Oxygen-release microspheres capable of releasing oxygen in response to environmental oxygen level to improve stem cell survival and tissue regeneration in ischemic hindlimbs

Article abstract of DOI:10.1016/j.jconrel.2021.01.034

Stem cell transplantation has been extensively explored to promote ischemic limb vascularization and skeletal muscle regeneration. Yet the therapeutic efficacy is low due to limited cell survival under low oxygen environment of the ischemic limbs. Therefore, continuously oxygenating the transplanted cells has potential to increase their survival. During tissue regeneration, the number of blood vessels are gradually increased, leading to the elevation of tissue oxygen content. Accordingly, less exogenous oxygen is needed for the transplanted cells. Excessive oxygen may induce reactive oxygen species (ROS) formation, causing cell apoptosis. Thus, it is attractive to develop oxygen-release biomaterials that are responsive to the environmental oxygen level. Herein, we developed oxygen-release microspheres whose oxygen release was controlled by oxygen-responsive shell. The shell hydrophilicity and degradation rate decreased as the environmental oxygen level increased, leading to slower oxygen release. The microspheres were capable of directly releasing molecular oxygen, which are safer than those oxygen-release biomaterials that release hydrogen peroxide and rely on its decomposition to form oxygen. The released oxygen significantly enhanced mesenchymal stem cell (MSC) survival without inducing ROS production under hypoxic condition. Co-delivery of MSCs and microspheres to the mouse ischemic limbs ameliorated MSC survival, proliferation and paracrine effects under ischemic conditions. It also significantly accelerated angiogenesis, blood flow restoration, and skeletal muscle regeneration without provoking tissue inflammation. The above results demonstrate that the developed microspheres have potential to augment cell survival in ischemic tissues, and promote ischemic tissue regeneration in a safer and more efficient manner.

Full text of DOI:10.1016/j.jconrel.2021.01.034

Journal of Controlled Release 331 (2021) 376–389
Contents lists available at ScienceDirect
Journal of Controlled Release
journal homepage: www.elsevier.com/locate/jconrel
Oxygen-release microspheres capable of releasing oxygen in response to
environmental oxygen level to improve stem cell survival and tissue
regeneration in ischemic hindlimbs
a,
b
,
1
a,
b,
1
a,
b
a,
b
a,b,*
Ya Guan
, Ning Gao
, Hong Niu , Yu Dang , Jianjun Guan
a
Department of Mechanical Engineering and Materials Science, Washington University in St. Louis, St. Louis, MO 63130, USA
Department of Materials Science and Engineering, The Ohio State University, Columbus, OH 43210, USA
b
A R T I C L E I N F O
A B S T R A C T
Keywords:
Stem cell transplantation has been extensively explored to promote ischemic limb vascularization and skeletal
muscle regeneration. Yet the therapeutic efficacy is low due to limited cell survival under low oxygen envi-
ronment of the ischemic limbs. Therefore, continuously oxygenating the transplanted cells has potential to in-
crease their survival. During tissue regeneration, the number of blood vessels are gradually increased, leading to
the elevation of tissue oxygen content. Accordingly, less exogenous oxygen is needed for the transplanted cells.
Excessive oxygen may induce reactive oxygen species (ROS) formation, causing cell apoptosis. Thus, it is
attractive to develop oxygen-release biomaterials that are responsive to the environmental oxygen level. Herein,
we developed oxygen-release microspheres whose oxygen release was controlled by oxygen-responsive shell. The
shell hydrophilicity and degradation rate decreased as the environmental oxygen level increased, leading to
slower oxygen release. The microspheres were capable of directly releasing molecular oxygen, which are safer
than those oxygen-release biomaterials that release hydrogen peroxide and rely on its decomposition to form
oxygen. The released oxygen significantly enhanced mesenchymal stem cell (MSC) survival without inducing
ROS production under hypoxic condition. Co-delivery of MSCs and microspheres to the mouse ischemic limbs
ameliorated MSC survival, proliferation and paracrine effects under ischemic conditions. It also significantly
accelerated angiogenesis, blood flow restoration, and skeletal muscle regeneration without provoking tissue
inflammation. The above results demonstrate that the developed microspheres have potential to augment cell
survival in ischemic tissues, and promote ischemic tissue regeneration in a safer and more efficient manner.
Critical limb ischemia
Stem cell therapy
Oxygenation
Angiogenesis
Skeletal muscle regeneration
1
. Introduction
Peripheral artery disease (PAD), which affects more than 200 million
acceleration of muscle repair represent the major goals for CLI
treatment.
Conventional treatments for CLI include arterial bypass and endo-
vascular therapy. However, these therapeutic approaches are rather
invasive, and are not suitable for CLI patients with severe comorbidity or
sepsis [5,6]. There has been extensive interest in alternative approaches
to address current limitations, such as proangiogenic growth factor
therapy [7], gene therapy [8], and stem cell therapy [9]. Compared to
growth factor and gene therapies, stem cell therapy has advantages. The
delivered stem cells may secrete a wide range of growth factors that are
necessary for the blood vessel formation and/or myogenesis [10,11].
The stem cells may also have the capacity of differentiating into
people worldwide, is a progressive atherosclerotic disorder induced
mainly by the lipid deposition in the vascular bed as a result of hard-
ening and narrowing of arteries [1]. Approximately 10% of the PAD
patients develop critical limb ischemia (CLI), with 30% of them gain
limited outcomes from conservative therapies and have no choice but
leg amputation [2]. CLI is characterized by low blood perfusion, severe
tissue ischemia and degenerated skeletal muscle, which lead to non-
healing wounds, ulcers and necrosis in the terminal stage [3,4]. Thus,
quick restoration of blood perfusion to rescue existing tissue and
*
Corresponding author at: Department of Mechanical Engineering and Materials Science, Washington University in St. Louis, 303F Jubel Hall (Office), 340
Whitaker Hall (Lab), Campus Box 1185, One Brookings Drive, St. Louis, MO 63130, United States of America.
E-mail address: jguan22@wustl.edu (J. Guan).
1
These authors contributed equally to this work.
https://doi.org/10.1016/j.jconrel.2021.01.034
Received 13 August 2020; Received in revised form 20 January 2021; Accepted 21 January 2021
Available online 27 January 2021
0
168-3659/© 2021 Elsevier B.V. All rights reserved.

