Aug 23, 2017 - vivo degradation of injectable hydrogels that are mixed with CNDs. ..... fluorescence dyes, semiconducting quantum dots and upconversion nanoparticles were ...... The CNDs encapsulated inside hydrogels did not diffuse.
Accepted Manuscript Visual in vivo degradation of injectable hydrogel by real-time and non-invasive tracking using carbon nanodots as fluorescent indicator Lei Wang, Baoqiang Li, Feng Xu, Ying Li, Zheheng Xu, Daqing Wei, Yujie Feng, Yaming Wang, Dechang Jia, Yu Zhou PII:
S0142-9612(17)30558-6
DOI:
10.1016/j.biomaterials.2017.08.039
Reference:
JBMT 18240
To appear in:
Biomaterials
Received Date: 31 March 2017 Revised Date:
23 August 2017
Accepted Date: 26 August 2017
Please cite this article as: Wang L, Li B, Xu F, Li Y, Xu Z, Wei D, Feng Y, Wang Y, Jia D, Zhou Y, Visual in vivo degradation of injectable hydrogel by real-time and non-invasive tracking using carbon nanodots as fluorescent indicator, Biomaterials (2017), doi: 10.1016/j.biomaterials.2017.08.039. This is a PDF file of an unedited manuscript that has been accepted for publication. As a service to our customers we are providing this early version of the manuscript. The manuscript will undergo copyediting, typesetting, and review of the resulting proof before it is published in its final form. Please note that during the production process errors may be discovered which could affect the content, and all legal disclaimers that apply to the journal pertain.
ACCEPTED MANUSCRIPT
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Visual in vivo Degradation of Injectable Hydrogel by Real-time and Non-invasive Tracking Using Carbon Nanodots as Fluorescent Indicator
ABSTRACT Visual in vivo degradation of hydrogel by fluorescence-related tracking and monitoring is crucial for quantitatively depicting the degradation profile of hydrogel in a real-time and
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non-invasive manner. However, the commonly used fluorescent imaging usually encounters limitations, such as intrinsic photobleaching of organic fluorophores and uncertain perturbation of degradation induced by the change in molecular structure of hydrogel. To address these problems,
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we employed photoluminescent carbon nanodots (CNDs) with low photobleaching, red emission
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and good biocompatibility as fluorescent indicator for real-time and non-invasive visual in vitro/in vivo degradation of injectable hydrogels that are mixed with CNDs. The in vitro/in vivo toxicity results suggested that CNDs were nontoxic. The embedded CNDs in hydrogels did not diffuse outside in the absence of hydrogel degradation. We had acquired similar degradation kinetics
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(PBS-Enzyme) between gravimetric and visual determination, and established mathematical equation to quantitatively depict in vitro degradation profile of hydrogels for the predication of in vivo hydrogel degradation. Based on the in vitro data, we developed a visual platform that could
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quantitatively depict in vivo degradation behavior of new injectable biomaterials by real-time and
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non-invasive fluorescence tracking. This fluorescence-related visual imaging methodology could be applied to subcutaneous degradation of injectable hydrogel with down to 7 mm depth in small animal trials so far. This fluorescence-related visual imaging methodology holds great potentials for rational design and convenient in vivo screening of biocompatible and biodegradable injectable hydrogels in tissue engineering.
Keywords: In vivo degradation; visualization; injectable hydrogel; real-time and non-invasive; fluorescence tracking 2
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1. INTRODUCTION Biocompatible and biodegradable hydrogels with three dimensional polymeric structure have served as water-swollen gels and received significant attention for biomedical applications such as
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controlled drug delivery and tissue engineering during past decades [1-5]. As for their tissue engineering applications, a quantitative assessment of in vivo degradation of hydrogels is of great importance especially for design of customized hydrogel with controllable degradation rate that
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can match with the regeneration rate of newly generated tissues [6-8]. However, it is difficult to
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reveal the in vivo degradation of hydrogels by in vitro degradation models due to the complex in vivo microenvironment (synergetic biodegradation by a variety of enzymes and cells). Currently, the commonly used and reliable technique for quantitatively assessing the in vivo degradation behavior of hydrogels is mainly based on gravimetric/volume determination. However, such
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technique needs to sacrifice lots of animals, which limits its wide application in determining the degradation of hydrogels [9-11]. Therefore, to minimize the uncontrollable parameters and intricate variability and to reduce the amount of required animals, there is an urgent need to
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develop strategies for real-time and non-invasively monitoring in vivo degradation. To date,
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non-invasive imaging techniques, including magnetic resonance imaging [6, 12-14], X-ray computed tomography [15], ultrasound imaging [16-18] and fluorescence imaging [8, 19-22], provide a reliable and efficient measurement for determining the in vivo degradation of hydrogels or a platform for continuously and noninvasively monitoring in vitro cell viability, proliferation and chemosensitivity [23, 24]. Although magnetic resonance imaging and X-ray computed tomography are intriguing for real-time and non-invasively monitoring in vivo degradation of hydrogels due to their high spatial resolution, they need radiopaque contrast agents for imaging 3
ACCEPTED MANUSCRIPT and complicated instrument. Hence fluorescence-related imaging remains the most widely employed technique for in vivo tracking and monitoring of hydrogel recently. To achieve this goal, various fluorescent probes
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such as organic fluorescent molecules and rare-earth upconversion nanoparticles have been developed to label hydrogels. For example, the strategies of covalently immobilizing fluorescein to the biomaterials have been adopted to track and quantify the in vitro or in vivo degradation of
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hydrogels [7, 8, 19, 21]. Fluorescein-5-carboxyamido hexanoic acid was covalently attached to
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model materials for in vivo tracking erosion of PEG/dextran hydrogel [8]. Similarly, rhodamine B and IR-Dye 800CW maleimide were employed to chemically label PEG hydrogel and hyaluronan hydrogels, respectively, for monitoring in vivo degradation of hydrogels [7, 19]. However, the strategy based on covalently immobilized fluorescein to hydrogel suffers from some intractable
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issues, including photobleaching when undergoing long exposure and uncertain perturbation of degradation due to the change in molecular structure of hydrogels. Recently, silica-coated lanthanide-doped rare-earth nanoparticles have been directly embedded into hydrogel to track
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hydrogel degradation in live tissues non-invasively, avoiding the need for chemical bonding with
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hydrogels [22]. However, the potential biotoxicity of rare-earth nanoparticles due to long-term retention in the liver and spleen system remains uncertain [25-27]. Inspiring by our previous finding that hydrogel degradation behaviors could be well reflected by the release of magnetic nanoparticle incorporated within hydrogel matrix [28], we attempted to explore another strategy of directly mixing biocompatible and luminescent nanoparticles with low photobleaching for real-time and non-invasively monitoring the in vivo degradation of hydrogels. Serving as novel carbon nanomaterials, carbon nanodots (CNDs) have offered potential for 4
ACCEPTED MANUSCRIPT widespread applications in bioimaging fields, mainly owing to their intriguing luminescent property, low photobleaching and good biocompatibility. These outstanding properties make CNDs promising luminescent nanomaterials for in vivo imaging compared to conventionally
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employed organic dyes and rare-earth nanoparticles [29-35]. Moreover, the large stokes shift can provide CNDs with red emission wavelengths, allowing better penetration ability for in vivo fluorescence bioimaging compared to UV or visible lights that could be largely absorbed by
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biomolecules in tissues [36, 37].
