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. Author manuscript; available in PMC: 2020 Oct 2.
Published in final edited form as: Acta Biomater. 2020 Aug 25;115:333–342. doi: 10.1016/j.actbio.2020.08.031

Photoluminescent oxygen-release microspheres to image the oxygen release process in vivo

Ya Guan a,b, Hong Niu a,b, Yu Dang a,b, Ning Gao a,b, Jianjun Guan a,b,*
PMCID: PMC7530143  NIHMSID: NIHMS1631126  PMID: 32853800

Abstract

Cell therapy is a promising strategy to treat ischemic diseases, but the efficacy is limited due to high rate of cell death under low oxygen environment of the ischemic tissues. Sustained release of oxygen to continuously oxygenate the transplanted cells may augment cell survival and improve therapeutic efficacy. We have shown previously that oxygen released from oxygen-release microspheres stimulated cell survival in ischemic tissue [1]. To understand how oxygen is released in vivo and duration of release, it is attractive to image the process of oxygen release. Herein, we have developed photoluminenscent oxygen-release microspheres where the in vivo oxygen release can be non-invasively and real-time monitored by an In Vivo Imaging System (IVIS). In the oxygen-release microspheres, a complex of polyvinylpyrrolidone, H2O2 and a fluorescent drug hypericin (HYP) was used as core, and poly(N-isopropylacrylamide-co-acrylate-oligolactide-co-hydroxyethyl methacrylate-co-N-acryloxysuccinimide) conjugated with catalase was used as shell. To distinguish fluorescent signal change for different oxygen release kinetics, the microspheres with various release profiles were developed by using the shell with different degradation rates. In vitro, the fluorescent intensity gradually decreased during the 21-day oxygen release period, consistent with oxygen release kinetics. The released oxygen significantly augmented mesenchymal stem cell (MSC) survival under hypoxic condition. In vivo, the oxygen release rate was faster. The fluorescent signal can be detected for 17 days for the microspheres with the slowest oxygen release kinetics. The implanted microspheres did not induce substantial inflammation. The above results demonstrate that the developed microspheres have potential to monitor the in vivo oxygen release.

Keywords: Ischemic vascular disease, Oxygen release, Photoluminescence, In vivo imaging, Cell therapy

1. Introduction

Ischemic vascular diseases (IVDs), such as heart attack, stroke and critical limb ischemia, affect more than 20 million people in the United States, with the highest mortality rate among all diseases [2,3]. IVD occurs when accumulated plaques clog the blood vessels and restrict the blood flow. The poor blood perfusion creates an ischemic microenvironment, where the oxygen and nutrients supply is limited, leading to cell death and tissue necrosis [4–6]. Thus, the goal of treating IVD is to restore the blood flow, and promote therapeutic angiogenesis and tissue regeneration. Cell therapy has been considered as a promising approach to treat IVD [7–10]. Various cell types, such as mesenchymal stem cells (MSCs), endothelial progenitor cells and hematopoietic stem cells, have been confirmed to improve blood perfusion, and stimulate muscle repair in mice by differentiation into vascular cell types and/or paracrine effects [7–11]. Despite the promising results from animal studies, clinical trials with cell transplantation have shown limited efficacy [12,13]. One of the key problems is the poor cell survival and engraftment in ischemic environment.

Several approaches have been explored to enhance the cell survival after transplantation. Genetic modification of the cells with anti-apoptotic and prosurvival genes has been shown to increase cell survival after transplantation into the infarcted heart [14,15]. However, direct genetic modification may raise safety concerns and is associated with a potential risk of cancer [16]. Transplantation of cells with angiogenic growth factors is another approach to enhance cell survival in ischemic tissues [17–19]. The angiogenic growth factors act to stimulate tissue angiogenesis. The regenerated vessels provide oxygen and nutrient for cell survival. Yet this approach experiences low cell survival before angiogenesis is established [20]. Hydrogels that mimic properties of the extracellular matrices have been used to augment cell survival in ischemic tissues [21–25]. These hydrogels improve cell-matrix adhesion, and avoid anoikis-mediated death [21,23]. However, long-term survival of the transplanted cells remains a problem, because the low oxygen environment compromises normal cell metabolism. To address the above hurdles, one of the strategies is to use hydrogels capable of releasing oxygen to deliver cells. The hydrogels will enhance cell-matrix interaction to increase cell survival, while released oxygen will oxygenate the cells to enhance their survival.