Y. Guan et al.
Journal of Controlled Release 331 (2021) 376–389
endothelial cells for vascularization [10,12,13]. Therefore, stem cell
therapy possesses greater potentials than the other approaches in pro-
moting ischemic tissue regeneration [6].
survival under hypoxia in vitro. We further transplanted the oxygen-
release microspheres and MSCs into ischemic limbs and evaluated the
survival and proliferation of MSCs, tissue angiogenesis, skeletal muscle
regeneration, and tissue inflammatory response.
Various stem cell types including adipose-derived stem cells [14,15],
mesenchymal stem cells (MSCs) [16–18], iPSC-derived mesenchymal
stem cells [19], ESC-derived stem cells [20,21] have been tested for
ischemic limb regeneration. Among them, MSCs are considered as
potent biofactories, which improve the angiogenesis and myogenesis by
secreting various types of angiogenic and myogenic factors [22]. These
include vascular endothelial growth factor (VEGF), platelet-derived
growth factor (PDGF), basic fibroblast growth factor (bFGF), hepato-
cyte growth factor (HGF), and insulin-like growth factor-1 (IGF-1).
These growth factors are also of particular interest in the growth factor
therapy [23]. Another key advantage of MSCs is that they perform
immunoregulatory function via producing soluble factors that regulate
immune response, such as PEG2 and IL-6 [24]. In addition, MSCs pro-
mote angiogenesis by acting as pericytes [25–27], and/or providing an
appropriate microenvironment for cell types like endothelial cells to
form blood vessels [28].
2. Materials and methods
2.1. Materials
All chemicals were purchased from Sigma-Aldrich unless otherwise
stated. N-isopropylacrylamide (NIPAAm, TCI) was recrystallized in
hexane for 3 times before use. 2-Hydroxyl methacrylate (HEMA, Alfa
Aesar) was passed through inhibitor removers. Hydrogen peroxide (30
wt% in water), PVP (40 kDa, Fisher Scientific), bovine liver catalase
(2000–5000 units/mg), tris(4,7-diphenyl-1,10-phenanthroline) ruthe-
nium (II) dichloride (Ru(Ph
received.
2 3 2
phen )Cl , GFS chemicals) were used as
2.2. Synthesis of 2-(2-nitroimidazolyl) ethanamine (NIEM)
Although MSCs exhibit promising potentials in accelerating angio-
genesis and myogenesis, clinical trials only showed low efficiency in the
improvement of blood perfusion and muscle repair [29]. One of the key
causes is the poor cell survival and engraftment under low oxygen
condition of the ischemic limbs [30,31]. Accordingly, we hypothesized
that co-delivery of oxygen-release biomaterials and MSCs may increase
cell survival and engraftment by continuously oxygenating the delivered
cells. The released oxygen may also support host cells to survive and
proliferate. The enhanced cell survival would promote ischemic tissue
vascularization and myogenesis.
NIEM was synthesized in order to conjugate it to the microspheres so
as to impart the microsphere shell with oxygen sensitivity (Fig. 2A).
Briefly, 2-bromoethylamine hydrobromide (1.0 equiv), di-tert-butyl
dicarbonate (Boc O, 1.1 equiv) and 4-dimethylaminopyridine (DMAP,
2
◦
0.2 equiv) were dissolved in dichloromethane (DCM) at 0 C. Triethyl-
amine (TEA, 1.1 equiv) was added dropwise to the solution. After
overnight stirring at room temperature, the mixture was rinsed subse-
quently with saturated NH Cl, NaHCO and NaCl. The organic layer was
4
3
dried over anhydrous Na SO4 and evaporated to obtain tert-butyl (2-
2
To improve cell survival under ischemia by continuous supply of
oxygen, various oxygen-release systems have been developed. They
bromoethyl) carbonate. Then, tert-butyl (2-bromoethyl) carbonate (1.1
equiv) and 2-nitroimidazole (1.0 equiv) were dissolved in dime-
thylformamide in the presence of K CO (1.5 equiv) and NaI (0.2 equiv).
were based on H
2 2 2
O [32,33], CaO [34] and fluorinated molecules
2
3
◦
[
35,36]. Current oxygen release systems typically release oxygen for less
The reaction was conducted at 80 C for 4 h followed by overnight
stirring at room temperature. The product was dissolved in ethyl acetate,
than two weeks [32–36], thus are unable to support long-term cell
survival in ischemic limbs [37,38], since the establishment of angio-
genesis needs more than 3 weeks [39,40]. To address this limitation, we
have recently developed novel oxygen-release microspheres (ORM)
rinsed sequentially with saturated NaHCO and NaCl, dried over anhy-
3
drous Na SO , and filtered. The filtrate was concentrated and purified
2
4
by flash chromatography using ethyl acetate/hexane (1/1). Finally, the
product was dissolved in DCM/trifluoroacetic acid, and stirred over-
night at room temperature to deprotect the amine group and obtain
based on polyvinylpyrrolidone (PVP)/H
glycolide) (PLGA) and catalase [41–43]. We have shown that the high-
molecular-weight PVP/H can gradually release from the PLGA shell
2 2
O complex, poly(lactide-co-
1
2
O
2
NIEM. The chemical structure of NIEM was confirmed by H NMR
and convert to molecular oxygen by catalase. The duration of the oxygen
release can reach up to 4 weeks, which has been proved to augment
cardiac cell survival when transplanted into the ischemic hearts [41].
While these findings are promising, the current oxygen release systems
cannot release oxygen in response to tissue oxygen level [41–43]. Dur-
ing tissue regeneration, the number of blood vessels are gradually
increased, leading to the elevation of tissue oxygen content. Accord-
ingly, less exogenous oxygen is needed for the transplanted cells.
Excessive oxygen may induce reactive oxygen species (ROS) formation,
causing cell apoptosis [44–47]. Thus, it is attractive to develop oxygen-
release systems that are responsive to the environmental oxygen level.
In this work, we developed new microspheres capable of releasing
oxygen depending on the environmental oxygen level, i.e., releasing
faster at a lower environmental oxygen level, while releasing slower at a
higher environmental oxygen level. The microspheres had an oxygen-
responsive shell that contained 2-nitroimidazole whose hydrophilicity
increases when the environmental oxygen level decreases [48,49]. This
enabled the shell to have higher hydrophilicity and degradation rate,
leading to faster oxygen release. The shell was also conjugated with
catalase on the surface, allowing the microspheres to directly release
molecular oxygen. These microspheres are safer than those existing
oxygen-release biomaterials that release hydrogen peroxide and rely on
its decomposition to form oxygen [32,33]. Hydrogen peroxide may
damage the cells when it is not timely decomposed [50]. We investi-
gated the oxygen release kinetics at different environmental oxygen
content, and the efficacy of the released oxygen in enhancing MSC
((CD ) SO): δ 7.64 (d, 1H), 7.23 (d, 1H), 4.62–4.65 (t, 2H), 3.32–3.35 (t,
3
2
2H)).
2.3. Synthesis of microsphere shell
The microsphere shell was based on NIPAAm, HEMA, acrylate-
oligolactide (AOLA) and N-acryloxysuccinimide (NAS). AOLA and NAS
were synthesized as previously reported [41,42,51]. The polymer poly
(NIPAAm-co-HEMA-co-AOLA-co-NAS) (abbreviated as PNHAN) was
synthesized by free radical polymerization using benzoyl peroxide as an
initiator (Fig. 2B) [41,51–53]. The feed ratio of NIPAAm/HEMA/AOLA/
◦
NAS was 50/5/25/20. The reaction was performed at 70 C for 20 h
under the protection of nitrogen. The polymer was precipitated in hex-
ane, and purified three times by dissolving in tetrahydrofuran and
precipitating in ethyl ether. The hydrogel poly(NIPAAm-co-HEMA-co-
AOLA) (abbreviated as PNHA) was synthesized using the same poly-
merization method with a feed ratio of NIPAAm/HEMA/AOLA = 86/
10/4. This hydrogel was used to deliver microspheres and MSCs into
ischemic limbs.
2.4. Conjugation of NIEM with PNHAN
NIEM was conjugated to the polymer by reacting with the succini-
mide group in NAS component (Fig. 2B). Briefly, PNHAN, NIEM, and
TEA were dissolved in dimethylformamide and mixed together. The
◦
reaction was conducted at 60 C overnight. After the solvent was
3
77