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Here we explored CNDs as fluorescent indicator for visual in vitro/in vivo degradation of hydrogel by fluorescence imaging in a real-time and non-invasive manner. The in vitro/in vivo toxicity of CNDs was evaluated by in vitro MTT assay and in vivo histopathological assay. The direct embedding of photoluminescent CNDs allowed low photobleaching, red emission and good
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biocompatibility and avoided the need for chemically bonding. The in vitro degradation kinetics of hydrogels was investigated by gravimetric and visual determination, respectively. The CNDs embedded in hydrogels did not diffuse outside in the absence of hydrogel degradation. We had
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established a mathematical equation for quantitatively depicting hydrogel degradation behavior.
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With help of the established mathematical equation we acquired similar constant k value among gravimetric and visual determination, which offered the feasibility in obtaining quantitative degradation behavior of hydrogels, thus representing conventional gravimetric determination. We also applied the strategy of fluorescent imaging for in vivo degradation of injectable hydrogels by real-time and non-invasively fluorescence monitoring, which quantitatively depicts the degradation behavior of biodegradable hydrogel.
5
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2. EXPERIMENTAL SECTION 2.1. Chemicals and materials Chitosan (CS, viscosity average molecular weight Mη = 3.4×105, degree of deacetylation =
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91.4%) was purchased from Qingdao Hecreat Bio-tech company Ltd (China). Porcine skin gelatin, sodium alginate, methacrylic anhydride (MA, 94%), photoinitiator (Irgacure® 2959, I2959), lysozyme (chicken egg white) and fluorescein isothiocyanate (FITC) were purchased from
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Sigma-Aldrich (USA). Citric acid monohydrate and formamide were supplied by Sinopharm
Da) were supplied by Solarbio (USA).
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Chemical Reagent Co (China). Dialysis bags (retained molecular weight 500 and 8,000~14,000
2.2. Synthesis and characterization of CNDs
CNDs were synthesized from citric acid and formamide following a solvothermal method.
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Briefly, citric acid monohydrate (1.19 g) was dissolved in formamide (20 mL), and the mixed solution was then transferred into a Teflon-lined autoclave (50 mL). After heating at 180°C for 4 h, the obtained dark red solution was centrifuged at 8000 rpm for 30 min to remove large or
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agglomerated deposit. The black CNDs powder (~50 mg) was obtained by dialyzing against
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deionized water using a dialysis bag and lyophilization. UV-vis absorption spectrum of CNDs was exhibited on TU 1901 spectrophotometer and the spectrum was collected from 200 nm to 500 nm. Photoluminescence (PL) spectra were measured on a Hitachi F-4600 fluorometer equipped with Xe lamp at ambient conditions. Transmission electron microscopy (TEM) images of CNDs were pictured from a transmission electron microscope (FEI Tecnai G2 F30). Fourier transform infrared (FTIR) spectra were obtained on a Perkin-Elmer Spectrum One ranging from 4000 to 500 cm-1. X-ray photoelectron spectroscopy (XPS) spectra were performed on an ESCALAB 250Xi X-ray 6
ACCEPTED MANUSCRIPT photoelectron spectrometer with Al/K α as the source, and the energy step size was set as 0.1 eV. Atomic force microscopy (AFM) was performed on Bruker Dimension Icon.
2.3. In vitro cytotoxicity assay and cell bioimaging of CNDs
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The cytotoxicity was evaluated by tetrazolium-based MTT assay against NIH/3T3 cells. In 96 well plates, 100 µL suspension of NIH/3T3 cells (1 × 104 cells/mL) in Dulbecco’s Modified Eagle’s Medium supplemented with 10% (v/v) fetal bovine serum, penicillin (50
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U/mL)/streptomycin (50 µg/mL) were added to each well and incubated in 5% CO2 humidified
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atmosphere at 37°C for 24 h. The CNDs with different concentrations (200, 400, 600, 800 µg/mL) were introduced into the wells and incubated for another 24h, 36h, 72h, respectively. At the designated time intervals, the medium was removed and cells were washed with phosphate-buffered saline. Then, 20 µL of 5 mg/mL MTT solution was added to each well. The
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96-well plates were further incubated for 4 h, followed by removing the culture medium with MTT, and then 200 µL of DMSO was added. The optical density of the mixtures at 490 nm was measured using microplate reader. Cell viability was expressed as percentage of absorbance
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relative to control (i.e., without CNDs). Experiment was performed in triplicates, with nine
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replicate wells for each sample and control per assay. NIH/3T3 cells with concentration of 1 × 104 cells/mL were seeded in each well of 96-well
plates and cultured at 37°C for 24 h. DMEM medium solution of CNDs (200 µg/mL) was filtered through a 0.22 µm membrane. The filtered fluorescent culture medium then added to plates. After an incubation of 6 h, the medium was removed and the cells were washed three times with PBS and kept in PBS for bioimaging. The fluorescence images were carried out on fluorescence microscope (Nikon Eclipse Ti S). 7
ACCEPTED MANUSCRIPT 2.4. In vivo toxicity assessment of CNDs All animal experimental protocols were approved by the animal care and use regulations (Ethics Committee of Xi’an Jiaotong University), and the experiments were carried out under
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control of the University’s Guidelines for Animal Experimentation. Kunming mice were 8 weeks of age, weighed 20~23g and acclimatized for 5 days after arrival. Then mice were subcutaneously injected with CNDs (500 µL of 1000 µg/mL solution for each mouse, i.e., a dose of ~23 mg/kg).
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Mice treated with normal saline solution (without CNDs) were used as control group. Mice were
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weighted every 3 days for 3 weeks. Three animals from each group were sacrificed at pre-set time points of 1 and 14 days after injection, and various organs (including liver, spleen, kidney, heart and lung) were collected and scanned for the fluorescence imaging. The fluorescence intensity of each organ was quantified using the Carestream MI software. The histopathological analysis of
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various organs such as liver, spleen, kidney, heart and lung was performed by hematoxylin and eosin (H&E) staining. Major organs were fixed in 4% paraformaldehyde buffer solution overnight, followed by dehydration with 70% ethanol, and then paraffin-embedded. Paraffin embedded
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tissues were cut (5 mm), stained with H&E and examined under a microscope.
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2.5. Visualization of CNDs hybrid hydrogel using CNDs as fluorescent indicator Injectable N-methacryloyl chitosan (N-MAC) with degree of substitution (DS) of 19%, 25%
and 28% (N-MAC 19, N-MAC 25, N-MAC 28) were synthesized according to our previous work [38]. To optimize the concentration of CNDs in hybrid hydrogel for in vitro and in vivo visualization, N-MAC phosphate buffer saline (PBS) solution with different CNDs concentrations (0, 20, 50, 200, 500, 1000, and 1500 µg/mL) were prepared in PDMS mold and irradiated under Omni Cure® S2000 spot curing system (EXFO Inc, Canada) with an intensity of 10 mW/cm2 for 8
ACCEPTED MANUSCRIPT 30 s. Fluorescence imaging (Pseudo-color images, 590 nm excitation wavelength with 700 nm emission wavelength) was acquired using a small animal in vivo fluorescence imaging system (In-Vivo FxPro; Carestream, MI, USA). The mean fluorescence intensity was quantified using the
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Carestream MI software. To assess in vitro visualization of CNDs hybrid hydrogel, patterned microgels (i.e., concentric ring and convex) were prepared by photolithographic method and imaged by fluorescence microscope (Nikon Eclipse Ti S).