Different compounds have been explored for controlled oxygen release. These include MgO2 [26], CaO2 [27], H2O2 [28,29], and fluorinated molecules [30,31]. The methods of using MgO2, CaO2 and H2O2 rely on H2O2 decomposition to generate oxygen. One of the disadvantages of these methods is that the released free H2O2 may damage the cells if it is not quickly decomposed. MgO2 and CaO2 will also release Mg2+ or Ca2+. These ions may lead to abnormal ion transient in muscular tissues [32]. Fluorinated molecules can capture oxygen, and provide it to the transplanted cells [30,33]. Overall, current oxygen release approaches have relatively short oxygen release duration, typically < 2 weeks [26–31]. Therefore, they cannot support long-term cell survival. To address this limitation, we have previously used polyvinylpyrrolidone (PVP) to bind to H2O2 to form a high molecular weight complex. The complex was able to gradually release from poly(lactide-co-glycolide) microspheres for 4 weeks, and the H2O2 in the complex was converted into oxygen by catalase [1,34]. One of the disadvantages of this oxygen release approach is that the released PVP/H2O2 relies on free catalase in the microsphere carrier to convert into oxygen. The release of catalase from the carrier may lead to incomplete conversion of H2O2 causing damages to cells.

When the oxygen-release system and cells are transplanted into ischemic tissues, it is attractive to monitor the release process so as to evaluate duration of the oxygen release, and whether it is necessary to apply extra oxygen-release biomaterials into the ischemic tissues to augment long-term cell survival. Hence, it is necessary to develop an approach capable of monitoring the release process in real-time. Current approaches, such as transcutaneous oximetry [35], pulse oximetry [36], fiber optic O2 probes [37], polarographic needle O2 electrode [38], and magnetic resonance-based methods magnetic resonance imaging and electron paramagnetic resonance [39–42], can be used to determine tissue oxygen content. Yet they cannot readily distinguish endogenous and exogenous oxygen. It is therefore necessary to develop a direct approach that only monitors oxygen released from the biomaterials.

In this work, we developed our microspheres where the molecular oxygen instead of H2O2 can be directly released out, and the release can be monitored by fluorescent signal change using an In Vivo Imaging System (IVIS). The microspheres were based on photoluminescent core PVP/H2O2/hypericin (HYP) complex, and biodegradable shell immobilized with catalase (Scheme 1). The catalase allows the H2O2 in the released complex to be timely converted into oxygen at the shell surface. The release of complex from the microspheres leads to fluorescent signal intensity decrease. HYP is a photo-stable molecule with fluorescence emission in orange/red regions and minimal toxicity [43]. It is also recognized as an antiviral, antibacterial, and antineoplastic photosensitizer [44,45]. A family of polymer shells with different degradation rates was synthesized to modulate oxygen release kinetics. The fluorescent signal change during the oxygen release both in vitro and in vivo, cytotoxicity of the core and shell material, biocompatibility of the microsphere were evaluated. The efficacy of oxygen-release microspheres in promoting cell survival under hypoxic condition was determined.

Scheme 1.

Scheme 1.

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Design of the oxygen-release microspheres that release molecular oxygen and can be monitored after in vivo implantation.

2. Materials and method

2.1. Materials

All materials were purchased from Sigma-Aldrich unless otherwise stated. N-Isopropyl acrylamide (NIPAAm, TCI) was recrystallized in hexane 3 times before polymerization. 2-Hydroxyethyl methacrylate (HEMA, Alfa Aesar) was passed through a column filled with inhibitor removers. Polyvinylpyrrolidone (PVP, 40 kDa, Fisher Scientific), hypericin (Beantown chemical), bovine liver catalase (2000–5000 units/mg), hydrogen peroxide (30% w/w in water, Fisher Scientific) and tris-(2,2’-bipyridine) ruthenium (II) chloride (Ru(bpy)2Cl2, GFS chemicals) were used as received.

2.2. Polymer synthesis

The degradable vinyl monomer acrylate-oligolactide (AOLA) was synthesized as previously reported [1]. The resulting monomer had four lactide units as determined from 1H-NMR. N-Acryloxysuccinimide (NAS) was synthesized by reacting N-hydroxysuccinimide (NHS) with acryloyl chloride in the presence of trimethylamine (TEA). Both AOLA and NAS were lyophilized to remove residual water before polymerization. The polymers used as shell of the microspheres were poly(NIPAAm-co-AOLA-co-HEMA-co-NAS) (abbreviated as PNAHN). They were synthesized by free radical polymerization using benzoyl peroxide as initiator [1,33,46,47]. The reaction was performed at 70°C under the protection of nitrogen for 20 h. The polymer was pre-cipitated in hexane and purified 3 times using tetrahydrofuran and ethyl ether. Three polymers were synthesized. The feed ratios of NIPAAm/AOLA/HEMA/NAS were 50/25/5/20, 58/17/5/20 and 65/10/5/20, respectively. Hydrogel poly(NIPAAm-co-AOLA-co-HEMA) (abbreviated as PNAH), was synthesized using the same polymerization method. The feed ratio of NIPAAm/AOLA/HEMA was 86/4/10. This hydrogel was used to deliver microspheres into tissues.

To determine properties of the final degradation products of the microsphere shell, they were synthesized by free radical polymerization of NIPAAm, acrylic acid, HEMA and NAS. The feed ratios were 50/25/5/20, 58/17/5/20 and 65/10/5/20, respectively. The polymers were then reacted with glycine to quench the succinimide group.

