- Email: [email protected]
Evaluation of cellular attachment and proliferation on different surface charged functional cellulose electrospun nanofibers
Accepted Manuscript Title: Evaluation of cellular attachment and proliferation on different surface charged functional cellulose electrospun nanofibers Authors: Mortaza Golizadeh, Afzal Karimi, Soheila Gandomi-Ravandi, Manouchehr Vossoughi, Mona Khafaji, Mohammad Taghi Joghataei, Faezeh Faghihi PII: DOI: Reference:
S0144-8617(18)31462-0 https://doi.org/10.1016/j.carbpol.2018.12.028 CARP 14382
To appear in: Received date: Revised date: Accepted date:
23 June 2018 3 November 2018 10 December 2018
Please cite this article as: Golizadeh M, Karimi A, Gandomi-Ravandi S, Vossoughi M, Khafaji M, Joghataei MT, Faghihi F, Evaluation of cellular attachment and proliferation on different surface charged functional cellulose electrospun nanofibers, Carbohydrate Polymers (2018), https://doi.org/10.1016/j.carbpol.2018.12.028 This is a PDF file of an unedited manuscript that has been accepted for publication. As a service to our customers we are providing this early version of the manuscript. The manuscript will undergo copyediting, typesetting, and review of the resulting proof before it is published in its final form. Please note that during the production process errors may be discovered which could affect the content, and all legal disclaimers that apply to the journal pertain.
Evaluation of cellular attachment and proliferation on different surface charged functional cellulose electrospun nanofibers
Golizadeh1,2,
Afzal
Karimi1,3,*,
Soheila
Gandomi-Ravandi
1
,
Manouchehr
SC RI PT
Mortaza
Vossoughi2,4,**, Mona Khafaji4, Mohammad Taghi Joghataei3,5, Faezeh Faghihi3
1
Faculty of Advanced Technologies in Medicine, Iran University of Medical Sciences,
1449614535 Tehran, Iran
Chemical and Petroleum Engineering Department, Sharif University of Technology, Tehran,
U
2
Cellular and Molecular Research Center, Iran University of Medical Sciences, 1449614535
A
3
N
14588-89694, Iran.
4
M
Tehran, Iran
Institute for Nanoscience and Nanotechnology, Sharif University of Technology, 14588-89694
School of Medicine, Iran University of Medical Sciences, 1449614535 Tehran, Iran
TE
5
D
Tehran, Iran.
*Corresponding author: Afzal Karimi
EP
Email: [email protected]
CC
Phone: +98 2188622687 Fax: +98 2186704606
A
** Corresponding author: Manouchehr Vossoughi Email: [email protected] Phone: +98 2166165487 Fax: +98 2166005417
1
SC RI PT
Graphical abstract
Highlights
Cellulose nanofibers were prepared by cellulose acetate electrospun deacetylation.
Anionic and cationic cellulose nanofibers were fabricated by chemical modification.
Several techniques were employed to characterize the cellulose based nanofibers.
Cellular studies were carried out to determine the cells viability.
Cationic and anionic cellulose nanofibers are favorable for biomedical application.
EP
Abstract
TE
D
M
A
N
U
CC
Fabrication and characterization of different surface charged cellulose electrospun scaffolds including cellulose acetate (CA), cellulose, carboxymethyl cellulose (CMC) and quaternary
A
ammonium cationic cellulose (QACC) for biomedical applications have been reported in this research. Several instrumental techniques were employed to characterize the nanofibers. MTT assay and cell attachment studies were also carried out to determine the cytocompatibility, viability and proliferation of the scaffolds. Fabricated CA, cellulose, CMC and QACC nanofibers had 100-600 nm diameter, -9, -1.75, -12.8, +22 mV surface potential, 2.5, 4.2, 7.2, 7 2
MPa tensile strength, 122, 320, 515, 482 MPa Young modules, 430, 530, 670 and 642% water uptake and 92o, 58o, 45o, 47o contact angle respectively. The findings showed that cell adhesion and proliferation is strongly enhanced on the modified surfaces with quaternary ammonium and
SC RI PT
carboxymethyl groups. We believe the use of cationic and anionic surface modified cellulose electrospun nanofibers presents promising materials for biomedical applications.
Keywords: Electrospun nanofiber, Cellulose, Carboxymethyl cellulose, Quaternized cellulose, Scaffold, Surface charge.
1. Introduction
U
In recent years, research on developing sustainable, green and environmentally friendly and
N
biocompatible nanosized materials has gained much interest (Abdul Khalil, Bhat, & Ireana
A
Yusra, 2012). In this regard, great efforts have been made to develop renewable, biodegradable
M
and biomimetic materials based on polysaccharides (Teeri, Brumer, Daniel, & Gatenholm, 2007;
D
Zhou, Rutland, Teeri, & Brumer, 2007). Especially, biomass based polymers such as cellulose substances have become the hotspot of science due to their intrinsic properties. Cellulose has
TE
been evaluated by Pelton as “particularly protein and biomolecule friendly” (Pelton, 2009).
EP
Cellulose has been applied for fabricating three-dimensional scaffolds, which have provided good support for cell adhesion and growth. The clinical applications of cellulose-based materials
CC
as scaffolds are including repairing, reconstructing and regenerating of almost all type of tissues in the mammalian organism. In tissue engineering and cell delivery, cellulose supports, cover
A
wounds and release drugs into them, through inhibiting postoperative adhesions, hemodialysis, hemostasis, or via covering and filling different tissue defects (Bacakova, Novotna, Sopuch, & Havelk, 2015).
3
With the development of nanotechnology, cellulose nanofibers have been widely studied due to environmentally benign nature, relatively easy production and compatibility with biological systems (Menon, Selvakumar, & Ramakrishna, 2017). Processing of cellulose to produce
SC RI PT
nanofibers can be carried out by a variety of techniques such as steam explosion (Deepa et al., 2011), ultrasonication (Chen et al., 2013), homogenization, grinding, cryocrushing (Siró & Plackett, 2010), refining and electrospinning (Inukai, Kurokawa, & Hotta, 2018; Antczak, 2012). Among these, electro-static spinning or “electrospinning” is a versatile and efficient method in terms of its simplicity and cost-effective setup. Moreover it allows producing smooth, continuous
U
ultrafine and highly porous nanofibers with highly controllable physical properties (Huang,
N
Zhang, Kotaki, & Ramakrishna, 2003). Polymer-based electrospun nanofibers have been widely
A
applied in tissue engineering (skin, bone, nerve and blood vessel) as scaffolds (Ma, Kotaki, Inai,
M
& Ramakrishna, 2005) for improvement of cell growth, controlling the proliferation of cells or neurons and regulation of cell behavior (Yang et al., 2010). The application of cellulose scaffolds
D
in tissue engineering and cell therapies needs to be developed. For this purpose, cellulose
TE
materials can be modified by combinations of cellulose with other synthetic or natural polymers,
EP
metals, ceramics, carbon materials or various chemical groups by and loading with bioactive molecules such as drugs and growth factors. These modifications, control the physical and
CC
chemical properties of cellulose scaffolds for cell-material interaction (Bacakova, Novotna, Sopuch, & Havelk, 2015).
A
CMC is a natural anionic polyelectrolyte prepared by introducing carboxymethyl groups along the cellulose chain. Carboxymethylation of cellulose improves solubility, stability and hydrophilicity of suspension, decreases the crystallinity index, increases the breakdown of fibers to nano size, decreases the agglomeration of fibers and makes the surface negatively charged
4
through functionalization of cellulose by breaking the hydrogen bonds (Rezaei, Nasirpour, & Fathi, 2015). Over recent decades, QACC as a new environmental friendly material is found special
SC RI PT
technological interest. Cationic cellulose derivatives are large-scale commercial products due to hydrophilicity, adsorption capacity for anionic substances, ionic exchange capacity (Yamamoto, Iwata, Nishiwaki, Kinoshita, & Suzuki, 2015; Zarth, 2013) biocompatibility, low hemo- and cytotoxicity, biodegradability, lack of skin irritation, low corrosiveness, good environmental stability, bacteriostatic properties, excellent cell membrane penetration properties and
U
antimicrobial activity (Fei et al., 2018). Quaternized cellulose (QC), as a positively charged
N
polysaccharide, is commonly produced by etherification of alkali cellulose. In this process,
A
quaternary epoxides or quaternary halohydrins act as cationizing reagent that producing
M
quaternary ammonium groups on the hydroxyl groups of cellulose (Hasani, Cranston, Westman, & Gray, 2008; Song, Sun, Zhang, Zhou, & Zhang, 2008). Cytotoxicity of cationic cellulose was
D
related to the molecular weight (Mw) and degree of substitution (DS). Lower Mw and DS of
TE
cationic celluloses appeared to be less cytotoxic compared to those with higher values. It was
EP
reported that cationic celluloses with DS value between 0.2-0.6 displayed relatively low cytotoxicity so that cells viability was almost 100% (Song et al., 2010). Among cationizing
CC
agents, 3-chloro-2-hydroxypropyltrimethylammonium chloride (CHPTAC) has attracted much interest in cationization of cellulose because of commercially availability, low toxicity and good
A
reactivity (Hashem, 2006). The studies have shown that the presence of quaternary ammonium and numerous hydroxyl groups in QC backbone afford various electrostatic interactions with ionic and non-ionic therapeutic molecules (Huang, Yu, & Xiao, 2007). QCs have exhibited elevated potential in biomedicine fields as promising nonviral gene (Song et al., 2010) and
5
protein (Song, Zhou, Li, Guo, & Zhang, 2009; Song, Zhou, & Chen, 2012) carriers, in drug delivery systems (Song, Zhang, Gan, & Zhou, 2011) and wound-dressing applications (Zahedi, Rezaeian, Ranaei-Siadat, Jafari, & Supaphol, 2009).
SC RI PT
Here, we describe CA electrospinning and preparation of cellulose nanofibers through its deacetylation. Moreover, surface modification of electrospun cellulose nanofibers is carried out using monochloroacetic acid (MCAA) and CHPTAC to obtain CMC and QC nanofibers, respectively. In addition, we evaluated the potential application of the modified cellulose derivatives as biomedical materials. At last, the structural, morphological, mechanical, swelling,
N
U
wettability and the cell culture properties of the scaffolds were analyzed and compared.
