mice (n = 5) were anesthetized and intravenously injected with 20 μg of polymer dots. Each animal was imaged at 5 and 30 min after dosing. As shown in Figure 3a, as early as 5 min after intravenous treatment with MEH-PPV-COOH-30, strong fluorescence selectively accumulated in the interscapular, axillar, and inguinal regions, which coincided with the adipocyte distribution. The uptake of polymer dots in the axillar and inguinal regions was stronger than in the liver. For 30 min, fluorescence signals in the interscapular, axillar, and inguinal areas as well as the liver grew stronger, and the fluorescence in the BAT-related areas was still more intense than in the liver. Moreover, recognizable fluorescence signals were observed in the paravertebral area of mice, although the intensity was relatively low compared with the interscapular area (Figure S1). According to the literature, there exists little BAT in the paravertebral area in the E


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Figure 3. Fluorescence imaging of BAT in BALB/c nude mice at 5 and 30 min after an intravenous 20 μg dose of MEH-PPV-COOH-30 (a), MEH-PPV-COOH-50 (b), MEH-PPV-NH2-30 (c), and PFBTCOOH-30 (d). (e) ROI analysis of the iBAT-to-skin fluorescence intensity ratio of mice (n = 3) treated with four different polymer dots and blank control. (*, p < 0.05 when compared with the blank control group using Student’s t test).

Figure 2. Cell imaging and flow cytometry assay. (a) CLSM imaging of mADSCs incubated with 10 μg of MEH-PPV-COOH polymer dots or PFBT-COOH polymer dots for 24 h. (b) mADSCs and NCI-H292 cells costained with DAPI (blue), LysoTracker (magenta), and MitoTracker (green). (c) Flow cytometry plot of mADSC treated with MEH-PPV-COOH polymer dots with and without functionalization of PE-FA. (d) Cell labeling efficiency calculated from the flow cytometry results in (c).

relatively low fluorescence was also noted in the inguinal area. However, the axillar adipocytes could not be clearly identified, and the liver signal was intense. The results indicated that PSPEG-COOH-coated polymer dots have better BAT-targeting ability than MMA-NH2-coated polymer dots. In addition, we compared another fluorescent polymer, PFBT, to examine its imaging performance (Figure 3d). The imaging pattern showed that PFBT-COOH-30 produced selective localization in the interscapular, paravertebral, and inguinal depots. Comparatively weak signals were observed in the neck and axillar depot as well. However, the fluorescence of PFBT-COOH-30 in the liver and head skin was much stronger than that of MEH-PPVCOOH-30. An ROI analysis was carried out to calculate the iBAT-to-skin signal ratio. As shown in Figure 3e, the values in four polymer dot-treated groups showed different degrees of increase (higher than 2) compared with the blank group (1.47). The ratio value in the MEH-PPV-COOH-30 group reached 3.07 at 5 min and 2.86 at 30 min postinjection. The PFBT-COOH-30 group showed similar levels: 3.21 and 2.48, respectively. The ratio values of the two groups were approximately 2-fold higher than the blank mice, indicating that MEH-PPV-COOH-30 and

mammalian species.1 In addition, Yudasaka, et al. also reported that the paravertebral BAT could be imaged using single-walled carbon nanotubes.31 At 5 and 30 min postinjection, the MEH-PPV-COOH-50 polymer dots showed limited selective accumulation in the interscapular depots, and a strong liver signal was collected from the dorsal position (Figure 3b). In the face-up position, most signals also accumulated in the liver, and no distinct localization was observed in the axillar and inguinal depots. The results demonstrated that smaller-sized MEH-PPV-COOH-30 has a better BAT targeting ability compared with larger-sized MEH-PPV-COOH-50, whereas MEH-PPV-COOH-50 was mainly trapped in the liver and captured by the RES. To compare with the COOH-modified polymer dots mentioned above, we tested NH2-modified polymer dots, MEH-PPV-NH2-30 (Figure 3c). The iBAT was localized, and F


