Tough and cell compatible chitosan physical hydrogels for mouse bone mesenchymal stem cells in vitro

Research Article
www.acsami.org

Tough and Cell-Compatible Chitosan Physical Hydrogels for Mouse
Bone Mesenchymal Stem Cells in Vitro
Beibei Ding,†,⊥ Huichang Gao,‡,⊥ Jianhui Song,§ Yaya Li,† Lina Zhang,† Xiaodong Cao,*,‡ Min Xu,§
and Jie Cai*,†
†

College of Chemistry & Molecular Sciences, Wuhan University, Wuhan 430072, People’s Republic of China
School of Materials Science and Engineering, South China University of Technology, Guangzhou 510641, People’s Republic of
China
§
Department of Physics, Shanghai Key Laboratory of Magnetic Resonance, East China Normal University, Shanghai 200062, People’s
Republic of China
‡

S Supporting Information
*

ABSTRACT: Most hydrogels involve synthetic polymers and organic
cross-linkers that cannot simultaneously fulfill the mechanical and cellcompatibility requirements of biomedical applications. We prepared a new
type of chitosan physical hydrogel with various degrees of deacetylation
(DDs) via the heterogeneous deacetylation of nanoporous chitin hydrogels
under mild conditions. The DD of the chitosan physical hydrogels ranged
from 56 to 99%, and the hydrogels were transparent and mechanically
strong because of the extra intra- and intermolecular hydrogen bonding
interactions between the amino and hydroxyl groups on the nearby chitosan
nanofibrils. The tensile strength and Young’s modulus of the chitosan
physical hydrogels were 3.6 and 7.9 MPa, respectively, for a DD of 56% and
increased to 12.1 and 92.0 MPa for a DD of 99% in a swelling equilibrium

state. In vitro studies demonstrated that mouse bone mesenchymal stem
cells (mBMSCs) cultured on chitosan physical hydrogels had better
adhesion and proliferation than those cultured on chitin hydrogels. In particular, the chitosan physical hydrogels promoted the
differentiation of the mBMSCs into epidermal cells in vitro. These materials are promising candidates for applications such as
stem cell research, cell therapy, and tissue engineering.
KEYWORDS: chitosan, hydrogels, heterogeneous deacetylation, mechanical properties, cell-compatibility

■

INTRODUCTION
Hydrogels, which are three-dimensional polymeric materials
with high water contents and diverse physical properties, have
been extensively used in food, cosmetics, drug-delivery devices,
and other applications. 1 The emergence of potential
applications for hydrogels include stem cell and cancer research,
cell therapy, tissue engineering, immunomodulation, and in
vitro diagnostics.2−7 However, the disadvantage of traditional
hydrogels is poor mechanic properties due to high water
content and structural defects at swollen state. Therefore,
several new approaches have been introduced to fabricate
mechanically tougher hydrogels,8 including forming supermolecular interactions,9,10 chemical cross-linking,11,12 and a
double network structure.13−15
Unfortunately, most hydrogels involve synthetic polymers
and organic cross-linkers that cannot simultaneously fulfill the
mechanical and cell-compatibility requirements of biomedical
applications. Alternatively, natural polymers, such as alginate,
chitosan, hyaluronidase, and collagen, have been shown to be
promising biomaterials.16−18 Among them, chitosan, a natural
amino polysaccharide derived from chitin, which is the main
© 2016 American Chemical Society

component of the exoskeletons of crustaceans (e.g., crabs and
shrimp), has received considerable attention because of its
excellent biodegradability, biocompatibility, and bioactivity.19−24 The exploitation of chitosan hydrogels for biomaterials
is, however, limited by the poor solubility and mechanical
integrity, difficulty in fabrication, and requirement for organic
cross-linkers.25−31
In our previous works, nanoporous chitin hydrogels prepared
in aqueous NaOH/urea showed remarkable mechanical
strength and biocompatibility.32,33 These materials were
characterized as having a large interior space with a threedimensional open network structure, and thus, they may be
directly converted into chitosan physical hydrogels via reactions
with deacetylation reagents under mild conditions. In this work,
we demonstrate the toughness and excellent cell-compatibility
of chitosan physical hydrogels based on the in situ
heterogeneous deacetylation of nanoporous chitin hydrogels
Received: May 4, 2016
Accepted: July 13, 2016
Published: July 13, 2016
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solvent-exchanged with absolute ethanol and then dried from