A
2. Materials and methods
M
2.1. Materials
D
Cellulose acetate (average molecular weight 30 kDa g/mol, 39.8 wt% acetyl content; degree of
TE
acetyl substitution 2.4) was purchased from Aldrich Chemical Compound. The cationizing agent, 3-chloro-2-hydroxypropyltrimethylammonium chloride (CHPTAC, aqueous solution of 69 wt
EP
%) was obtained from Fluka. Other chemicals, namely, sodium hydroxide, hydrochloric acid, glutaraldehyde and MCAA were received from Sigma–Aldrich Chemicals. All solvents such as
CC
acetone, ethanol, isopropanol and N,N-dimethylformamide (DMF) were provided by Merck (Germany). All reagents and chemicals were of analytical grade and were used as received
A
without further purification. All experiments were performed using double-distilled water. For cell culture study, Human Bone Marrow Derived Mesenchymal Stem Cells (hMSCs) were obtained from the Royan Cell Bank, Tehran, Iran. MTT (3-(4,5-Dimethylthiazol-2-yl)-2,5diphenyl phenyltetrazolium bromide) reagent (98%), phosphate-buffered saline (PBS, pH 7.4),
6
Dulbecco’s modified eagle’s medium (DMEM)-F12, fetal bovine serum (FBS) and penicillinstreptomycin (pen-strip) were purchased from GIBCO Invitrogen (Carlsbad, CA, USA). 2.2. Fabrication and modification of electrospun cellulose nanofibers
SC RI PT
Electrospun CA nanofibers were synthesized following the procedure described in literature (Khatri, Mayakrishnan, Hirata, Wei, & Kim, 2013). Electrospinning apparatus (SBS LAB ES SBS, Nanoazma, Tehran, Iran) was applied to form cellulose nanofibers. The spinning solution was prepared by dissolving 17% w/v CA in a binary solvent system of DMF: acetone (2:1 w/w) under constant stirring at room temperature until the solution became transparent. Freshly
U
prepared CA solution was loaded into a 10 mL tip plastic syringe with a stainless steel needle as
N
the nozzle. The flow rate was controlled by the syringe pump at 1 mL/h. The supplied voltage
A
was 12.5 kV and the needle tip to collector distance was 15 cm. The rotational speed of the roller
M
during electrospinning was set at 100 rpm to get a uniform mesh. The obtained electrospun
D
nanofibers were detached from the collectors and dried in a vacuum oven at 60 ℃ for 5 h to
TE
remove residual solvent. The electrospun CA nanofibers were deacetylated in 0.05 M NaOH ethanol solution at room temperature for 24 h. After deacetylation, the cellulose nanofibers were
EP
rinsed thoroughly with an excess amount of deionized water to remove sodium and acetate ions. The alkaline solution was prepared by dissolving of 5 g NaOH in 5 mL distilled water and then
CC
mixed with 100 mL ethanol. The cellulose nanofibers were soaked into the solution for 1 h. The mercerization was continued for 1 h to alkalize the cellulose.
A
CMC nanofibers were synthesized by dissolving 0.75 g MCAA in the alkaline solution containing mercerized cellulose nanofibers at 60 ℃ for 2 h. The produced CMC nanofibers were soaked in distilled water for 3 h and were dried in a vacuum oven at 60 ℃ for 5 h.
7
The QC nanofibers were produced by dissolving 5 g CHPTAC solution in alkaline solution containing mercerized cellulose nanofibers at 80 ℃ for 2 h. The produced QC nanofibers were soaked in distilled water for 3 h and were dried in a vacuum oven at 60 ℃ for 5 h.
SC RI PT
2.3. Characterization of electrospun cellulose based nanofibers The surface morphology and distribution of the prepared nanofibers before and after cell culture were observed using Field Emission Scanning Electron Microscope (FESEM) (Tescan – mira 3, Czech Republic) operated at 15 kV accelerating voltage. Prior to the FESEM analysis, the samples were sputter coated with a thin layer of gold (6 nm) to avoid charge accumulation.
U
Chemical structures of the cellulose samples were characterized on a Nicolette 6700 Fourier
N
transform infrared (FT–IR) spectrometer (Thermo Fisher Scientific Co. Ltd., MA, USA) with the
A
wavenumber range of 4000–400 cm–1 using KBr pellets. Nanofibers were dried at 60 ℃ for 12 h
M
before the measurement.
D
DS value of CMC sample was determined by the standard method ASTM D1439 (Hong, 2013).
TE
The nitrogen content of the QC nanofibers was measured to determine the DS. Elemental analysis (CHNS Analyzer, FlashEA 1112 series, Thermo Finnigan, Italy) was employed to
EP
determine the presence and mass percentage of nitrogen in cationically modified cellulose nanofibers (Song et al. 2010). Contact angle measurements were performed to investigate the
CC
wettability properties of the nanofibers mat using a G10 goniometer from Krüss (Germany) at
A
room temperature. Briefly, deionized water was dropped onto the nanofibers mat from a needle. Picture of the drop was taken and analyzed. The contact angle was obtained after 0, 1, 2, 3, 4 and 5 s. The water uptake measurements were carried out using a simple method. The scaffolds were cut into small squares at dimensions of 1 × 1 cm and their weights were measured. This is the dry weight (Wd) of each sample. Then, each sample was immersed in 50 mL of deionized 8
water for 2 h at room temperature. To measure swelling behavior, after the immersion period, the samples were reweighed to record its wet weight (Ww). The amount of water uptake of the samples was calculated from equation (1) (Huang, Ren, Chen, Ren, & Zhou, 2008):
W w W d 100 Wd
(1)
SC RI PT
Water Uptake % =
The nanofiber mats surface zeta potential was estimated by streaming potential measurements, which were carried out using an electro kinetic analyzer (EKA, Anton Paar KG, Graz, Austria) at 25 ℃ (Yu et al. 2013). The mechanical properties of the scaffolds containing Young’s modulus
U
and tensile strength were measured on a BOSE mechanical testing machine (Model ELF 3200).
N
Tensile test was carried out at 35-40% relative humidity and 1 mm/min speed until the samples
A
were ruptured. The nanofibers were cut into 15 mm × 10 mm (L × W) strips. The average
M
thicknesses of electrospun nanofibers were 220±15 µm. 2.5. Cell culture studies
D
hMSCs were used to determine the cell viability on the cellulose scaffolds. The cells were
TE
cultured in (DMEM)-F12 medium supplemented with 10% FBS and 1% pen-strip. The cells
EP
were incubated at 37 °C in a humidified atmosphere containing 5% CO2. The nanofibers were cut into circular shape that fit into the 24-well plates. The standard cell culture treated 24-well
CC
plates used as a control. Cells in all the samples were cultured in the same growth medium and treated in the same manner. The nanofibrous scaffolds were immersed in 70 % v/v ethanol for 12
A
h followed by washing with sterile PBS three times for 20 min. The nanofibers were sterilized under UV light for 1 h on each side. Under sterile conditions, DMEM-F12 medium containing 10% FBS was added to each well and nanofibers scaffolds allowed to hydrate for 24 h at 37 ℃. The cultured cells were seeded on the samples with a density of 16,000 cells/cm2. In total, 1000
9
μL of culture medium was added to each well. Cells were incubated in 5% CO2 incubator at 37 ℃ and the culture medium was replaced every 2 days. After incubation, cell proliferation and scaffold cytotoxicity were studied by MTT assay.
SC RI PT
MTT assay was performed at 1, 3, and 5 days of cell-seeding. On the day of the test (days 1, 3, and 5), 500 µL of medium in the cell-loaded scaffold culture plate was removed and replaced by 50 μL of MTT solution (0.5 mg/mL) in PBS (pH 7.4). After 4 h incubation at 37 ℃ in the humid environment with 5% CO2, the MTT containing medium was gently removed. After that, 150 µL acidic isopropanol (0.1 N HCL and isopropanol) was added to dissolve purple formazan crystals
U
formed in live cells and kept in the dark at room temperature for 15 min. Then, 100 µL
N
supernatant solution of each sample (the dissolved formazan solution) transferred into individual
A
well of a 96-well plate. The absorbance of each well was measured at 570 nm using a microplate
M
reader and results were averaged from three replications. (Synergy HTX, Biotek).
D
The hMSCs were used to investigate the adhesion behavior of cells onto the scaffolds using a
TE
cell adhesion assay. After the cells were cultured on the nanofibers for 24 h, each sample was rinsed with PBS two times. Subsequently, 2.5% glutaraldehyde as a crosslinking reagent was
EP
added to the samples and left in 4 ℃. After 2 h, the glutaraldehyde was rinsed with PBS and deionized water. The fixed cells were dehydrated via exposure to gradient of ethanol (50%, 60%,
CC
70%, 80%, 90%, and 100% ethanol, respectively) for every 15 min and allowed to air dry at 4
A
°C. The cell images of seeded cells on the scaffolds were observed by FESEM. Data were statistically analyzed using SPSS v. 25.0 software. Multiple samples were evaluated by one-way ANOVA followed by Student–Newman–Keuls post hoc tests. Measures are expressed as means ± standard deviations and considered significant at p < 0.05.
10
3. Results and discussion 3.1. Electrospinning The processing of cellulose usually presents many challenges in electrospinning because of
SC RI PT
limited solubility in conventional solvents, tendency to aggregate or form gels and inability to melt as a result of its high crystallinity, supramolecular architecture and rigid backbone structure (Frey, 2008). Hence, CA nanofibers were produced through electrospinning process and then deacetylated to convert into electrospun cellulose nanofibers. Electrospun CA nanofibers were completely regenerated to cellulose nanofibers through alkaline hydrolyze in 0.05 M
U
NaOH/ethanol solution at ambient temperature. Then, cellulose nanofibers were chemically
N
modified with MCAA and CHPTAC in the presence of sodium hydroxide to introduce negative
A
and positive charges, respectively.
M
The mechanism of carboxymethylation of cellulose surface is suggested and discussed (Fig. 1).
D
CMC can be formed in a two-step reaction based on mercerization (alkalization) and
TE
etherification as follow: In general, the cellulose nanofibers are swollen in concentrated sodium hydroxide solution which has destroyed the crystalline aggregation of cellulose and increased the
EP
accessibility of fibers to chemicals by swelling. Then, through a nucleophilic substitution reaction between alkalized cellulose and MCAA, CMC has been synthesized (Ambjörnsson,
A
CC
Schenzel, & Germgård, 2013).
11
ONa
OH O HO
NaOH
OH
HO O
O
O
HO
n
OH
n
OH
O O
OH
-H2O
O
OH
OH
HO O
Alkali Cellulose
Cellulose
O ONa
O HO O
O
O
-NaCl
O
OH
HO
OH
HO O
O
O
OH
n
OH
OH
SC RI PT
O HO
ClCH2COOH
OH
OH
n
CMC
Alkali Cellulose
Fig. 1. Mechanism of preparation of carboxymethylated cellulose from cellulose in the presence of MCAA under alkali conditions
U
The cationizating creates positive charges on the surfaces of cellulose without changing the size
N
and shape, considerable increase on the mechanical properties and cell adhesion of the
A
nanofibers. Cellulose nanofibers were cationized via two sequential wet-on-wet baths process: 1)
M
CHPTAC instantly converted to 2,3-epoxypropyl trimethylammonium chloride (EPTMAC) and
D
2) subsequently EPTMAC reacted with cellulose nanofibers to form cationic cellulose nanofibers
TE
(Fig. 2). In the cationization of cellulose nanofibers, the base catalyst is required to form the epoxide ring and to react of ammonium epoxides with the hydroxyl groups of cellulose. The
EP
obtained amount of carboxyl content of CMC, DSCMC, nitrogen content of QACC and DSQACC
A
CC
were 7.682%, 0.23,1. 714% and 0.24 respectively.
12
ONa
OH
HO
OH HO O
HO O
HO
n
OH
n
OH
O O
OH
-H2O
O
OH
OH
O
O
Alkali Cellulose
Cellulose O
OH
O
NaOH
H2CCHCH2N(CH3)3Cl
H2CCHCH2N(CH3)3Cl
Cl
Cl
CH2
CHCH2N(CH3)3Cl
-NaCl
SC RI PT
O
NaOH
EPTMAC
CHPTAC
OH O ONa
CH2 O
HO
H2C CHCH2N(CH3)3Cl O
CHCH2N(CH3)3Cl
OH HO O OH
HO
NaOH, H2O
O
OH
OH
O
O
HO O
O
O
OH
n
OH
n
QC
Alkali Cellulose
N
alkaline solution
U
Fig. 2. Mechanism of heterogeneous quaternization of cellulose nanofibers with CHPTAC in
A
3.2. Characterization of nanofibers
M
The surface morphologies and the fibers diameter were observed using FESEM as presented in Fig. 3. FESEM images represent the smooth, continuous and regular surface of CA nanofibers
D
(Fig. 3a), while, for cellulose, CMC and cationic cellulose nanofibers show surface roughness
TE
and irregularities; may be owing to NaOH soaking (Fig. 3b-d). As shown in Fig. 3, the structure
EP
and nanofibers shape were well retained after deacetylation, carboxymethylation and cationization. But slight changes were appeared in nanofibers surface roughness. The average
CC
diameter of nanofibers was estimated from the FESEM images by image j software (Table 1). Compared with CA nanofibers, the average diameter of the cellulose nanofibers did not
A
significantly change after deacetylation and modification.