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ACS Applied Materials & Interfaces PFBT-COOH-30 polymer dots showed ideal signal-to-skin ratios in the interscapular depot. Comparatively, the MEHPPV-COOH-50 and MEH-PPV-NH2-30 groups showed a lower iBAT-to-skin signal ratio. These results demonstrated that the smaller-sized polymer dots, MEH-PPV-COOH-30 and PFBT-COOH-30, exhibited significant and selective accumulation in whole body BAT-related regions including the interscapular, axillar, and inguinal depots in a fast manner. We further compared the imaging efficacy of MEH-PPVCOOH-30 polymer dots with or without modification of PEFA. We found that PE-FA modified polymer dots showed a distinct retention in BAT at 30 min post i.v. injection (Figure 3a). Comparing with the corresponding control group (Figure S2), the result indicated that these probes were circulated into BAT in short time through blood transport and then could be steadily uptake into adipocytes. 3.4. Ex Vivo Fluorescence Imaging Analysis of Brown Adipocytes. To confirm the in vivo imaging results, we conducted an ex vivo imaging analysis of tissues dissected from the mice 30 min postinjection (Figure 4). Followed by fluorescence imaging, we exposed subcutaneous tissue on the back of the mice. As shown in Figure S4, the triangular-shaped iBAT displayed a natural brown color and little adjacent interscapular WAT (iWAT) was also visualized. Then, the mouse was imaged before and after the removal of the interscapular adipocytes (Figure 4a,b). The fluorescence area was consistent with the natural brown color area, and almost no fluorescence signal was observed on the mouse back after the iBAT/iWAT ablation. In Figure 4b, fluorescence detected in the iBAT was stronger over the adjacent iWAT. ROI analysis showed that the quantitative fluorescence intensity in the iBAT was 2.9-fold higher than that of the iWAT and 3.6-fold higher than that of the femur muscle (Figure S5). Shown in Figure 4c was the fluorescence-emitting adipocyte depots dissected from the mouse including iBAT and axillar and inguinal adipocytes. Strong fluorescence was mainly detected in iBAT and partial axillar adipocytes, where the brown adipose cells are distributed. We also observed the selective distribution of polymer dots in iBAT over iWAT under an inverted fluorescence microscope (Figure S10a). A significant intensity contrast was observed between the iBAT and iWAT, which was in accordance with the ex vivo ROI analysis. To investigate the fluorescence distribution of polymer dots in mice, the major organs, iBAT, iWAT, i.p. WAT (inter peritoneal WAT, taken from abdomen near to the spleen), and femur muscle were removed and imaged (Figure 4d). The polymer dots mainly accumulated within the liver and iBAT over the iWAT and i.p. WAT. The results suggested the selective BAT localization ability of MEH-PPV-COOH-30 and low off-target accumulation in other organs, except for the liver. The liver signal showed a distinct increase over in vivo imaging; this increase may be due to the different locations in depth between the liver and subcutaneous adipocytes. The harvested major organs were further frozen and sliced for confocal imaging (Figure S8a). NIR fluorescence was collected in the liver and spleen, and signals in the heart, lung, kidney, and limb muscle were very weak. 3.5. pCLE Imaging of Adipocytes in Living BALB/c Nude Mice and SD Rats. To gain deeper insight in the BAT at microscopic resolution, we applied the pCLE technique to monitor the in vivo behavior of polymer dots in a real-time and minimally invasive manner. The dot-like fluorescence signal of polymer dots started to distribute in the BAT-related areas at 5

Figure 4. Fluorescence imaging of the nude mouse 30 min post i.v. injection of 20 μg of MEH-PPV-COOH-30 polymer dots. (a,b) Fluorescence imaging of the nude mouse before and after the interscapular adipose tissue was removed. (c) Fluorescence imaging of the iBAT, axilla adipocyte, and inguinal adipocyte. (d) Fluorescence imaging of the major organs, iBAT, iWAT, i.p. WAT, inguinal adipocyte, and muscle tissue from the hind limb. (e) Biodistribution of MEH-PPVCOOH-30 and PFBT-COOH-30 polymer dots in organs taken from the mice (n = 3) that were sacrificed 30 min after intravenous injection. The intensities were calculated by ROI analysis based on the average mode.

min postinjection, and the signal intensity reached a relative peak up to 30 min as shown in Figure 5 (green channel). To provide a complementary contrast of the vascular networks, Evans blue dye was used as a blood vessel imaging agent (red channel). In general, a high heterogeneity of vascular networks inside the BAT was observed. These vasculatures exhibited variable size in length and diameter (∼5−60 μm) and cross-linked morphologies, stressing the necessity to monitor numerous FOVs for analysis. Three representative FOVs were displayed in each area, including iBAT and axillar and inguinal adipocytes. The fluorescence of MEH-PPV-COOH-30 (Figure 5a) and PFBT-COOH-30 (Figure 5b) was strongly detected in the blood vessels, capillaries, and adipocytes. We observed that the polymer dots were circulating along the vessels in adipocytes with some bright particles binding on the blood vessel wall. The G


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(∼150 g) is less than that of the BALB/c nude mice (∼20 g), which is mainly because the BAT distributes more in newborn animals. In the interscapular and axillar depots, a small amount of BAT was found to be covered by a mass of WAT and distributed at a deeper location. After administration of ∼600 μg of MEH-PPV-COOH-30 or PFBT-COOH-30, we monitored the same depots using pCLE. As expected, the circulation and extravasation as well as perfusion of polymer dots were observed in the iBAT and axillar and inguinal adipocytes. The fluorescence perfusion in the axillar and inguinal adipocytes was relatively weaker comparing with that in the iBAT. The results suggested that the prepared polymer dots possess a promising capability of BAT imaging in different rodents in spite of their different BAT masses. Evans blue, as a commonly used fluorescent tracer, visualizes the whole body’s vasculature. However, the small molecule dye leaks from blood vessels into the surrounding tissues within a few minutes after intravenous injection, thus reducing imaging contrast. This phenomenon can be found in some of the FOVs in Figure 5. We also examined whether Evans blue has selectivity to image the vessels in the BAT of mice using the imaging system (Figure S3). The signals of Evans blue were detected all over the mice body and the margins of BAT could not be figured. Ex vivo imaging showed that all the major organs and adipose tissues emitted strong fluorescence, especially the lung. The results suggested that Evans blue visualizes the blood vessel system in mice without distinct specificity. 3.6. Observation of the Adipose Tissue Heterogeneity via H&E and UCP-1 Immunohistochemical Staining. Next, we studied the cell types in fluorescence-emitting adipocytes via H&E and UCP-1 immunohistochemical staining (Figure 6). Most cells in the interscapular depot showed typical brown adipocyte morphologies: a compact cell arrangement with multiple small lipid droplets and high UCP-1 expression. In addition, a small amount of white fat cells was also noted in adjacent areas, which were indicated using blue arrows. Comparatively, the axillar depot showed mixed cell types. In addition to brown adipose cells, some cells with large lipid droplets and negative UCP-1 staining were observed, indicating the white adipose cells, whereas in the inguinal area, white adipose cells were the majority. We also noted that there exist abundant vessels in the iBAT and axillar and inguinal adipocytes. These vessels facilitate efficient heat dissipation from the BAT, and they provide the access of material transport including nanoprobes. Moreover, in inguinal area, the inguinal lymph node (LN) was observed to be surrounded by adipose tissue during the dissection and histology analysis (Figure S7).5 3.7. Polymer Dot Distribution in Adipocytes via Immunofluorescence Staining. To determine whether the polymer dots have selectivity of accumulation in BAT over WAT at a histological level, we analyzed the adipose tissues from polymer dot-injected nude mice using Z-stack confocal imaging (Figure 7). By costaining tissue sections with fluorophore-conjugated antibody labelling the UCP-1 and DAPI labelling nucleus, we observed that UCP-1 expression exhibited a cell type-dependent manner. This result is in accordance with the immunohistochemical results, reflecting that the brown cells are mainly distributed in the interscapular and partial axillar depots, whereas white cells in the inguinal as well as axillar area. Strong fluorescence from PFBT-COOH-30 was detected throughout the iBAT (Figure 7b) and the signals