Y. Guan et al.
Journal of Controlled Release 331 (2021) 376–389
evaporated, the product was purified 3 times by dissolving in tetrahy-
drofuran and precipitating in ethyl ether. PNHAN with different NIEM
contents were obtained by changing the ratio of PNHAN and NIEM. The
NIEM conjugated polymer was abbreviated as PNHAN/NIx where x
represents the ratio of 2-nitroimidazolyl (NI) group.
2.7. Characterization of oxygen release kinetics under different
environmental oxygen level
The oxygen release kinetics was tested under 1% or 5% oxygen
condition for 28 days. PDMS membrane encapsulated with an oxygen-
sensitive luminophore Ru(Ph
rhodamine B was placed in a 96-well plate [54]. Rhodamine B was used
as a reference. 200 L of DPBS was then added into each well. The plate
2 3 2
phen )Cl and an oxygen-insensitive dye
2
.5. Characterization of the hypoxia-sensitive polymer
μ
Chemical structures of the polymer before and after NIEM conjuga-
was placed under 1% or 5% oxygen condition for 24 h followed by
adding 50mg of dry microspheres into each well under corresponding
oxygen condition (n = 8 for each group). At each time point, the fluo-
tion were confirmed by 1H NMR. Polymer composition was calculated
from the spectra. Water content and degradation of PNHAN with
different NIEM contents were conducted in Dulbecco’s phosphate-
buffered saline (DPBS, Thermo Fisher) under normoxia (21% O
hypoxia (1% O ), respectively. Briefly, the polymer was dissolved in
DCM, and added into a 1.5 mL centrifuge tube. After solvent evapora-
tion, 200 L of DPBS was added. DPBS was supplemented with 1%
rescence intensity was measured for Ru(Ph
2
phen
3
)Cl
2
(λexcitation = 470
2
) and
nm, λemission = 610 nm) and rhodamine B (λexcitation = 543 nm, λemission
= 576 nm) using a fluorescent plate reader (Molecular Devices). The
2
2 3 2
oxygen level measured by Ru(Ph phen )Cl represented the oxygen
μ
content in PDMS membrane in real-time. A calibration curve was used to
penicillin/streptomycin (Thermo Fisher) to prevent the growth of bac-
convert the fluorescence intensity to oxygen level (Appendix. 1). The
◦
teria. After 12 h of incubation at 37 C, the hydrated weight of polymer
2 2
residual H O concentration in the release medium was determined by
film was measured (w
dry weight was measured (w
)/w
× 100%.
For degradation study, the polymer films were incubated in DPBS
under normoxia (21% O ) and hypoxia (1% O ) for 8 weeks, respec-
1
). The polymer film was then lyophilized and the
Pierce peroxide assay (Thermo Fisher).
2
). The water content was calculated as (w
1
-
To evaluate bioactivity of the conjugated catalase, oxygen release
from the microspheres with catalase conjugation, and from the micro-
spheres incubated in the release medium containing the same amount of
w
2
2
2
2
free catalase (260 gcatalase/mgmicrosphere) was measured, respectively.
μ
tively. At each time point, the samples (n = 4 for each group) 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.
2.8. MSC survival, ROS content, and cellular oxygen level measurement
Rat mesenchymal stem cells (Cell Applications) were cultured in
minimum essential medium alpha ( MEM, Thermo Fisher) supple-
α
2
.6. Fabrication and characterization of oxygen-release microspheres
mented with 10% fetal bovine serum (FBS, Corning) and 1% penicillin/
streptomycin. To investigate the effect of oxygen release on MSC sur-
vival under hypoxia, 50 mg/mL of microspheres and 2 million/mL MSCs
The oxygen-release microspheres were fabricated by coaxial elec-
◦
trospraying. NIEM conjugated PNHAN, and PVP/H
2
O
2
were used as
were encapsulated in 4 wt% PNHA hydrogel at 4 C. The mixture was
◦
shell and core, respectively. Blends of PNHAN and PNHAN/NI18 with
molar ratio of 0, 3/7 and 9/1 were dissolved in DCM at a concentration
of 5 wt%. The NI content in the polymer shell was 0, 5% and 16%,
placed in a 37 C incubator for gelation. Then, the supernatant was
replaced by serum-free MEM medium pre-incubated in 1% O2 condi-
α
tion. The constructs were cultured in a 1% O2 incubator. At each time
respectively. PVP was dissolved in 30% H
and repeating unit vinylpyrrolidone (VP) was controlled at 4.5/1. The
flow rates of the PVP/H complex and PNHAN solution were 0.2 and
mL/h, respectively. The coaxial device was charged at a voltage of
15 kV. The microspheres were collected on a rotating mandrel covered
2
O
2
. The molar ratio of H
2
O
2
point, the constructs (n = 4 for each group) were digested by papain
◦
solution at 60 C. The double stranded DNA (dsDNA) content was
2
O
2
measured using a PicoGreen dsDNA assay kit (Thermo Fisher). To
visualize the live cells, MSCs were pre-labeled with a live cell tracker,
CM-Dil (Thermo Fisher). To determine cell ROS expression, MSCs were
1
+
with aluminum foil charged at ꢀ 10 kV. Following fabrication, the mi-
pre-stained with CM-H DCFDA (Thermo Fisher). Fluorescent images
2
◦
crospheres were lyophilized and stored at ꢀ 20 C before use. To visu-
were taken using a confocal microscope.
alize the core-shell structure of the microspheres, rhodamine B and
fluorescein isothiocyanate (FITC) were added to the PNHAN solution
Electron paramagnetic resonance (EPR) was used to determine
intracellular oxygen content. Briefly, MSCs were incubated with lithium
and PVP/H
right after microsphere fabrication using
Olympus FV1200) to avoid the diffusion of fluorescent dyes. To char-
2
O
2
solution, respectively. The fluorescent images were taken
phthalocyanine (LiPc) nanoparticles (10 mg/mL in MEM) for 1 h to
α
a
confocal microscope
allow cellular uptake. The cells were then washed with DPBS for 3 times
to remove the free LiPc nanoparticles from cell surface. After trypsini-
zation, the cells (2 million/mL) and microspheres (50 mg/mL) were
encapsulated in the PNHA gel (4 wt%). The constructs were loaded into
EPR tubes (n = 3 for each group). The tubes were placed in a hypoxic
(
acterize morphology and size of the microspheres, scanning electron
microscopy (SEM) was used.
The microspheres were further conjugated with catalase so that the
◦
released PVP/H
2
O
2
can be timely converted into oxygen at the micro-
incubator (1% O , 37 C) for 48 h. The EPR tests were performed on an
2
sphere surface. In brief, 50 mg of oxygen-release microspheres were
X-band EPR instrument (Bruker), with 0.1 mW for microwave power,
1.0 dB for attenuation, 9.8 GHz for frequency, and 1.5 for modulation
amplitude [52]. The intracellular oxygen content was determined based
on the linewidth of samples, and calibration curve of linewidth vs. ox-
ygen content.
mixed with 6 mL catalase solution (5 mg/mL in DI water), and stirred for
◦
4
h at 4 C. The mixture was then centrifuged. The microspheres were
washed 3 times with DI water to remove un-conjugated catalase. To
confirm the conjugation, catalase was pre-labeled with FITC, and fluo-
rescent images were taken using a confocal microscope (Olympus
FV1200). The microspheres without catalase conjugation were used as
control. The catalase conjugation efficiency was determined by
measuring the fluorescent intensity of the microspheres and converting
the fluorescent intensity into catalase concentration using a calibration
curve.
2.9. Implantation of oxygen-release microspheres and MSCs into mouse
ischemic limbs
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 Com-
mittee of The Ohio State University. Wild type C57BL/6 mice (Charles
River Laboratories) at the age of 8–10 weeks were used. Ischemic sur-
gery was performed on one hindlimb, and the contralateral hindlimb
3
78

Y. Guan et al.
Journal of Controlled Release 331 (2021) 376–389
was used as control. To induce ischemia, unilateral femoral artery and
vein were ligated and cut (Fig. 1). Thirty minutes later, 4 wt% PNHA
solution (Gel group), or PNHA solution with 2 million cells/mL CMDil-
labeled MSCs (Gel/MSC group), or PNHA solution with 2 million cells/
mL CMDil-labeled MSCs and 50 mg/mL oxygen-release microspheres
isolated from the ischemic limbs using TRIzol. cDNA was synthesized at
1
μg/reaction using a high capacity cDNA reverse transcription kit
(Invitrogen). The primer sequences are listed in Appendix. 2. Real-time
RT-PCR was conducted using SYBR Green master mix (Invitrogen) on
Applied Biosystems™ QuantStudio™ 5 Real-Time PCR System (384-
well). The fold of change was calculated using a standard ΔΔCt method
(n ≥ 6 for each group).
(
Gel/MSC/ORM group) was injected to the gracilis muscle. Four in-
jections were made for each ischemic hindlimb (50 L/injection). Six
μ
mice were used for each group. At days 1, 7, 14, 21 and 28, the blood
perfusion of the normal and surgery hindlimbs was measured using a
laser Doppler perfusion imager (Perimed). The blood flow recovery
percentage was calculated by normalizing the blood perfusion intensity
in the ischemic hindlimb to the normal hindlimb in the same animal
2
.12. Statistical analysis
All results were presented as mean ± standard deviation. Compari-
sons among groups were performed using one-way ANOVA with post
[
55].
hoc Tukey test. Statistical significance was set as p < 0.05.
2
.10. Histology and immunohistochemical staining
3
. Results and discussion
Four weeks after surgery, the mice were euthanized. The hindlimbs
3
.1. Synthesis and characterization of oxygen-sensitive polymers
with and without surgery were collected. The tissues were fixed with 4%
paraformaldehyde for 48 h, embedded in paraffin, and sectioned into 5-
The oxygen-sensitive polymers were synthesized by conjugating
μ
m slices. The CM-Dil + MSCs were imaged using a confocal microscope.
PNHAM with NIEM. The PNHAM was made by free radical polymeri-
zation of NIPAAm, HEMA, AOLA and NAS (Fig. 2B). The molar ratio of
the four components was calculated as NIPAAm/HEMA/AOLA/NAS =
Cell density was then quantified from the images. For histological
assessment of skeletal muscle, hematoxylin and eosin (H&E) staining
was performed. Muscle fiber diameter, and density of central nucleus in
muscle fibers were quantified based on the H&E images. For immuno-
histochemical analysis, the tissue sections were first stained with anti-
5
2.5/4.4/22.3/20.8 (Fig. 2C), which was consistent with the feed ratio.
PNHAN was conjugated with NIEM via the succinimide-amine reaction
Fig. 2B). The conjugation was confirmed by 1H NMR spectrum where
(
alpha smooth muscle actin-alpha ( -SMA, Abcam), isolectin (Thermo
α
the characteristic peak of NAS at 2.8 ppm became unpronounced after
conjugation (Fig. 2C). Polymers with 9% and 18% of NI group were
synthesized (abbreviated as PNHAN/NI9 and PNHAN/NI18,
respectively).
Fisher), anti-Ki67 (Thermo Fisher), anti-myosin heavy chain (MHC,
R&D) and anti-F4/80 (Santa Cruz Biotechnology), respectively. The
tissue sections were then incubated with corresponding Dylight488-anti
mouse and 647-anti rabbit secondary antibodies. Cell nuclei were
stained with Hoechst 33342 or DRAQ5. Immunofluorescence images
were taken with a confocal microscope (Zeiss LSM700). Proliferating
MSCs were identified as those CMDil+ and Ki67+ cells. Blood vessels
were isolectin+ lumens. Mature blood vessels were lumens positive to
To investigate whether these polymers were responsive to the envi-
ronmental oxygen level, water content of PNHAN/NI9 and PNHAN/
2 2
NI18 at both normoxia (21% O ) and hypoxia (1% O ) conditions were
measured (Fig. 3A). The results demonstrate that PNHAN/NI9 and
PNHAN/NI18 exhibited a significantly higher water content under
hypoxia than normoxia (p < 0.01). This is largely due to the oxygen-
dependent reductive reaction which converts the hydrophobic nitro
group to the hydrophilic amine group under hypoxia [48,49]. With the
increase of NI content from 9% to 18%, the water content of the poly-
mers in hypoxia appeared to have increased, although statistical sig-
nificance was not reached. The reductive reaction requires a reductant
(i.e. an electron donor). In this experiment, glucose, a reducing sugar, in
DPBS served as reductant. In the studies with cells, many reductase
enzymes produced by cells can act as electron donors, such as xanthine
both isolectin and -SMA.
α
2
.11. Gene expression of angiogenic and prosurvival growth factors in
ischemic limbs
To determine angiogenic and prosurvival growth factor expression in
the ischemic limbs with or without implantation of MSCs and oxygen-
release microspheres, the expression of bFGF, PDGF-BB and HGF at
mRNA level was characterized using real time RT-PCR. RNA was
Fig. 1. Scheme of the design of hypoxia-sensitive oxygen-release microspheres and the animal model of critical limb ischemia (CLI).
79
3

