Fabrication of Electrospun Double Layered Biomimetic Collagen–Chitosan Polymeric Membranes with Zinc-Doped Mesoporous Bioactive Glass Additives

Abstract

Several therapeutic approaches have been developed to promote bone regeneration, including guided bone regeneration (GBR), where barrier membranes play a crucial role in segregating soft tissue and facilitating bone growth. This study emphasizes the importance of considering specific tissue requirements in the design of materials for tissue regeneration, with a focus on the development of a double-layered membrane to mimic both soft and hard tissues within the context of GBR. The hard tissue-facing layer comprises collagen and zinc-doped bioactive glass to support bone tissue regeneration, while the soft tissue-facing layer combines collagen and chitosan. The electrospinning technique was employed to achieve the production of nanofibers resembling extracellular matrix fibers. The production of nano-sized (~116 nm) bioactive glasses was achieved by microemulsion assisted sol-gel method. The bioactive glass-containing layers developed hydroxyapatite on their surfaces starting from the first week of simulated body fluid (SBF) immersion, demonstrating that the membranes possessed favorable bioactivity properties. Moreover, all membranes exhibited distinct degradation behaviors in various mediums. However, weight loss exceeding 50% was observed in all tested samples after four weeks in both SBF and phosphate-buffered saline (PBS). The double-layered membranes were also subjected to mechanical testing, revealing a tensile strength of approximately 4 MPa. The double-layered membranes containing zinc-doped bioactive glass demonstrated cell viability of over 70% across all tested concentrations (0.2, 0.1, and 0.02 g/mL), confirming the excellent biocompatibility of the membranes. The fabricated polymer bioactive glass composite double-layered membranes are strong candidates with the potential to be utilized in tissue engineering applications.

Full text

Citation: Altan, D.; Özarslan, A.C.;
Özel, C.; Tuzlako˘glu, K.; Sahin, Y.M.;
Yücel, S. Fabrication of Electrospun
Double Layered Biomimetic
Collagen–Chitosan Polymeric
Membranes with Zinc-Doped
Mesoporous Bioactive Glass
Additives. Polymers 2024,16, 2066.
https://doi.org/10.3390/
polym16142066
Academic Editor: Raffaella Striani
Received: 8 June 2024
Revised: 12 July 2024
Accepted: 14 July 2024
Published: 19 July 2024
Copyright: © 2024 by the authors.
Licensee MDPI, Basel, Switzerland.
This article is an open access article
distributed under the terms and
conditions of the Creative Commons
Attribution (CC BY) license (https://
creativecommons.org/licenses/by/
4.0/).
polymers
Article
Fabrication of Electrospun Double Layered Biomimetic
Collagen–Chitosan Polymeric Membranes with Zinc-Doped
Mesoporous Bioactive Glass Additives
Dilan Altan 1,2,* , Ali Can Özarslan 1,2 , Cem Özel 1,2 , Kadriye Tuzlako˘glu 3, Yesim Muge Sahin 4,5 and
Sevil Yücel 1,2
1Faculty of Chemical and Metallurgical Engineering, Department of Bioengineering, Yildiz Technical
University, 34220 Istanbul, Türkiye; alicanozarslan@gmail.com (A.C.Ö.); cemozel@yildiz.edu.tr (C.Ö.);
yuce.sevil@gmail.com (S.Y.)
2Health Biotechnology Joint Research and Application Center of Excellence, 34903 Istanbul, Türkiye
3Department of Polymer Engineering, Yalova University, 77200 Yalova, Türkiye; ktuzlakoglu@yalova.edu.tr
4Polymer Technologies and Composite Application and Research Center, Istanbul Arel University,
34537 Istanbul, Türkiye; ymugesahin@arel.edu.tr
5Faculty of Engineering, Department of Biomedical Engineering, Istanbul Arel University,
34537 Istanbul, Türkiye
*Correspondence: dylanaltan@gmail.com
Abstract: Several therapeutic approaches have been developed to promote bone regeneration, includ-
ing guided bone regeneration (GBR), where barrier membranes play a crucial role in segregating soft
tissue and facilitating bone growth. This study emphasizes the importance of considering specific
tissue requirements in the design of materials for tissue regeneration, with a focus on the development
of a double-layered membrane to mimic both soft and hard tissues within the context of GBR. The
hard tissue-facing layer comprises collagen and zinc-doped bioactive glass to support bone tissue
regeneration, while the soft tissue-facing layer combines collagen and chitosan. The electrospinning
technique was employed to achieve the production of nanofibers resembling extracellular matrix
fibers. The production of nano-sized (~116 nm) bioactive glasses was achieved by microemulsion
assisted sol-gel method. The bioactive glass-containing layers developed hydroxyapatite on their
surfaces starting from the first week of simulated body fluid (SBF) immersion, demonstrating that the
membranes possessed favorable bioactivity properties. Moreover, all membranes exhibited distinct
degradation behaviors in various mediums. However, weight loss exceeding 50% was observed in all
tested samples after four weeks in both SBF and phosphate-buffered saline (PBS). The double-layered
membranes were also subjected to mechanical testing, revealing a tensile strength of approximately
4 MPa. The double-layered membranes containing zinc-doped bioactive glass demonstrated cell
viability of over 70% across all tested concentrations (0.2, 0.1, and 0.02 g/mL), confirming the excellent
biocompatibility of the membranes. The fabricated polymer bioactive glass composite double-layered
membranes are strong candidates with the potential to be utilized in tissue engineering applications.
Keywords: collagen; chitosan; bioactive glass; electrospinning; guided bone regeneration; zinc
1. Introduction
Guided bone regeneration (GBR) has become a standard in clinics for treating jawbone
defects by employing membranes to prevent soft tissue from infiltrating bone defects [
1
].
This technique facilitates bone tissue growth, particularly in cases where there is not
sufficient volume for dental implants, by creating an isolated area for bone regeneration.
Gore-Tex, a non-degradable material based on expanded polytetrafluoroethylene (e-PTFE)
fluoropolymer, has been extensively utilized as a GBR membrane. Despite its advantageous
properties such as biocompatibility, ability to create space, and stability, the need for a
second surgery to remove the material after recovery has been a significant drawback
Polymers 2024,16, 2066. https://doi.org/10.3390/polym16142066 https://www.mdpi.com/journal/polymers