Figure 5. pCLE imaging of adipocytes in living BALB/c nude mice. Interscapular, axillar, and inguinal adipocytes were imaged with a laser probe 30 min post i.v. injection of 80 μg of MEH-PPV-COOH-30 (left) or PFBT-COOH-30 (right) polymer dots without NIR775 doping. The polymer dots channel (green channel, for both MEHPPV and PFBT) is excited at 488 nm and collected from 505 to 700 nm. The Evans blue channel (red channel, visualizing blood vessels) is excited at 660 nm and collected from 680 to 900 nm.

binding polymer dots sketched out the vessel structure, which colocalized with Evans blue in the merged channel. In the adipose depots, a stronger fluorescence intensity of the polymer dots channel over Evans blue channel was observed, indicating that the polymer dots could not only accumulate but also extravasate effectively from the capillaries and then perfuse into the brown fat tissues. The extravasation and perfusion of polymer dots proceeded steadily so that the intense fluorescence in the BAT filled most of the FOV. At the same time point, we also tested the fluorescence signals in the major organs (such as heart, liver, spleen, lung, and kidney) and limb muscles (Figure S8b) through the inserted probe; strong fluorescence was detected in the liver and spleen and weak signals in the heart, lung, and muscle. The results were in accordance with the ex vivo fluorescence imaging analysis. Some signals were detected in the kidney, indicating that fractional polymer dots may be excreted through renal clearance. Moreover, we utilized the protocol on SD rats to investigate the imaging efficacy in different rodent species (Figure S9). We found that the BAT mass of SD rats over their body mass H


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Figure 6. Histology of bright fluorescence emitting tissues in mice. Adipose tissues were taken from interscapular, axillar, and inguinal areas of MEHPPV-COOH-30 treated mice at 30 min postinjection. Staining: H&E (left) and UCP-1 (right). Small amounts of adjacent white adipose cells in the interscapular and axillar areas are marked by blue arrows.

Accordingly, large-scale studies employing tens or hundreds of animals appear to be cumbersome. Therefore, fast imaging methods of BAT are greatly needed for high-throughput preclinical research. Comparatively, NIR fluorescence imaging is a safe, simple, and cost-effective technique to address these problems. The NIR signal in the window of 650−800 nm where the tissues are optically thin could penetrate the BAT because of its relative shallow location. A fluorescence imaging system scans the whole body within seconds. Currently, optical probe-based imaging of BAT has typically relied on three classes of agents: (1) bare small fluorescent molecules or small fluorescent molecule-doped micelles, (2) fluorescent dye conjugated peptides, and (3) carbon nanotubes.29−31 In this present study, we introduced the fluorescence imaging of BAT based on a series of NIR polymer dots for the first time. These NIR polymer dots displayed unique features for BAT imaging compared with the reported optical agents. Among these polymer dots, MEH-PPV-COOH-30 and PFBT-COOH-30, which have a hydrodynamic size of ∼30 nm, showed the distinct capacity for whole body BAT imaging. iBAT, axillar BAT, and inguinal adipocytes were localized as early as 5 min after i.v. injection. The imaging time is much shorter than with PET, MRI, and other reported methods using optical probes. The reported micelle probe consisting of a small molecule (IR 786 and SRFluor 680) need at least 1 h for the probe to accumulate at the iBAT.30 The reported IRDye 800 conjugated peptide (PEP3) achieved endothelium targeting ability and showed a preferred distribution in the blood vessels of iBAT at 1 h postinjection. In addition, these small molecule probes, micelles, or peptides are limited to image BAT only in the interscapular area rather than whole body BAT imaging. So far, only NIR photoluminescent carbon nanotubes have been reported for whole-body BAT imaging based on an optical method.31 Approximately 1 to 5 h are needed for the maximum probe accumulation, which is much longer than our reported profile based on NIR polymer dots. Comparing with 18F-FDG-