2.3. Characterization of polymers

Structure of the polymers was confirmed by 1H-NMR. Polymer composition was calculated from the spectra. The degradation of PNAHN was conducted in Dulbecco’s phosphate-buffered saline (DPBS) for 6 weeks. Briefly, the polymer was dissolved in dichloromethane, and added into a 1.5 mL centrifuge tube. After solvent evaporation, 200 μL of DPBS was added. The tube was placed in a 37°C water bath. At each time point, the samples (n = 4) were taken out, washed with DI water, and lyophilized. The sample weight was then measured. The percentage of weight remaining was calculated as weight at each time point normalized to the weight before degradation.

Cytotoxicity of the final degradation products of the microsphere shell was tested in terms of their capability of preserving mesenchymal stem cell (MSC) viability. Rat bone marrow-derived MSCs were cultured in minimal essential medium alpha (αMEM) supplemented with 10% fetal bovine serum (FBS) and 1% penicillin-streptomycin at 37°C under 21% O2 and 5% CO2. The cells were seeded in a 96-well tissue culture plate at a density of 105 cells/mL. After 24 h, the final degradation product solution was added to each well. The final concentration was 10 or 20 mg/mL (n = 5 for each concentration). The wells without final degradation product were used as control. After culturing for 48 h, cell viability was determined by MTT assay [48].

2.4. Microsphere fabrication and characterization

The oxygen-release microspheres were fabricated using PVP/H2O2/HYP complex as core, and PNAHN as shell. The shell was further conjugated with catalase so as to timely convert H2O2 in the released PVP/H2O2/HYP complex into molecular oxygen. PVP/H2O2/HYP complex was formed by first mixing H2O2 and PVP under vigorous stirring at 4°C overnight. The ratio of H2O2/VP was 4.5. HYP was then added to the PVP/H2O2 solution at a concentration of 5, 10, or 30 mgHYP/gPVP. The mixture was stirred overnight in a 4°C refrigerator. The mixture was then dialyzed in DI water for 3 days to remove un-complexed HYP. The water was changed every 12 h. To confirm the formation of PVP/H2O2/HYP complex, emission bands were determined using SpectraMax iD3 spectrometer (Molecular Devices). The excitation wavelength of 555 nm was used. Following catalase conversion of H2O2 in the PVP/H2O2/HYP complex into molecular oxygen, PVP/HYP complex is left. To determine cytotoxicity of the PVP/HYP complex, different concentration (5, 10, or 30 mgHYP/gPVP) of complex was added into the MSC culture medium. Cell viability was determined by MTT assay following the method described above (n =5 for each concentration).

To fabricate microspheres, a coaxial electrospraying technique was used [1,34]. In the coaxial device, the inner stream was PVP/H2O2/HYP complex with H2O2/VP ratio of 4.5, and HYP/PVP ratio of 30 mgHYP/gPVP. The outer stream was 5 wt% PNAHN in dichloromethane. The two streams were charged at a voltage of +15 kV. The flow rates for the PVP/H2O2/HYP complex and PNAHN solution were 0.2 and 1 mL/h, respectively. The microspheres were collected on a rotating mandrel covered with an aluminum foil, and charged at a voltage of −10 kV. The microspheres were lyophilized, and stored at −20°C before use. To confirm the core-shell structure, the PNAHN solution was mixed with rhodamine B. The fluorescent images were taken by a confocal microscope (Olympus FV1000). Scanning electron microscopy (SEM) was used to determine morphology of the microspheres.

Catalase was immobilized onto the surface of the microspheres by reacting with the NHS groups in NAS component of the shell polymer. The microspheres were suspended in DPBS at a concentration of 10 mg/mL. Catalase solution (5 mg/mL) was then added to the suspension. The mixture was stirred at 4°C for 4 h. After centrifugation, the collected microspheres were washed with DI water followed by lyophilization. To confirm the conjugation of catalase on the microspheres, the catalase was pre-labeled by fluorescein isothiocyanate (FITC). The fluorescent images of the microspheres were taken by a confocal microscope.

2.5. Oxygen release kinetics

The microsphere oxygen release was tested in 37°C DPBS under 1% O2 condition for 21 days [1,34]. Briefly, the oxygen-release microspheres were suspended in DPBS under 1% O2 environment. The final concentration was 50 mg/mL. 200 μL of mixture was then added into a 96-well plate (n = 8 for each group). The bottom of well was covered by a polydimethylsiloxane (PDMS) membrane loaded with an oxygen-sensitive luminophore Ru(bpy)2Cl2, and an oxygen-insensitive dye rhodamine-B. The latter dye was used as a reference. The highly oxygen permeable PDMS membrane allows the released oxygen to diffuse into the membrane and interact with Ru(bpy)2Cl2. The 96-well plate was sealed. The oxygen release was conducted in a 1% O2 incubator at 37°C. At each time point, the fluorescent intensities were measured for both Ru(bpy)2Cl2 and rhodamine B using a fluorescent plate reader (Molecular Devices). A calibration curve was used to convert the fluorescence intensity to oxygen level. The measured oxygen content at each time point was the real-time oxygen content.