supercritical CO2 to give dried gels.
Fourier transform infrared spectroscopy (FT-IR) analyses were
carried out on a FT-IR spectrometer (Nicolet 5700 FTIR
Spectrometer, MA). The powdered samples and KBr were mixed
and loaded into the sample holder. The spectra in the range of 400−
4000 cm−1 were collected in 32 scans at 4 cm−1 resolution.
The polymorphisms of the crystals in the chitin and chitosan gels
were determined by X-ray diffraction (XRD, D8-Advance, Bruker,
USA) over the 2θ range from 5° to 40° with 40 kV and 40 mA Nifiltered CuKα radiation. The powdered samples were used to eliminate
the effect of the crystalline orientation. The peak position and
crystallinity (χc) of the chitin and chitosan gels were estimated from
multipeak fitting of the XRD profiles.
Solid-state cross-polarization/magnetic angle spinning (CP/MAS)
13
C nuclear magnetic resonance (NMR) spectra were collected on a
Bruker AVANCE-300 Spectrometer (13C frequency = 75.4 MHz) with
a standard 4 mm rotor at ambient temperature. The spinning rate was
kept at 5.0 kHz. The contact time and relaxation time were 1.0 ms and
4.0 s, respectively. Two thousand scans were collected for each sample.
The light transmittance of the chitin and chitosan physical
hydrogels was determined by ultraviolet−visible (UV−vis) spectroscopy (UV-6, Mapada, China) at wavelengths ranging from 400 to 800
nm.
Dynamic mechanical analysis (DMA) temperature sweep was
performed on a DMA Q800 (TA Instruments, USA) under oscillatory
stress in tensile mode from −50 to 300 °C. The heating rate and
frequency were 5 °C min−1 and 1 Hz, respectively. The width of the
samples was approximately 5 mm.
Thermogravimetric analysis (TGA) was conducted using a STA
449C (Netzsch, German) from 25 to 600 °C at a heating rate of 10 °C
min−1 under nitrogen.

The hydrogels were subjected to tension tests on a universal tensile
tester (CMT 6503, SANS, China). The hydrogel was stretched at a 2
mm min−1 stretch speed at ambient temperature. The modulus was
calculated from the initial linear regions of the stress−strain curves.
Mouse bone mesenchymal stem cells (mBMSCs, ATCC, CRL12424) were propagated in Dulbecco’s Modified Eagle’s Medium
(DMEM, Gibco) supplemented with 10% fetal bovine serum (FBS,
Gibco). Chitin and chitosan physical hydrogels were placed in 96-well
plates and then sterilized in 75% (v/v) aqueous ethanol for 2 h
followed by three rinses with sterilized phosphate-buffered saline
(PBS). Subsequently, the hydrogels were prewetted with culture
medium for 12 h. After removing the culture medium, 200 μL of the
mBMSCs suspension (1 × 104 cells well−1) was seeded on the
hydrogels and then incubated at 37 °C in a humidified incubator at 5%
CO2. The Cell Counting Kit-8 (CCK-8, Dojindo Laboratories, Japan)
was used to evaluate the cell proliferation on the hydrogels after 1, 3, 5,
and 7 days of culture. Briefly, at each time point, the culture medium
was removed, and the CCK-8 working solution was added at 37 °C for
2 h. Subsequently, the supernatant medium was extracted to determine
the absorbance at 450 nm using a Thermo 3001 microplate reader
(Thermo, USA) (n = 5). The cell viability and morphology on the
hydrogels were characterized using a Live/Dead assay kit (Dojindo
Laboratories, Japan). The cell-seeded hydrogels were thoroughly
washed with PBS. Subsequently, the hydrogels were incubated in
standard working solution for 30 min. After washing again with PBS,
the hydrogels were imaged using an Eclipse Ti−U fluorescence
microscope (Nikon, Japan). SPSS 12.0 software (SPSS, USA) was
used to analyze the results with one-way analysis of variance
(ANOVA). The data are presented as the mean-standard deviation.
To compare the differentiation of the mBMSCs into epidermal cells
on the hydrogels, the mBMSCs were seeded onto the chitin and