13
SC RI PT U N A M D TE EP CC A Fig. 3. Representative FESEM images of (a) electrospun CA nanofibers, (b) regenerated cellulose nanofibers, (c) CMC nanofibers and (d) QC nanofibers
14
Table 1. Average diameter of nanofibers Item Average Diameter Standard Deviation
CA 491.52nm 253.99 nm
C 495.05 nm 204.85 nm
CMC 506.07 nm 201.69 nm
QC 503.99 nm 182.96nm
SC RI PT
FTIR spectroscopy was used to identify the change in the chemical structure of CA nanofibers during deacetylation and to investigate the presence of functional groups on the surface of chemically modified cellulose. Fig. 4 presents the FTIR spectra in the range of 400– 4000 cm–1. As can be observed, the vibrations of the acetate group from the CA sample were observed at 1735 cm–1 (C=O, carbonyl stretching), 1367 and 1430 cm–1 (C–CH3, symmetric and
N
U
antisymmetric bending of methyl) and 1217 cm–1 (C–O–C, alkoxyl stretching of the ester). These peaks disappeared from the cellulose spectrum and a broad absorption band ascribed to the
M
A
stretching vibration modes of O–H group at 3319 cm–1 was observed, which indicates the presence of more hydroxyl groups in the regenerated cellulose. FTIR spectrum of CMC shows
D
additional absorption bands at 1595 and 1414 cm–1 that can be assigned to asymmetrical and
TE
symmetric stretching of the carboxylate group of –COO, respectively. These FTIR results confirm that the cellulose was successfully modified into CMC. In the QC spectrum, new bands
EP
were observed at 1045 and 1116 cm–1, assign to the C–N stretching vibration of the quaternary
CC
ammonium groups. Moreover, the peaks at 1475 and 2920 cm–1 were ascribed to C–H bending and stretching vibration modes of the methyl groups on the
A
quaternary ammonium, respectively. This results demonstrated the introduction of quaternary ammonium groups on the cellulose backbone.
15
SC RI PT U N A M D TE
EP
Fig. 4. FTIR spectra of (a) electrospun CA nanofibers, (b) deacetylated CA nanofibers, (c) CMC nanofibers and (d) QC nanofibers
CC
The electrokinetic zeta potential of each material is indicative of the interaction of the surface charge with the exposure media. Fig. 5 shows the change in zeta potential of the nanofibers
A
versus pH ranging from 2 to 10. The cationic surface functionalization of cellulose nanofibers resulted in a positive zeta potential, while all other nanofibers presented negative zeta potential. Indeed, cationization increases zeta potential and changes the nanofibers surface charge. The results of zeta potential measurements of CMC nanofibers showed that considerable number of
16
negatively charged groups has been introduced to cellulose structure. CMC had the most negative zeta potential (-12.8 mV at pH 7) due to carboxyl groups on the surface, whereas the CA and cellulose samples had zeta potentials slightly negative and much closer to zero at pH 7,
SC RI PT
respectively. The zeta potential of CMC and cationic cellulose nanofibers were -12.8 and +22 mV at pH 7 respectively, which confirms the presence of negative and positive charges in the
TE
D
M
A
N
U
modified cellulose nanofibers.
Fig. 5. Zeta potential of cellulose based nanofibers versus pH
EP
The nanofibers were further characterized by investigating their swelling behavior in distilled water. The water uptake ability of the nanofibers was evaluated after 2 h and was shown in Fig.
CC
6. After about 2 h, an equilibrium water uptake was found which was confirmed without further
A
increase in weight upon contacting with water. Among the various samples, CA is hydrophobic, which exhibited the lowest water uptake ability. This is described owing to the lower availability of hydroxyl groups to form hydrogen bond network and consequently inhibiting the interaction of the polymer chains with water. Cellulose nanofibers provide some higher water uptake properties than CA nanofibers. The water uptake ratio was remarkably enhanced by chemical
17
modification of cellulose nanofibers. The results revealed that by increasing the time to 2 h, the swelling ratio of cationic cellulose and CMC nanofibers reached 642 % and 670%, respectively. This high water uptake ability is due to the chemical structure of these modified nanofibers,
SC RI PT
which are hydrophilic materials and exhibited improved water absorption capacity. CMC revealed the highest swelling among these scaffolds. Indeed, the degree of swelling is influenced by the acidic groups of cellulose, which carboxymethylation is the most extensively used process
TE
D
M
A
N
U
to introduce acidic groups into the cellulose network.
EP
Fig. 6. Water uptake ability of as-electrospun nanofibers
CC
Figure 7a compares the contact angles and surface wettability of prepared nanofibers after contacting with water drop at the final time (5 s). The change in contact angle over time (the
A
initial 5 s) after water drop deposition is illustrated in Fig. 7b. CA nanofibers were hydrophobic with water contact angle of 92o due to the presence of acetyl groups. With alkaline hydrolysis of acetyl groups in CA nanofibers, wettability of obtained cellulose nanofibers is dramatically improved. They became much more hydrophilic, reaching water contact angle of 58o. The differences in the water contact angle of samples can be corresponded to the changes of the 18
surface chemistry and roughness of the electrospun nanofibers during deacetylation. From water contact angle measurements, it was observed that there was no significant difference in the wettability of the cationic cellulose nanofibers and CMC nanofibers and both showed
SC RI PT
hydrophilic properties. The initial contact angle of water measured on cationically modified cellulose nanofibers was 47o; while for CMC nanofibers, it was 45o. The contact angle measurement results confirmed that when electrospun nanofibrous samples were modified, they
CC
EP
TE
D
M
A
N
U
became much more hydrophilic.
A
Fig. 7. Water contact angle measurements for samples (a) at final time, (b) over time
The mechanical characteristics in terms of the tensile strength and Young’s modulus were investigated and presented in Fig.8. When comparing mechanical properties of the nanofibers, it was obvious that surface modification improved the tensile strength and Young’s modulus of the scaffolds. A similar increasing trend was found for tensile strength of CMC and cationic 19
cellulose nanofibers, but Young’s modulus of CMC is significantly higher than those of produced from cationic cellulose nanofibers. It can be seen that the cationic cellulose and CMC nanofibers had tensile strength of 7 and 7.2 MPa, and Young’s modulus of 482 and 515 MPa,
SC RI PT
respectively. The mechanical properties exhibited direct correlation with interaction between the nanofibers. Obviously, cationization and anionization of nanofibers plays the main role in improving mechanical properties of scaffolds compared to unmodified nanofibers. In addition, tensile strength and Young’s modulus were enhanced via the heat and alkaline treatment process, consequently, dispersion and the structural integrity of nanofibers and the mechanical properties
U
were dramatically improved (Ma, Kotaki, & Ramakrishna, 2005). The improvement in tensile
N
strength was mainly due to cross-linking between the nanofibers after heat treatment led to
A
bonding between nanofibers at the crossover points. The improvement of mechanical properties
M
of modified nanofibers may have corresponded to the formation of rigid hydrogen bonds network between nanofibers and electrostatic interaction between introduced groups that cause a
D
good stress transfer from network to nanofibers (Alemdar & Sain, 2008a,b). It seems that
A
CC
EP
TE
interfiber hydrogen bonding and network structure result in improved mechanical strength.
20
SC RI PT U N A M D TE EP
CC
Fig. 8. Mechanical properties of nanofibers: (a) Tensile strength (MPa) and (b) Young’s modulus (MPa) of prepared nanofibers
A
3.3. Cellular proliferation, viability and cytotoxicity studies The biocompatibility of the nanofibrous scaffolds was investigated by evaluating cellular adhesion and viability of hMSCs cultured on the scaffolds. hMSCs were cultured at 37 °C in a humidified atmosphere (5% CO2). Then, cells were seeded on the nanofibers and cultured for 5
21
days. The standard test method of MTT assay was performed to quantitatively evaluate cell viability, proliferation and cytotoxicity onto the fibrous scaffolds. The colorimetric MTT assay, in fact measures the reduction of the tetrazolium molecule of MTT by viable cells to blue
SC RI PT
formazan product which reflects the normal function of mitochondria (He et al., 2013). In hence, the level of the reduction of MTT into formazan component can reveal the level of cellular metabolism. Fig. 9 illustrates the results of MTT assay after 1, 3 and 5 days of cell culture on the nanofibers in comparison to control. The optical densities (OD) of samples and control were measured at a wavelength of 570 nm by a microplate reader.
U
According to p values there is no significant difference between the control, CA and cellulose in
N
day 5, 3, 1 (p>0.05), only QC sample shows the significant increase in day 1, 3, 5, comparing to
A
control (p<0.001), for CMC there is no difference comparing to control in day 1, 5 (p>0.05), but
M
the increase of cell viability in day 3 was significant (p<0.05). As can be seen from Fig. 9, all nanofibers showed better cell proliferation after 5 day of cell culture compared to days of 1 and
D
3. It is clear that cells cultured onto the modified nanofibers scaffolds have the best viability at
TE
all the time points. The results of cell proliferation assay showed high viability for modified
EP
samples in comparison to control. Estimation of cellular growth using MTT assay indicated the modified nanofibers cause more increment in cell proliferation after 5 days. This is likely due to
CC
the fact that hydrophilicity and biocompatibility of modified nanofibrous scaffolds. Successful proliferation of the hMSCs on produced nanofibers describes potential applicability of the
A
scaffold for cell growth. The MTT assay tests showed that the scaffold did not produce any toxic effect on the hMSCs, proving their potential to support cell proliferation and utility for tissue engineering applications. Besides, the MTT assay results indicate that QC nanofibers are cell biocompatible and nontoxic to hMSCs with higher growth and proliferation ability.
22
SC RI PT U N A
M
Fig. 9. MTT assay for cell viability and proliferation of hMSCs on scaffolds and control after in vitro culture for days of 1, 3 and 5
D
3.4. Cell adhesion studies using FESEM
TE
The physicochemical properties and surface changes of nanofibers significantly influence the
EP
cell behavior, adhesion, proliferation and morphology (Stevens & George, 2005). FESEM images were studied to understand cell adhesion, proliferation and morphology on the scaffolds.