from MEH-PPV-COOH-30 radiated from the blood vessel to the adjacent iBAT, implying that these polymer dots could accumulate in the vessels and then have a significant retention in BAT. In the axillar area, polymer dots and UCP-1 showed a distinct colocalization, which suggested that polymer dots have a selective targeting for BAT over WAT. Comparatively, fluorescence was also noted in the inguinal adipocytes, although UCP-1 expression was negative. According to the literature, some “brown-like adipocytes” which called “beige cells” or “brite cells” are preferentially located in the “segmentable lobule area” of the inguinal fat pad.5 These beige cells can be activated in cold exposure known as the process of “browning”. The expression of UCP-1 would be greatly up-regulated compared with its level at room temperature. This may explain why the UCP-1 was negative in the inguinal areas because all the experiments are conducted at room temperature. Channel separated images with same magnification and scale bars are shown in Figure S11. The CD31 staining showed that fractional fluorescence signals of polymer dots colocalized with the blood vessels and much fluorescence were also detected beyond the blood vessels (Figure 7c). These findings confirmed the results of the in vivo/ex vivo fluorescence imaging, pCLE imaging, and BAT targeting capacity of both MEH-PPV-COOH-30 and PFBT-COOH-30.

4. DISCUSSION Currently, PET/CT and MRI are the most widely used methods for BAT localization. PET imaging using 18F-FDG requires pretreatment such as cold stimulation, which would enhance the metabolic activity for nuclear probe uptake.20 Thus, PET imaging can identify only metabolically active BAT and is prone to false-positive signals from other metabolically active tissues.29 MRI can image BAT by exploiting the water-tofat ratio, but it shows poor tissue signal contrast from surrounding tissues. These modalities are suboptimal for high throughput preclinical studies because they are generally too expensive and need expensive devices and a long scan time. I


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Figure 7. Z-stack confocal immunofluorescence analysis of adipose tissue sections from BALB/c nude mice 30 min after intravenous injection of 80 μg of MEH-PPV-COOH-30 (a) or PFBT-COOH-30 (b). Channel color for either MEH-PPV or PFBT uses red, DAPI uses blue, and UCP-1 uses green to achieve ideal merge contrast. Insets (high magnification) display the polymer dot distribution around the vessels. (c) Immunofluorescence stains for CD31 (red) indicated the colocalization of blood vessels with PFBT-COOH-30 (yellow).

based PET imaging, our imaging experiments were conducted at room temperature and do not require pretreatment such as cold stimulation or noradrenaline treatment to activate BAT metabolism. In addition, fluorescence imaging is radical-free compared with PET, thus avoiding related safety concerns. The possible mechanisms of the selective accumulation of fluorescence probes in BAT have been discussed in the reported studies. Smith et al. conducted the FRET efficiency study to deduce that SRFluor680 could irreversibly translocate into the lipid environment because of the hydrophobic small molecule dye.35 Kataura et al. proposed that the single-walled nanotube may behave like biogenic lipid compounds and interact with apolipoproteins, thus realizing the targeting ability in BAT.31 In this study, we applied pCLE to monitor the transport, extravasation, and retention of the NIR polymer dots in the BAT, and the results were further confirmed by immunofluorescence assay. Using pCLE enables the possibility for real-time imaging of the microvasculature with minimal invasiveness and at microscopic resolution.51,52 We observed the real-time process of the prepared polymer dots’ accumulation in the BAT vasculature and the continuous extravasation out of the vasculature as well as BAT perfusion. Some polymer dots bound on the vessel walls and sketched out the vessel structure. On the basis of this, we generated the

hypothesis that the prepared polymer dots could reach the BAT because of their highly vascularized networks, which were reported in the literature and observed in the frozen slices.1−3 In some research, some proteins of the BAT vasculature were reported to play an important role in BAT browning or other metabolic activities.53−55 The functionalized nanoprobes may have some interactions with these proteins. Further studies are necessary to evaluate the hypothesis. Although the precise mechanism remains elusive, it is worth investing more efforts to develop the NIR polymer dots, considering their excellent performance in BAT imaging. Recently, many studies demonstrated that the lymphatic system plays an essential role in lipid metabolism and excessive adipose tissue may result in dysfunction of lymphatic system.56−68 We also found that inguinal LN was surrounded by the adipocytes during the tissue dissection, ex vivo imaging, and histology analysis (Figure S7). Notable fluorescence signals of polymer dots were detected inside of the LN and its outer membrane (Figure S10b). In the future work, the fluorescence polymer dots may be developed for the study of the adipocytes−lymphatic system interaction and obesity-associated therapy via lymphatic approaches. J