chitosan physical hydrogels at a density of 1.5 × 104 cells cm−2. After
culture for 7 d in the presence of 30 ng mL−1 recombinant human
EGF (PeproTech, USA) and 50 ng mL−1 recombinant murine IGF-1
(PeproTech, USA), the nuclei were stained by DAPI and the cell
morphology were observed by laser scanning confocal microscopy
(Leica, Germany). Additionally, reverse transcriptase polymerase chain
reaction (RT-PCR) was used to evaluate the expression of the

under mild conditions and show that these properties can be
attributed to the formation of extra intra- and inter-molecular
hydrogen bonding interactions between the amino and
hydroxyl groups on the nearby chitosan nanofibrils. Unlike
the existing chitosan chemical hydrogels described in the
literature, our method allows for the formation of tough and
cell-compatible chitosan physical hydrogels that are suited for
stem cell culture and promote differentiation into epithelial
cells.

■

EXPERIMENTAL SECTION

Materials. The raw chitin powder was purchased from GoldenShell Biochemical Co. Ltd. (Zhejiang, China). The raw chitin powder
was purified with 0.1 mol L−1 aqueous NaOH at ambient temperature
overnight and combined with 0.3% (w/w) aqueous NaClO2 buffered
to pH 4.7 with acetate buffer at 80 °C for 3.5 h. Washing with
deionized water was performed after each step to remove any residual
proteins and chemical regents. The purification procedures were
repeated twice. The purified chitin powder was finally freeze-dried, and
its viscosity-average molecular weight (Mη) was calculated to be 10.7 ×

104 in 5% (w/w) LiCl/N,N-dimethylacetamide (DMAc) at 25 ± 0.02
°C by viscometry.34
Fabrication of Chitin Hydrogels. The purified chitin powder was
dispersed in aqueous 11% NaOH-4% urea (w/w) and then frozen at
−30 °C overnight. Subsequently, after thawing at 5 °C, the chitin was
dissolved completely and used to form a transparent and viscous 7%
(w/w) chitin solution according to our previous method.32 The chitin
solution was centrifuged at 5 °C for 15 min to prevent gelation and
remove air bubbles. The resultant chitin solution was spread on a glass
plate as a 1.0 mm-thick layer and then immersed in ethanol at 5 °C for
1 h to produce the chitin gels. The gels were then thoroughly washed
with deionized water to create the chitin hydrogels.
Heterogeneous Deacetylation of Chitin Hydrogels. Typically,
chitin hydrogels were immersed in 35% (w/w) aqueous NaOH at 60
°C for 6 h. The deacetylation was stopped by removing the hydrogels
from the aqueous NaOH and then immersing them into 50% (v/v)
aqueous ethanol. The degree of deacetylation (DD) of the hydrogels
can be controlled by the number of heterogeneous deacetylation
cycles. The deacetylated chitin hydrogels, that is, the chitosan
hydrogels coded as S1, S2, S3, and S4, were obtained by one, two,
three, and four heterogeneous deacetylation cycles, respectively.
Characterization. The weight-average molecular weight (Mw) of
the chitosan was performed on a size exclusion chromatography
combined with multiangle laser light scattering (SEC-LLS) (DAWN
EOS, Wyatt, USA) equipped with a He−Ne laser (λ = 632.8 nm). A
p100 pump equipped with a TSK GEL G6000 and G4000 PWXL
column (MicroPak, TSK) and an Optilab refractometer (Wyatt, USA)
was combined with the instrument. The fluent was 0.1 M NaAc/HAc
buffer (pH = 2.8) with a flow rate of 0.6 mL min−1.
The DDs of the chitin and chitosan physical hydrogels was

calculated by potentiometry. The hydrogels were cut into small pieces
and freeze-dried from t-BuOH. The quantitative dried gel was
accurately weighed and added to 0.1 M aqueous HCl solution.
Subsequently, the mixture was titrated with 0.1 M NaOH, with the
standard substance KH5C8O4 used for calibration. The DD value was
calculated as follows:35
DD =