Polymers 2024,16, 2066 2 of 18
associated with this class of materials. Another type of non-degradable material used
as a GBR membrane is titanium mesh [
2
]. However, despite their superior mechanical
properties like e-PTFE, titanium meshes also need to be removed after recovery. The
removal procedure can increase the risk of morbidity and is burdensome for the patient [
3
].
Another disadvantage associated with non-degradable membranes has been reported as
recurrent infections occurring during membrane use, which may adversely affect new bone
formation [4].
Synthetic and natural biodegradable polymers are being increasingly used as mem-
brane materials in clinical practice to prevent the necessity of a second surgery with
non-degradable membranes. The gradual degradation of degradable membranes allows
for the development of new bone in areas of defects [
5
]. Aliphatic esters like polylactic acid
(PLA), polyglycolic acid (PGA), and polycaprolactone (PCL), as well as their numerous
combinations, might be noted as synthetic biodegradable polymers used for tissue engineer-
ing purposes. Although these materials demonstrate satisfactory mechanical properties,
their usage is limited by the acidic degradation products they produce [
6
]. These products
can lead to inflammation in adjacent tissues due to the enzymatic activities involved in the
process [
7
]. Natural biodegradable polymers like collagen [
8
], gelatin [
9
], chitosan [
10
], silk
fibroin [
11
], and chitin [
12
] are frequently utilized in the fabrication of GBR membranes.
Collagen, one of the constituents of the extracellular matrix, stands out as one of the most
used natural polymers in membrane fabrication, owing to its excellent cell affinity and
biocompatibility. One of the most appealing characteristics of collagen for bone tissue
engineering is its structural integrity that allows the accumulation of calcium phosphate
and calcium carbonate minerals [
13
]. Chitosan is an important naturally occurring poly-
mer with a polysaccharide structure that is produced by deacetylating chitin. Chitosan’s
hemostatic, antimicrobial, biodegradable, and biocompatible properties have led to its
widespread use in various tissue engineering applications [
14
,
15
]. In addition to its afore-
mentioned advantageous properties, chitosan’s structural similarity to glycosaminoglycan,
the space filling component of the natural extracellular matrix (ECM) [
16
], suggests that the
collagen–chitosan complex used in scaffold construction for tissue engineering is expected
to mimic the constituents of the native ECM.
Introducing or enhancing bioactive properties can be achieved by incorporating bioac-
tive compounds into the membrane structure during fabrication of membranes. Bioactive
glasses are widely utilized in tissue engineering applications for their unique properties.
This category of materials, also recognized as biodegradable glass materials, is extensively
researched. They demonstrate bioactivity by facilitating the formation of apatite on their
surface [
17
]. The bone-stimulating properties of bioactive glasses can be enhanced with
various elemental additions, such as magnesium (Mg), zinc (Zn), strontium (Sr), and silver
(Ag) [18].
In this study, zinc-doped bioactive glasses produced by the microemulsion assisted
sol-gel method were added to membranes to enhance their bioactivity and stimulate bone
development. Bioactive glasses synthesized through the microemulsion technique exhibit
reduced susceptibility to agglomeration in contrast to other bioactive glasses, owing to their
high surface areas [
19
,
20
], which is an important feature for uniform distribution within
the membrane structure during production, allowing the membrane to exhibit similar
properties and bioactivity throughout. Although found in small amounts in the body
(75–125
µ
g/dL in serum), zinc performs various functions related to the immune system,
cell division, fertility, and growth, and is necessary for the activity of over 300 enzymes [
21
].
Zinc has been proven to be an essential element for the formation, mineralization, devel-
opment, and maintenance of healthy bones [
22
]. There are studies showing zinc’s (Zn)
beneficial effects on bone formation in both
in vivo
and
in vitro
settings, and an osteoclast
impairing effect of zinc has been shown [23].
Various techniques such as solvent casting/particle leaching, phase separation, and
electrospinning are utilized in the production of dental membranes. However, electrospin-
ning stands out as a simple, cost-effective, and efficient technique due to its applicability to