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(5) Barreau, C.; Labit, E.; Guissard, C.; Rouquette, J.; Boizeau, M.-L.; Jeanson, Y.; Berger-Müller, S.; Koumassi, S. G.; Carrière, A.; Casteilla, L. Regionalization of Browning Revealed by Whole Subcutaneous Adipose Tissue Imaging. Obesity 2016, 24, 1081−1089. (6) Chi, J.; Wu, Z.; Choi, C. H. J.; Marchildon, F.; Tessier-lavigne, M.; Cohen, P. Three-Dimensional Adipose Tissue Imaging Reveals Regional Variation in Beige Fat Biogenesis and PRDM16-Dependent Sympathetic Neurite Density. Cell Metab. 2018, 27, 226−236. (7) Jiang, H.; Ding, X.; Cao, Y.; Wang, H.; Zeng, W. Short Article Dense Intra-Adipose Sympathetic Arborizations Are Essential for Cold-Induced Beiging of Mouse White Adipose Tissue. Cell Metab. 2017, 26, 686−692. (8) Xue, Y.; Xu, X.; Zhang, X.-Q.; Farokhzad, O. C.; Langer, R. Preventing Diet-Induced Obesity in Mice by Adipose Tissue Transformation and Angiogenesis Using Targeted Nanoparticles. Proc. Natl. Acad. Sci. U.S.A. 2016, 113, 5552−5557. (9) Sun, L.; Xie, H.; Mori, M. A.; Alexander, R.; Yuan, B.; Hattangadi, S. M.; Liu, Q.; Kahn, C. R.; Lodish, H. F. Mir-193b-365, a Brown Fat Enriched MicroRNA Cluster, is Essential for Brown Fat Differentiation. Nat. Cell Biol. 2012, 13, 958−965. (10) Vickers, S. P.; Jackson, H. C.; Cheetham, S. C. Cardiovascular and Metabolic Disease: Scientific Discoveries and New Therapies, 1st ed.; RSC, 2015; Vol. 45. (11) Nawrocki, A. R.; Scherer, P. E. Keynote Review: The Adipocyte as a Drug Discovery Target. Drug Discovery Today 2005, 18, 1219− 1230. (12) Lin, Y.; Li, X.; Zhang, L.; Zhang, Y.; Zhu, H.; Zhang, Y.; Xi, Z.; Yang, D. Inhaled SiO2 Nanoparticles Blunt Cold-Exposure-Induced WAT-Browning and Metabolism Activation in White and Brown Adipose Tissue. Toxicol. Res. 2016, 5, 1106−1114. (13) Borensztein, M.; Viengchareun, S.; Montarras, D.; Journot, L.; Binart, N.; Lombès, M.; Dandolo, L. Double Myod and Igf2 Inactivation Promotes Brown Adipose Tissue Development by Increasing Prdm16 Expression. FASEB J. 2012, 26, 4584−4591. (14) Won, Y.-W.; Adhikary, P. P.; Lim, K. S.; Kim, H. J.; Kim, J. K.; Kim, Y.-H. Oligopeptide Complex for Targeted Non-viral Gene Delivery to Adipocytes. Nat. Mater. 2014, 13, 1157−1164. (15) Giordano, A.; Frontini, A.; Cinti, S. Convertible Visceral Fat as a Therapeutic Target to Curb Obesity. Nat. Rev. Drug Discovery 2016, 15, 405−424. (16) Sakurai, Y.; Kajimoto, K.; Harashima, H. Biomaterials Science Systems to Tumors and Adipose Tissue Vasculature. Biomater. Sci. 2015, 3, 1253−1265. (17) Harms, M. J.; Ishibashi, J.; Wang, W.; Lim, H.-W.; Goyama, S.; Sato, T.; Kurokawa, M.; Won, K.-J.; Seale, P. Article Prdm16 Is Required for the Maintenance of Brown Adipocyte Identity and Function in Adult Mice. Cell Metab. 2014, 19, 593−604. (18) Urano, T.; Shiraki, M.; Sasaki, N.; Ouchi, Y.; Inoue, S. Largescale Analysis Reveals a Functional Single-nucleotide Polymorphism in the 5’-flanking Region of PRDM16 Gene Associated with Lean Body Mass. Aging Cell 2014, 13, 739−743. (19) Sun, L.; Yan, J.; Sun, L.; Velan, S. S.; Leow, M. K. S. A Synopsis of Brown Adipose Tissue Imaging Modalities for Clinical Research. Diabetes Metab. 2017, 43, 401−410. (20) Marzola, P.; Boschi, F.; Moneta, F.; Sbarbati, A.; Zancanaro, C. Preclinical In Vivo Imaging for Fat Tissue Identification, Quantification, and Functional Characterization. Front. Pharmacol. 2016, 7, 336. (21) Bauwens, M.; Wierts, R.; Van Royen, B. Molecular Imaging of Brown Adipose Tissue in Health and Disease. Eur. J. Nucl. Med. Mol. Imaging 2014, 41, 776−791. (22) Sampath, S. C.; Sampath, S. C.; Cypess, A. M.; Bredella, M. A.; Torriani, M. Imaging of Brown Adipose Tissue: State of the Art. Radiology 2016, 280, 4−19. (23) Berry, R.; Church, C. D.; Gericke, M. T.; Jeffery, E.; Colman, L.; Rodeheffer, M. S. Imaging of Adipose Tissue, 1st ed.; Elsevier Inc., 2014; Vol. 537. (24) Paulus, A.; Van Marken Lichtenbelt, W.; Mottaghy, F. M.; Bauwens, M. Brown Adipose Tissue and Lipid Metabolism Imaging. Methods 2017, 1, 105−113.

5. CONCLUSIONS In summary, the prepared fluorescence polymer dots, termed MEH-PPV-COOH-30 and PFBT-COOH-30, display promising optical properties and have a distinct capacity for wholebody BAT imaging. To evaluate the imaging efficacy, we conducted a study longitudinally at multiple scales from the whole-body level to the tissue level to the cellular level. The iBAT and axillar and inguinal adipocytes of living mice could be localized as early as 5 min postinjection. The real-time in vivo behavior of the polymer dots in BALB/c mice and SD rats was monitored using pCLE imaging at microscopic resolution. The preferred distribution of polymer dots in the BAT over WAT was measured by ROI analysis, and all these results were confirmed by a series of histological analyses. This study may provide a promising tool for generating high-quality images of BAT using a noninvasive optical method. We anticipate that fluorescence imaging combined with other imaging modalities (such as photoacoustic imaging70) can be developed as a new paradigm for BAT-related research.