(V2 − V1) × C × 0.016
0.0994 × W

(1)

where C is the accurate concentration of aqueous NaOH solution
(mol L−1), V1 is the volume of aqueous NaOH solution (mL) at the
first titration jump, V2 is the volume of aqueous NaOH solution at the
second titration jump, W is the sample weight (g), 0.016 is the molar
mass weight of NH2 (kg mol−1), and 0.0994 is the theoretical NH2
percentage in chitosan.
Scanning electron microscopy (SEM) images of the cross sections
of the chitin and chitosan hydrogels was carried out on a Hitachi S4800 instrument. The chitin and chitosan physical hydrogels were
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Figure 1. (a) Schematic representation of the creation of a chitosan physical hydrogel from a nanoporous chitin hydrogel. (b) Photographs of the
square chitin hydrogel (S0) and chitosan physical hydrogels (S1−S4) at each heterogeneous deacetylation cycle. (c) Macroscopic views of the
chitosan physical hydrogels (S4) under stretching, torsional and rolling loading.

Table 1. Physical Properties of the Chitin Hydrogel and Chitosan Physical Hydrogelsa
samples

DD, %

WH2O, %

M × 10−4, g/mol

crys. %

σb, MPa

εb, %

E, MPa

S0
S1
S2
S3
S4

8
56

80
91
99

84
61
58
51
50

10.7
4.9
4.0
3.9
3.1

50
35
34
36
43

1.7 ± 0.1
3.6 ± 0.5
10.5 ± 0.5
12.1 ± 1.1
12.1 ± 0.7

56 ± 4
67 ± 5

106 ± 14
67 ± 5
57 ± 4

4.5
7.9
34.5
57.3
92.0

a

The DD and WH2O are the degree of deacetylation and the water content, respectively, of the chitin hydrogel (S0) and chitosan physical hydrogels
(S1−S4).The viscosity-average molecular weight (Mη) of sample S0 was determined by viscometry, and the weight-average molecular weights (Mw)
of samples S1−S4 were determined by SEC-LLS. Crys. is the degree of crystalline of the dried gels. The σb, εb, and E are the tensile strength,
elongation at break, and Young’s modulus of the hydrogels, respectively.
epidermal cell differentiation marker gene K18 and K19. The following
primer sequences were used: K18 gene: forward, 5′-AAGGCTGCAGCTGGAGACAGA-3′; reverse, 3′-TGGGCTTCCAGACCTTGGAC-5′; K19 gene: forward, 5′-TGACCTGGAGATGCAGATTGAGA-3′; reverse, 3′- TGGAATCCACCTCCACACTGAC-5′; and
GAPDH gene: forward, 5′-TGTGTCCGTCGTGGATCTGA-3′;
reverse, 3′-TTGCTGTTGAAGTCGCAGGAG-5′. The relative quantification of the target gene was normalized to GAPDH and
determined using the 2−ΔΔCt method. At the end of each PCR, the
melting curve profiles were generated to identify the specific
transcription of the amplification. To evaluate the in vitro degradation
of the chitosan hydrogels, certain weights of the chitin hydrogel and
chitosan hydrogels were immersed in 4 mg/mL three-times-recrystallized egg white lysozyme (TSZ, USA) in 0.1 M PBS at pH 7.4 and 37
°C.20 After specific time intervals, the hydrogels were removed from
the lysozyme solution, thoroughly washing with double distilled water
and freeze-dried. The extent of the in vitro degradation was calculated
from the percentage of the weight of the dried hydrogels before and
after the lysozyme treatment.