2.6. In vitro fluorescence measurement

To determine the HYP signal change within the microspheres during oxygen release, fluorescent intensity was measured, and fluorescent images were taken at each time point. The microspheres were suspended in DPBS at a concentration of 50 mg/mL under 1% O2 condition. 200 μL of suspension was added into a 96-well plate (n = 5 for each group). The plates were placed in a 37°C, 1% O2 incubator. At each time point, the supernatant was removed, and the fluorescent intensity of HYP in the microspheres was measured by a fluorescent plate reader. Fluorescent images of the microspheres were taken by a confocal microscope.

2.7. Effect of oxygen release on cell survival under hypoxia

To investigate the efficacy of released oxygen in promoting cell survival under low oxygen condition, the oxygen-release microspheres and bone marrow-derived MSCs were encapsulated in PNAH hydrogel and cultured under 1% O2 condition for 2 weeks. MSCs were chosen because they have the potential to stimulate angiogenesis and muscle repair in ischemic tissues [10,15,25,49]. In brief, 50 mg/mL of microspheres and 2 million/mL of MSCs were mixed with 6 wt% PNAH hydrogel. The mixture was placed at 37°C for gelation for 1 h. αMEM without FBS was then added. The constructs were cultured in a 37°C incubator with 1% O2 (n = 4 for each group). At days 0, 3, 7 and 14, the MSCs were digested by pa-pain solution. The double stranded DNA (dsDNA) content was measured by Picogreen dsDNA assay (Invitrogen). To visualize the live cells, MSCs were pre-labeled with a live cell tracker CM-Dil (Invitrogen) before encapsulation into the hydrogel. Fluorescent images were taken by a confocal microscope.

2.8. Injection of oxygen-release microspheres, MSCs, and hydrogel into thigh muscles

All animal care and experiment procedures were conducted in accordance with the National Institutes of Health guidelines. The animal protocol was approved by the Institutional Animal Care and Use Committee of the Ohio State University. To determine microsphere fluorescent intensity change after implantation, 100 μL of 6 wt% PNAH hydrogel solution with or without 50 mg/mL microspheres was injected into thigh muscles of 8-week-old NCr nude mice (Taconic Biosciences. n =4 for each group). At days 0, 3, 7, 10, 14, 17 and 21, the mice were anesthetized with isoflurane, and imaged using an In Vivo Imaging System (IVIS) Lumina II (PerkinElmer) with DsRed emission filter.

To determine biocompatibility of the microspheres, 200 μL of 6 wt% PNAH hydrogel solution with or without 50 mg/mL microspheres was injected into thigh muscles of 8-week-old C57BL/6 mice (Jackson laboratory. n =4 for each group). The muscles were harvested 3 weeks after the injection. The tissues were fixed with 4% paraformaldehyde overnight, processed and embedded in paraffin. Tissue sections of 5-μm thickness were cut, and stained with mouse monoclonal anti-F4/80 (Santa Cruz Biotechnology). Nuclei were stained by DRAQ5. The cells were imaged using a confocal microscope. F4/80 positive cell density was quantified by ImageJ.

2.9. Statistical analysis

All results were presented as mean ± standard deviation. Comparison among groups was performed using two-way ANOVA with post hoc Tukey test. Difference was considered statistically significant when p<0.05.

3. Results

3.1. Synthesis and characterization of the polymers used as microsphere shell

The PNAHN polymers for microsphere shell were based on NIPAAm, AOLA, HEMA and NAS (Scheme 2). The rationale of using NIPAAm, AOLA and HEMA is that hydrogels based on NIPAAm, AOLA and HEMA had good biocompatibility [1,25,46,50]. MSCs can proliferate within the hydrogel. The degradation products were not toxic even when the concentration was as high as 50 mg/mL. The NAS component imparted the polymers with conjugation capability, allowing for immobilizing catalase on the shell to timely convert H2O2 into molecular oxygen. A family of PNAHN polymers were synthesized with different AOLA content (Table 1). The chemical structure of the polymers was confirmed by 1H-NMR spectra (representative spectrum shown in Fig. 1A). The composition of the polymers was calculated from the characteristic peaks of each component in the NMR spectra (a for NIPAAm, d for AOLA, g for HEMA, and h for NAS). The composition of each polymer was consistent with initial feed ratio (Table 1).

Scheme 2.

Scheme 2.

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Synthesis of poly(NIPAAm-co-AOLA-co-HEMA-co-NAS) (PNAHN), catalase conjugation and degradation.

Table 1.

Feed ratio and actual composition of poly(NIPAAm-co-AOLA-co-HEMA-co-NAS).