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ASSOCIATED CONTENT

S Supporting Information *

The Supporting Information is available free of charge on the ACS Publications website at DOI: 10.1021/acsami.8b06094. Chemical structure of MMA-NH2; ingredient table of NIR polymer dot synthesis; excitation and emission filter settings of in vivo fluorescence imaging; in vivo fluorescence imaging; ROI analysis; frozen adipose tissue images of the treated nude mouse; confocal and pCLE imaging; inverted fluorescence microscopy images; and Z-stack confocal immunofluorescence analysis (PDF)

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AUTHOR INFORMATION

Corresponding Author

*E-mail: [email protected]. ORCID

Liqin Xiong: 0000-0001-8611-9691 Notes

The authors declare no competing financial interest.

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ACKNOWLEDGMENTS This study was supported by grants from the National Key R&D Program of China (2016YFC1303100), the National Natural Science Foundation of China (81671738, 81301261, and 21374059), and the Shanghai Pujiang Project (13PJ1405000).We sincerely thank Prof. Zhijun Zhang and Dr. Xiaowen Hu for their offering of the mASDCs. We also appreciate Dr. Kai Li and his team for their professional and exquisite design of the scheme picture.

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REFERENCES

(1) Cannon, B.; Nedergaard, J. Brown Adipose Tissue: Function and Physiological Significance. Physiol. Rev. 2004, 84, 277−359. (2) Lim, S.; Honek, J.; Xue, Y.; Seki, T.; Cao, Z.; Andersson, P.; Yang, X.; Hosaka, K.; Cao, Y. Cold-Induced Activation of Brown Adipose Tissue and Adipose Angiogenesis in Mice. Nat. Protoc. 2012, 7, 606− 615. (3) Kolonin, M. G. How Brown Is Brown Fat That We Can See? Adipocyte 2017, 3, 155−159. (4) Nedergaard, J.; Cannon, B. Perspective The Browning of White Adipose Tissue: Some Burning Issues. Cell Metab. 2014, 20, 396−407. K


Research Article

ACS Applied Materials & Interfaces (25) Yang, J.; Yang, J.; Wang, L.; Moore, A.; Liang, S. H.; Ran, C. Synthesis-Free PET Imaging of Brown Adipose Tissue and TSPO via Combination of Disulfiram and 64CuCl2. Sci. Rep. 2017, 7, 8298−8306. (26) Mirbolooki, M. R.; Constantinescu, C. C.; Pan, M.-L.; Mukherjee, J. Quantitative Assessment of Brown Adipose Tissue Metabolic Activity and Volume Using 18F-FDG PET/CT and β3Adrenergic Receptor Activation. EJNMMI Res. 2011, 1, 30−41. (27) Sophie, C.; Jung, L.; Heine, M.; Freund, B.; Reimer, R.; Koziolek, E. J.; Kaul, M. G.; Kording, F.; Schumacher, U. Quantitative Activity Measurements of Brown Adipose Tissue at 7 T Magnetic Resonance Imaging After Application of Triglyceride-Rich Lipoprotein 59 Fe-Superparamagnetic Iron Oxide Nanoparticle. Invest. Radiol. 2016, 51, 194−202. (28) Branca, R. T.; Mccallister, A.; Yuan, H.; Aghajanian, A.; Faber, J. E.; Weimer, N. Accurate Quantification of Brown Adipose Tissue Mass by Xenon-Enhanced Computed Tomography. Proc. Natl. Acad. Sci. U.S.A. 2017, 115, 174−179. (29) Azhdarinia, A.; Daquinag, A. C.; Tseng, C.; Ghosh, S. C.; Ghosh, P.; Amaya-manzanares, F.; Sevick-muraca, E.; Kolonin, M. G. A Peptide Probe for Targeted Brown Adipose Tissue Imaging. Nat. Commun. 2013, 4, 2472. (30) Nakayama, A.; Bianco, A. C.; Zhang, C.-Y.; Lowell, B. B.; Frangioni, J. V. Quantitation of Brown Adipose Tissue Perfusion in Transgenic Mice Using Near-Infrared Fluorescence Imaging. Mol. Imaging 2003, 2, 37−49. (31) Yudasaka, M.; Yomogida, Y.; Zhang, M.; Tanaka, T.; Nakahara, M.; Kataura, H. Near-Infrared Photoluminescent Carbon Nanotubes for Imaging of Brown Fat. Sci. Rep. 2017, 7, 44760. (32) Dinish, U. S.; Wong, C. L.; Sriram, S.; Ong, W. K.; Balasundaram, G. Diffuse Optical Spectroscopy and Imaging to Detect and Quantify Adipose Tissue Browning. Sci. Rep. 2017, 7, 41357. (33) Zhang, X.; Tian, Y.; Zhang, H.; Kavishwar, A.; Lynes, M. Curcumin Analogues as Selective Fluorescence Imaging Probes for Brown Adipose Tissue and Monitoring Browning. Sci. Rep. 2015, 3, 13116. (34) Zhang, X.; Kuo, C.; Moore, A.; Ran, C.; Elmer, P. Cerenkov Luminescence Imaging of Interscapular Brown Adipose Tissue. J. Visualized Exp. 2014, e51790. (35) Rice, D. R.; White, A. G.; Matthew Leevy, W.; Smith, B. D. Fluorescence Imaging of Interscapular Brown Adipose Tissue in Living Mice. J. Mater. Chem. B 2015, 3, 1979−1989. (36) Zhao, Q.; Zhou, X.; Cao, T.; Zhang, K. Y.; Yang, L. Conjugated Polymer Dots for Hypoxia Bioimaging. Chem. Sci. 2015, 6, 1825− 1831. (37) Sun, T.; Dou, J.-H.; Liu, S.; Wang, X.; Zheng, X.; Wang, Y.; Pei, J.; Xie, Z. Second Near-Infrared Conjugated Polymer Nanoparticles for Photoacoustic Imaging and Photothermal Therapy. ACS Appl. Mater. Interfaces 2018, 10, 7919−7926. (38) Cao, F.; Xiong, L. Folic Acid Functionalized PFBT Fluorescent Polymer Dots for Tumor Imaging. Chin. J. Chem. 2016, 34, 570−575. (39) Xiong, L.; Cao, F.; Cao, X.; Guo, Y.; Zhang, Y.; Cai, X. LongTerm-Stable Near-Infrared Polymer Dots with Ultrasmall Size and Narrow-Band Emission for Imaging Tumor Vasculature in Vivo. Bioconjugate Chem. 2015, 26, 817−821. (40) Xiong, L.; Guo, Y.; Zhang, Y.; Cao, F. Highly Luminescent and Photostable near-Infrared Fluorescent Polymer Dots for Long-Term Tumor Cell Tracking in Vivo. J. Mater. Chem. B 2016, 4, 202−206. (41) Xiong, L.; Shuhendler, A. J.; Rao, J. Self-luminescing BRETFRET Near-infrared Dots for In Vivo Lymph-node Mapping and Tumour Imaging. Nat. Commun. 2012, 3, 1193. (42) Geng, J.; Li, K.; Ding, D.; Zhang, X.; Qin, W.; Liu, J. Lipid-PEGFolate Encapsulated Nanoparticles with Aggregation Induced Emission Characteristics: Cellular Uptake Mechanism and TwoPhoton Fluorescence Imaging. Small 2012, 23, 3655−3663. (43) Jeong, K.; Park, S.; Lee, Y.-D.; Lim, C.-K.; Kim, J.; Chung, B. H.; Kwon, I. C.; Park, C. R.; Kim, S. Conjugated Polymer/Photochromophore Binary Nanococktails: Bistable Photoswitching of NearInfrared Fluorescence for In Vivo Imaging. Adv. Mater. 2013, 25, 5574−5580.