Figure 2. Top: SEM images of the surfaces of the chitin hydrogel S0
(a) and the chitosan physical hydrogels S1 (b), S2 (c), and S4 (d).
Bottom: SEM images of the inner parts of the chitin hydrogel S0 (e)
and the chitosan physical hydrogel S1 (f), S2 (g), and S4 (h) (scale bar
=500 nm).

chitosan physical hydrogel under mild conditions. The chitin
hydrogel was subjected to in situ heterogeneous deacetylation
in 35% (w/w) aqueous NaOH at 60 °C for 6 h. Then, the
NaOH was almost fully removed using a 50/50 ethanol/water
mixture followed by the addition of deionized water, and a
chitosan physical hydrogel containing free amine groups
formed. The −NHCOCH3 sites were deacetylated to give
−NH2 groups, leading to the disappearance of hydrophobic
interactions between the polymeric chains, which favored
physical cross-links corresponding to hydrogen bonding
interactions. This procedure was repeated four times to obtain
chitosan physical hydrogels with 4 different DDs, which, as
expected, ranged from 56 to 99% (Table 1). The weight-

■

RESULTS AND DISCUSSION
The preparation of the chitosan physical hydrogels from the
nanoporous chitin hydrogel by in situ heterogeneous
deacetylation cycles is described in Figure 1. The chitin
hydrogel is a transparent nanoporous material, which forms via
the hydrogen bonding of the chitin−NaOH−urea aqueous
solution by a sol−gel transition in ethanol without an external

cross-linker. It has a large interior space that reacts with the
deacetylation reagents and can thus be converted into a
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average molecular weight (Mw) of the chitosan physical
hydrogels decreased from 5.6 × 104 to 4.7 × 104 g mol−1. At
the swelling equilibrium state, the water content of the
hydrogels decreased gradually from 84% for the chitin hydrogel
(S0) to 50% for the chitosan physical hydrogel (S4). The gross
visual appearance of the chitosan physical hydrogels showed
that the chitin hydrogel underwent significant volume changes
after washing with aqueous ethanol, mainly because of the
diffusion of ethanol, which disturbed the hydrophobic and
hydrogen bonding interactions between the chitosan chains and
thereby influenced the final density of the physical cross-linking
and water content of the hydrogels (Figure 1b).19 After four
heterogeneous deacetylation cycles, the volume of the chitosan
physical hydrogel was approximately one-third of that of the
pristine chitin hydrogel, generating chitosan physical hydrogels
with maximum physical cross-linking density. Thus, the
chitosan physical hydrogels demonstrated good mechanical
integrity under stretching, torsional and rolling loading (Figure

1c).
The SEM images of the surface and inner part of the chitin
gel (Figure 2a and e) show an open nanoporous network
structure composed of interconnected chitin nanofibrils. The
typical diameter of the chitin nanofibrils was approximately 10
nm, which is in good agreement with the Brunauer−Emmett−
Teller (BET) surface area of 364 m2 g−1, as determined by
nitrogen adsorption and desorption isotherms (see Supporting
Information, Figure S1), which corresponds to a fibril width of
7 nm. Moreover, the SEM images and the nitrogen
adsorption−desorption isotherms of the chitosan physical gels
(Figure 2b−h, Figure S1) show features of a smaller porous
structure and surface area after the heterogeneous deacetylation. The fibrils that comprise the networks of the chitosan
physical hydrogels seem to gradually thicken, and therefore,
chitosan likely sticks together to create a close network
structure in the fourth deacetylation cycle (S4).
The FT-IR spectrum of the chitin gel (Figure 3, S0) shows
the characteristic peaks of α-chitin, including broad OH
stretching absorption peaks at 3447 and 3268 cm−1, a CH3
stretching absorption peak at 3100 cm−1, and splitting of the
CO stretching absorption peak at 1660 and 1627 cm−1 for
amide I and 1560 cm−1 for amide II.36−38 After the
heterogeneous deacetylation cycles, the CH 3 stretching
absorption peak of the chitosan physical gel was nearly absent,
the amide I and II stretching absorption peaks had gradually
weakened, and the new N−H bending absorption peak at 1596
cm−1 was enhanced (Figure 3, S1−S4), indicating the
prevalence of NH2 groups and the successful formation of
the chitosan physical hydrogels from chitin hydrogel. Moreover,
the OH stretching absorption peak of the chitosan physical gels