Abbreviation of polymer Feed ratioa Compositiona
N50A25 50:25:5:20 53:21:4:22
N58A17 58:17:5:20 57:16:5:22
N65A10 65:10:5:20 68:8:4:19
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a

Molar ratio of NIPAAm:AOLA:HEMA:NAS

Fig. 1.

Fig. 1.

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Characterizations of the polymer shell. (A) Representative 1H-NMR spectrum of poly(NIPAAm-co-AOLA-co-HEMA-co-NAS); (B) Degradation of poly(NIPAAm-co-AOLA-co-HEMA-co-NAS) with different AOLA content; (C) Viability of MSCs cultured in the medium treated with the final degradation product of the shell. Cell viability was normalized to the control group without treatment. ** p<0.01, ***p<0.001.

The 3 polymers were insoluble in DPBS when tested at temperatures ≥ 4°C. To determine degradation property, the polymers with different AOLA content were incubated in 37°C DPBS. After 6 weeks, the weight loss was 16–29% (Fig. 1B). The weight loss was dependent on the AOLA content. The increase of AOLA content augmented the weight loss. At week 6, the polymer (N50A25) with the highest AOLA content had significantly greater weight loss than the polymers (N58A17 and N65A10) with lower AOLA content (p<0.01).

To test cytotoxicity of the final degradation products, poly(NIPAAm-co-acrylic acid-co-HEMA-co-NAS) with 3 NIPAAm/acrylic acid/HEMA/NAS ratios were first synthesized. The rationale of using acrylic acid is that AOLA component forms acrylic acid component after degradation of oligolactide (Scheme 2). The 3 polymers were further reacted with glycine to quench NHS group in NAS component. The polymers were added to the MSC culture medium with final concentrations of 10 and 20 mg/mL, respectively. MTT assay demonstrated that all groups had similar cell viability (p>0.05. Fig. 1C). These results indicate that the 3 final degradation products were nontoxic even at a relatively high concentration, 20 mg/mL.

3.2. Preparation and characterization of PVP/H2O2/HYP complex

To allow H2O2 release to be sustained and imageable, it was complexed with PVP together with photoluminescent HYP. Both H2O2 and HYP can form hydrogen bonding with PVP (Scheme 1). To verify the complexing, fluorescence emission spectra of HYP and the PVP/H2O2/HYP complex were measured. Free HYP showed two emission peaks at 615 nm and 670 nm, respectively (Fig. 2A). After forming a complex with PVP and H2O2, the peak at 615 nm was shifted to 625 nm, while the peak at 670 nm became less pronounced. These results demonstrate that the wavelength 625 nm can be used to detect the complex.

Fig. 2.

Fig. 2.

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Characterizations of the PVP/H2O2/HYP complex. (A) Fluorescence emission spectra of HYP and PVP/HYP/H2O2 complex; (B) Viability of MSCs cultured in the medium treated with PVP or HYP/PVP complex. Cell viability was normalized to the control group without treatment.

After PVP/H2O2/HYP complex interacts with catalase, it becomes PVP/HYP complex. To evaluate cytotoxicity of PVP/HYP, MTT assay was performed. Three concentrations, 5, 10, and 30 mgHYP/gPVP, were used. After incubating with MSCs for 48 h, all groups had similar cell viability as the group added with PVP only, and the group without adding PVP or PVP/HYP (p>0.05 for all groups. Fig. 2B). These results demonstrate that PVP/HYP was non-toxic.

3.3. Fabrication of oxygen-release microspheres, and characterization of oxygen release kinetics

The oxygen-release microspheres were fabricated by coaxial electrospraying, with PVP/H2O2/HYP complex as core, and PNAHN as shell. The microspheres assumed core-shell structure (Fig. 3A). Morphology of the microspheres was further characterized by SEM (Fig. 3B). The microspheres exhibited average size of 4 μm. To confirm that catalase was conjugated onto the polymer shell, FITC-labeled catalase was used. Confocal image showed that catalase was conjugated to the microspheres (Fig. 3C).

Fig. 3.

Fig. 3.

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Characterizations of the oxygen-release microspheres. (A) Fluorescent images of the oxygen-release microspheres. The yellow fluorescence is HYP in the core and the red fluorescence is the rhodamine added to the shell. Scale bar = 5 μm; (B) SEM images of the oxygen-release microsphere; (C) Fluorescent image of the oxygen-release microsphere after conjugation with FITC-labelled catalase. Scale bar = 5 μm; (D) Calibration curve for calculating the oxygen release kinetics. I0 is the fluorescence intensity of rhodamine. I is the fluorescence intensity of Ru(bpy)2Cl2; (E) Oxygen release kinetics of the microspheres with different shells for 3 weeks. ***p<0.001.