(44) Miao, Q.; Xie, C.; Zhen, X.; Lyu, Y.; Duan, H.; Liu, X.; Jokerst, J. V.; Pu, K. Molecular Afterglow Imaging with Bright, Biodegradable Polymer Nanoparticles. Nat. Biotechnol. 2017, 35, 1102−1110. (45) Jin, Y.; Ye, F.; Zeigler, M.; Wu, C.; Chiu, D. T. Near-Infrared Fluorescent Dye-Doped Semiconducting Polymer Dots. ACS Nano 2011, 5, 1468−1475. (46) Pu, K.; Shuhendler, A. J.; Rao, J. Semiconducting Polymer Nanoprobe for In Vivo Imaging of Reactive Oxygen and Nitrogen Species. Angew. Chem., Int. Ed. Engl. 2013, 52, 10325−10329. (47) Li, J.; Rao, J.; Pu, K. Recent Progress on Semiconducting Polymer Nanoparticles for Molecular Imaging and Cancer Phototherapy. Biomaterials 2018, 155, 217−235. (48) Zhen, X.; Xie, C.; Pu, K. Temperature-Correlated Afterglowofa Semiconducting Polymer Nanococktailfor Imaging-Guided Photothermal Therapy. Angew. Chem., Int. Ed. 2018, 57, 3938−3942. (49) Lyu, Y.; Zeng, J.; Jiang, Y.; Zhen, X.; Wang, T.; Qiu, S.; Lou, X.; Gao, M.; Pu, K. Enhancing Both Biodegradability and Efficacy of Semiconducting Polymer Nanoparticles for Photoacoustic Imaging and Photothermal Therapy. ACS Nano 2018, 12, 1801−1810. (50) Mri, T. Ultra-Large-Scale Production of Ultrasmall Superparamagnetic Iron Oxide Nanoparticles for T1-weighted MRI. RSC Adv. 2016, 6, 22575−22585. (51) Laser, C.; Dietrich, A.; Stewart, J.; Huether, M.; Helm, M.; Schuetze, C.; Schnittler, H.-J.; Jaffray, D. A.; Kunz-schughart, L. A. Macromolecule ExtravasationXenograft Size Matters: A Systematic Study Using Probe-Based. Mol. Imaging Biol. 2013, 15, 693−702. (52) Trottmann, M.; Stepp, H.; Sroka, R.; Heide, M.; Liedl, B.; Reese, S.; Becker, A. J.; Stief, C. G.; Kölle, S. Probe-Based Confocal Laser Endomicroscopy ( pCLE ) − a New Imaging Technique for in Situ Localization of Spermatozoa. J. Biophotonics 2015, 8, 415−421. (53) Rosenwald, M.; Perdikari, A.; Rülicke, T.; Wolfrum, C. BiDirectional Interconversion of Brite and White Adipocytes. Nat. Cell Biol. 2013, 15, 659−667. (54) Sanchez-Gurmaches, J.; Hung, C.-M.; Sparks, C. A.; Tang, Y.; Li, H.; Guertin, D. A. PTEN Loss in the Myf5 Lineage Redistributes Body Fat and Reveals Subsets of White Adipocytes That Arise from Myf5 Precursors. Cell Metab. 2012, 16, 348−362. (55) Ussar, S.; Lee, K. Y.; Dankel, S. N.; Boucher, J.; Haering, M.-F.; Kleinridders, A.; Thomou, T.; Xue, R.; Macotela, Y.; Cypess, A. M. ASC-1, PAT2, and P2RX5 Are Cell Surface Markers for White, Beige, and Brown Adipocytes. Sci. Transl. Med. 2014, 6, 247ra103. (56) Souza-smith, F. M.; Siggins, R. W.; Molina, P. E. Mesenteric Lymphatic-perilymphatic Adipose Crosstalk: Role in Alcohol-induced Perilymphatic Adipose Tissue Inflammation. Alcohol.: Clin. Exp. Res. 2016, 39, 1380−1387. (57) Blum, K. S.; Karaman, S.; Proulx, S. T.; Ochsenbein, A. M.; Luciani, P.; Leroux, J.-C.; Wolfrum, C.; Detmar, M. Chronic High-Fat Diet Impairs Collecting Lymphatic Vessel Function in Mice. PLoS One 2014, 9, e94713. (58) Dixon, J. B. Lymphatic Lipid Transport: Sewer or Subway? Trends Endocrinol. Metab. 2011, 21, 480−487. (59) Escobedo, N.; Oliver, G. The Lymphatic Vasculature: Its Role in Adipose Metabolism and Obesity. Cell Metab. 2017, 26, 598−609. (60) Escobedo, N.; Proulx, S. T.; Karaman, S.; Dillard, M. E.; Johnson, N.; Detmar, M.; Oliver, G. Restoration of Lymphatic Function Rescues Obesity in Prox1-Haploinsufficient Mice. JCI Insight 2016, 1, e85096. (61) Haemmerle, M.; Keller, T.; Egger, G.; Schachner, H.; Steiner, C. W.; Stokic, D.; Neumayer, C.; Brown, M. K.; Kerjaschki, D.; Hantusch, B. Enhanced Lymph Vessel Density, Remodeling, and Inflammation Are Reflected by Gene Expression Signatures in Dermal Lymphatic Endothelial Cells in Type 2 Diabetes. Diabetes 2013, 62, 2509−2529. (62) Kuan, E. L.; Ivanov, S.; Bridenbaugh, E. A.; Victora, G.; Wang, W.; Childs, E. W.; Platt, A. M.; Jakubzick, C. V.; Mason, R. J.; Gashev, A. A. Collecting Lymphatic Vessel Permeability Facilitates Adipose Tissue Inflammation and Distribution of Antigen to Lymph Node − Homing Adipose Tissue Dendritic Cells. J. Immunol. 2015, 194, 5200− 5210. L