Y. Guan et al.
Journal of Controlled Release 331 (2021) 376–389
Fig. 2. Synthesis of a hypoxia-sensitive and degradable polymer. (A) Synthesis route of 2-(2-nitroimidazolyl) ethanamine (NIEM); (B) Synthesis of poly(NIPAAm-co-
1
HEMA-co-AOLA-co-NAS) (PNHAN), conjugation with NIEM and its degradation mechanism; (C) H NMR spectrum of PNHAN and PNHAN/NI18.
◦
oxidase and aldehyde dehydrogenase [48].
37 C. The degraded polymer can thus be eliminated from the body by
urinary system in vivo.
We further studied the degradation of oxygen-sensitive polymer
(
PNHAN/NI9) under normoxia and hypoxia conditions (Fig. 3B). Under
aqueous condition, the ester groups in oligolactide can hydrolyze
Fig. 2B). The polymer had 30.7% weight loss after 8 weeks in normoxic
3.2. Fabrication and characterizations of oxygen-release microspheres
(
environment. The weight loss was significantly increased to 39.2% in
hypoxic condition (p < 0.001). The higher degradation rate under
hypoxia can be attributed to faster hydrolysis caused by enhanced hy-
drophilicity of the polymer in response to the low oxygen environment.
After complete degradation, the resulting polymer was soluble in DPBS
The oxygen-release microspheres were fabricated by co-axial elec-
trospraying. The microspheres assumed core-shell structure as
confirmed by confocal images of the shell and core (Fig. 4A). The
diameter was ~5 m (Fig. 4B). The shell was a blend of PNHAN and
μ
PNHAN/NI18. The NHS group in the PNHAN was used to conjugate
catalase on the microspheres. The PNHAN/NI18 imparted the
◦
◦
at 37 C, as its gelation temperature was increased to 63 C, greater than
3
80

Y. Guan et al.
Journal of Controlled Release 331 (2021) 376–389
Fig. 3. Characterizations of the polymer under different oxygen content. (A) Water content of the polymer with different NI conjugation ratios under normoxia and
◦
hypoxia. The samples were incubated in 37 C DPBS for 12 h; (B) Degradation of the PNHAN/NI9 polymer under normoxia and hypoxia for 8 weeks. *p < 0.05, **p <
0
.01, ***p < 0.001.
Fig. 4. Characterizations of the oxygen-release microspheres. (A) Fluorescent images of the oxygen-release microspheres. The green fluorescence is FITC added to the
m; (C) Fluorescent image of the
core and the red fluorescence is the rhodamine added to the shell; (B) SEM images of the oxygen-release microsphere. Scale bar = 5
μ
oxygen-release microsphere before and after conjugation with FITC-labeled catalase. (For interpretation of the references to colour in this figure legend, the reader is
referred to the web version of this article.)
microspheres with oxygen sensitivity. The ratio of PNHAN and PNHAN/
NI18 were modulated to allow the shell to have NI content of 5% and
conditions, respectively. The oxygen was able to gradually release from
the microspheres during the 4-week experimental period (Fig. 5). The
release rate was dependent on the NI content in the microspheres, and
environment oxygen level. For the microspheres without NI (PNHAN/
NI0/CAT), the oxygen release rate was similar during the first 5 days
regardless of the environmental oxygen level (Fig. 5). The release at 1%
1
6%, respectively. The core of the microspheres was PVP/H
2
O
2
com-
is
plex. The rationale of using PVP/H complex instead of pure H
2
O
2
2
O
2
that the small molecule H diffuses too fast to achieve long-term ox-
2
O
2
ygen release. Binding H O with a high molecular weight PVP can
2 2
decrease the diffusion rate [41,42], resulting in continuous oxygen
release. Long-term continuous oxygen supply is critical for the survival
of transplanted cells in ischemic limbs since angiogenesis typically re-
quires more than 3 weeks to establish [37,56,57]. The microspheres
were further immobilized with catalase. Successful conjugation was
confirmed by confocal image where FITC-labeled catalase was in the
2
and 5% O conditions peaked at day 14 and day 7 respectively, followed
by slower release till day 28. These results demonstrate that the mi-
crospheres without NI cannot release oxygen in response to environ-
mental oxygen level. In contrast, the microspheres with NI (PNHAN/
NI5/CAT and PNHAN/NI16/CAT) were able to release higher amount of
2 2
oxygen under 1% O than under 5% O during the 4-week period (Fig. 5,
shell (Fig. 4C). The catalase loading efficiency was 260
μg
catalase
/
Appendix. 3). The microspheres with higher NI content released more
mgmicrosphere
.
oxygen at both oxygen conditions. In PNHAN/NI16/CAT group, the
Oxygen was released from the microspheres after the PVP/H
complex diffused to the shell was converted by the catalase immobilized
on the shell. The oxygen release study was performed at 1% and 5% O
2
O
2
2 2
released oxygen level was significantly higher under 1% O than 5% O
starting from day 3 (p < 0.001). When comparing the oxygen release
2
kinetics of the microspheres with different NI content under the same
3
81

Y. Guan et al.
Journal of Controlled Release 331 (2021) 376–389
Fig. 5. Oxygen release kinetics. The oxygen release of the microspheres with different NI contents was tested for 28 days under 1% and 5% oxygen content.
oxygen condition, it was found that the release kinetics was similar
among the microspheres with different NI content under 5% O condi-
tion (Appendix. 3). However, under 1% O condition, the oxygen
for ischemic limb regeneration [9,62]. To investigate whether the oxy-
gen released from microspheres can improve MSC survival under hyp-
oxia, we loaded MSCs and oxygen-release microspheres together in the
PNHA gel. This hydrogel has been confirmed to be injectable and
biocompatible [41,51]. The constructs were cultured under hypoxic
2
2
release rate was dependent on the NI content. A higher NI content led to
a greater level of released oxygen during the 4-week study period. These
results reveal that the microspheres with NI can release oxygen in
response to environmental oxygen level. The oxygen release rate for
individual NI-containing microsphere type is consistent with water
content and degradation results where both were increased under lower
oxygen conditions (Fig. 3). The higher water content and faster degra-
2
condition (1% O ) without supplement of FBS. In the constructs without
oxygen-release microspheres (Gel), MSCs experienced extensive death
as the dsDNA content largely decreased during the 28-day culture period
(Fig. 6A). At day 28, dsDNA content was less than 25% of that at day 1.
In the constructs supplemented with oxygen-release microspheres (Gel/
PNHAN/NI0/CAT, Gel/PNHAN/NI5/CAT and Gel/PNHAN/NI16/CAT),
the released oxygen rescued the cells and even supported their prolif-
eration (Fig. 6A). The dsDNA content did not decrease and/or increased
over 4 weeks. This is likely due to the elevation in cellular oxygen
content by released oxygen. The constructs containing microspheres
with lower NI content (Gel/PNHAN/NI5/CAT) exhibited substantially
higher dsDNA content than those containing microspheres without NI
(Gel/PNHAN/NI0/CAT). Further increase of NI content significantly
increased dsDNA content at each time point (p < 0.001, Gel/PNHAN/
NI5/CAT vs. Gel/PNHAN/NI16/CAT). The live cell images confirmed
the dsDNA results (Fig. 6B). These results are in accordance with the
oxygen release kinetics where higher oxygen release amount more
greatly improved cell survival.
dation rate allowed PVP/H
faster oxygen release. For the microspheres with different NI content
PNHAN/NI5/CAT and PNHAN/NI16/CAT), the increased water con-
2 2
O complex to diffuse out quicker, leading to
(
tent likely stimulated polymer degradation. The combined effects of
water content and degradation rate possibly resulted in accelerated
release of oxygen. The hypoxia-sensitive oxygen release is crucial for in
vivo applications because the cells in the more hypoxic environment
require greater amount of oxygen for their basic metabolism [58]. For
the cells in the less ischemic condition, moderate oxygen supply is
adequate for their survival and proliferation. An oxygen-release system
without hypoxia-sensitivity will cause a universal elevation in oxygen
level among all microenvironments. Excessive oxygen will likely cause
overproduction of ROS that may have detrimental effect on cells [59].
In this work, the catalase was directly immobilized on microsphere
surface (Fig. 4C). To determine whether catalase conjugation affected its
bioactivity, the oxygen release from microspheres conjugated with
catalase, and from microspheres incubated with free catalase was
measured respectively. The microspheres with catalase conjugation
released similar amount of oxygen as the microspheres incubated with
free catalase on days 1 and 3 (Appendix. 4, p > 0.05), indicating that
To understand the mechanism that released oxygen enhanced MSC
survival under hypoxia, oxygen content in MSCs was measured using
EPR. The MSCs encapsulated in the PNHA gel without oxygen-release
microspheres had an oxygen content of 4.2% after 48 h of culture
2
under 1% O . The oxygen content was significantly increased to 10.4%
when the MSCs were encapsulated in the PNHA gel with oxygen-release
microspheres (Fig. 6C, p < 0.001). These results demonstrate that the
enhanced cell survival under hypoxia was resulted from increased
intracellular oxygen content. It remains to be investigated whether the
released oxygen changes cell metabolic behaviors, such as oxygen con-
sumption rate, glycolysis rate and fatty acid oxidation under hypoxia.
One of the concerns for released oxygen is that it may increase ROS
content in MSCs leading to cell death. To determine ROS content in the
2 2
the conjugation did not affect catalase bioactivity in converting H O
into molecular oxygen. One of the advantages of the microspheres with
conjugated catalase is that molecular oxygen can be timely and directly
released from the microspheres. This avoids released, relatively high
concentration of H
them. Most of current oxygen release biomaterials release H
and the oxygen release is dependent on H decomposition [32,33].
The undecomposed H may interact with cells causing cell death.
2
O
2
to contact with transplanted cells and damage
2 2
O
first,
2
O
2
2
cells, CM-H DCFDA staining was performed (Appendix. 5). At days 1,
2
O
2
14 and 28, the groups with oxygen-release microspheres exhibited
similar ROS content (Fig. 6D). Interestingly, the ROS content was
significantly lower than that in the group without oxygen release (p <
0.05 or 0.001). These results demonstrate that the released oxygen did
not raise the oxidative stress in MSCs. There was no significant differ-
These microspheres were also advantageous than our previously
developed oxygen release system where catalase was mixed with the
oxygen-release microspheres in the hydrogel [41,42]. Catalase may
2 2
release from the hydrogel. The released H O may not be timely con-
verted into oxygen when the amount of catalase in the hydrogel is low.
In this work, the immobilized catalase efficiently converted the released
2
ence between O -sensitive and insensitive microspheres in ROS pro-
duction under hypoxia (p > 0.05). It is possible that the oxygen released
from these microspheres was consumed by metabolic-demanding MSCs
for survival under hypoxia instead of generating ROS.
H
2
O
2
into oxygen. During the 4-week release study period, the release
medium contained less than 10 M H as measured by a peroxide
assay kit. This concentration is too low to trigger cell apoptosis [60,61].
μ
2 2
O
Since PNHAN/NI16/CAT microspheres had the fastest oxygen
release, most significantly promoted MSC survival under ischemic con-
dition, and did not induce oxidative stress, they were used in vivo to
investigate whether the implantation of MSCs and oxygen-release mi-
crospheres can promote cell survival and tissue regeneration in ischemic
limbs.
3
.3. Effect of oxygen-release microspheres on MSC survival under hypoxic
condition
The survival of transplanted MSCs in ischemic environment is pivotal
3
82