In vitro oxygen release kinetics was tested using an oxygen-sensitive fluorescence dye Ru(bpy)2Cl2 embedded in PDMS membrane. PDMS is highly oxygen permeable, allowing the oxygen released from the microspheres to readily diffuse inside, and quench the luminescence of the dye [51]. Therefore, a relationship can be built between the fluorescent intensity and the oxygen content (Fig. 3D). Oxygen was able to continuously release from the microspheres (Fig. 3E). In all groups, the microspheres exhibited a burst release in the first 7 days followed by a relatively steady release till day 21. After 4 days of release, the oxygen level reached 10%, and remained above this level till day 21. The release kinetics was depended on the AOLA content in the shell. The increase of AOLA content led to release of greater amount of oxygen during day 2 and day 21.

3.4. In vitro fluorescence tracking of the oxygen-release microspheres

To investigate how the fluorescent signal in microsphere changed in response to oxygen release, the fluorescent images were taken, and the fluorescent intensity of HYP was monitored during the 21-day release period. The fluorescent images demonstrated that the fluorescent signal became weaker over time (Fig. 4A). This is confirmed by the trend of fluorescent intensity change (Fig. 4B). It was also consistent with oxygen release kinetics (Fig. 3E). The microspheres that released more oxygen exhibited weaker fluorescent signal. After 21 days, the microspheres made from N50A25, N58A17, and N65A10 retained 25%, 48%, and 53% of the initial fluorescent intensity, respectively. The microspheres based on N50A25 exhibited significantly lower fluorescent intensity than those based on N58A17 and N65A10 (p<0.01). These results provide the foundation of using fluorescence imaging to track oxygen release in vivo.

Fig. 4.

Fig. 4.

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In vitro fluorescence tracking of the oxygen-release microspheres. (A) Fluorescent images of the oxygen-release microspheres at each time point for 21 days. Scale bar = 5 μm; (B) Fluorescence intensity change of the oxygen-release microspheres in 21 days. **p<0.01.

3.5. MSC survival under hypoxia in response to oxygen released from microspheres

The purpose of fabrication of oxygen-release microspheres is to increase cell survival in ischemic tissues. To determine whether the oxygen released from microspheres can promote cell survival under hypoxia, MSCs and oxygen-release microspheres were encapsulated in PNAH hydrogel (Fig. 5A). The constructs were cultured under 1% O2 condition, and supplemented with αMEM without serum. dsDNA content was measured to quantify the cell survival (Fig. 5B). The results demonstrated that MSCs died quickly in the hydrogel without oxygen-release microspheres. More than 85% of cells died in two weeks as a result of deficient oxygen and nutrients. However, in the presence of oxygen-release microspheres, the MSCs survived and proliferated during the 14-day culture period. N50A25 group, which showed fastest oxygen release between day 2 and day 14, had the highest MSC survival rate. The live cell images at day 14 were consistent with the dsDNA results, where the N50A25 group had the highest cell density (Fig. 5C).

Fig. 5.

Fig. 5.

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Imageable oxygen-release microspheres enhanced MSC survival under hypoxia. (A) Scheme of the experimental design of the cell survival assay; (B) dsDNA content of MSCs encapsulated in PNAH hydrogel loaded with oxygen-release microspheres during 14 days of culture; (C) Live cell images of MSCs encapsulated in PNAH hydrogel loaded with oxygen-release microspheres during 14 days of culture. MSCs were pre-labeled with a live cell tracker CM-Dil (red). *p<0.05, **p<0.01, ***p<0.001.

3.6. In vivo monitoring of the oxygen-release microspheres

To monitor the release process in vivo, the oxygen-release microspheres were encapsulated into PNAH hydrogel, and injected into thigh muscles of nude mouse. PNAH hydrogel was injectable at 4°C, and solidified within 10 s at body temperature [1,34,46]. The use of fast gelation hydrogel can increase microsphere retention in tissues [1].The IVIS imaging was performed for 21 days. The DsRed emission filter with wavelengths ranging from 575 nm to 650 nm was used to detect the fluorophore. The images showed that the PNAH hydrogel retained the microspheres at the injection site (Fig. 6). The fluorescent signal became weaker for all groups with microspheres as the PVP/H2O2/HYP complex gradually released out of microspheres. Gel/N65A10 group, of which the oxygen release was the slowest, maintained the highest fluorescent signal at each time point. The signal was still visible 17 days after the implantation. The signal became invisible at day 21. In contrast, the groups with faster oxygen release (Gel/N58A17 and Gel/N50A25) more quickly decreased signal intensity. The signal for both groups were invisible at day 14. These results demonstrate that the incorporation of HYP together with PVP/H2O2 allowed the oxygen release behavior to be monitored in real-time. In addition, oxygen release was faster in vivo than in vitro.

Fig. 6.

Fig. 6.

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In vivo monitoring of the oxygen-release microspheres. IVIS images of the mouse limbs implanted with PNAH gel loaded with or without oxygen-release microspheres. DsRed was used as the emission filter.