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To determine the photobleaching, the CNDs hybrid hydrogel or FITC hybrid hydrogel were
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prepared and irradiated using a 30W xenon excitation source of 365~405 nm. The fluorescence pseudo-color images of hybrid hydrogels were acquired using a small animal in vivo fluorescence imaging system with respect to time. The fluorescence intensity was quantified using the Carestream MI software.
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A laser scanning confocal microscope (PerkinElmer Ultra VIEW system, USA) was used to examine the homogeneity of CNDs in hydrogels. In typical experiments, z planes covering 100 µm thick sections of hydrogels were chosen for imaging. The x-y plane images at different depth
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across the thickness of 100 µm were captured in 10 µm increments and the fluorescence emission
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intensity was measured. The red field images were excited with a 568 nm Argon/Krypton laser with 400 µW of power (exposure time: 0.2 s), and emissions were filtered with 605 nm band-pass filter.
2.6. In
vitro
degradation
of
CNDs
hybrid
hydrogel
by
gravimetric
determination In vitro degradation of CNDs hybrid hydrogels was conducted in 10 mL of phosphate buffer saline (PBS, pH=7.4) solution with or without lysozyme (0.2 mg/mL) at 37 °C. The 9
ACCEPTED MANUSCRIPT PBS-lysozyme solution was refreshed daily to ensure continuous enzyme activity. At pre-set time intervals, the residual CNDs hybrid hydrogels samples were removed from medium, gently washed with distilled water and weighed. The weight loss (WL) was defined as: WL= (W0 − )⁄W0 ×100%
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(1)
where W0 and Wt are the weights of samples at initial time and time t during degradation, respectively.
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Synchronously the in vitro accumulative release profile of CNDs in hydrogels with or without
nm) from degradation medium.
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lysozyme was estimated by using UV–vis spectrophotometer (measuring the absorbance at 270
2.7. Visual in vitro degradation of CNDs hybrid hydrogel by fluorescence tracking
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Visual in vitro degradation of CNDs hybrid hydrogel (N-MAC 19, N-MAC 25, and N-MAC 28) was performed based on the relative grayscale change in fluorescent images. Typically, thin cuboid-shaped (5 × 8 × 1 mm) CNDs hybrid hydrogels was immersed in 10 mL of PBS solution
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with or without lysozyme (0.2 mg/mL) at 37 °C. At pre-set time intervals, the fluorescent images
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of CNDs hybrid hydrogels were captured using a fluorescence microscope (Nikon Eclipse Ti S) with fixed exposure time (500 ms). The fluorescence reduction (FR) was defined as: FR= (IOD0 − )⁄IOD0 ×100%
(2)
where IOD0 and IODt are integrated optical density (analyzed by Image Pro 6) of the samples at initial time and time t during degradation, respectively.
2.8. Tissue penetration evaluation of CNDs at wavelength of 590 nm Fresh slices of chicken chip (breast meat) with different thickness (2 mm, 5 mm and 7 mm) 10
ACCEPTED MANUSCRIPT were prepared. The CNDs hybrid hydrogel was placed on the top of the chicken chip. The excitation light was located beneath the chicken chip for evaluating tissue penetration. The fluorescence images (pseudo-color images) were obtained via small animal in vivo fluorescence
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imaging system. The fluorescence intensity was quantified using the Carestream MI software. The excitation wavelength, emission wavelength and exposure time were set as 590 nm, 700 nm and 20 s, respectively.
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non-invasive fluorescence tracking
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2.9. Visual in vivo degradation of CNDs hybrid hydrogel by real-time and
All animal experimental protocols were approved by the local animal care and use regulations (Ethics Committee of Xi’an Jiaotong University), and the experiments were carried out under control of the University’s Guidelines for Animal Experimentation. Kunming mice were
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8 weeks of age, weighed 20~25g and acclimatized for 5 days after arrival. The CNDs (1000 µg/mL) and N-MAC (N-MAC 19, N-MAC 25, and N-MAC 28) was directly dissolved in PBS to form a homogeneous solution (sterilized by filtration; 0.22 µm). The Kunming mice were
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randomly distributed in three groups treated with CNDs hybrid hydrogels (N-MAC 19, N-MAC
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25 and N-MAC 28). The transdermal curing hydrogels were carried out via subcutaneous injection of mice. After anesthetization, 500 µL of CNDs hybrid solution was injected into subcutaneous space of mice back through syringe with 25G needle and then sequentially crosslinked by UV irradiation for 30 s. Mice treated with free CNDs (CNDs solution without hydrogel) were used as control group. At pre-set time intervals, the mice were anesthetized and the fluorescence images (pseudo-color images) were obtained via small animal in vivo fluorescence imaging system. The fluorescence intensity was quantified using the Carestream MI software. The excitation 11
ACCEPTED MANUSCRIPT wavelength, emission wavelength and fixed exposure time were set as 590 nm, 700 nm and 20s, respectively. In order to prove the reliability of the visual determination method, the mice in the parallel control group were euthanized at the designed time intervals and the remained hydrogels
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were collected, washed with PBS and then weighted to estimate the percentage of degradation. To confirm the feasibility that this visual determination method can be applicable to various biomaterial systems, CNDs hybrid gelatin hydrogel and alginate hydrogel were injected into
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real-time and non-invasive fluorescence tracking.
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subcutaneous space of mice back using the same procedure for visual in vivo degradation by
2.10. Histological observation of CNDs hybrid hydrogel
To assess the biocompatibility of CNDs hybrid hydrogel, the mice were sacrificed by intraperitoneal injection of excess chloral hydrate at the designated time intervals of 72, 120, 192,
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and 288 h. The hydrogel samples adjacent with surrounding skin was resected and fixed immediately in 4% paraformaldehyde buffer solution. The samples were dehydrated with 70% ethanol, and then paraffin-embedded. Paraffin embedded tissues were cut (5 mm), stained with
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H&E and examined under a digital microscope.