Y. Guan et al.
Journal of Controlled Release 331 (2021) 376–389
Fig. 6. Effect of oxygen-release micro-
spheres on MSC survival and ROS produc-
tion under hypoxia. (A) dsDNA content of
MSCs encapsulated in PNHA gel with and
without oxygen-release microspheres during
2
8 days of culture; (B) Fluorescent images of
live MSCs encapsulated in PNHA gel with
and without oxygen-release microspheres
during 28 days of culture; (C) Cellular oxy-
gen content measurement of MSCs encap-
sulated in PNHA gel with and without
microspheres after 48 h; (D) ROS content
expressed by MSCs encapsulated in PNHA
gel with and without oxygen-release micro-
spheres during 28 days of culture. *p < 0.05,
**p < 0.01, ***p < 0.001.
3
.4. Effect of oxygen release on MSC survival and paracrine effects in
results that the released oxygen increased MSC intracellular oxygen
content (Fig. 6C), it is likely that the oxygen released in vivo also
augmented intracellular oxygen content in the transplanted MSCs,
leading to enhanced cell survival. The oxygen content either in the MSCs
or ischemic tissues remains to be measured since current techniques
cannot non-invasively and long-term detect tissue oxygen content.
To determine whether the released oxygen enhanced MSC prolifer-
ation in the ischemic limbs, Ki67 staining was performed (Fig. 7C). The
Ki67+/CMDil+ cells were identified as proliferating MSCs. The cell
density in the Gel/MSC/ORM group was raised more than 4-fold
compared to that in the Gel/MSC group (p < 0.001, Fig. 7D),
ischemic limbs
To determine whether the oxygen-release microspheres (ORM) can
increase MSC survival in ischemic limbs, the microspheres and live cell
tracker CM-Dil-labeled MSCs were encapsulated in the PNHA gel, and
injected into thigh muscles. Four weeks after implantation, live MSCs
were imaged. The number of CMDil+ MSCs was apparently much higher
in the Gel/MSC/ORM group than in the Gel/MSC group (Fig. 7A&B).
The quantitative live MSC density of the Gel/MSC/ORM group was more
than 3 times of the Gel/MSC group (p < 0.001). Based on the in vitro
3
83

Y. Guan et al.
Journal of Controlled Release 331 (2021) 376–389
Fig. 7. In vivo MSC survival, proliferation and paracrine effect. (A) Fluorescent images of MSCs in ischemic limbs 28 days after implantation. MSCs were pre-labeled
with live cell track CM-Dil (red) before implantation. Scale bar = 50 m; (B) MSC density in ischemic limbs 28 days after implantation; (C) Immunohistochemical
staining of Ki67 (green) of limb tissue sections 28 days after CLI surgery. Nuclei were stained with Hoechst 33342 (blue) and MSCs were pre-labeled with CM-Dil
μ
(
red) before implantation; (D) Quantification of overall Ki67+ cell density in ischemic limbs. The Ki67+ cells were proliferating cells; (E) Quantification of Ki67+/
CM-Dil + cell density. The Ki67+/CM-Dil + cells were proliferating MSCs that were transplanted into ischemic limbs; (F) Expression of bFGF, PDGF-BB and HGF in
ischemic limb tissues 28 days after surgery. *p < 0.05, **p < 0.01, ***p < 0.001. (For interpretation of the references to colour in this figure legend, the reader is
referred to the web version of this article.)
demonstrating that the released oxygen promoted MSC proliferation
besides survival. The Gel/MSC/ORM group also had significantly
greater overall proliferating cell density than the Gel/MSC group (p <
MSCs are known to promote cell survival and tissue regeneration
mainly by paracrine effects [63]. To determine how MSC paracrine ef-
fects were expressed in the ischemic limbs, the prosurvival growth factor
expressions were quantified at the mRNA level at day 28. Among
different prosurvival growth factors, the bFGF and PDGF-BB expressions
were significantly increased in the Gel/MSC and Gel/MSC/ORM groups
compared with Surgery and Gel groups (Fig. 7F). It is possible that these
growth factors promoted host cell survival. The Gel/MSC/ORM group
exhibited significantly higher bFGF and PDGF-BB expressions than the
Gel/MSC group (p < 0.001). Besides bFGF and PDGFBB, the Gel/MSC/
0
.001, Fig. 7E). It is possible that the released oxygen also improved host
cell survival. This is consistent with our previous study where delivery of
oxygen into ischemic hearts ameliorated cardiac cell survival [41].
Notably, the groups with MSCs exhibited more abundant Ki67 expres-
sion than the groups without MSCs (Surgery and Gel groups, Fig. 7 C &
D), illuminating that the survived MSCs played an important role to the
host cell survival.
3
84

Y. Guan et al.
Journal of Controlled Release 331 (2021) 376–389
ORM group also had significantly greater HGF expression (p < 0.001),
whereas the Gel/MSC, Surgery and Gel groups had similar expression (p
and paracrine effects, and host cell survival accelerated tissue
vascularization.
>
0.05). The above results reveal that the released oxygen significantly
To visualize the blood vessels in ischemic limbs, isolectin and SMA
α
promoted MSC paracrine effects. Stem cell paracrine effects are depen-
dent on the oxygen level. Studies have shown that a relatively low ox-
ygen condition (5–20%) may increase cell paracrine effects [64–68].
However, an extremely low oxygen condition (<1%) largely decreases
paracrine effects [69]. Our results demonstrate that the amount of
released oxygen was in the range that increases MSC paracrine effects.
stainings were performed for tissues harvested at day 28 (Fig. 8C).
Consistent with blood perfusion results, the implantation of Gel/MSC
significantly increased the blood vessel density than the Surgery and Gel
groups (p < 0.001, Fig. 8D). The implantation of Gel/MSC/ORM further
increased vessel density to 90% of that of the normal limbs (Fig. 8D).
The co-delivery of MSCs and oxygen-release microspheres also largely
promoted vessel maturation. The isolectin+/αSMA+ mature vessel
density was significantly higher than the delivery of MSCs only (Fig. 8E).
3
.5. Efficacy of oxygen-release microspheres and MSCs in promoting
As the transplanted MSCs did not differentiate into isolectin+ endo-
vascularization in ischemic limb
thelial cells or
αSMA+ smooth muscle cells, the enhanced vasculariza-
tion is likely resulted from upregulated bFGF, PDGF-BB, and HGF
expressions (Fig. 7F). These growth factors have angiogenic effect
To determine whether enhanced MSC survival and paracrine effect,
and increased host cell survival accelerated tissue vascularization, blood
perfusion in the ischemic limbs was monitored by Laser Doppler, and
blood vessels were characterized by immunohistology. The blood
perfusion in the Surgery group was slowly increased during the 4-week
experimental period with 41% blood perfusion at week 4 (Fig. 8 A & B).
The injection of PNHA gel only (Gel group) did not significantly enhance
the blood perfusion. In contrast, the encapsulation of MSCs in the
hydrogel (Gel/MSC group) significantly increased blood perfusion
compared to Gel group at days 14, 21 and 28 (p < 0.05). The fastest
blood flow recovery was found for the Gel/MSC/ORM group where both
MSCs and oxygen-release microspheres were encapsulated in the
hydrogel (Fig. 8 A & B). The blood perfusion was ~90% of that of the
normal limb at day 28. The blood flow result proved that the incorpo-
ration of microspheres could enhance the survival of MSCs and host cells
under ischemia. These results demonstrate that enhanced MSC survival
[
7,55,70]. The upregulated PDGF-BB can also recruit smooth muscle
cells to stimulate vessel maturation. Besides paracrine effects, MSCs may
also stabilize blood vessels by acting as pericytes [25–27]. It remains to
be investigated whether delivery of MSCs and oxygen-release micro-
spheres promoted the formation of functional vessels in ischemic limbs,
which can be determined by performing vascular perfusion with
fluorescent-labeled lectin following by visualizing the functional blood
vessels.
3.6. Effect of oxygen-release microspheres and MSCs on skeletal muscle
regeneration and tissue inflammation in ischemic limbs
Vascularization and myogenesis are two goals for ischemic tissue
regeneration. Previous studies have demonstrated that myogenesis
Fig. 8. Delivery of oxygen-release microspheres and MSCs promoted tissue angiogenesis. (A) Laser Doppler perfusion images of non-surgery group and surgery
groups with different treatments for 28 days; (B) Blood perfusion of ischemic limbs for 28 days. The results were normalized to the non-surgery limb for each animal;
(
C) Immunohistochemical staining of isolectin (green) and
α
-SMA (red) of limb tissue sections 28 days after CLI surgery. Nuclei were stained with Hoechst 33342
(
blue); (D) Quantification of total blood vessel density; (E) Quantification of mature blood vessel density. *p < 0.05, ***p < 0.001. (For interpretation of the ref-
erences to colour in this figure legend, the reader is referred to the web version of this article.)
85
3