shifted from 3447 to 3416 cm−1, indicating a lower-order
structure of polymeric chains, which is consistent with the XRD
patterns of the chitin and chitosan physical gels.
In the XRD patterns, the chitin gel (Figure 4, S0) shows
characteristic peaks at 9.4°, 12.8°, 19.3°, 20.8°, 23.4°, and 26.4°,
corresponding to the (020), (021), (110), (120), (130), and
(013) reflections, respectively, of an α-chitin crystal.37,39,40 The
XRD patterns of the chitosan physical gels (Figure 4, S1−S4)
show near-systematic superposition with those of pure α-chitin
and chitosan (DD of 100%), indicating a homogeneous
distribution of the two components in the structure resulting
from the in situ heterogeneous deacetylation cycles. Moreover,
the intensities of the (020), (021), (130), and (013) reflections
of the chitosan physical gels decreased gradually, and the

Figure 3. FT-IR spectra of the chitin hydrogel (S0) and chitosan
physical hydrogels (S1−S4).

Figure 4. XRD patterns of the chitin hydrogel (S0) and chitosan
physical hydrogels (S1−S4).

Figure 5. Solid-state CP/MAS 13C NMR spectra of the chitin
hydrogel (S0) and chitosan physical hydrogels (S1−S4).

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Table 2. Chemical Shifts of the Chitin Hydrogel and Chitosan Physical Hydrogel Determined by CP/MAS 13C NMR
chemical shift/ppm
samples

C7 (CO)

C1

C4

C5

C3

C6

C2

C8 (CH3)

S0
S1
S2
S3
S4

174.1
174.3
174.3

104.5
104.5
105.5
105.3
105.7

83.6
83.2
82.9
82.9
82.7

76.1
75.9
75.9
75.7
75.9

74.2
75.9
75.9
75.7
75.9

61.8
61.1

61.2
61.7
61.5

55.9
58.2
58.0
58.5
58.3

23.3
23.5
23.0
24.0

Figure 9. Morphology of the mBMSCs (a−e) and the nuclei staining
by DAPI (f−j) on the chitin hydrogel (S0) and chitosan physical
hydrogels (S1−S4) after a 7-day differentiation period (scale bar =100
μm).
Figure 6. Typical stress−strain curves of the chitin hydrogel (S0) and
chitosan physical hydrogels (S1−S4). The inset is the stress−strain
curve of the chitosan film dried from sample S3.

cycles increased. The crystallinities estimated from multipeak
fitting were 50% for the chitin gel and between 34 and 43% for
the chitosan physical gels (Table 1). Additionally, the crystallite
sizes evaluated based on the (110) reflection using the Scherrer
equation were 4.4 nm for the chitin gel (S0) and 3.7 nm for the
chitosan physical gel (S4). These results are also consistent
with the higher optical transmittance of the chitosan physical

hydrogels relative to the chitin hydrogel (94% vs 81% at 800
nm) (see Supporting Information, Figure S2).
The structure change of the chitin hydrogel caused by the
heterogeneous deacetylation was confirmed by the solid-state
CP/MAS 13C NMR analysis (Figure 5). The corresponding
chemical shifts are listed in Table 2. The spectrum of the chitin
gel shows the characteristic eight resonances of α-chitin: C1
(104.5 ppm), C2 (55.9 ppm), C3 (74.2 ppm), C4 (83.6 ppm),
C5 (76.1 ppm), C6 (61.8 ppm), CH3 (23.3 ppm), and CO
(174.1 ppm).41−44 Compared with the spectrum of the chitin
gel, the C1 and C4 resonances of the chitosan physical
hydrogels became weak and broad and shifted slightly,
indicating a loosely packed structure and altered internal
torsion angles of the polymeric chains.45,46 Moreover, the
signals of C3 and C5 merged into a single resonance centered
at 75.9 ppm, and the signal intensities of the methyl and
carbonyl carbons of the chitosan physical gels decreased
gradually, disappearing after the final deacetylation cycle.
Simultaneously, all of the resonances of the deacetylated C2,
which are primarily involved in hydrogen bonding with
glucosamine units, of the chitosan physical gels shifted
downfield. These spectral and morphological characteristics
indicate that nanoscale heterogeneous deacetylation was
achieved in the nanoporous chitin hydrogel; that is, the Nacetylglucosamine units on the surface of the interconnected
chitin nanofibrils were removed, and the resultant amino
groups interacted with the hydroxyl groups on the nearby
nanofibrils to form extra intra- and inter-molecular hydrogen
bonds.
The tensile strength (σb), elongation at break (εb), and
Young’s modulus (E) of the chitin hydrogel (Figure 6, S0;