3.7. Biocompatibility of the hydrogel and microspheres

The in vivo biocompatibility of the oxygen-release microspheres and PNAH hydrogel was evaluated 21 days after the implantation. F4/80 staining for macrophages was used to evaluate the inflammatory response (Fig. 7A). The implantation of microspheres and PNAH hydrogel did not initiate substantial inflammation. No significant difference of F4/80 positive cell density was found among untreated group, PNAH implanted group (Gel), and PNAH and microsphere implanted group (Gel/N65A10) (Fig. 7B). These results confirmed that the microspheres and PNAH hydrogel possess good in vivo biocompatibility.

Fig. 7.

Fig. 7.

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Biocompatibility of the PNAH gel and the oxygen-release microspheres. (A) F4/80 staining (green) images of the limb tissues with or without the injection of PNAH gel and oxygen-release microspheres. Nuclei were stained with DRAQ5. Scale bar = 50 μm; (B) Quantification of F4/80 positive cells.

4. Discussion

The objective of this work is to develop photoluminescent oxygen-release microspheres that not only release molecular oxygen to promote cell survival under hypoxic condition, but also allow to real-time image the oxygenation process so that duration of oxygen release in vivo can be informed. Oxygen is needed for the survival of endogenous cells in the ischemic tissues and exogenous cells transplanted into the ischemic tissues. Angiogenesis is an approach to oxygenate cells. Yet cell survival is limited before angiogenesis can be established. Direct supply of exogenous oxygen to the cells has potential to promote early stage cell survival [52]. To better understand how oxygen is released and duration of oxygen release, real-time monitoring of the release process is necessary. Yet this is challenging using currently available oxygenation measurement approaches such as transcutaneous oximetry [35], pulse oximetry [36], fiber optic O2 probes [37]and polarographic needle O2 electrode [38]. Transcutaneous oximetry and pulse oximetry are used to monitor the oxygen level in skin and blood, and are not well-suited to detect oxygen in deeper tissues. Optical oxygen sensors and polarographic needle O2 electrode can perform point oxygenation measurement, but cannot specifically monitor the oxygen release process. Unlike point oxygenation measurement, oxygenation imaging techniques such as positron emission tomography [53], magnetic resonance imaging (MRI) [39–41]and electron paramagnetic resonance imaging [42]provide real-time, non-invasive mapping of the tissue oxygen content. However, these approaches only detect the oxygen in tissues, but cannot readily distinguish the oxygen released exogenously from the oxygen in the local environment [54,55]. Use of oxygen-sensitive fluorescent dyes such as transition metal polypyridyl complexes may allow to image exogenous oxygen [51]. However, the poor solubility and high toxicity of the dyes restrict their in vivo applications.

In this work, we developed a new approach where the oxygen-generating material itself can be imaged in real-time. The PVP/H2O2/HYP complex was used as the oxygen-generating material. In the complex, HYP is photoluminescent. When the complex is released from the shell, the H2O2 is converted by catalase immobilized on the shell to generate oxygen. The remaining PVP/HYP complex is water soluble, and can thus dissolve in body fluid to diffuse away. The release of PVP/H2O2/HYP complex leads to decrease of fluorescent intensity in the core of microsphere. The oxygen release behavior can therefore be tracked accordingly. PVP/H2O2/HYP complex is stable as PVP/H2O2 complex and PVP/HYP complex are stable [34,47,56]. The complex exhibited one major emission peak in the fluorescent emission spectrum, 625 nm. The peak was slightly shifted compared with that of HYP (Fig. 2A), likely due to the change in chemical environment. The emission peak at 625 nm is suitable for in vivo detection using DsRed emission filter (575 nm-650 nm) equipped in most IVIS systems. One of the concerns when using HYP is that it may generate toxic singlet oxygen during IVIS imaging like during photodynamic therapy. Yet this is unlikely to occur as the fluence rate of IVIS (10−10 – 10−9 mW/cm2 [57]) is ~1011 times lower than photodynamic therapy (100 mW/cm2 [58]). Besides HYP, other oxygen sensitive dyes that can form stable complex with PVP may be used to monitor the oxygen release process.

The shell of the microspheres was based on poly(NIPAAm-co-AOLA-co-HEMA-co-NAS) (Scheme 2). NAS was used to conjugate catalase on the shell. These microspheres are advantageous over previously reported PVP/H2O2-based oxygen-release microspheres without catalase on the shell [1, 34], as the H2O2 in the released PVP/H2O2/HYP complex can be timely converted into oxygen to avoid potential toxic effect from H2O2. In the shell polymer, AOLA is the degradable component. In aqueous environment, the oligolatide can be degraded by hydrolysis of the ester group. Upon degradation, the resulting polymers became highly hydrophilic as they were soluble in DPBS at 37°C. In vivo, the polymers are expected to dissolve in body fluid and be eliminated from the body by urinary system.