Research Article

ACS Applied Materials & Interfaces (63) Rutkowski, J. M.; Davis, K. E.; Scherer, P. E. Mechanism of Obesity and Ralated Pathologies: The Macro- and Microcirculation of Adipose Tissue. FEBS J. 2009, 276, 5738−5746. (64) Sawane, M.; Kajiya, K.; Kidoya, H.; Takagi, M.; Muramatsu, F.; Takakura, N. Apelin Inhibits Diet-Induced Obesity by Enhancing Lymphatic and Blood Vessel Integrity. Diabetes 2013, 62, 1970−1980. (65) Scallan, J. P.; Hill, M. A.; Davis, M. J. Lymphatic Vascular Integrity Is Disrupted in Type 2 Diabetes due to Impaired Nitric Oxide Signalling. Cardiovasc. Res. 2015, 107, 89−97. (66) Torrisi, J. S.; Hespe, G. E.; Cuzzone, D. A.; Savetsky, I. L.; Nitti, M. D.; Gardenier, J. C.; Nores, G. D. G.; Jowhar, D.; Kataru, R. P.; Mehrara, B. J. Inhibition of Inflammation and iNOS Improves Lymphatic Function in Obesity. Sci. Rep. 2016, 6, 19817. (67) Trevaskis, N. L.; Kaminskas, L. M.; Porter, C. J. H. From Sewer to SaviourTargeting the Lymphatic System to Promote Drug Exposure and Activity. Nat. Rev. Drug Discovery 2015, 14, 781−803. (68) Vuorio, T.; Nurmi, H.; Moulton, K.; Pikkarainen, J.; Robciuc, M. R.; öhman, M.; Heinonen, S. E.; Samaranayake, H.; Heikura, T.; Alitalo, K.; Yla-Herttuala, S. Lymphatic Vessel Insufficiency in Hypercholesterolemic Mice Alters Lipoprotein Levels and Promotes Atherogenesis. Arterioscler., Thromb., Vasc. Biol. 2015, 34, 1162−1170. (69) Heo, J. H.; Yoon, J. A.; Ahn, E. K.; Oh, C. O.; Yu, B. C.; Lee, S. J. Intraperitoneal Administration of Adipose Tissue-derived Stem Cells for the Rescue of Retinal Degeneration in a Mouse Model via Indigenous CNTF up-regulation by IL-6. J. Tissue Eng. Regener. Med. 2008, 12, e1370−e1382. (70) Jiang, Y.; Pu, K. Advanced Photoacoustic Imaging Applications of Near-Infrared Absorbing Organic Nanoparticles. Small 2017, 13, 1700710.

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