Figure 7. Proliferation of the mBMSCs cultured on the surfaces of the
chitin hydrogel (S0) and chitosan physical hydrogels (S1−S4).

Figure 8. Viability and morphology of the mBMSCs cultured on the
chitin hydrogel (S0) and chitosan physical hydrogels (S1−S4) for 48
h. The green bright spots represent the living mBMSCs stained by
Calcein-AM. The red spots indicate the dead mBMSCs stained by PI
(scale bar =100 μm).

diffraction angle of the (110) reflection increased, suggesting
that the crystallinity and crystallite size of the chitosan physical
gels decreased as the number of heterogeneous deacetylation
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Figure 10. Relative gene expression of the K18 (a) and K19 (b) as epidermal cell differentiation markers by RT-PCR.

Cell viability was also examined by a Live/Dead assay kit. As
shown in Figure 8, high cell viability (green) was achieved for
all samples, but more cells were visible on the chitosan physical
hydrogels, indicating increased cell proliferation, consistent
with the cell proliferation results. Compared with the synthetic

polymer hydrogels51,52 and chitosan chemical hydrogels,53 the
chitosan physical hydrogels showed a greater cell proliferation
rate, improved mechanical properties, and less cytotoxicity.
Moreover, the chitosan physical hydrogels showed a lower
degradation rate than the chitin hydrogel (see Supporting
Information, Figure S5). These results are, in part, attributable
to the nanoporous structure and saturated positively charged
amino acids of the chitosan physical hydrogels resulting from
the heterogeneous deacetylation, which allow for stronger
electrostatic interactions with glycosaminoglycan. In addition,
the surface properties of the chitosan physical hydrogels
promote cell growth and proliferation.20,54,55 Furthermore, after
culturing the cells for 7 d in the presence of EGF and IGF-1,
the mBMSCs cultured on the chitin hydrogel and chitosan
physical hydrogels took on a cobblestone morphology under
the light microscope with the cell nucleolus in the middle of
each cell (Figure 9), which is characteristic of epidermal
cells.56,57 RT-PCR analysis of the epidermal cell differentiation
marker genes K18 and K19 were performed to verify the
epidermal cell differentiation of the BMSCs cultured on the
hydrogels (Figure 10). The results demonstrated that the chitin
hydrogel (S0) and chitosan physical hydrogels (S3 and S4)
induced the differentiation of the mBMSCs into epidermal cells
in cooperation with EGF and IGF-1 in vitro. Thus, the chitosan
physical hydrogels constructed via the heterogeneous deacetylation of nanoporous chitin hydrogel have excellent mechanical
properties and good cell-compatibility with potential applications in stem cell research and tissue engineering.

Table 1) were 1.7 MPa, 56% and 4.5 MPa, respectively. In
contrast, the tensile behavior of the chitosan physical hydrogels
(Figure 6, S1−S4; Table 1) showed remarkable strengthening