CC
As can be observed from the FESEM images (Fig. 10), the hMSCs attached and spread on the surface of scaffolds after culture. After incubation, attached cells start spreading on the scaffolds
A
and then cells spreading became more noticeable. The cells were observed in a flat, elongated and rounded morphology with monolayer like structure on the scaffold surface and a good cell adhesion to the scaffold was obvious from cell surface. The cell adhesion and proliferation are significantly influenced by the hydrophilicity of the scaffolds (Altankov & Groth, 1994). Moreover, the nanofibers surface roughness created by the presence functional groups could 23
promote cell adhesion to the nanofibers since cells generally display enhanced proliferation and bioactivity on rough surfaces (Kunzler, Drobek, Schuler, & Spencer, 2007). The results indicate that the reported nanofibrous scaffolds support cell adhesion/attachment and proliferation and
SC RI PT
hence are suitable for tissue engineering applications. The hydrophilic nature of modified
EP
TE
D
M
A
N
U
nanofibers promotes uniform adhesion of cells on the surface nanofibers.
nanofibers (c) CMC nanofibers and (d) QC nanofibers
A
CC
Fig. 10. FESEM images of hMSCs adhered on the surfaces of (a) CA nanofibers, (b) cellulose
4. Conclusions The novel, simple and inexpensive methods were reported to produce high performance advanced cellulosic materials with a wide range of functionalities makes them suitable for 24
valuable applications. The purpose of this study is to improve an effective and eco-friendly procedure in aqueous suspension for the surface modification of electrospun cellulose nanofibers using CHPTAC and MCAA. The FESEM images of CA nanofibers indicated narrow
SC RI PT
distribution, uniform structures and smooth morphology. After chemical modification no significant changes in shape or morphology were observed except for surface roughness and irregularities. The zeta potentials of CA and cellulose nanofibers at pH 7 were slightly negative and almost zero, respectively, indicating the deacetylation only resulted in a slight enhancement in the zeta potential compared to starting material. Quaternization produced a positive zeta
U
potential due to an increase in the surface charge and carboxymethylation created the most
N
negative zeta potential due to carboxyl group on the surfaces. Among the different samples, CA
A
(hydrophobic) showed the lowest swelling, because of the lower availability of hydroxyl group
M
to form hydrogen bonding resulted in preventing the interaction of the cellulose chains with water. When cellulose nanofibers were modified, the hydrophilicity was increased, the contact
D
angle was decreased with time, and the surface swelling was improved. It was revealed from
TE
MTT assay that the prepared nanofibers did not produce any cytotoxic effect on hMSCs.
EP
Furthermore, the modified nanofibers presented high cell viability. The cell adhesion is influenced by the surface chemistry of the nanofibers. When hMSCs were cultured on the
CC
surface of the modified nanofibers, favorable cell adhesion was observed owing to the hydrophilicity of surface. The findings showed that cell adhesion is strongly enhanced on the
A
modified surfaces with carboxymethyl and quaternary ammonium groups. The studies showed biocompatibility and non-cytotoxicity of nanofibers and so these nanofibers are favorable for cell adhesion and proliferation and hence are promising materials for biomedical applications. The results indicated that the produced scaffolds support cell adhesion and proliferation and hence
25
they are suitable for tissue engineering applications. We believe the use of cellulose derivatives to produce nanofibers using the electrospinning process presents an excessive opportunity for better application of cellulose nanofibers and development of novel presentations.
SC RI PT
Acknowledgments The authors are gratefully acknowledged the National Elites Foundation, Iran University of Medical Sciences and Sharif University of Technology, for their assistance and financial support. References
Abdul Khalil, H. P. S., Bhat, A. H., Ireana Yusra, A. F. (2012). Green composites from
U
sustainable cellulose nanofibrils: A review. Carbohydrate Polymers, 87, 963–979.
N
Alemdar, A., Sain, M. (2008a). Biocomposites from wheat straw nanofibers: Morphology,
A
thermal and mechanical properties. Composite Science and Technology, 68(2), 557–565.
M
Alemdar, A., Sain, M. (2008b). Isolation and characterization of nanofibers from agricultural residues–Wheat straw and soy hulls. Bioresource and Technology, 99(6), 1664–1671.
D
Altankov, G., Groth, T. (1994). Reorganization of substratum-bound fibronectin on hydrophilic
TE
and hydrophobic materials is related to biocompatibility. Journal of Materials Science:
EP
Materials in Medicine. 5(9-10), 732–737. Ambjörnsson, H. A., Schenzel, K., Germgård, U. (2013). Carboxymethyl cellulose produced at
CC
different mercerization conditions and characterized by NIR FT Raman spectroscopy in combination with multivariate analytical methods. BioResources, 8(2), 1918–1932.
A
Antczak, T. (2012). Nanotechnology-methods of manufacturing cellulose nanofibers, Fibres & Textiles in Eastern Europe, 20 (2), 91.
26
Bacakova, L., Novotna, K., Sopuch, T., Havelk, P. (2015). Cell interaction with cellulose-based scaffolds for tissue engineering: a review. In: Cellulose and Cellulose Composites Editor: Md. Ibrahim H. Mondal, Nova Science Publishers, Inc. 341–376.
SC RI PT
Chen, P., Yu, H., Liu, Y., Chen, W., Wang, X., Ouyang, M. (2013). Concentration effects on the isolation and dynamic rheological behavior of cellulose nanofibers via ultrasonic processing, Cellulose 20, 149–157.
Deepa, B., Abraham, E., Cherian, B. M., Bismarck, A., Blaker, J. J., Pothan, L. A., Leao, A. L., de Souza, S. F., Kottaisamy, M. (2011). Structure, morphology and thermal characteristics of
U
banana nano fibers obtained by steam explosion, Bioresource Technology, 102, 1988–1997.
N
Fei, P., Liao, L., Meng, J., Cheng, B., Hu, X., Song, J. (2018). Non-leaching antibacterial
A
cellulose triacetate reverse osmosis membrane via covalent immobilization of quaternary
M
ammonium cations. Carbohydrate polymers, 181, 1102–1111. Frey, M. (2008). Electrospinning cellulose and cellulose derivatives, Polymer Reviews, 48 378–
D
391.
TE
Hasani, M., Cranston, E. D., Westman, G., Gray, D. G. (2008). Cationic surface
EP
functionalization of cellulose nanocrystals. Soft Matter, 4(11), 2238–2244. Hashem, M. M. (2006). Development of a one‐stage process for pretreatment and cationisation
CC
of cotton fabric. Coloration technology, 122(3), 135–144.
He, X., Xiao, Q., Lu, C., Wang, Y., Zhang, X., Zhao, J., Zhang, W., Zhang, X., Deng, Y. (2014).
A
Uniaxially Aligned Electrospun All-Cellulose Nanocomposite Nanofibers Reinforced with Cellulose Nanocrystals: Scaffold for Tissue Engineering. Biomacromolecules 15 (2), 618– 627.
27
Hebeish, A. A., El-Rafie, M. H., Abdel-Mohdy, F. A., Abdel-Halim, E. S., Emam, H. E. (2010). Carboxymethyl cellulose for green synthesis and stabilization of silver nanoparticles, Carbohydrate Polymers, 82, 933–941.
SC RI PT
Hong, K. M. (2013). Preparation and characterization of Carboxymethyl cellulose from sugarcane bagasse, Universiti Tunku Abdul Rahman.
Huang, Y. X., Ren, J., Chen, C., Ren, T. B., Zhou, X. Y. (2008). Preparation and properties of poly(lactide-co-glycolide) (PLGA)/nano-hydroxyapatite (NHA) scaffolds by thermally induced phase separation and rabbit MSCs culture on scaffolds. Journal of Biomaterial
U
Applied Science, 22, 409–432.
N
Huang, Y., Yu, H., Xiao, C. (2007). pH-sensitive cationic guar gum/poly (acrylic acid)
A
polyelectrolyte hydrogels: swelling and in vitro drug release. Carbohydrate Polymers, 69,
M
774–783.
Inukai, S., Kurokawa, N., & Hotta, A. (2018). Annealing and saponification of electrospun
D
cellulose-acetate nanofibers used as reinforcement materials for composites. Composites Part
TE
A: Applied Science and Manufacturing, 113, 158-165.
EP
Khatri, Z., Mayakrishnan, G., Hirata, Y., Wei, K., Kim, I. (2013). Cationic-cellulose nanofibers: Preparation and dyeability with anionic reactive dyes for apparel application. Carbohydrate
CC
polymers 91 (1), 434–443. Kunzler, T. P., Drobek, T., Schuler, M., Spencer, N. D. (2007). Systematic study of osteoblast
A
and fibroblast response to roughness by means of surface-morphology gradients. Biomaterials 28, 2175–2182.
Ma, Z., Kotaki, M., Inai, R., Ramakrishna, S. (2005). Potential of nanofiber matrix as tissue engineering scaffolds. Tissue engineering, 11, 101.
28
Ma, Z., Kotaki, M., Ramakrishna, S. (2005). Electrospun cellulose nanofber as afnity membrane. Journal of Membrane Science, 265 (1-2), 115–123, Menon, M. P., Selvakumar, R., Ramakrishna, S. (2017). Extraction and modification of cellulose
SC RI PT
nanofibers derived from biomass for environmental application, RSC Advances, 7(68), 42750–42773.
Pelton, R. (2009). Bioactive paper provides a low-cost platform for diagnostics, TrAC Trends in Analytical Chemistry, 28 925–942.
Rezaei, A., Nasirpour, A., Fathi, M. (2015). Application of cellulosic nanofibers in food science
U
using electrospinning and its potential risk. Comprehensive Reviews in Food Science and
N
Food Safety, 14(3), 269–284.
A
Siró, I., Plackett, D. (2010). Microfibrillated cellulose and new nanocomposite materials: a
M
review, Cellulose 17, 459–494.
Science, 310, 1135–1138.
D
Stevens, M. M., George, J. H. (2005). Exploring and engineering the cell surface interface.
TE
Song, Y., Sun, Y., Zhang, X., Zhou, J., Zhang, L. (2008). Homogeneous quaternization of
2264.
EP
cellulose in NaOH/urea aqueous solutions as gene carriers. Biomacromolecules 9(8), 2259–
CC
Song, Y., Wang, H., Zeng, X., Sun, Y., Zhang, X., Zhou, J., Zhang, L. (2010). Effect of molecular weight and degree of substitution of quaternized cellulose on the efficiency of
A
gene transfection, Bioconjugate Chemistry, 21, 1271–1279.
Song, Y., Zhang, L., Gan, W., Zhou, (2011). Self-assembled micelles based on hydrophobically modified quaternized cellulose for drug delivery, Colloids and Surfaces B: Biointerfaces, 83, 313–320.
29
Song, Y., Zhou, J., Li, Q., Guo, Y., Zhang, L. (2009). Preparation and characterization of novel quaternized cellulose nanoparticles as protein carriers. Macromolecular Bioscience, 9, 857– 863.
SC RI PT
Song, Y., Zhou, Y., Chen, L. (2012). Wood cellulose-based polyelectrolyte complex nanoparticles as protein carriers. Journal of Materials Chemistry, 22, 2512–2519.
Teeri, T. T., Brumer, H., Daniel, G., Gatenholm, P. (2007). Biomimetic engineering of cellulosebased materials. Trends in Biotechnology, 25, 299–306.
Yamamoto, S., Iwata, T., Nishiwaki, K., Kinoshita, M., & Suzuki, S. (2015). Synthesis of
N
Electrophoresis. Chromatography, 36(3), 93-98.
U
Quaternary Ammonium Derivatives of Cellulose as the Coating Reagents for Capillary
A
Yang, W. J., Fu, J., Wang, D. X., Wang, T., Wang, H., Jin, S. G., He, N. Y. (2010). Study on
M
chitosan/polycaprolactone blending vascular scaffolds by electrospinning. Journal of Biomedical Nanotechnology, 6, 254.
D
Yu, S., Cheng, Q., Huang, C., Liu, J., Peng, X., Liu, M., & Gao, C. (2013). Cellulose acetate
TE
hollow fiber nanofiltration membrane with improved permselectivity prepared through
EP
hydrolysis followed by carboxymethylation. Journal of membrane science, 434, 44-54. Zahedi, P., Rezaeian, I., Ranaei-Siadat, S., Jafari, S., Supaphol, P. (2009). A review on wound
CC
dressings with an emphasis on electrospun nanofibrous polymeric bandages, Polymers for Advanced Technologies, 21, 77–95.