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2.11. Statistical analysis
All the data were expressed as means ± standard deviation of at least triplicate samples. The
statistically significant difference was evaluated by Student's T-test, and statistical significance was considered for p value 3.1. Morphology, Optical and in vitro/in vivo toxicity of CNDs 13
ACCEPTED MANUSCRIPT We synthesized CNDs via solvothermal method and characterized the morphology, surface properties and optical properties via TEM, AFM, XPS and PL spectra. As illustrated in Figure 1A, the morphology of CNDs was examined using HR-TEM. The mean diameter of CNDs was
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5.1±1.2 nm without aggregation, supporting that CNDs were uniformly spherical morphology. Fast Fourier transform revealed characteristic hexagonal diffraction pattern of graphite, further supporting the formation of graphitic structure [42]. As shown in AFM images, the densely and
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well-dispersed CNDs appeared on the silicon substrate with particle heights information about 3~6
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nm, which also consistent with particle size from HR-TEM image. As illustrated in Figure 1B, the CNDs absorption spectrum exhibited a characteristic absorption peak at 270 nm, which are assigned to typical absorption of π→π∗ electronic transition of aromatic system (suggestive of sp2 carbon network). The well-dispersed suspension showed transparent under day light and exhibited
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bright blue luminescence under UV excitation (insert in Figure 1B). Three dominant peaks at 286, 400 and 531 eV appeared in XPS survey spectrum, suggesting composition of carbon/C1s, nitrogen/N1s, and oxygen/O1s elements in CNDs (Figure S1A). The high-resolution of C1s
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spectrum (Figure S1B) could be fitted to three peaks at 284.6, 285.8 and 287.9 eV, corresponding
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to C=C, C–N/C–OH and C=O bonds. The FTIR spectrum showed characteristic peaks at 3210, 1680 and 1600 which correspond to σ (stretching vibration) O–H/N-H, σ C=O and NH2 band, respectively (Figure S2), indicating the presence of hydroxyl, carboxyl and amino groups. The presence of abundant functional groups imparted excellent solubility in water without further chemical modification. The PL emission wavelength shifted to longer wavelength as excitation wavelengths increased from 300 to 460 nm. The strongest fluorescence emission was observed with a peak at 460 nm when using 375 nm excitation wavelengths (Figure 1C). The absolute 14
ACCEPTED MANUSCRIPT fluorescence quantum yield of CNDs was measured to be 17.2%, which was comparable to previous reports [43, 44]. Notably, the large stokes shift provided CNDs with significant benefits (red emission) for in vivo fluorescence bioimaging, as red emission wavelengths would provide a
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deeper penetration ability, minimal autofluorescence and high signal-to-noise ratios [45, 46]. The desired fluorescent indicator for visual in vivo degradation of hydrogel should be non-toxic. Therefore, the in vitro cytotoxicity of CNDs with different concentrations (0, 200, 400,
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600 and 800 µg/mL) was evaluated by culturing with NIH/3T3 cells for 24, 36 and 72h using
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MTT method, respectively. The NIH/3T3 cells viability with various concentrations maintained all above 90% even at 72 h (Figure 1D). In addition, no statistically significant differences were observed between CNDs groups and control group in cell viability, which indicated that CNDs exhibited good biocompatibility, potentially for live cell imaging and visual in vivo degradation of
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hydrogel. The application of CNDs as live cell imaging probe was demonstrated. As depicted in Figure 1E, cell uptake of CNDs was clearly observed with bright red fluorescence at fluorescence (540~560 nm) and bright field after incubation with 200 µg/mL CNDs for 6 h. The result implied
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that CNDs can serve as fluorescent probes for live cell bioimaging. Moreover, no autofluorescence
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emerged from cells of control group in fluorescence field, which confirmed that bright red fluorescence was ascribed to the cell uptake of CNDs and being internalized into the cells. More importantly, no morphological damage of the cells was observed upon incubation with the CNDs further demonstrating their good biocompatibility. The primary challenge of any fluorescent indicator is to achieve high photostability and possess low cytotoxicity. So in this regard, organic fluorescence dyes, semiconducting quantum dots and upconversion nanoparticles were extensively studied as bioimaging probes. However, the inherent limitations include poor photostability of 15
ACCEPTED MANUSCRIPT organic fluorescence dyes and potential toxicity of heavy metal containing semiconducting quantum dots/upconversion nanoparticles [47-49]. Hence, for in vivo applications, CNDs could serve as an exciting alternative as they were nontoxic.
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To determine the clearance and in vivo biocompability of CNDs as fluorescent indicator, mice were sacrificed after 1 and 14 days of injection and isolated liver, spleen, kidney, heart and lung
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were scanned for the fluorescence imaging. As depicted in Figure 2A, the observed fluorescence
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from control group was so low and typical autofluorescence of tissues [50]. Importantly, the same marginal values of fluorescence were also observed after 1 and 14 days of CNDs injection. No statistically significant differences were observed between CNDs group and control group in fluorescence intensity of various organs (Figure 2B). Based on the results of fluorescence
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measurements, we could infer that the CNDs using as fluorescent indicator would be completely removed from the body after 14 days of CNDs injection, and long-term retention in various organs could not occur. Over a period of 3 weeks, neither death nor significant body weight drop was
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noted in the CNDs group (Figure S3), indicating that the mice of CNDs group could continue to
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mature without any significant toxic effects. As the injection of CNDs may induce subsequent damage in the organs related to nanoparticle clearance, in vivo toxicity of CNDs was further examined in various organs (including liver, spleen, kidney, heart and lung) after 1 and 14 days of CNDs injection, using CNDs-untreated rats as the control group. The key organs from both CNDs injection and control groups had integrated tissue structure without edema, inflammation and abnormal defects (Figure 2C), similar with previous researches about in vivo toxicity of CNDs [51, 52]. Histopathologically, no significant alterations occurred in these tissues relative to the 16
ACCEPTED MANUSCRIPT control group, demonstrating that no significant in vivo toxicity was observed when employing CNDs as fluorescent indicator. In both CNDs injection and control groups, cardiac myocytes were clear and arrayed in order without any inflammatory exudate, hemorrhage, hypertrophy or
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necrosis. The structure of liver lobules with central vein was clearly delineated, and there were no inflammatory infiltrates. Additionally, the splenic corpuscle structure of spleen was normal and clearly delineated. No pathological changes were found in spleen sinus, the white pulp, and red
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pulp of spleen. The tissue structure of lung was normal, and there were no bronchioles and alveoli
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ectasia or collapse, alveolar epithelial denaturation, interstitial hyperemia, or inflammatory cell infiltration surrounding the bronchus. From micrograph of mice kidney, the renal glomerulus and various kidney tubes displayed normal shape, without degeneration, bleeding or inflammatory exudate. All above histopathological results suggest that CNDs were nontoxic in vivo. Thus CNDs
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could serve as a promising fluorescent indicator for visual in vivo degradation of hydrogel.
3.2. Visualization of CNDs hybrid hydrogel using CNDs as fluorescent indicator
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The fluorescence pseudo-color images of CNDs hybrid hydrogel with different
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concentrations of CNDs were acquired on a small animal in vivo fluorescence imaging system. The quantitative relationship of fluorescence signal intensity versus CNDs concentration and the pseudo-color images were shown in Figure 3A. The fluorescence signal intensity was positively correlated to CNDs concentration. When CNDs concentration increased to 1000 µg/mL, fluorescence signal intensity does not increase anymore. It was suggested that CNDs concentration of 1000 µg/mL could give a detectable and strong enough fluorescence signal for visualization in vitro/in vivo. The evaluation of CNDs hybrid hydrogel for in vivo bioimaging was 17
ACCEPTED MANUSCRIPT investigated using subcutaneous injection into mice back and then sequentially applied 30s UV irradiation. The fluorescent region signals of CNDs hybrid hydrogel could be obviously observed with a distinct boundary between CNDs hybrid hydrogel and surrounding skin tissue, which
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showed that CNDs hybrid hydrogel are capable of in vivo bioimaging with long emission wavelengths (λem: 700 nm) as shown in Figure S4. Importantly, the excitation-dependent PL behavior allowed CNDs with crucial benefits for in vivo fluorescence bioimaging owing to red
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emission wavelengths with stronger penetration ability. The patterning CNDs hybrid microgels
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were fabricated via UV lithography with help of photomask. The images of patterned microgel, such as concentric ring pattern (left) and convex pattern (right), exhibited green and red fluorescence under different excitation (460~490 and 540~560 nm), and highly tallied with photomask which demonstrated that CNDs could be dispersed homogeneously in patterned
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microgels (Figure 3B).