and toughening effects after the heterogeneous deacetylation.
The σb, εb, and E values of the chitosan physical hydrogels were
3.6 MPa, 67% and 7.9 MPa, respectively, for sample S1 (DD of
56%) and increased to 12.1 MPa, 57% and 92.0 MPa for sample
S4 (DD of 99%) at the swelling equilibrium state. These values
were much higher than those of chitosan hydrogels obtained by
neutralization or chemical cross-linking of their acidic solutions
(σb, from 0.1 to 2.9 MPa; E, from 31 kPa to 4 MPa).22,24,47−50
Interestingly, the chitosan physical hydrogel (sample S2) had a
moderate σb of 10.5 MPa and, remarkably, an εb of 106%,
probably because of the lower degree of crystallinity and
physical cross-linking density in the chitosan hydrogel at a DD
of 80%. Moreover, upon drying, the σb and E of the
heterogeneous-deacetylated chitosan film were 107.1 and
3053 MPa, respectively, confirming that the good mechanical
properties of the chitosan physical hydrogels are attributable to
hydrogen bonding interactions between the amino and
hydroxyl groups of the chitosan chains. The DMA of the
chitosan film (see Supporting Information, Figure S3) revealed
typical behavior of a semicrystalline polymer. The tensile
storage modulus (E′) of the chitosan film was reduced from 58
to 24 MPa at temperatures from −50 to 200 °C, demonstrating
significant mechanical stability. Moreover, in the TGA of the
chitosan physical gels, decomposition was observed between
230 and 400 °C, regardless of the DD value (see Supporting
Information, Figure S4). As evidenced by the tensile, DMA, and
TGA results, the chitosan physical hydrogels demonstrated
strong mechanical properties and sufficient thermal adaptivity
for applications in biomaterials after heterogeneous deacetylation.
In this study, we aimed to exploit chitosan physical hydrogels

in tissue engineering repair, especially as adaptive substrates for
stem cells. For this application, we chose mBMSCs as model
cells to assess the biological performance of our chitin and
chitosan physical hydrogels. The proliferation and viability of
the mBMSCs cultured on the hydrogels were studied in vitro.
Figure 7 shows that the heterogeneous deacetylation of the
chitin hydrogels enhanced the adhesion and proliferation of the
mBMSCs on the surface of the resultant chitosan physical
hydrogels. As the culture time increased, the mBMSCs
gradually proliferated in the chitin and chitosan physical
hydrogels. Significant differences for all of the samples were
observed after 5 days. The proliferation rate of mBMSCs on the
surface of the chitosan physical hydrogels was superior to that
on the chitin hydrogel and was independent of the DD value of
the chitosan physical hydrogels.

■

CONCLUSIONS
In summary, we developed a novel chitosan physical hydrogels
cross-linked by hydrogen bonding with considerable technical
and commercial importance. The heterogeneous deacetylation
of the nanoporous chitin hydrogel offers a facile approach to
synthesize chitosan physical hydrogels with excellent mechanical properties and cell compatibility. In vitro studies showed
that mBMSCs cultured on these chitosan physical hydrogels
exhibited good adhesion and proliferation. Furthermore, the
chitosan physical hydrogels also induced the differentiation of
mBMSCs into epidermal cells in cooperation with EGF and
IGF-1 in vitro. The simplicity of the process and the widely
tunable properties of the chitosan physical hydrogels make

them promising candidates for potential biomedical applications in stem cell culture and differentiation.
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DOI: 10.1021/acsami.6b05302
ACS Appl. Mater. Interfaces 2016, 8, 19739−19746


Research Article

ACS Applied Materials & Interfaces

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ASSOCIATED CONTENT

S Supporting Information
*

The Supporting Information is available free of charge on the
ACS Publications website at DOI: 10.1021/acsami.6b05302.
Nitrogen adsorption and desorption, UV−visible spectra,
DMA, and TGA (PDF)

■

AUTHOR INFORMATION

Corresponding Authors

*E-mail: ; Telephone: +86-27-6878-9321.
*E-mail:
Author Contributions
⊥

B.D. and H.G. contributed equally to this work.

Notes

The authors declare no competing financial interest.

■

ACKNOWLEDGMENTS
This work was supported by the National Natural Science
Foundation of China (21422405, 51373125, 21574045) and
the Major Program of National Natural Science Foundation of
China (21334005). The authors thank the facility support of
the Natural Science Foundation of Hubei Province and the
Fundamental Research Funds for the Central Universities.

■

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DOI: 10.1021/acsami.6b05302
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