A
Zarth, C. S. P. (2013). Novel cationic esters derived from cellulose and dextran: synthesis, structure analysis and properties. LAP LAMBERT Academic Publishing.
Zhou, Q., Rutland, M., Teeri, T., Brumer, H. (2007). Xyloglucan in cellulose modification. Cellulose, 14, 625–641.
30
S0144-8617(18)31462-0 https://doi.org/10.1016/j.carbpol.2018.12.028 CARP 14382
To appear in: Received date: Revised date: Accepted date:
23 June 2018 3 November 2018 10 December 2018
Please cite this article as: Golizadeh M, Karimi A, Gandomi-Ravandi S, Vossoughi M, Khafaji M, Joghataei MT, Faghihi F, Evaluation of cellular attachment and proliferation on different surface charged functional cellulose electrospun nanofibers, Carbohydrate Polymers (2018), https://doi.org/10.1016/j.carbpol.2018.12.028 This is a PDF file of an unedited manuscript that has been accepted for publication. As a service to our customers we are providing this early version of the manuscript. The manuscript will undergo copyediting, typesetting, and review of the resulting proof before it is published in its final form. Please note that during the production process errors may be discovered which could affect the content, and all legal disclaimers that apply to the journal pertain.
Evaluation of cellular attachment and proliferation on different surface charged functional cellulose electrospun nanofibers
Golizadeh1,2,
Afzal
Karimi1,3,*,
Soheila
Gandomi-Ravandi
1
,
Manouchehr
SC RI PT
Mortaza
Vossoughi2,4,**, Mona Khafaji4, Mohammad Taghi Joghataei3,5, Faezeh Faghihi3
1
Faculty of Advanced Technologies in Medicine, Iran University of Medical Sciences,
1449614535 Tehran, Iran
Chemical and Petroleum Engineering Department, Sharif University of Technology, Tehran,
U
2
Cellular and Molecular Research Center, Iran University of Medical Sciences, 1449614535
A
3
N
14588-89694, Iran.
4
M
Tehran, Iran
Institute for Nanoscience and Nanotechnology, Sharif University of Technology, 14588-89694
School of Medicine, Iran University of Medical Sciences, 1449614535 Tehran, Iran
TE
5
D
Tehran, Iran.
*Corresponding author: Afzal Karimi
EP
Email: [email protected]
CC
Phone: +98 2188622687 Fax: +98 2186704606
A
** Corresponding author: Manouchehr Vossoughi Email: [email protected] Phone: +98 2166165487 Fax: +98 2166005417
1
SC RI PT
Graphical abstract
Highlights
Cellulose nanofibers were prepared by cellulose acetate electrospun deacetylation.
Anionic and cationic cellulose nanofibers were fabricated by chemical modification.
Several techniques were employed to characterize the cellulose based nanofibers.
Cellular studies were carried out to determine the cells viability.
Cationic and anionic cellulose nanofibers are favorable for biomedical application.
EP
Abstract
TE
D
M
A
N
U
CC
Fabrication and characterization of different surface charged cellulose electrospun scaffolds including cellulose acetate (CA), cellulose, carboxymethyl cellulose (CMC) and quaternary
A
ammonium cationic cellulose (QACC) for biomedical applications have been reported in this research. Several instrumental techniques were employed to characterize the nanofibers. MTT assay and cell attachment studies were also carried out to determine the cytocompatibility, viability and proliferation of the scaffolds. Fabricated CA, cellulose, CMC and QACC nanofibers had 100-600 nm diameter, -9, -1.75, -12.8, +22 mV surface potential, 2.5, 4.2, 7.2, 7 2
MPa tensile strength, 122, 320, 515, 482 MPa Young modules, 430, 530, 670 and 642% water uptake and 92o, 58o, 45o, 47o contact angle respectively. The findings showed that cell adhesion and proliferation is strongly enhanced on the modified surfaces with quaternary ammonium and
SC RI PT
carboxymethyl groups. We believe the use of cationic and anionic surface modified cellulose electrospun nanofibers presents promising materials for biomedical applications.
Keywords: Electrospun nanofiber, Cellulose, Carboxymethyl cellulose, Quaternized cellulose, Scaffold, Surface charge.
1. Introduction
U
In recent years, research on developing sustainable, green and environmentally friendly and
N
biocompatible nanosized materials has gained much interest (Abdul Khalil, Bhat, & Ireana
A
Yusra, 2012). In this regard, great efforts have been made to develop renewable, biodegradable
M
and biomimetic materials based on polysaccharides (Teeri, Brumer, Daniel, & Gatenholm, 2007;
D
Zhou, Rutland, Teeri, & Brumer, 2007). Especially, biomass based polymers such as cellulose substances have become the hotspot of science due to their intrinsic properties. Cellulose has
TE
been evaluated by Pelton as “particularly protein and biomolecule friendly” (Pelton, 2009).
EP
Cellulose has been applied for fabricating three-dimensional scaffolds, which have provided good support for cell adhesion and growth. The clinical applications of cellulose-based materials
CC
as scaffolds are including repairing, reconstructing and regenerating of almost all type of tissues in the mammalian organism. In tissue engineering and cell delivery, cellulose supports, cover
A
wounds and release drugs into them, through inhibiting postoperative adhesions, hemodialysis, hemostasis, or via covering and filling different tissue defects (Bacakova, Novotna, Sopuch, & Havelk, 2015).
3
With the development of nanotechnology, cellulose nanofibers have been widely studied due to environmentally benign nature, relatively easy production and compatibility with biological systems (Menon, Selvakumar, & Ramakrishna, 2017). Processing of cellulose to produce
SC RI PT
nanofibers can be carried out by a variety of techniques such as steam explosion (Deepa et al., 2011), ultrasonication (Chen et al., 2013), homogenization, grinding, cryocrushing (Siró & Plackett, 2010), refining and electrospinning (Inukai, Kurokawa, & Hotta, 2018; Antczak, 2012). Among these, electro-static spinning or “electrospinning” is a versatile and efficient method in terms of its simplicity and cost-effective setup. Moreover it allows producing smooth, continuous
U
ultrafine and highly porous nanofibers with highly controllable physical properties (Huang,
N
Zhang, Kotaki, & Ramakrishna, 2003). Polymer-based electrospun nanofibers have been widely
A
applied in tissue engineering (skin, bone, nerve and blood vessel) as scaffolds (Ma, Kotaki, Inai,
M
& Ramakrishna, 2005) for improvement of cell growth, controlling the proliferation of cells or neurons and regulation of cell behavior (Yang et al., 2010). The application of cellulose scaffolds
D
in tissue engineering and cell therapies needs to be developed. For this purpose, cellulose
TE
materials can be modified by combinations of cellulose with other synthetic or natural polymers,
EP
metals, ceramics, carbon materials or various chemical groups by and loading with bioactive molecules such as drugs and growth factors. These modifications, control the physical and
CC
chemical properties of cellulose scaffolds for cell-material interaction (Bacakova, Novotna, Sopuch, & Havelk, 2015).
A
CMC is a natural anionic polyelectrolyte prepared by introducing carboxymethyl groups along the cellulose chain. Carboxymethylation of cellulose improves solubility, stability and hydrophilicity of suspension, decreases the crystallinity index, increases the breakdown of fibers to nano size, decreases the agglomeration of fibers and makes the surface negatively charged
4
through functionalization of cellulose by breaking the hydrogen bonds (Rezaei, Nasirpour, & Fathi, 2015). Over recent decades, QACC as a new environmental friendly material is found special
SC RI PT
technological interest. Cationic cellulose derivatives are large-scale commercial products due to hydrophilicity, adsorption capacity for anionic substances, ionic exchange capacity (Yamamoto, Iwata, Nishiwaki, Kinoshita, & Suzuki, 2015; Zarth, 2013) biocompatibility, low hemo- and cytotoxicity, biodegradability, lack of skin irritation, low corrosiveness, good environmental stability, bacteriostatic properties, excellent cell membrane penetration properties and
U
antimicrobial activity (Fei et al., 2018). Quaternized cellulose (QC), as a positively charged
N
polysaccharide, is commonly produced by etherification of alkali cellulose. In this process,
A
quaternary epoxides or quaternary halohydrins act as cationizing reagent that producing
M
quaternary ammonium groups on the hydroxyl groups of cellulose (Hasani, Cranston, Westman, & Gray, 2008; Song, Sun, Zhang, Zhou, & Zhang, 2008). Cytotoxicity of cationic cellulose was
D
related to the molecular weight (Mw) and degree of substitution (DS). Lower Mw and DS of
TE
cationic celluloses appeared to be less cytotoxic compared to those with higher values. It was
EP
reported that cationic celluloses with DS value between 0.2-0.6 displayed relatively low cytotoxicity so that cells viability was almost 100% (Song et al., 2010). Among cationizing
CC
agents, 3-chloro-2-hydroxypropyltrimethylammonium chloride (CHPTAC) has attracted much interest in cationization of cellulose because of commercially availability, low toxicity and good
A
reactivity (Hashem, 2006). The studies have shown that the presence of quaternary ammonium and numerous hydroxyl groups in QC backbone afford various electrostatic interactions with ionic and non-ionic therapeutic molecules (Huang, Yu, & Xiao, 2007). QCs have exhibited elevated potential in biomedicine fields as promising nonviral gene (Song et al., 2010) and
5
protein (Song, Zhou, Li, Guo, & Zhang, 2009; Song, Zhou, & Chen, 2012) carriers, in drug delivery systems (Song, Zhang, Gan, & Zhou, 2011) and wound-dressing applications (Zahedi, Rezaeian, Ranaei-Siadat, Jafari, & Supaphol, 2009).
SC RI PT
Here, we describe CA electrospinning and preparation of cellulose nanofibers through its deacetylation. Moreover, surface modification of electrospun cellulose nanofibers is carried out using monochloroacetic acid (MCAA) and CHPTAC to obtain CMC and QC nanofibers, respectively. In addition, we evaluated the potential application of the modified cellulose derivatives as biomedical materials. At last, the structural, morphological, mechanical, swelling,
N
U
wettability and the cell culture properties of the scaffolds were analyzed and compared.
A
2. Materials and methods
M
2.1. Materials
D
Cellulose acetate (average molecular weight 30 kDa g/mol, 39.8 wt% acetyl content; degree of
TE
acetyl substitution 2.4) was purchased from Aldrich Chemical Compound. The cationizing agent, 3-chloro-2-hydroxypropyltrimethylammonium chloride (CHPTAC, aqueous solution of 69 wt
EP
%) was obtained from Fluka. Other chemicals, namely, sodium hydroxide, hydrochloric acid, glutaraldehyde and MCAA were received from Sigma–Aldrich Chemicals. All solvents such as
CC
acetone, ethanol, isopropanol and N,N-dimethylformamide (DMF) were provided by Merck (Germany). All reagents and chemicals were of analytical grade and were used as received
A
without further purification. All experiments were performed using double-distilled water. For cell culture study, Human Bone Marrow Derived Mesenchymal Stem Cells (hMSCs) were obtained from the Royan Cell Bank, Tehran, Iran. MTT (3-(4,5-Dimethylthiazol-2-yl)-2,5diphenyl phenyltetrazolium bromide) reagent (98%), phosphate-buffered saline (PBS, pH 7.4),
6
Dulbecco’s modified eagle’s medium (DMEM)-F12, fetal bovine serum (FBS) and penicillinstreptomycin (pen-strip) were purchased from GIBCO Invitrogen (Carlsbad, CA, USA). 2.2. Fabrication and modification of electrospun cellulose nanofibers
SC RI PT
Electrospun CA nanofibers were synthesized following the procedure described in literature (Khatri, Mayakrishnan, Hirata, Wei, & Kim, 2013). Electrospinning apparatus (SBS LAB ES SBS, Nanoazma, Tehran, Iran) was applied to form cellulose nanofibers. The spinning solution was prepared by dissolving 17% w/v CA in a binary solvent system of DMF: acetone (2:1 w/w) under constant stirring at room temperature until the solution became transparent. Freshly
U
prepared CA solution was loaded into a 10 mL tip plastic syringe with a stainless steel needle as
N
the nozzle. The flow rate was controlled by the syringe pump at 1 mL/h. The supplied voltage
A
was 12.5 kV and the needle tip to collector distance was 15 cm. The rotational speed of the roller
M
during electrospinning was set at 100 rpm to get a uniform mesh. The obtained electrospun
D
nanofibers were detached from the collectors and dried in a vacuum oven at 60 ℃ for 5 h to
TE
remove residual solvent. The electrospun CA nanofibers were deacetylated in 0.05 M NaOH ethanol solution at room temperature for 24 h. After deacetylation, the cellulose nanofibers were
EP
rinsed thoroughly with an excess amount of deionized water to remove sodium and acetate ions. The alkaline solution was prepared by dissolving of 5 g NaOH in 5 mL distilled water and then
CC
mixed with 100 mL ethanol. The cellulose nanofibers were soaked into the solution for 1 h. The mercerization was continued for 1 h to alkalize the cellulose.