Y. Guan et al.
Journal of Controlled Release 331 (2021) 376–389
largely depends on vascularization [71]. To determine myogenesis in
the ischemic limbs, H&E and MHC stainings were used (Fig. 9 A & B). In
the Surgery and Gel groups, the skeletal muscle was degenerated as the
muscle fibers were highly isolated (Fig. 9 A & B). The injection of Gel/
MSC and Gel/MSC/ORM promoted skeletal muscle regeneration, as the
muscle fiber diameter was significantly increased compared to the
Surgery and Gel groups (p < 0.001. Fig. 9C). The Gel/MSC/ORM group
exhibited significantly larger muscle fiber diameter than the Gel/MSC
group (p < 0.001. Fig. 9C). In addition, the blood vessel distribution in
the muscle fibers was similar to that in the healthy hindlimbs (Fig. 9B).
Regenerating skeletal muscle contains central nucleus [72]. Consistent
with muscle fiber results, the central nucleus density was remarkably
greater in the Gel/MSC/ORM group than other groups (Fig. 9D). The
above results demonstrate that co-delivery of MSCs and oxygen-release
microspheres significantly promoted skeletal muscle regeneration. It is
speculated that the enhanced MSC paracrine effects, and improved
survival of skeletal muscle cells by released oxygen accelerated myo-
genesis. It is also possible that the MSC paracrine effects activated
muscle resident stem cells to proliferate and differentiate to form new
myofibers [73].
(Fig. 10). Compared with the Non-surgery group, the Surgery group had
a significantly higher density of F4/80+ macrophages. The injection of
Gel group, Gel/MSC group, and Gel/MSC/ORM group did not augment
tissue inflammation as these groups had similar density of F4/80+
macrophages as the Surgery group (p > 0.05). These results demonstrate
that the PNHA and oxygen-release microspheres had excellent
biocompatibility.
The above results demonstrate that the oxygen-release microspheres
with hypoxia sensitivity were able to release oxygen in response to
environmental oxygen level. The released oxygen increased cell survival
under ischemia while avoiding excessive oxygenation that may cause
ROS formation and inflammation. Various studies have shown that
excessive oxygenation leads to ischemia/reperfusion injury, character-
ized by high oxidative stress and inflammation [74–80]. The oxygen-
release microspheres in this work have the potential to reduce
ischemia/reperfusion injury.
4. Conclusion
In this study, oxygen-release microspheres whose oxygen release
kinetics was responsive to environmental oxygen level were developed.
The microspheres released oxygen faster in the lower oxygen environ-
ment, and slower in the higher oxygen environment. The released oxy-
gen improved MSC survival without elevating oxidative stress in vitro.
In the ischemic limbs, the oxygen-release microspheres significantly
The implantation of MSCs, oxygen-release microspheres and PNHA
hydrogel may lead to immune response and inflammation. Excessive
inflammation has been shown to inhibit the activation of the quiescent
population of muscle-resident stem cells for skeletal muscle regenera-
tion [73]. F4/80 staining was used to investigate tissue inflammation
Fig. 9. Delivery of oxygen-release microspheres and MSCs enhanced skeletal muscle regeneration. (A) H&E staining of limb tissue sections 28 days after CLI surgery.
m; (B) Immunohistochemical staining of MHC (red) and isolectin (green) of limb tissue sections 28 days after CLI surgery. Nuclei were stained with
Scale bar = 50
μ
Hoechst 33342 (blue); (C) Quantification of muscle fiber diameter; (D) Quantification of central nucleus density. *p < 0.05, ***p < 0.001. (For interpretation of the
references to colour in this figure legend, the reader is referred to the web version of this article.)
3
86

Y. Guan et al.
Journal of Controlled Release 331 (2021) 376–389
Fig. 10. Biocompatibility of the oxygen-release microspheres. Immunohistochemical staining of F4/80 (green) of limb tissue sections 28 days after CLI surgery and
the quantification of F4/80 positive cell percentage. Nuclei were stained with DRAQ5 (blue). Scale bar = 50
μ
m. (For interpretation of the references to colour in this
figure legend, the reader is referred to the web version of this article.)
improved MSC survival, proliferation and paracrine effects, and host cell
survival, leading to accelerated tissue vascularization and skeletal
muscle regeneration. The microspheres possessed excellent biocompat-
ibility. These microspheres provided a smart way of delivering oxygen to
ischemic tissues to enhance exogenous and endogenous cell survival.
The environment-responsive oxygen release can avoid excessive oxygen
release that may increase oxidative stress and cause cell death.
[3] W. Leppert, R. Zajaczkowska, J. Wordliczek, J. Dobrogowski, J. Woron,
M. Krzakowski, Pathophysiology and clinical characteristics of pain in most
common locations in cancer patients, J. Physiol. Pharmacol. 67 (2016) 787–799.
[
4] C.N. Hess, L. Norgren, G.M. Ansel, W.H. Capell, J.P. Fletcher, F.G.R. Fowkes, et al.,
A structured review of antithrombotic therapy in peripheral artery disease with a
focus on revascularization: a TASC (InterSociety consensus for the Management of
Peripheral Artery Disease) initiative, Circulation 135 (2017) 2534–2555.
[
[
5] L. Uccioli, M. Meloni, V. Izzo, L. Giurato, S. Merolla, R. Gandini, Critical limb
ischemia: current challenges and future prospects, Vasc. Health Risk Manag. 14
(
2018) 63–74.
6] M. Hassanshahi, S. Khabbazi, Y. Peymanfar, A. Hassanshahi, Z. Hosseini-Khah, Y.
W. Su, et al., Critical limb ischemia: current and novel therapeutic strategies,
J. Cell. Physiol. 234 (2019) 14445–14459.
Credit author statement
[
[
7] G.O. Ouma, B. Zafrir, E.R. Mohler III, M.Y. Flugelman, Therapeutic angiogenesis in
critical limb ischemia, Angiology 64 (2013) 466–480.
YG, NG and JG developed ideas. YG, NG, HN and YD performed
experiments. YG, NG and JG wrote manuscript.
8] F. Sanada, Y. Taniyama, J. Azuma, I.-I. Yuka, Y. Kanbara, M. Iwabayashi, et al.,
Therapeutic angiogenesis by gene therapy for critical limb ischemia: choice of
biological agent, in: Immunology, Endocrine & Metabolic Agents in Medicinal
Chemistry (Formerly Current Medicinal Chemistry-Immunology, Endocrine and
Metabolic Agents) 14, 2014, pp. 32–39.
Acknowledgement
[
9] M. Qadura, D.C. Terenzi, S. Verma, M. Al-Omran, D.A. Hess, Concise review: cell
therapy for critical limb ischemia: an integrated review of preclinical and clinical
studies, Stem Cells 36 (2018) 161–171.
This work was supported by US National Institutes of Health
(
R01HL138175, R01HL138353, R01EB022018, R01AG056919, and
[
10] E. Tateishi-Yuyama, H. Matsubara, T. Murohara, U. Ikeda, S. Shintani, H. Masaki,
et al., Therapeutic angiogenesis for patients with limb ischaemia by autologous
transplantation of bone-marrow cells: a pilot study and a randomised controlled
trial, Lancet 360 (2002) 427–435.
R01AR077616), and National Science Foundation (1922857).
Appendix A. Supplementary data
[
[
11] R. Al-Rifai, P. Nguyen, N. Bouland, C. Terryn, L. Kanagaratnam, G. Poitevin, et al.,
In vivo efficacy of endothelial growth medium stimulated mesenchymal stem cells
derived from patients with critical limb ischemia, J. Transl. Med. 17 (2019) 261.
12] M.G. MacAskill, J. Saif, A. Condie, M.A. Jansen, T.J. MacGillivray, A.A. Tavares, et
al., Robust revascularization in models of limb ischemia using a clinically
translatable human stem cell-derived endothelial cell product, Mol. Ther. 26
(2018) 1669–1684.
Supplementary data to this article can be found online at https://doi.
org/10.1016/j.jconrel.2021.01.034.
References
[
[
13] K.J. Portalska, A. Leferink, N. Groen, H. Fernandes, L. Moroni, C. van Blitterswijk,
et al., Endothelial differentiation of mesenchymal stromal cells, PLoS One 7 (2012),
e46842.
[
[
1] F. Sanada, Y. Taniyama, J. Muratsu, R. Otsu, H. Shimizu, H. Rakugi, et al., Gene-
therapeutic strategies targeting angiogenesis in peripheral artery disease,
Medicines 5 (2018) 31.
14] K. Zhi, Z. Gao, J. Bai, Y. Wu, S. Zhou, M. Li, et al., Application of adipose-derived
stem cells in critical limb ischemia, Frontiers in bioscience 19 (2014) 768–776
2] P. Bar ´c , M. Antkiewicz, B. Sliwa, D. Baczynska, W. Witkiewicz, J.P. Skora,
Treatment of critical limb ischemia by pIRES/VEGF165/HGF Administration, Ann.
Vasc. Surg. 60 (2019) 346–354.
´
´
´
(
Landmark edition).
3
87