In order to take one picture, it includes several procedures (such as pre-scan, ROI position adjustment and taking picture) and the time required to irradiate is about 120s. Specifically, it
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usually takes several quarters required to irradiate for long-term in vivo hydrogel tracking. So the
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photostability of fluorescent indicator indeed was a non-negligible problem, especially for the biomaterial with slow degradation. The photobleaching of CNDs was compared with FITC under a continuous 365 nm UV lamp illumination. The images of CNDs hybrid hydrogel (Figure 3C, up) and FITC hybrid hydrogel (Figure 3C, down) showed a clearly fluorescence signal. An obvious descending of fluorescence intensity of FITC hybrid hydrogel could be observed, while the CNDs hybrid hydrogel’s fluorescence intensity changed slightly. According to the vividly contrast (Figure 3D), the decay curve of FITC descended (decrease by 50%) with a much larger amplitude 18
ACCEPTED MANUSCRIPT than the curve of CNDs (decrease by less than 10%). It was elucidated that CNDs as fluorescent indicator were much more photostable than fluorescent dye (such as FITC or RhB) [53, 54]. To demonstrate the homogeneity of CNDs in hydrogels, a cuboid region (100×300×300 µm)
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was chosen in hydrogels, and the x-y plane images at different depth in the vertical cross-sections of the regions were captured. The fluorescence intensity profiles were examined using confocal microscopy (Figure 3E). The representative x-z plane image exhibited homogeneous and bright
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red fluorescence, suggesting that the CNDs were evenly distributed in horizontal cross-section.
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The fluorescence intensity at different depth displays relatively consistent, further confirming the homogeneity of CNDs in horizontal cross-sections. Additionally the representative x-y plane images at different depth also exhibited homogeneous and bright red fluorescence, demonstrating that the CNDs were evenly distributed in vertical cross-sections. Thus the homogeneity of CNDs vertical
cross-sections
three-dimensional space.
and
horizontal
cross-section
proved
the
homogeneity
in
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in
Fluorescence-related imaging technique has seen revolutionary advancements in both
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sensitivity and resolution over the past decade. However, photoinduced degradation
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(photobleaching) of the commonly used organic fluorescent molecule remains a key obstacle that limits the temporal and spatial resolutions of imaging [55]. In addition, photoinduced fluorophore toxicity (phototoxicity) may also lead to unwanted perturbations to the biological system that can obscure the signal of interest [56]. Conversely, CNDs exhibit high resistance to photobleaching and possess better biocompability compared to organic fluorescent molecule and traditional semiconductor quantum dots and much more suitable for visualization research [57].
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ACCEPTED MANUSCRIPT 3.3. Visual In Vitro Degradation of CNDs Hybrid Hydrogel To explore the correlation between CNDs release and hydrogel degradation by gravimetric determination in vitro, we evaluated the degradation of CNDs hybrid hydrogels in PBS and
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PBS-lysozyme solution over 500 h by synchronously recording the weight loss of hydrogel and amount of released CNDs at regular intervals. Almost no weight loss (no degradation) was observed during the 500 h for all CNDs hybrid hydrogels (N-MAC 19, N-MAC 25, and N-MAC
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28) in absence of lysozyme (Figure 4A), which could specifically break glycosidic bonds of
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chitosan backbone linkages [58]. More importantly, the CNDs release behavior followed a similar trend: CNDs (1%~ 3% releases) were hardly detected during 500 h. It was suggested that CNDs release was not controlled by diffusion but is to be related to hydrogel degradation (Figure 4B). That was similar with our previous research about magnetic nanoparticles release profile: only
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degradation of hydrogels would allow the release of the magnetic nanoparticles [28]. It had been confirmed that there were abundant functional groups (including hydroxyl, carboxyl and amino groups) on the surface of CNDs (Figure S1&Figure S2). And chitosan is a biodegradable and
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natural polymer with amino groups. Thus, CNDs in hydrogels would be immobilized to the
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chitosan polymer chain due to the hydrogen bonds [59, 60] formed between carboxyl groups (-COOH) of CNDs and amino groups (-NH2) of chitosan in the absence of hydrogel degradation. The CNDs are retained in the hydrogel by non-covalent interactions (partially ascribed to hydrogen bonds) [61]. The CNDs immobilized to the chitosan polymer chain could diffuse outside from hydrogel when degradation of chitosan hydrogel occurred. The degradation of hydrogels is mostly ascribed to breakage of polymer backbone in the presence of lysozyme [62, 63]. The degradation of CNDs hybrid hydrogels by gravimetric determination was followed for 500 h. The 20
ACCEPTED MANUSCRIPT CNDs hybrid hydrogels (N-MAC 19, N-MAC 25, and N-MAC 28) ultimately degraded with 48%, 59% and 85% after 500 h in the presence of lysozyme, indicating that increase of chemical cross-linked networks density could result in a slower in vitro degradation profile with an identical
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enzyme concentration (Figure 4A). On the other hand, the release behaviors of CNDs embedded in hydrogels also exhibited a very similar trend: the accumulative release amount of CNDs in hybrid hydrogels (N-MAC 19, N-MAC 25, and N-MAC 28) ultimately reached 43%, 57%
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and 82% after 500 h in the presence of lysozyme. Furthermore, following the first-order enzymatic
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hydrolysis kinetics [64-66], a mathematical equation for in vitro degradation of hybrid hydrogel was developed via exponential fitting for evaluating the correlation between CNDs release and hydrogel degradation in vitro in the presence of enzyme: -dmE/dt = kmE, upon integration and deformation
% = 1 − × 100% = 1 − exp (− )
(3)
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Where WL is the weight loss, WE and WE0 is the undegraded weight and initial weight of hydrogel, k is the enzymatic hydrolysis constant related to the polymeric structure, t is degradation
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time. Thus, 1-mE/mE0 represents the fraction of degraded hydrogel. Using Eq. 3 to fit the
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degradation curves shown in Figure 4A, we had acquired the following fitted equations, respectively: N-MAC 19 (R2=0.96), N-MAC 28
WL =
WL
1 ‒ exp(-0.00527t) (R2=0.98), N-MAC 25
WL =
1 ‒ exp(-0.00245t)
= 1 ‒ exp(-0.001t) (R2=0.94). Furthermore, we also found that there
existed an analogous and classical Lagergren pseudo first order enzyme adsorption kinetics [67, 68]: dq/dt = ka (qe-q), upon integration and deformation
= 1 − exp (− )
(4)
Where qe and q is the equilibrium adsorption mass and adsorption mass of enzyme, ka is the 21
ACCEPTED MANUSCRIPT pseudo first order adsorption rate coefficient. Based on the close correlation between hydrogel degradation and enzyme adsorption, the degradation mechanism of CNDs hybrid hydrogels could be explained that the enzyme adsorption and contact with hydrogel internal/external surface
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gradually triggered the breakage of chitosan polymer backbone. It was demonstrated that the embedded CNDs were released from the hydrogels as CNDs hybrid hydrogels degraded and a positive correlation between CNDs release and hydrogel degradation was achieved. All these
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results indicated that it was possible to predict in vitro degradation kinetics of injectable
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biomaterials through employing CNDs release profile. To confirm this, we also fit the CNDs release curve using the established mathematical equation and acquired the following fitted equations, respectively: N-MAC 19
AR
= 1 ‒ exp(-0.00518t) (R2=0.97), N-MAC 25
AR
= 1 ‒
exp(-0.0025t) (R2=0.95), N-MAC 28 AR = 1 ‒ exp(-0.00127t) (R2=0.93). It should be noted that we
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acquired similar k value between gravimetric determination (5.27×10-3, 2.45××10-3 and 1×10-3) and CNDs release (5.18×10-3, 2.5×10-3 and 1.27×10-3), which further confirmed that the positive correlation between CNDs release and hydrogel degradation. Only by degradation of CNDs
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hybrid hydrogels can CNDs embedded in hydrogel be released.