A
CMC nanofibers were synthesized by dissolving 0.75 g MCAA in the alkaline solution containing mercerized cellulose nanofibers at 60 ℃ for 2 h. The produced CMC nanofibers were soaked in distilled water for 3 h and were dried in a vacuum oven at 60 ℃ for 5 h.
7
The QC nanofibers were produced by dissolving 5 g CHPTAC solution in alkaline solution containing mercerized cellulose nanofibers at 80 ℃ for 2 h. The produced QC nanofibers were soaked in distilled water for 3 h and were dried in a vacuum oven at 60 ℃ for 5 h.
SC RI PT
2.3. Characterization of electrospun cellulose based nanofibers The surface morphology and distribution of the prepared nanofibers before and after cell culture were observed using Field Emission Scanning Electron Microscope (FESEM) (Tescan – mira 3, Czech Republic) operated at 15 kV accelerating voltage. Prior to the FESEM analysis, the samples were sputter coated with a thin layer of gold (6 nm) to avoid charge accumulation.
U
Chemical structures of the cellulose samples were characterized on a Nicolette 6700 Fourier
N
transform infrared (FT–IR) spectrometer (Thermo Fisher Scientific Co. Ltd., MA, USA) with the
A
wavenumber range of 4000–400 cm–1 using KBr pellets. Nanofibers were dried at 60 ℃ for 12 h
M
before the measurement.
D
DS value of CMC sample was determined by the standard method ASTM D1439 (Hong, 2013).
TE
The nitrogen content of the QC nanofibers was measured to determine the DS. Elemental analysis (CHNS Analyzer, FlashEA 1112 series, Thermo Finnigan, Italy) was employed to
EP
determine the presence and mass percentage of nitrogen in cationically modified cellulose nanofibers (Song et al. 2010). Contact angle measurements were performed to investigate the
CC
wettability properties of the nanofibers mat using a G10 goniometer from Krüss (Germany) at
A
room temperature. Briefly, deionized water was dropped onto the nanofibers mat from a needle. Picture of the drop was taken and analyzed. The contact angle was obtained after 0, 1, 2, 3, 4 and 5 s. The water uptake measurements were carried out using a simple method. The scaffolds were cut into small squares at dimensions of 1 × 1 cm and their weights were measured. This is the dry weight (Wd) of each sample. Then, each sample was immersed in 50 mL of deionized 8
water for 2 h at room temperature. To measure swelling behavior, after the immersion period, the samples were reweighed to record its wet weight (Ww). The amount of water uptake of the samples was calculated from equation (1) (Huang, Ren, Chen, Ren, & Zhou, 2008):
W w W d 100 Wd
(1)
SC RI PT
Water Uptake % =
The nanofiber mats surface zeta potential was estimated by streaming potential measurements, which were carried out using an electro kinetic analyzer (EKA, Anton Paar KG, Graz, Austria) at 25 ℃ (Yu et al. 2013). The mechanical properties of the scaffolds containing Young’s modulus
U
and tensile strength were measured on a BOSE mechanical testing machine (Model ELF 3200).
N
Tensile test was carried out at 35-40% relative humidity and 1 mm/min speed until the samples
A
were ruptured. The nanofibers were cut into 15 mm × 10 mm (L × W) strips. The average
M
thicknesses of electrospun nanofibers were 220±15 µm. 2.5. Cell culture studies
D
hMSCs were used to determine the cell viability on the cellulose scaffolds. The cells were
TE
cultured in (DMEM)-F12 medium supplemented with 10% FBS and 1% pen-strip. The cells
EP
were incubated at 37 °C in a humidified atmosphere containing 5% CO2. The nanofibers were cut into circular shape that fit into the 24-well plates. The standard cell culture treated 24-well
CC
plates used as a control. Cells in all the samples were cultured in the same growth medium and treated in the same manner. The nanofibrous scaffolds were immersed in 70 % v/v ethanol for 12
A
h followed by washing with sterile PBS three times for 20 min. The nanofibers were sterilized under UV light for 1 h on each side. Under sterile conditions, DMEM-F12 medium containing 10% FBS was added to each well and nanofibers scaffolds allowed to hydrate for 24 h at 37 ℃. The cultured cells were seeded on the samples with a density of 16,000 cells/cm2. In total, 1000
9
μL of culture medium was added to each well. Cells were incubated in 5% CO2 incubator at 37 ℃ and the culture medium was replaced every 2 days. After incubation, cell proliferation and scaffold cytotoxicity were studied by MTT assay.
SC RI PT
MTT assay was performed at 1, 3, and 5 days of cell-seeding. On the day of the test (days 1, 3, and 5), 500 µL of medium in the cell-loaded scaffold culture plate was removed and replaced by 50 μL of MTT solution (0.5 mg/mL) in PBS (pH 7.4). After 4 h incubation at 37 ℃ in the humid environment with 5% CO2, the MTT containing medium was gently removed. After that, 150 µL acidic isopropanol (0.1 N HCL and isopropanol) was added to dissolve purple formazan crystals
U
formed in live cells and kept in the dark at room temperature for 15 min. Then, 100 µL
N
supernatant solution of each sample (the dissolved formazan solution) transferred into individual
A
well of a 96-well plate. The absorbance of each well was measured at 570 nm using a microplate
M
reader and results were averaged from three replications. (Synergy HTX, Biotek).
D
The hMSCs were used to investigate the adhesion behavior of cells onto the scaffolds using a
TE
cell adhesion assay. After the cells were cultured on the nanofibers for 24 h, each sample was rinsed with PBS two times. Subsequently, 2.5% glutaraldehyde as a crosslinking reagent was
EP
added to the samples and left in 4 ℃. After 2 h, the glutaraldehyde was rinsed with PBS and deionized water. The fixed cells were dehydrated via exposure to gradient of ethanol (50%, 60%,
CC
70%, 80%, 90%, and 100% ethanol, respectively) for every 15 min and allowed to air dry at 4
A
°C. The cell images of seeded cells on the scaffolds were observed by FESEM. Data were statistically analyzed using SPSS v. 25.0 software. Multiple samples were evaluated by one-way ANOVA followed by Student–Newman–Keuls post hoc tests. Measures are expressed as means ± standard deviations and considered significant at p < 0.05.
10
3. Results and discussion 3.1. Electrospinning The processing of cellulose usually presents many challenges in electrospinning because of
SC RI PT
limited solubility in conventional solvents, tendency to aggregate or form gels and inability to melt as a result of its high crystallinity, supramolecular architecture and rigid backbone structure (Frey, 2008). Hence, CA nanofibers were produced through electrospinning process and then deacetylated to convert into electrospun cellulose nanofibers. Electrospun CA nanofibers were completely regenerated to cellulose nanofibers through alkaline hydrolyze in 0.05 M
U
NaOH/ethanol solution at ambient temperature. Then, cellulose nanofibers were chemically
N
modified with MCAA and CHPTAC in the presence of sodium hydroxide to introduce negative
A
and positive charges, respectively.
M
The mechanism of carboxymethylation of cellulose surface is suggested and discussed (Fig. 1).
D
CMC can be formed in a two-step reaction based on mercerization (alkalization) and
TE
etherification as follow: In general, the cellulose nanofibers are swollen in concentrated sodium hydroxide solution which has destroyed the crystalline aggregation of cellulose and increased the
EP
accessibility of fibers to chemicals by swelling. Then, through a nucleophilic substitution reaction between alkalized cellulose and MCAA, CMC has been synthesized (Ambjörnsson,
A
CC
Schenzel, & Germgård, 2013).
11
ONa
OH O HO
NaOH
OH
HO O
O
O
HO
n
OH
n
OH
O O
OH
-H2O
O
OH
OH
HO O
Alkali Cellulose
Cellulose
O ONa
O HO O
O
O
-NaCl
O
OH
HO
OH
HO O
O
O
OH
n
OH
OH
SC RI PT
O HO
ClCH2COOH
OH
OH
n
CMC
Alkali Cellulose
Fig. 1. Mechanism of preparation of carboxymethylated cellulose from cellulose in the presence of MCAA under alkali conditions
U
The cationizating creates positive charges on the surfaces of cellulose without changing the size
N
and shape, considerable increase on the mechanical properties and cell adhesion of the
A
nanofibers. Cellulose nanofibers were cationized via two sequential wet-on-wet baths process: 1)
M
CHPTAC instantly converted to 2,3-epoxypropyl trimethylammonium chloride (EPTMAC) and
D
2) subsequently EPTMAC reacted with cellulose nanofibers to form cationic cellulose nanofibers
TE
(Fig. 2). In the cationization of cellulose nanofibers, the base catalyst is required to form the epoxide ring and to react of ammonium epoxides with the hydroxyl groups of cellulose. The
EP
obtained amount of carboxyl content of CMC, DSCMC, nitrogen content of QACC and DSQACC
A
CC
were 7.682%, 0.23,1. 714% and 0.24 respectively.
12
ONa
OH
HO
OH HO O
HO O
HO
n
OH
n
OH
O O
OH
-H2O
O
OH
OH
O
O
Alkali Cellulose
Cellulose O
OH
O
NaOH
H2CCHCH2N(CH3)3Cl
H2CCHCH2N(CH3)3Cl
Cl
Cl
CH2
CHCH2N(CH3)3Cl
-NaCl
SC RI PT
O
NaOH
EPTMAC
CHPTAC
OH O ONa
CH2 O
HO
H2C CHCH2N(CH3)3Cl O
CHCH2N(CH3)3Cl
OH HO O OH
HO
NaOH, H2O
O
OH
OH
O
O
HO O
O
O
OH
n
OH
n
QC
Alkali Cellulose
N
alkaline solution
U
Fig. 2. Mechanism of heterogeneous quaternization of cellulose nanofibers with CHPTAC in
A
3.2. Characterization of nanofibers
M
The surface morphologies and the fibers diameter were observed using FESEM as presented in Fig. 3. FESEM images represent the smooth, continuous and regular surface of CA nanofibers
D
(Fig. 3a), while, for cellulose, CMC and cationic cellulose nanofibers show surface roughness
TE
and irregularities; may be owing to NaOH soaking (Fig. 3b-d). As shown in Fig. 3, the structure
EP
and nanofibers shape were well retained after deacetylation, carboxymethylation and cationization. But slight changes were appeared in nanofibers surface roughness. The average
CC
diameter of nanofibers was estimated from the FESEM images by image j software (Table 1). Compared with CA nanofibers, the average diameter of the cellulose nanofibers did not
A
significantly change after deacetylation and modification.