Y. Guan et al.
Journal of Controlled Release 331 (2021) 376–389
[
[
[
15] X. Wang, J. Zhang, W. Cui, Y. Fang, L. Li, S. Ji, et al., Composite hydrogel modified
by IGF-1C domain improves stem cell therapy for limb ischemia, ACS Appl. Mater.
Interfaces 10 (2018) 4481–4493.
[47] J. Xiang, C. Wan, R. Guo, D. Guo, Is hydrogen peroxide a suitable apoptosis inducer
for all cell types? Biomed. Res. Int. 2016 (2016).
[48] S. Kizaka-Kondoh, H. Konse-Nagasawa, Significance of nitroimidazole compounds
and hypoxia-inducible factor-1 for imaging tumor hypoxia, Cancer Sci. 100 (2009)
1366–1373.
16] L. Debin, J. Youzhao, L. Ziwen, L. Xiaoyan, Z. Zhonghui, C. Bing, Autologous
transplantation of bone marrow mesenchymal stem cells on diabetic patients with
lower limb ischemia, J. Med. Coll. PLA 23 (2008) 106–115.
[49] J. Yu, Y. Zhang, Y. Ye, R. DiSanto, W. Sun, D. Ranson, et al., Microneedle-array
patches loaded with hypoxia-sensitive vesicles provide fast glucose-responsive
insulin delivery, Proc. Natl. Acad. Sci. 112 (2015) 8260–8265.
[50] M. Gülden, A. Jess, J. Kammann, E. Maser, H. Seibert, Cytotoxic potency of H2O2
in cell cultures: impact of cell concentration and exposure time, Free Radic. Biol.
Med. 49 (2010) 1298–1305.
17] P.A. Tebebi, S.J. Kim, R.A. Williams, B. Milo, V. Frenkel, S.R. Burks, et al.,
Improving the therapeutic efficacy of mesenchymal stromal cells to restore
perfusion in critical limb ischemia through pulsed focused ultrasound, Sci. Rep. 7
(
2017) 41550.
[
[
18] A. Liew, T. O’Brien, Therapeutic potential for mesenchymal stem cell
transplantation in critical limb ischemia, Stem Cell Res Ther 3 (2012) 28.
19] Q. Lian, Y. Zhang, J. Zhang, H.K. Zhang, X. Wu, Y. Zhang, et al., Functional
mesenchymal stem cells derived from human induced pluripotent stem cells
attenuate limb ischemia in mice, Circulation 121 (2010) 1113–1123.
20] A. Abdal Dayem, S.B. Lee, K. Kim, K.M. Lim, Jeon T-i, J. Seok, et al., Production of
Mesenchymal Stem Cells through Stem Cell Reprogramming, International journal
of molecular sciences 20 (2019) 1922.
[51] Z. Fan, Z. Xu, H. Niu, Y. Sui, H. Li, J. Ma, et al., Spatiotemporal delivery of basic
fibroblast growth factor to directly and simultaneously attenuate cardiac fibrosis
and promote cardiac tissue vascularization following myocardial infarction,
J. Control. Release 311 (2019) 233–244.
[
[
[52] H. Niu, C. Li, Y. Guan, Y. Dang, X. Li, Z. Fan, et al., High oxygen preservation
hydrogels to augment cell survival under hypoxic condition, Acta Biomaterialia
105 (2020) 56–67.
21] P. Yan, Q. Li, L. Wang, P. Lu, K. Suzuki, Z. Liu, et al., FOXO3-engineered human
ESC-derived vascular cells promote vascular protection and regeneration, Cell stem
cell 24 (2019) 447–461, e8.
[53] H. Niu, X. Li, H. Li, Z. Fan, J. Ma, J. Guan, Thermosensitive, fast gelling,
photoluminescent, highly flexible, and degradable hydrogels for stem cell delivery,
Acta Biomater. 83 (2019) 96–108.
[
[
[
[
22] Y. Fujita, A. Kawamoto, Stem cell-based peripheral vascular regeneration, Adv.
Drug Deliv. Rev. 120 (2017) 25–40.
[54] M. Quaranta, S.M. Borisov, I. Klimant, Indicators for optical oxygen sensors,
Bioanal. Rev. 4 (2012) 115–157.
23] V. Gorenoi, M.U. Brehm, A. Koch, A. Hagen, Growth factors for angiogenesis in
peripheral arterial disease, in: Cochrane Database of Systematic Reviews, 2017.
24] M.E. Bernardo, W.E. Fibbe, Mesenchymal stromal cells: sensors and switchers of
inflammation, Cell Stem Cell 13 (2013) 392–402.
[55] Y. Xu, M. Fu, Z. Li, Z. Fan, X. Li, Y. Liu, et al., A prosurvival and proangiogenic stem
cell delivery system to promote ischemic limb regeneration, Acta Biomater. 31
(2016) 99–113.
[56] A. Uccelli, T. Wolff, P. Valente, N. Di Maggio, M. Pellegrino, L. Gürke, et al.,
Vascular endothelial growth factor biology for regenerative angiogenesis, Swiss
Med. Wkly. 149 (2019).
25] K. Gaengel, G. Genov ´e , A. Armulik, C. Betsholtz, Endothelial-mural cell signaling in
vascular development and angiogenesis, Arterioscler. Thromb. Vasc. Biol. 29
(
2009) 630–638.
[57] J.E. Markkanen, T.T. Rissanen, A. Kivel a¨ , S. Yl a¨ -Herttuala, Growth factor-induced
therapeutic angiogenesis and arteriogenesis in the heart–gene therapy, Cardiovasc.
Res. 65 (2005) 656–664.
[
[
26] A. Blocki, Y. Wang, M. Koch, P. Peh, S. Beyer, P. Law, et al., Not all MSCs can act as
pericytes: functional in vitro assays to distinguish pericytes from other
mesenchymal stem cells in angiogenesis, Stem Cells Dev. 22 (2013) 2347–2355.
27] M. Crisan, S. Yap, L. Casteilla, C.W. Chen, M. Corselli, T.S. Park, et al.,
A perivascular origin for mesenchymal stem cells in multiple human organs, Cell
Stem Cell 3 (2008) 301–313.
[58] G. Solaini, A. Baracca, G. Lenaz, G. Sgarbi, Hypoxia and mitochondrial oxidative
metabolism, Biochimica et Biophysica Acta (BBA)-Bioenergetics 1797 (2010)
1171–1177.
[59] M. Redza-Dutordoir, D.A. Averill-Bates, Activation of apoptosis signalling
pathways by reactive oxygen species. Biochimica et Biophysica Acta (BBA)-
molecular, Cell Res. 1863 (2016) 2977–2992.
[
[
28] D.P. Sieveking, M.K. Ng, Cell therapies for therapeutic angiogenesis: back to the
bench, Vasc. Med. 14 (2009) 153–166.
29] F. Hached, C. Vinatier, C. Le Visage, H. Gond ´e , J. Guicheux, G. Grimandi, et al.,
Biomaterial-assisted cell therapy in osteoarthritis: from mesenchymal stem cells to
cell encapsulation, Best Pract. Res. Clin. Rheumatol. 31 (2017) 730–745.
30] N.K. Gupta, E.J. Armstrong, S.A. Parikh, The current state of stem cell therapy for
peripheral artery disease, Curr. Cardiol. Rep. 16 (2014) 447.
[60] A. Cossu, A.M. Posadino, R. Giordo, C. Emanueli, A.M. Sanguinetti, A. Piscopo, et
al., Apricot melanoidins prevent oxidative endothelial cell death by counteracting
mitochondrial oxidation and membrane depolarization, PLoS One 7 (2012),
e48817.
[
[
[
[61] V. Houot, P. Etienne, A.S. Petitot, S. Barbier, J.P. Blein, L. Suty, Hydrogen peroxide
induces programmed cell death features in cultured tobacco BY-2 cells, in a dose-
dependent manner, J. Exp. Bot. 52 (2001) 1721–1730.
31] X.L. Aranguren, C.M. Verfaillie, A. Luttun, Emerging hurdles in stem cell therapy
for peripheral vascular disease, J. Mol. Med. 87 (2009) 3.
32] S.E. Bae, J.S. Son, K. Park, D.K. Han, Fabrication of covered porous PLGA
microspheres using hydrogen peroxide for controlled drug delivery and
regenerative medicine, J. Control. Release 133 (2009) 37–43.
[62] S. Lee, E. Choi, M.-J. Cha, K.-C. Hwang, Cell adhesion and long-term survival of
transplanted mesenchymal stem cells: a prerequisite for cell therapy, Oxidative
Med. Cell. Longev. 2015 (2015).
[
33] S.-M. Ng, J.-Y. Choi, H.-S. Han, J.-S. Huh, J.O. Lim, Novel microencapsulation of
potential drugs with low molecular weight and high hydrophilicity: hydrogen
peroxide as a candidate compound, Int. J. Pharm. 384 (2010) 120–127.
34] S.H. Oh, C.L. Ward, A. Atala, J.J. Yoo, B.S. Harrison, Oxygen generating scaffolds
for enhancing engineered tissue survival, Biomaterials 30 (2009) 757–762.
35] K. Chin, S.F. Khattak, S.R. Bhatia, S.C. Roberts, Hydrogel-perfluorocarbon
composite scaffold promotes oxygen transport to immobilized cells, Biotechnol.
Prog. 24 (2008) 358–366.
[63] H.M. Kwon, S.-M. Hur, K.-Y. Park, C.-K. Kim, Y.-M. Kim, H.-S. Kim, et al., Multiple
paracrine factors secreted by mesenchymal stem cells contribute to angiogenesis,
Vasc. Pharmacol. 63 (2014) 19–28.
[
[
[64] H. Thangarajah, I.N. Vial, E. Chang, S. El-Ftesi, M. Januszyk, E.I. Chang, et al.,
IFATS collection: Adipose stromal cells adopt a proangiogenic phenotype under the
influence of hypoxia, Stem cells (Dayton, Ohio) 27 (2009) 266–274.
[65] J. Rehman, D. Traktuev, J. Li, S. Merfeld-Clauss, C.J. Temm-Grove, J.E. Bovenkerk,
et al., Secretion of angiogenic and antiapoptotic factors by human adipose stromal
cells, Circulation 109 (2004) 1292–1298.
[
36] A. Wijekoon, N. Fountas-Davis, N.D. Leipzig, Fluorinated methacrylamide chitosan
hydrogel systems as adaptable oxygen carriers for wound healing, Acta Biomater. 9
[66] C.M. Cameron, F. Harding, W.S. Hu, D.S. Kaufman, Activation of hypoxic response
in human embryonic stem cell-derived embryoid bodies, Experimental Biology and
Medicine (Maywood, NJ) 233 (2008) 1044–1057.
(
2013) 5653–5664.
[
[
[
37] J. Chal, O. Pourqui ´e , Making muscle: skeletal myogenesis in vivo and in vitro,
Development 144 (2017) 2104–2122.
[67] S.C. Hung, R.R. Pochampally, S.C. Chen, S.C. Hsu, D.J. Prockop, Angiogenic effects
of human multipotent stromal cell conditioned medium activate the PI3K-Akt
pathway in hypoxic endothelial cells to inhibit apoptosis, increase survival, and
stimulate angiogenesis, Stem cells (Dayton, Ohio) 25 (2007) 2363–2370.
[68] K. Tamama, H. Kawasaki, S.S. Kerpedjieva, J. Guan, R.K. Ganju, C.K. Sen,
Differential roles of hypoxia inducible factor subunits in multipotential stromal
cells under hypoxic condition, J. Cell. Biochem. 112 (2011) 804–817.
[69] Y. Xu, M. Fu, Z. Li, Z. Fan, X. Li, Y. Liu, et al., A prosurvival and proangiogenic stem
cell delivery system to promote ischemic limb regeneration, Acta Biomater. 31
(2016) 99–113.
38] B.H. Annex, Therapeutic angiogenesis for critical limb ischaemia, Nat. Rev.
Cardiol. 10 (2013) 387.
39] I.E. Hoefer, N. van Royen, I.R. Buschmann, J.J. Piek, W. Schaper, Time course of
arteriogenesis following femoral artery occlusion in the rabbit, Cardiovasc. Res. 49
(
2001) 609–617.
[
40] P.I. Makarevich, M.A. Boldyreva, E.V. Gluhanyuk, A.Y. Efimenko, K.V. Dergilev, E.
K. Shevchenko, et al., Enhanced angiogenesis in ischemic skeletal muscle after
transplantation of cell sheets from baculovirus-transduced adipose-derived stromal
cells expressing VEGF165, Stem Cell Research Therapy 6 (2015) 204.
41] Z. Fan, Z. Xu, H. Niu, N. Gao, Y. Guan, C. Li, et al., An injectable oxygen release
system to augment cell survival and promote cardiac repair following myocardial
infarction, Sci. Rep. 8 (2018) 1371.
[
[
[
[70] R. Cao, E. Bråkenhielm, R. Pawliuk, D. Wariaro, M.J. Post, E. Wahlberg, et al.,
Angiogenic synergism, vascular stability and improvement of hind-limb ischemia
by a combination of PDGF-BB and FGF-2, Nat. Med. 9 (2003) 604–613.
[71] C. Christov, F. Chr ´e tien, R. Abou-Khalil, G. Bassez, G. Vallet, F.-J. Authier, et al.,
Muscle satellite cells and endothelial cells: close neighbors and privileged partners,
Mol. Biol. Cell 18 (2007) 1397–1409.
42] Z. Li, X. Guo, J. Guan, An oxygen release system to augment cardiac progenitor cell
survival and differentiation under hypoxic condition, Biomaterials 33 (2012)
5
914–5923.
43] Y. Guan, H. Niu, Y. Dang, N. Gao, J. Guan, Photoluminescent oxygen-release
microspheres to image the oxygen release process in vivo, Acta Biomater. 115
[72] B. Cadot, V. Gache, E.R. Gomes, Moving and positioning the nucleus in skeletal
muscle - one step at a time, Nucleus (Austin, Tex) 6 (2015) 373–381.
[73] A.J. Wagers, I.M. Conboy, Cellular and molecular signatures of muscle
regeneration: current concepts and controversies in adult myogenesis, Cell 122
(2005) 659–667.
(
2020) 333–342.
[
[
44] T.M. Buttke, P.A. Sandstrom, Oxidative stress as a mediator of apoptosis,
Immunology Today 15 (1994) 7.
45] H. Fujita, I. Morita, Murota S-i, Hydrogen peroxide induced apoptosis of
endothelial cells concomitantly with cycloheximide, J. Atheroscler. Thromb. 7
[74] J. Lutz, K. Thürmel, U. Heemann, Anti-inflammatory treatment strategies for
ischemia/reperfusion injury in transplantation, J. Inflamm. 7 (2010) 1–8.
[75] W. Yao, L.W. Tai, Y. Liu, Z. Hei, H. Li, Oxidative Stress and Inflammation
Interaction in Ischemia Reperfusion Injury: Role of Programmed Cell Death,
Hindawi, 2019.
(
2000) 209–215.
[
46] M. Singh, H. Sharma, N. Singh, Hydrogen peroxide induces apoptosis in HeLa cells
through mitochondrial pathway, Mitochondrion 7 (2007) 367–373.
3
88