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To test the feasibility of CNDs serving as fluorescent indicator for monitoring in vitro
degradation of hydrogel via the strategy of fluorescent imaging, the fluorescence images acquired with 700 nm emission (Figure 5A) and quantified fluorescence reduction at 0, 54, 140, 180, 300, 504 hour was shown in Figure 5B. In absence of enzyme, all CNDs hybrid hydrogels maintained their initial shape and fluorescence intensity during the whole degradation process. Slight fluorescence reduction (FR) (2%~5%) occurred (Figure 5B), indicating that almost no 22
ACCEPTED MANUSCRIPT degradation occurred, and the CNDs did not diffuse outside of the hydrogels. On the other hand, in the presence of lysozyme the shape of CNDs hybrid hydrogels diminished with different degree of enzymatic degradation, indicating the decrease of chemical cross-linked networks density could
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result in faster degradation. It was quantified that FL of CNDs hybrid hydrogels (N-MAC 19, N-MAC 25, and N-MAC 28) ultimately reached a different extent: 39%, 52% and 79% after 500 h as shown in Figure 5B. An exponential fitting was also established for evaluating the dependence
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between gravimetric degradation and visual in vitro degradation in the presence of enzyme. We
(R2=0.97), N-MAC 25
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have acquired the following fitted equations, respectively: N-MAC 19
= 1 ‒ exp(-0.00201t) (R2=0.96), N-MAC 28
FR
FR
= 1 ‒ exp(-0.00455t)
FR
= 1 ‒ exp(-0.00125t)
(R2=0.94). We acquired similar k value between gravimetric determination (5.27×10-3, 2.45××10-3 and 1×10-3) and visual determination (4.55×10-3, 2.01××10-3 and 1.25×10-3), which confirmed a
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close correlation between the strategy of fluorescent imaging and gravimetric degradation. The strategy of fluorescent imaging would potentially provide a reliable platform to predict degradation kinetics of injectable biomaterials.
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To demonstrate the feasibility that the strategy of fluorescent imaging indeed reflected real
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hydrogel degradation profile, we compared the correlation of classical gravimetric degradation, CNDs release and visual determination (Figure 5C). It was noteworthy that hydrogel degradation from gravimetric degradation coincided with CNDs release profile and visual determination for all time points (from 54 to 504 h). No statistically significant differences were observed in gravimetric degradation, CNDs release and visual determination for all time points. These matching trends between gravimetric degradation and CNDs release suggested that CNDs release could be used as an indication of hydrogel degradation. The close correlation between visual 23
ACCEPTED MANUSCRIPT determination and gravimetric degradation further demonstrate that the strategy of fluorescent imaging would potentially provide a reliable platform to monitor hydrogel degradation in vivo by real-time and non-invasive fluorescence tracking.
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3.4. Visual in vivo Degradation of CNDs Hybrid Hydrogel by Real-time and Non-invasive Fluorescence Tracking
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Fluorescence is a direct method that allows the imaging of a fluorescent probe in tissue in
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vivo. In vivo fluorescence imaging is possible up to a depth of several centimeters, limited by photon absorption, scattering and diffusion. The use of red emission wavelengths allows deeper penetration than other wavelengths. Our experiments were performed in shaved mice to further minimize autofluorescence background from fur (including keratin, porphyrins, collagen and
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elastin) [69]. To evaluate the tissue penetration ability of CNDs at wavelength of 590 nm, the fluorescent images of CNDs hybrid hydrogel on the top of the chicken chip were obtained by allowing 590 nm light pass though chicken chip with different thickness (Figure 6A). Bright
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fluorescent derived from CNDs hybrid hydrogel was observed before placing chicken chip (0 mm)
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under the excitation of 590 nm light. Strong fluorescent emission could still be observed with the tissue coverage of 2 or 5 mm thickness (reduced by 15~30%), demonstrating relatively deep tissue penetration capability of CNDs at wavelength of 590 nm. However the fluorescent sharply reduced by 60% with the tissue coverage of 7 mm thickness (Figure 6B). The extinction coefficient (ε) of CNDs was determined as 1.36 cm2·g-1 according to Beer-Lambert Law (Figure S5). Overall, the good biocompatibility, low photobleaching and relatively deep tissue penetration would enable CNDs to serve as promising fluorescent indicator for long-term in vivo hydrogel 24
ACCEPTED MANUSCRIPT tracking.
To investigate visualization and quantitation of in vivo degradation, CNDs hybrid hydrogels
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different DS were subcutaneously injected and transdermally UV cured to distinguish in vivo fate of hydrogels. The CNDs provided necessary contrast between embedded hybrid hydrogels and surrounding tissues to document shape and location of hydrogel implant during degradation
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process. Mice subcutaneously injected with CNDs hybrid hydrogels were analyzed via qualitative
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(visualization) and quantitative (fluorescence reduction) strategy. To evaluate the effect of CNDs on UV light penetration (UV crosslinking ability), hydrogels with or without CNDs were prepared. UV curing depth of hydrogel with or without CNDs was 1.4 cm and 1.8 cm, respectively (Figure S6). It suggested that although the UV light penetration was indeed inhibited, the UV curing depth
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of 1.4 cm could meet the requirement of transdermal curing. Based on the above in vitro visual determination that provided a reliable and quantitative relationship between fluorescence reduction and hydrogel degradation, we further applied this visual determination in degradation of
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hydrogel in vivo. As depicted in Figure 7A, all CNDs hybrid hydrogels remained localized to the
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site of subcutaneous injection. A distinct decay of the fluorescence signal in vivo was observed from the hydrogels over degradation time. Since the CNDs embedded in hydrogels as the fluorescent indicator were low photobleaching (almost no fluorescence reduction during irradiation), the signal attenuation in area of interest (AOI) should be ascribe to CNDs release induced by hydrogel degradation and rapid diffusion away from the site of AOI. Qualitative analysis of in vivo fluorescence images with different CNDs hybrid hydrogels indicated the differences in degradation kinetics and more signal attenuation in AOI with lower DS: the 25
ACCEPTED MANUSCRIPT decrease of chemical cross-linked networks density resulted in faster degradation. However the fluorescence signal from mice treated with CNDs solution sharply reduced and even disappeared after short time (72 h) of CNDs injection, suggesting that free CNDs or released CNDs from
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hybrid hydrogels could rapidly diffuse away from the subcutaneous injection site and could not make noise signal for long-term visual in vivo degradation of hydrogel. Furthermore quantitative fluorescence reduction over time would provide reliable assessment of hydrogels degradation as
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shown in Figure 7B. It was clearly demonstrated that N-MAC 19 exhibited a faster degradation
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compared with N-MAC 25 and N-MAC 28, which consistent with the trends of in vitro degradation. After 288 hours, about 87%, 63% and 58% of the initial hydrogels was biodegraded in the case of N-MAC 19, N-MAC 25 and N-MAC 28, respectively. It was well known that in body environment, the implantation of biomaterials would activate foreign body response, which
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is correlated to the release of various enzymes (including collagenase, lipases and lysozyme) and other bio-reactive intermediates that could accelerate degradation of the implanted biomaterial [70, 71]. It should be noted that the quantitative in vivo fluorescence reduction demonstrated a close
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correlation with in vitro degradation behavior (including gravimetric and visual determination) but
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with faster rate of degradation probably due to synergetic degradation with a variety of enzymes in vivo. These degradation results were consistent with previous literatures [8] that a higher chemical cross-linked networks density limits the in vivo bioresorption and degradation rate and lower cross-linked networks density may account for disproportionately fast bioresorption kinetics. To further confirm the feasibility and veracity of visual method for assessment of in vivo degradation, mice were sacrificed after 0 h, 72 h and 288 h and injection sites were surgically dissected to exposure the CNDs hybrid hydrogel with adjacent skin (Figure 7C, up). The gross 26
ACCEPTED MANUSCRIPT appearance demonstrated that CNDs hybrid hydrogels were significantly visible and did not reveal a series of alterations in the microvasculature including redness, swelling and bleeding [72], indicating serious inflammatory response was hardly observed near the hybrid hydrogels injection
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site. The fluorescence pseudo-color images of CNDs hybrid hydrogel with adjacent skin were shown in (Figure 7C, down). It was found that the fluorescence signal gradually attenuated over the time of in vivo degradation, which is consistent with the in vivo optical imaging results shown
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in (Figure 7C, up). It should be noted that the intense fluorescence signal demonstrated a well
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overlap with optical imaging and exhibited an obvious distinction with the surrounding skin tissue. In order to verify that the visual determination method indeed reflected real in vivo hydrogel degradation profile, the correlation between classical gravimetric degradation and visual determination was compared parallelly (Figure 7D). The weight loss results were plotted as
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percentage of the hydrogel degradation versus time and correlated with the fluorescence reduction results obtained from the visual determination. It was noteworthy that results from hydrogel weight loss coincided with that from visual determination for all time points (72, 192 and 288 h).