The sustained oxygen release was achieved during the gradual degradation of the shell (Scheme 1). Compared with other H2O2-based oxygen-release systems [28,29], the PVP/H2O2-based oxygen-release microspheres can release oxygen in a more sustained manner (at least 3 weeks. Fig. 3E), because binding H2O2 to a high molecular weight PVP decreased its diffusivity. The sustained oxygen release is crucial for the survival of transplanted or host cells under ischemia since angiogenesis typically needs more than 2 weeks to establish [59,60]. The oxygen release kinetics was dependent on the degradation rate of the shell. The microspheres based on N50A25 that had the greatest degradation rate, exhibited the highest level of oxygen content among all groups from day 2 to day 21. In contrast, the microspheres fabricated using N65A10 that had slowest degradation rate, showed least oxygen release. Oxygen release kinetics in the ischemic tissues may be different from that in vitro. This is because (1) the oxygen content in ischemic tissues may increase during therapy while the in vitro study was conducted at 1% O2 condition; (2) the released oxygen may diffuse quickly into tissues; and (3) it is possible that oxygen release is quicker in vivo due to faster degradation. The oxygen released from microspheres significantly promoted the survival of MSCs under ischemia (Fig. 5). The increased MSC viability is likely due to the elevated cellular oxygen content. MSC has been reported to promote angiogenesis and muscle repair in various disease models [10,15,25,49]. In our future studies, we will deliver MSCs and oxygen-release microspheres into ischemic tissues such as ischemic limb and heart, and evaluate the in vivo oxygen release, MSC survival, and tissue regeneration. The released oxygen may not only promote MSC survival, but also increase the survival of host cells after diffusion into thigh muscles.

The microsphere photoluminescent emission intensity gradually decreased during the 3-week study period (Fig. 4 A and B). There is a correlation among the remaining fluorescent intensity of the microspheres, the degradation rate of the shell, and the oxygen release rate. The microspheres whose polymer shell degraded faster (Fig. 1B), released larger amount of oxygen (Fig. 3E), and exhibited weaker fluorescent signal (Fig. 4A and B). This provides the foundation for monitoring of oxygen release process based on the change of microsphere photoluminescent emission intensity. After the microspheres were implanted into thigh muscles, the fluorescent signal can be monitored by IVIS (Fig. 6). Similar to the in vitro results, the fluorescent intensity of the microspheres progressively decreased during the 21-day experimental period. The microspheres with faster degradation rate more quickly weakened the signal intensity. At day 14, the signal in two types of microspheres (N50A25 and N58A17) with faster degradation rate became invisible. The signal intensity decrease was much faster than that in vitro. As the HYP complex has low photobleaching and can be detected even after 28 days of in vivo implantation [47], the faster signal intensity reduction can be attributed to quicker microsphere degradation and oxygen release in vivo. The above in vitro and in vivo studies demonstrate that the oxygen release process and duration of release can be real-time monitored using the developed photoluminescent microspheres. While these results are promising, this work has limitations. First, the photoluminescent microspheres allow to qualitatively instead of quantitatively monitor the oxygen release in vivo. Second, detection of oxygen release in deep tissues using IVIS may be challenging due to scattering loss of the fluorescent signals.

To determine biocompatibility of the photoluminescent microspheres, the final degradation products of the microsphere shell were tested in terms of their cytotoxicity. The 3 final degradation products did not show cytotoxicity to MSCs even when the concentration was 20 mg/mL. To further evaluate cytotoxicity, the microspheres were implanted into thigh muscles of wild-type mice. F4/80 staining images at day 21 demonstrate that the microspheres did not provoke substantial inflammation (Fig. 7). The macrophage density was similar to that of the healthy muscles (Fig. 7B). The complementary in vitro and in vivo studies confirm that the photoluminescent microspheres had good biocompatibility.

5. Conclusion

In this work, we developed microspheres that not only released oxygen to augment cell survival under low oxygen condition, but also were photoluminescent for real-time monitoring of oxygen release process in vivo. The oxygen release kinetics was modulated by shell polymer degradation rate. The fluorescent intensity of the microspheres was negatively correlated with the oxygen release rate. In addition, the microspheres possessed good biocompatibility. These results demonstrate that the developed microspheres have potential to be used for cell transplantation for ischemic tissue regeneration.

Statement of significance.

Current cell therapy for ischemic tissue regeneration has unsatisfied efficacy largely due to low cell survival under hypoxic condition of the tissues. We have shown previously that sustained release of oxygen can improve cell survival in ischemic tissue. To understand how oxygen is released in vivo and duration of release, it is attractive to image the process of oxygen release. In this report, we have developed oxygen-release microspheres where the oxygen release can be tracked non-invasively and real-time using an In Vivo Imaging System (IVIS).

Acknowledgement

This work was supported by US National Institutes of Health (R01HL138175, R01HL138353, R01EB022018, R01AG056919), and National Science Foundation (1708956).

Footnotes

Declaration of Competing Interest

The authors declare no conflict of interest.

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