13
SC RI PT U N A M D TE EP CC A Fig. 3. Representative FESEM images of (a) electrospun CA nanofibers, (b) regenerated cellulose nanofibers, (c) CMC nanofibers and (d) QC nanofibers
14
Table 1. Average diameter of nanofibers Item Average Diameter Standard Deviation
CA 491.52nm 253.99 nm
C 495.05 nm 204.85 nm
CMC 506.07 nm 201.69 nm
QC 503.99 nm 182.96nm
SC RI PT
FTIR spectroscopy was used to identify the change in the chemical structure of CA nanofibers during deacetylation and to investigate the presence of functional groups on the surface of chemically modified cellulose. Fig. 4 presents the FTIR spectra in the range of 400– 4000 cm–1. As can be observed, the vibrations of the acetate group from the CA sample were observed at 1735 cm–1 (C=O, carbonyl stretching), 1367 and 1430 cm–1 (C–CH3, symmetric and
N
U
antisymmetric bending of methyl) and 1217 cm–1 (C–O–C, alkoxyl stretching of the ester). These peaks disappeared from the cellulose spectrum and a broad absorption band ascribed to the
M
A
stretching vibration modes of O–H group at 3319 cm–1 was observed, which indicates the presence of more hydroxyl groups in the regenerated cellulose. FTIR spectrum of CMC shows
D
additional absorption bands at 1595 and 1414 cm–1 that can be assigned to asymmetrical and
TE
symmetric stretching of the carboxylate group of –COO, respectively. These FTIR results confirm that the cellulose was successfully modified into CMC. In the QC spectrum, new bands
EP
were observed at 1045 and 1116 cm–1, assign to the C–N stretching vibration of the quaternary
CC
ammonium groups. Moreover, the peaks at 1475 and 2920 cm–1 were ascribed to C–H bending and stretching vibration modes of the methyl groups on the
A
quaternary ammonium, respectively. This results demonstrated the introduction of quaternary ammonium groups on the cellulose backbone.
15
SC RI PT U N A M D TE
EP
Fig. 4. FTIR spectra of (a) electrospun CA nanofibers, (b) deacetylated CA nanofibers, (c) CMC nanofibers and (d) QC nanofibers
CC
The electrokinetic zeta potential of each material is indicative of the interaction of the surface charge with the exposure media. Fig. 5 shows the change in zeta potential of the nanofibers
A
versus pH ranging from 2 to 10. The cationic surface functionalization of cellulose nanofibers resulted in a positive zeta potential, while all other nanofibers presented negative zeta potential. Indeed, cationization increases zeta potential and changes the nanofibers surface charge. The results of zeta potential measurements of CMC nanofibers showed that considerable number of
16
negatively charged groups has been introduced to cellulose structure. CMC had the most negative zeta potential (-12.8 mV at pH 7) due to carboxyl groups on the surface, whereas the CA and cellulose samples had zeta potentials slightly negative and much closer to zero at pH 7,
SC RI PT
respectively. The zeta potential of CMC and cationic cellulose nanofibers were -12.8 and +22 mV at pH 7 respectively, which confirms the presence of negative and positive charges in the
TE
D
M
A
N
U
modified cellulose nanofibers.
Fig. 5. Zeta potential of cellulose based nanofibers versus pH
EP
The nanofibers were further characterized by investigating their swelling behavior in distilled water. The water uptake ability of the nanofibers was evaluated after 2 h and was shown in Fig.
CC
6. After about 2 h, an equilibrium water uptake was found which was confirmed without further
A
increase in weight upon contacting with water. Among the various samples, CA is hydrophobic, which exhibited the lowest water uptake ability. This is described owing to the lower availability of hydroxyl groups to form hydrogen bond network and consequently inhibiting the interaction of the polymer chains with water. Cellulose nanofibers provide some higher water uptake properties than CA nanofibers. The water uptake ratio was remarkably enhanced by chemical
17
modification of cellulose nanofibers. The results revealed that by increasing the time to 2 h, the swelling ratio of cationic cellulose and CMC nanofibers reached 642 % and 670%, respectively. This high water uptake ability is due to the chemical structure of these modified nanofibers,
SC RI PT
which are hydrophilic materials and exhibited improved water absorption capacity. CMC revealed the highest swelling among these scaffolds. Indeed, the degree of swelling is influenced by the acidic groups of cellulose, which carboxymethylation is the most extensively used process
TE
D
M
A
N
U
to introduce acidic groups into the cellulose network.
EP
Fig. 6. Water uptake ability of as-electrospun nanofibers
CC
Figure 7a compares the contact angles and surface wettability of prepared nanofibers after contacting with water drop at the final time (5 s). The change in contact angle over time (the
A
initial 5 s) after water drop deposition is illustrated in Fig. 7b. CA nanofibers were hydrophobic with water contact angle of 92o due to the presence of acetyl groups. With alkaline hydrolysis of acetyl groups in CA nanofibers, wettability of obtained cellulose nanofibers is dramatically improved. They became much more hydrophilic, reaching water contact angle of 58o. The differences in the water contact angle of samples can be corresponded to the changes of the 18
surface chemistry and roughness of the electrospun nanofibers during deacetylation. From water contact angle measurements, it was observed that there was no significant difference in the wettability of the cationic cellulose nanofibers and CMC nanofibers and both showed
SC RI PT
hydrophilic properties. The initial contact angle of water measured on cationically modified cellulose nanofibers was 47o; while for CMC nanofibers, it was 45o. The contact angle measurement results confirmed that when electrospun nanofibrous samples were modified, they
CC
EP
TE
D
M
A
N
U
became much more hydrophilic.
A
Fig. 7. Water contact angle measurements for samples (a) at final time, (b) over time
The mechanical characteristics in terms of the tensile strength and Young’s modulus were investigated and presented in Fig.8. When comparing mechanical properties of the nanofibers, it was obvious that surface modification improved the tensile strength and Young’s modulus of the scaffolds. A similar increasing trend was found for tensile strength of CMC and cationic 19
cellulose nanofibers, but Young’s modulus of CMC is significantly higher than those of produced from cationic cellulose nanofibers. It can be seen that the cationic cellulose and CMC nanofibers had tensile strength of 7 and 7.2 MPa, and Young’s modulus of 482 and 515 MPa,
SC RI PT
respectively. The mechanical properties exhibited direct correlation with interaction between the nanofibers. Obviously, cationization and anionization of nanofibers plays the main role in improving mechanical properties of scaffolds compared to unmodified nanofibers. In addition, tensile strength and Young’s modulus were enhanced via the heat and alkaline treatment process, consequently, dispersion and the structural integrity of nanofibers and the mechanical properties
U
were dramatically improved (Ma, Kotaki, & Ramakrishna, 2005). The improvement in tensile
N
strength was mainly due to cross-linking between the nanofibers after heat treatment led to
A
bonding between nanofibers at the crossover points. The improvement of mechanical properties
M
of modified nanofibers may have corresponded to the formation of rigid hydrogen bonds network between nanofibers and electrostatic interaction between introduced groups that cause a
D
good stress transfer from network to nanofibers (Alemdar & Sain, 2008a,b). It seems that
A
CC
EP
TE
interfiber hydrogen bonding and network structure result in improved mechanical strength.
20
SC RI PT U N A M D TE EP
CC
Fig. 8. Mechanical properties of nanofibers: (a) Tensile strength (MPa) and (b) Young’s modulus (MPa) of prepared nanofibers
A
3.3. Cellular proliferation, viability and cytotoxicity studies The biocompatibility of the nanofibrous scaffolds was investigated by evaluating cellular adhesion and viability of hMSCs cultured on the scaffolds. hMSCs were cultured at 37 °C in a humidified atmosphere (5% CO2). Then, cells were seeded on the nanofibers and cultured for 5
21
days. The standard test method of MTT assay was performed to quantitatively evaluate cell viability, proliferation and cytotoxicity onto the fibrous scaffolds. The colorimetric MTT assay, in fact measures the reduction of the tetrazolium molecule of MTT by viable cells to blue
SC RI PT
formazan product which reflects the normal function of mitochondria (He et al., 2013). In hence, the level of the reduction of MTT into formazan component can reveal the level of cellular metabolism. Fig. 9 illustrates the results of MTT assay after 1, 3 and 5 days of cell culture on the nanofibers in comparison to control. The optical densities (OD) of samples and control were measured at a wavelength of 570 nm by a microplate reader.
U
According to p values there is no significant difference between the control, CA and cellulose in
N
day 5, 3, 1 (p>0.05), only QC sample shows the significant increase in day 1, 3, 5, comparing to
A
control (p<0.001), for CMC there is no difference comparing to control in day 1, 5 (p>0.05), but
M
the increase of cell viability in day 3 was significant (p<0.05). As can be seen from Fig. 9, all nanofibers showed better cell proliferation after 5 day of cell culture compared to days of 1 and
D
3. It is clear that cells cultured onto the modified nanofibers scaffolds have the best viability at
TE
all the time points. The results of cell proliferation assay showed high viability for modified
EP
samples in comparison to control. Estimation of cellular growth using MTT assay indicated the modified nanofibers cause more increment in cell proliferation after 5 days. This is likely due to
CC
the fact that hydrophilicity and biocompatibility of modified nanofibrous scaffolds. Successful proliferation of the hMSCs on produced nanofibers describes potential applicability of the
A
scaffold for cell growth. The MTT assay tests showed that the scaffold did not produce any toxic effect on the hMSCs, proving their potential to support cell proliferation and utility for tissue engineering applications. Besides, the MTT assay results indicate that QC nanofibers are cell biocompatible and nontoxic to hMSCs with higher growth and proliferation ability.
22
SC RI PT U N A
M
Fig. 9. MTT assay for cell viability and proliferation of hMSCs on scaffolds and control after in vitro culture for days of 1, 3 and 5
D
3.4. Cell adhesion studies using FESEM
TE
The physicochemical properties and surface changes of nanofibers significantly influence the
EP
cell behavior, adhesion, proliferation and morphology (Stevens & George, 2005). FESEM images were studied to understand cell adhesion, proliferation and morphology on the scaffolds.