Y. Guan et al.
Journal of Controlled Release 331 (2021) 376–389
[
[
76] M. Sasaki, T. Joh, Inflammation and ischemia-reperfusion injury in gastrointestinal
tract and antioxidant, protective agents, J. Clin. Biochem. Nutr. 40 (2007) 1–12.
77] D.K. de Vries, K.A. Kortekaas, D. Tsikas, L.G. Wijermars, C.J. van Noorden, M.-
T. Suchy, et al., Oxidative damage in clinical ischemia/reperfusion injury: a
reappraisal, Antioxid. Redox Signal. 19 (2013) 535–545.
[79] R.S. Ferrari, C.F. Andrade, Oxidative stress and lung ischemia-reperfusion injury,
Oxidative Med. Cell. Longev. 2015 (2015).
[80] M. Kishimoto, J. Suenaga, H. Takase, K. Araki, T. Yao, T. Fujimura, et al., Oxidative
stress-responsive apoptosis inducing protein (ORAIP) plays a critical role in
cerebral ischemia/reperfusion injury, Sci. Rep. 9 (2019) 1–12.
[
78] G.A. Kurian, R. Rajagopal, S. Vedantham, M. Rajesh, The role of oxidative stress in
myocardial ischemia and reperfusion injury and remodeling: revisited, Oxidative
Med. Cell. Longev. 2016 (2016).
3
89