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No statistically significant differences were observed between gravimetric degradation and visual
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determination, further confirmed the reliability and accuracy using CNDs as fluorescent indicator for long-term visual in vivo degradation of hydrogel.
To verify that this visual determination method could be applicable to various biomaterial,
CNDs hybrid gelatin hydrogel and alginate hydrogel was injected into subcutaneous space using the above procedure for visual in vivo degradation. As depicted in Figure S7, the CNDs hybrid gelatin hydrogel could almost completely degrade after 150 h, however, the CNDs hybrid alginate 27
ACCEPTED MANUSCRIPT hydrogel could completely degrade after 350 h. It was clearly demonstrated that different fluorescence reduction time between gelatin hydrogel and alginate hydrogel indeed depended on the property of material itself.
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Recently fluorescence-related imaging technique has been employed for non-invasive in vivo tracking of biomaterial degradation (Table S1) [7, 8, 19, 22, 73-75]. For example, the strategies of covalently bonding organic fluorescent molecule (such as Fluorescein-5-carboxyamido hexanoic
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acid [8], FITC [75], Rhodamine B [7], Alexa Fluor 546 [74] and IR-Dye 800CW maleimide [19])
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to the biomaterials have been adopted to track and quantify the in vivo degradation of hydrogels or fibrin sealant. However, the strategy based on covalently immobilized fluorescent molecule to biomaterials suffers from some intractable issues, including high photobleaching for long-term in vivo hydrogel tracking and uncertain perturbation of degradation profile due to the change in
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molecular structure of hydrogels. There were some reports of tracking hydrogel degradation non-invasively though directly embedding lanthanide-doped rare-earth upconversion nanoparticles (such as silica-coated LiYF4: Yb/Tm UCNPs [22] and polyacrylic acid-coated NaYF4: Yb/Tm
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UCNPs [73]) into hydrogel, avoiding the need for chemical bonding with hydrogels matrix.
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However, the synthesized UCNPs had to be further decorated with oleic acid and silica (or polyacrylic acid) to improve dispersion and biocompatibility, respectively. And the potential in vivo biotoxicity of UCNPs nanoparticles due to long-term retention in the liver and spleen system remains uncertain and more systematic investigations are still required [48, 76]. Thus, these outstanding properties of low photobleaching and good biocompatibility make CNDs promising luminescent nanomaterials for in vivo imaging compared to conventionally employed organic dyes and rare-earth nanoparticles. 28
ACCEPTED MANUSCRIPT In order to evaluate the in vivo degradation and biocompatibility of the CNDs hybrid hydrogel, histopathological analysis of the host tissues surrounding hydrogels was performed. For gross observation of CNDs hybrid hydrogel in vivo, the injection sites were surgically dissected to
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exposure the CNDs hybrid hydrogel with adjacent skin. The gross appearance of CNDs hybrid hydrogel demonstrated that the CNDs hybrid hydrogel was tightly adhered to the injection regional skin and did not show acute inflammation, redness, bleeding and swelling on the skin.
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The results of H&E staining of the surrounding skin tissues at 72, 120, 192 and 288 h after
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subcutaneous injection were shown in Figure 8 and CNDs hybrid hydrogel was indicated by eosin-staining (marked with asterisks). At the first 72 and 120 h post-injection, increased number of inflammatory cells was observed, indicating an acute inflammatory reaction surrounding the eosinophilic hydrogel in the initial stage. Hence, the results indicated that the CNDs hybrid
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hydrogel displayed acceptable biocompatibility in vivo, suggesting that the degradation of hydrogel was consistent with visual determination result. Overall the CNDs can be used as indicator for the monitoring of degradation of hydrogels, because the CNDs were characterized by
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red fluorescence emission, low photobleaching and good biocompatibility. In addition, good
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homogeneity of CNDs in hydrogels and the feature that the embedded CNDs in hydrogels did not diffuse outside in the absence of hydrogel degradation, also enable CNDs to be used as an indicator for the monitoring of degradation of hydrogels by real-time and non-invasive tracking (Figure S8).
In vivo fluorescence imaging of hydrogel degradation would enable the reduction of animal numbers in comparison with conventional invasive methods in which tissue sample at each time-point depended on the sacrifice of larger numbers of animals. Therefore, in vivo fluorescence 29
ACCEPTED MANUSCRIPT imaging was in accordance with the ‘3Rs’ principle of replacement, reduction and refinement in animal studies. So a universal method was established to follow and quantify degradation of injectable hydrogel by non-invasive tracking using CNDs as fluorescent indicator. The use of in
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vivo fluorescence imaging was a simple and cost-effective method which allowed continuous analysis of the process of hydrogel degradation and further allowed the selection and optimization
factors or the combination with other biomaterials.
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of injectable hydrogel in tissue engineering, including the addition of cells as well as growth
4. CONCLUSION
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We explored CNDs as fluorescent indicator for real-time and non-invasively tracking in vitro/in vivo degradation of hydrogels by fluorescence imaging. The direct embedding of
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photoluminescent CNDs allowed low photobleaching, red emission and good biocompatibility and avoid the need for chemically bonding. The CNDs embedded in hydrogels did not diffuse outside in the absence of hydrogel degradation. We established mathematical equation of in vitro
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degradation for quantitatively depicting degradation profile of hydrogel and a close correlation
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(similar degradation kinetics) between visual determination and gravimetric determination was acquired. This in vitro visual determination could also be expanded to in vivo degradation of hydrogels by real-time and non-invasive fluorescence monitoring. A visual platform which quantitatively depicts the degradation profile of biodegradable hydrogel has been developed and can be applied to subcutaneous degradation of injectable hydrogel with down to 7 mm depth in small animal trials. This fluorescence-related visual imaging methodology would hold great potential for tracking and location in live tissue, providing a multifunctional theranostic platform. 30
The authors thank the financial support from National Science Foundation of China (51372051, 51621091), State Key Laboratory of Urban Water Resource and Environment of
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Harbin Institute of Technology (2016TS03), HIT Environment and Ecology Innovation Special Funds (HSCJ201623), Innovation Talents of Harbin Science and Engineering (2013RFLXJ023) and Fundamental Research Funds for Central Universities (HIT.IBRSEM.201302). F.X. was
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11532009).