CC
As can be observed from the FESEM images (Fig. 10), the hMSCs attached and spread on the surface of scaffolds after culture. After incubation, attached cells start spreading on the scaffolds
A
and then cells spreading became more noticeable. The cells were observed in a flat, elongated and rounded morphology with monolayer like structure on the scaffold surface and a good cell adhesion to the scaffold was obvious from cell surface. The cell adhesion and proliferation are significantly influenced by the hydrophilicity of the scaffolds (Altankov & Groth, 1994). Moreover, the nanofibers surface roughness created by the presence functional groups could 23
promote cell adhesion to the nanofibers since cells generally display enhanced proliferation and bioactivity on rough surfaces (Kunzler, Drobek, Schuler, & Spencer, 2007). The results indicate that the reported nanofibrous scaffolds support cell adhesion/attachment and proliferation and
SC RI PT
hence are suitable for tissue engineering applications. The hydrophilic nature of modified
EP
TE
D
M
A
N
U
nanofibers promotes uniform adhesion of cells on the surface nanofibers.
nanofibers (c) CMC nanofibers and (d) QC nanofibers
A
CC
Fig. 10. FESEM images of hMSCs adhered on the surfaces of (a) CA nanofibers, (b) cellulose
4. Conclusions The novel, simple and inexpensive methods were reported to produce high performance advanced cellulosic materials with a wide range of functionalities makes them suitable for 24
valuable applications. The purpose of this study is to improve an effective and eco-friendly procedure in aqueous suspension for the surface modification of electrospun cellulose nanofibers using CHPTAC and MCAA. The FESEM images of CA nanofibers indicated narrow
SC RI PT
distribution, uniform structures and smooth morphology. After chemical modification no significant changes in shape or morphology were observed except for surface roughness and irregularities. The zeta potentials of CA and cellulose nanofibers at pH 7 were slightly negative and almost zero, respectively, indicating the deacetylation only resulted in a slight enhancement in the zeta potential compared to starting material. Quaternization produced a positive zeta
U
potential due to an increase in the surface charge and carboxymethylation created the most
N
negative zeta potential due to carboxyl group on the surfaces. Among the different samples, CA
A
(hydrophobic) showed the lowest swelling, because of the lower availability of hydroxyl group
M
to form hydrogen bonding resulted in preventing the interaction of the cellulose chains with water. When cellulose nanofibers were modified, the hydrophilicity was increased, the contact
D
angle was decreased with time, and the surface swelling was improved. It was revealed from
TE
MTT assay that the prepared nanofibers did not produce any cytotoxic effect on hMSCs.
EP
Furthermore, the modified nanofibers presented high cell viability. The cell adhesion is influenced by the surface chemistry of the nanofibers. When hMSCs were cultured on the
CC
surface of the modified nanofibers, favorable cell adhesion was observed owing to the hydrophilicity of surface. The findings showed that cell adhesion is strongly enhanced on the
A
modified surfaces with carboxymethyl and quaternary ammonium groups. The studies showed biocompatibility and non-cytotoxicity of nanofibers and so these nanofibers are favorable for cell adhesion and proliferation and hence are promising materials for biomedical applications. The results indicated that the produced scaffolds support cell adhesion and proliferation and hence
25
they are suitable for tissue engineering applications. We believe the use of cellulose derivatives to produce nanofibers using the electrospinning process presents an excessive opportunity for better application of cellulose nanofibers and development of novel presentations.
SC RI PT
Acknowledgments The authors are gratefully acknowledged the National Elites Foundation, Iran University of Medical Sciences and Sharif University of Technology, for their assistance and financial support. References
Abdul Khalil, H. P. S., Bhat, A. H., Ireana Yusra, A. F. (2012). Green composites from
U
sustainable cellulose nanofibrils: A review. Carbohydrate Polymers, 87, 963–979.
N
Alemdar, A., Sain, M. (2008a). Biocomposites from wheat straw nanofibers: Morphology,
A
thermal and mechanical properties. Composite Science and Technology, 68(2), 557–565.
M
Alemdar, A., Sain, M. (2008b). Isolation and characterization of nanofibers from agricultural residues–Wheat straw and soy hulls. Bioresource and Technology, 99(6), 1664–1671.
D
Altankov, G., Groth, T. (1994). Reorganization of substratum-bound fibronectin on hydrophilic
TE
and hydrophobic materials is related to biocompatibility. Journal of Materials Science:
EP
Materials in Medicine. 5(9-10), 732–737. Ambjörnsson, H. A., Schenzel, K., Germgård, U. (2013). Carboxymethyl cellulose produced at
CC
different mercerization conditions and characterized by NIR FT Raman spectroscopy in combination with multivariate analytical methods. BioResources, 8(2), 1918–1932.
A
Antczak, T. (2012). Nanotechnology-methods of manufacturing cellulose nanofibers, Fibres & Textiles in Eastern Europe, 20 (2), 91.
26
Bacakova, L., Novotna, K., Sopuch, T., Havelk, P. (2015). Cell interaction with cellulose-based scaffolds for tissue engineering: a review. In: Cellulose and Cellulose Composites Editor: Md. Ibrahim H. Mondal, Nova Science Publishers, Inc. 341–376.
SC RI PT
Chen, P., Yu, H., Liu, Y., Chen, W., Wang, X., Ouyang, M. (2013). Concentration effects on the isolation and dynamic rheological behavior of cellulose nanofibers via ultrasonic processing, Cellulose 20, 149–157.
Deepa, B., Abraham, E., Cherian, B. M., Bismarck, A., Blaker, J. J., Pothan, L. A., Leao, A. L., de Souza, S. F., Kottaisamy, M. (2011). Structure, morphology and thermal characteristics of
U
banana nano fibers obtained by steam explosion, Bioresource Technology, 102, 1988–1997.
N
Fei, P., Liao, L., Meng, J., Cheng, B., Hu, X., Song, J. (2018). Non-leaching antibacterial
A
cellulose triacetate reverse osmosis membrane via covalent immobilization of quaternary
M
ammonium cations. Carbohydrate polymers, 181, 1102–1111. Frey, M. (2008). Electrospinning cellulose and cellulose derivatives, Polymer Reviews, 48 378–
D
391.
TE
Hasani, M., Cranston, E. D., Westman, G., Gray, D. G. (2008). Cationic surface
EP
functionalization of cellulose nanocrystals. Soft Matter, 4(11), 2238–2244. Hashem, M. M. (2006). Development of a one‐stage process for pretreatment and cationisation
CC
of cotton fabric. Coloration technology, 122(3), 135–144.
He, X., Xiao, Q., Lu, C., Wang, Y., Zhang, X., Zhao, J., Zhang, W., Zhang, X., Deng, Y. (2014).
A
Uniaxially Aligned Electrospun All-Cellulose Nanocomposite Nanofibers Reinforced with Cellulose Nanocrystals: Scaffold for Tissue Engineering. Biomacromolecules 15 (2), 618– 627.
27
Hebeish, A. A., El-Rafie, M. H., Abdel-Mohdy, F. A., Abdel-Halim, E. S., Emam, H. E. (2010). Carboxymethyl cellulose for green synthesis and stabilization of silver nanoparticles, Carbohydrate Polymers, 82, 933–941.
SC RI PT
Hong, K. M. (2013). Preparation and characterization of Carboxymethyl cellulose from sugarcane bagasse, Universiti Tunku Abdul Rahman.
Huang, Y. X., Ren, J., Chen, C., Ren, T. B., Zhou, X. Y. (2008). Preparation and properties of poly(lactide-co-glycolide) (PLGA)/nano-hydroxyapatite (NHA) scaffolds by thermally induced phase separation and rabbit MSCs culture on scaffolds. Journal of Biomaterial
U
Applied Science, 22, 409–432.
N
Huang, Y., Yu, H., Xiao, C. (2007). pH-sensitive cationic guar gum/poly (acrylic acid)
A
polyelectrolyte hydrogels: swelling and in vitro drug release. Carbohydrate Polymers, 69,
M
774–783.
Inukai, S., Kurokawa, N., & Hotta, A. (2018). Annealing and saponification of electrospun
D
cellulose-acetate nanofibers used as reinforcement materials for composites. Composites Part
TE
A: Applied Science and Manufacturing, 113, 158-165.
EP
Khatri, Z., Mayakrishnan, G., Hirata, Y., Wei, K., Kim, I. (2013). Cationic-cellulose nanofibers: Preparation and dyeability with anionic reactive dyes for apparel application. Carbohydrate
CC
polymers 91 (1), 434–443. Kunzler, T. P., Drobek, T., Schuler, M., Spencer, N. D. (2007). Systematic study of osteoblast
A
and fibroblast response to roughness by means of surface-morphology gradients. Biomaterials 28, 2175–2182.
Ma, Z., Kotaki, M., Inai, R., Ramakrishna, S. (2005). Potential of nanofiber matrix as tissue engineering scaffolds. Tissue engineering, 11, 101.
28
Ma, Z., Kotaki, M., Ramakrishna, S. (2005). Electrospun cellulose nanofber as afnity membrane. Journal of Membrane Science, 265 (1-2), 115–123, Menon, M. P., Selvakumar, R., Ramakrishna, S. (2017). Extraction and modification of cellulose
SC RI PT
nanofibers derived from biomass for environmental application, RSC Advances, 7(68), 42750–42773.
Pelton, R. (2009). Bioactive paper provides a low-cost platform for diagnostics, TrAC Trends in Analytical Chemistry, 28 925–942.
Rezaei, A., Nasirpour, A., Fathi, M. (2015). Application of cellulosic nanofibers in food science
U
using electrospinning and its potential risk. Comprehensive Reviews in Food Science and
N
Food Safety, 14(3), 269–284.
A
Siró, I., Plackett, D. (2010). Microfibrillated cellulose and new nanocomposite materials: a
M
review, Cellulose 17, 459–494.
Science, 310, 1135–1138.
D
Stevens, M. M., George, J. H. (2005). Exploring and engineering the cell surface interface.
TE
Song, Y., Sun, Y., Zhang, X., Zhou, J., Zhang, L. (2008). Homogeneous quaternization of
2264.
EP
cellulose in NaOH/urea aqueous solutions as gene carriers. Biomacromolecules 9(8), 2259–
CC
Song, Y., Wang, H., Zeng, X., Sun, Y., Zhang, X., Zhou, J., Zhang, L. (2010). Effect of molecular weight and degree of substitution of quaternized cellulose on the efficiency of
A
gene transfection, Bioconjugate Chemistry, 21, 1271–1279.
Song, Y., Zhang, L., Gan, W., Zhou, (2011). Self-assembled micelles based on hydrophobically modified quaternized cellulose for drug delivery, Colloids and Surfaces B: Biointerfaces, 83, 313–320.
29
Song, Y., Zhou, J., Li, Q., Guo, Y., Zhang, L. (2009). Preparation and characterization of novel quaternized cellulose nanoparticles as protein carriers. Macromolecular Bioscience, 9, 857– 863.
SC RI PT
Song, Y., Zhou, Y., Chen, L. (2012). Wood cellulose-based polyelectrolyte complex nanoparticles as protein carriers. Journal of Materials Chemistry, 22, 2512–2519.
Teeri, T. T., Brumer, H., Daniel, G., Gatenholm, P. (2007). Biomimetic engineering of cellulosebased materials. Trends in Biotechnology, 25, 299–306.
Yamamoto, S., Iwata, T., Nishiwaki, K., Kinoshita, M., & Suzuki, S. (2015). Synthesis of
N
Electrophoresis. Chromatography, 36(3), 93-98.
U
Quaternary Ammonium Derivatives of Cellulose as the Coating Reagents for Capillary
A
Yang, W. J., Fu, J., Wang, D. X., Wang, T., Wang, H., Jin, S. G., He, N. Y. (2010). Study on
M
chitosan/polycaprolactone blending vascular scaffolds by electrospinning. Journal of Biomedical Nanotechnology, 6, 254.
D
Yu, S., Cheng, Q., Huang, C., Liu, J., Peng, X., Liu, M., & Gao, C. (2013). Cellulose acetate
TE
hollow fiber nanofiltration membrane with improved permselectivity prepared through
EP
hydrolysis followed by carboxymethylation. Journal of membrane science, 434, 44-54. Zahedi, P., Rezaeian, I., Ranaei-Siadat, S., Jafari, S., Supaphol, P. (2009). A review on wound
CC
dressings with an emphasis on electrospun nanofibrous polymeric bandages, Polymers for Advanced Technologies, 21, 77–95.
A
Zarth, C. S. P. (2013). Novel cationic esters derived from cellulose and dextran: synthesis, structure analysis and properties. LAP LAMBERT Academic Publishing.
Zhou, Q., Rutland, M., Teeri, T., Brumer, H. (2007). Xyloglucan in cellulose modification. Cellulose, 14, 625–641.
30