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supported by the National Natural Science Foundation of China (11372243, 11522219, and
Appendix A. Figures with essential colour discrimination Certain figures in this article, particularly Figures 1~8 are difficult to interpret in black and white. The full colour images can be found in the online version.
Appendix B. Supplementary data Supplementary data related to this article can be found at…… 31
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Figures Captions:
Scheme 1. Schematic illustration of low photobleaching, red fluorescence emission and good
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biocompability CNDs for visual in vitro/in vivo degradation of injectable hydrogel by real-time and non-invasive fluorescence tracking. The visual determination (replacing conventional gravimetric determination) using CNDs as fluorescent indicator was performed for monitoring in
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vitro degradation of hydrogel. This visual determination could also be expanded to quantitatively
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assessment of in vivo degradation of hydrogels by real-time and non-invasive fluorescence tracking.
Figure 1. Characterization and in vitro cytotoxicity of CNDs. (A) TEM images of the CNDs
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(mean diameter = 5.1 ± 1.2 nm) with high-magnification HR-TEM images, the insert displays the fast-Fourier-transformed diffraction pattern. AFM topography image of CNDs on a silicon substrate, with the height profile along the line in the topographic image; (B) UV-vis absorbance
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spectra of CNDs diluted suspension. Inset shows photographs of CNDs suspension under day light
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(left) and UV light at 365 nm (right); (C) Excitation-dependent photoluminescence spectra of CNDs. PL emission wavelength shifts to longer wavelength (from 430 to 580 nm) as excitation wavelength increased from 300 to 460 nm. The maximum emission is 450 nm when using 375 nm excitation; (D) Cell viability of NIH/3T3 cells after incubation with different concentrations CNDs for 24, 36 and 48 h, determined by MTT assay. The cells viability with various concentrations maintains all above 90%; (E) Merge image (fluorescence and bright field) of NIH/3T3 cells treated with CNDs. 37
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Figure 2. In vivo toxicity assessment of CNDs. (A) Fluorescence imaging of various organs (including liver, spleen, kidney, heart and lung) at 1 and 14 days. Mice treated with normal saline
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solution (without CNDs) were control group; (B) Quantification of the fluorescence intensity from various organs (including liver, spleen, kidney, heart and lung); (C) H&E stained tissue slices (liver, spleen, kidney, heart and lung) of mice injected with CNDs solution (dose of 23 mg/kg) at 1
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Figure 3. CNDs serving as fluorescent indicator for visualization of CNDs hybrid hydrogel. (A) Quantitative fluorescence intensity of CNDs hybrid hydrogel versus CNDs concentration. Inset shows fluorescent pseudo-colored image of hydrogel with different CNDs concentrations; (B)
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Fluorescence images of patterning CNDs hybrid microgels of concentric ring pattern (left) and convex pattern (right), exhibiting green and red fluorescence under different excitation (460~490 and 540~560 nm); (C) Photobleaching experiment of CNDs hybrid hydrogel. Fluorescent
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pseudo-colored image taken in every 5 min interval showed photobleaching of CNDs hybrid
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hydrogel or FITC hybrid hydrogel when exposed to 100W Xenon excitation source; (D) A specific region of interest (ROI) was selected for quantitative calculation of fluorescence intensity. CNDs as the fluorescent indicator were much more photostable than fluorescent dye (such as FITC and RhB) and much more suitable for in vivo visualization research; (E) Confocal microscopy images of CNDs hybrid hydrogel: the representative x-z plane image, the representative x-y plane images and fluorescence intensity profile at different depth. The red field was excited with a 568 nm laser and emissions were filtered with a 605 nm band-pass filter. 38
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Figure 4. In vitro degradation of CNDs hybrid hydrogel. (A) Gravimetric degradation of CNDs hybrid hydrogel and (B) release behavior of CNDs in the absence of lysozyme or in the presence
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of lysozyme over a period of 500 hours. The CNDs encapsulated inside hydrogels did not diffuse outside in the absence of hydrogel degradation (NO degradation and NO CND release). In vitro
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mathematical equation was established to quantitatively depict degradation profile.
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Figure 5. In vitro degradation of CNDs hybrid hydrogel by visual determination. (A) Representative fluorescent image of in vitro degradation of CNDs hybrid hydrogel (N-MAC 19, N-MAC 25 and N-MAC 28) in the absence of lysozyme or in the presence of lysozyme over a period of 500 hours; (B) Quantitative fluorescence reduction of hybrid hydrogels (N-MAC 19,
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N-MAC 25, and N-MAC 28) in vitro degradation in the absence of lysozyme or in the presence of lysozyme as a function of time; (C) Correlation graphs of degradation of CNDs hybrid hydrogel
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Figure 6. Tissue penetration evaluation of CNDs at wavelength of 590 nm for visual in vivo degradation of hydrogel. (A) Illustration of the experimental setup used to estimate the tissue penetration ability of CNDs at wavelength of 590 nm. The CNDs hybrid hydrogel was placed on the top of chicken chip with varying thickness (2~7 mm); (B) The fluorescence images (pseudo-color images) of CNDs hybrid hydrogel and quantitative fluorescence intensity. The excitation wavelength, emission wavelength and fixed exposure time were set as 590 nm, 700 nm and 20s, respectively. 39
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Figure 7. Visual in vivo degradation of CNDs hybrid hydrogel by real-time and non-invasive fluorescence tracking. (A) Representative in vivo pseudo-colored images of CNDs hybrid
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hydrogel (N-MAC 19, N-MAC 25, and N-MAC 28) via subcutaneous injection over 288 hours. A gradual attenuation of fluorescence signal was observed in all hybrid hydrogels with different rates. Mice treated with free CNDs (CNDs solution without hydrogel) were control group; (B)
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Quantitative in vivo degradation via fluorescence reduction of CNDs hybrid hydrogels (N-MAC
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19, N-MAC 25, and N-MAC 28) as a function of time; (C) The gross appearance (up) and corresponding fluorescence pseudo-colored images (down) of CNDs hybrid hydrogel with adjacent skin after 0 h, 72 h and 288 h. The CNDs hybrid hydrogels were visible and did not reveal redness, swelling and bleeding. Fluorescence signal demonstrated a well overlap with
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optical imaging and exhibited an obvious distinction with surrounding skin tissue; (D) Correlation of CNDs hybrid hydrogel degradation quantified by applying non-invasive visual determination
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Figure 8. The Gross appearance and H&E stained histological analysis of subcutaneous tissue with the CNDs hybrid hydrogel at 72, 120, 192 and 288 h post-injection and the CNDs hybrid hydrogel was indicated by eosin-staining (marked with asterisks).
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Highlight We first employed photoluminescent carbon nanodots (CNDs) as fluorescent indicator with low photobleaching, red emission and good biocompatibility for visualization and quantitation of in vitro/in vivo degradation of hydrogels by real-time and non-invasive fluorescence tracking.
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Statement of Significance As to the field of tissue engineering, a quantitative assessment of in vivo degradation of engineered hydrogels is of great importance especially for advisable design of molecule structure
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and matching up with regeneration rate of newly generated tissue. However, fluorescence-related visual imaging usually encounters some challenge such as intrinsic photobleaching of fluorophores and uncertain perturbation of degradation induced by change in molecular structure.
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The present work is devoted to employed photoluminescent carbon nanodots (CNDs) as
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fluorescent indicator with low photobleaching, red emission and good biocompatibility for visualization and quantitation of in vitro/in vivo degradation of hydrogels by real-time and non-invasive tracking. Also in vitro mathematical equation of hydrogel degradation kinetics had been established which would open the possibility to develop predictive in vivo models for tissue