Polymers 2024,16, 2066 3 of 18
both natural and synthetic polymers. This technique enables the production of nanofibrous
structures with dimensions and high surface area resembling those of the natural ECM
(5 to 500 nm) [
24
]. Electrospinning enables the fabrication of polymer structures with
different morphologies by utilizing an electric field. During this procedure, a solution
contained within a syringe is propelled along a metal needle linked to a high-voltage
power source. Following solvent evaporation, the resultant nanofibers are deposited on a
grounded collector surface in the form of nonwoven mats or membranes [25].
Mechanical properties and stability of nonwoven membranes can be increased with
crosslinking. There several techniques such as chemical EDC (1-Ethyl-3-(3-dimethylaminopropyl)
carbodiimide), NHS (N-hydroxy succinimide), glutaraldehyde (GA), biological (Genipin), phys-
ical (ultraviolet irradiation (UV), and dehydrothermal (DHT)) used for these purpose [
26
].
Despite concerns about the toxicity of GA, its low cost, extensive crosslinking capabilities, and
short time required for crosslinking make it one of the most effective methods available [
27
].
The toxicity of crosslinked membranes can be reduced by using vapor crosslinking techniques,
and post-treatments such as high pressure and rinsing with glycine [28].
In tissue engineering, it is crucial to utilize materials customized for the specific
requirements of each tissue. Therefore, a method involving the production of layered
membranes is available to address the unique needs of each tissue. This study focuses
on the production of double-layered collagen-based membranes with an electrospinning
technique to be used as GBR membranes. The bone-facing layer of the material is designed
to mimic bone tissue by incorporating nanosized bioactive glass into the collagen fiber
structure. This approach is inspired by the composite nature of bone, which consists of
protein (collagen fibers) and mineral (hydroxyapatite), and the soft tissue-facing layer also
aims to mimic the structure of the soft tissue, specifically targeting the protein (collagen)
and glucosamine glucan (chitosan) composition by combining collagen and chitosan. In this
study, the physiochemical and structural characteristics of double-layered membranes were
comprehensively examined. The impact of incorporating zinc-doped bioactive glass on the
bioactivity and biocompatibility behaviors of these membranes was assessed and discussed.
2. Materials and Methods
2.1. Materials
The reagents used in glass synthesis, including tetraethyl orthosilicate (TEOS), cal-
cium nitrate tetrahydrate (Ca(NO
3
)
2·
4H
2
O), nitric acid (HNO
3
), zinc nitrate hexahydrate
Zn(NO
3
)
2·
6H
2
O, hexadecyltrimethylammonium bromide (CTAB), and ethyl acetate, were
obtained from Sigma Aldrich. The solvent for electrospinning, 1,1,1,3,3,3-hexafluoro-2-
propanol (HFIP), was purchased from Sigma Aldrich (Steinheim, Germany); Trifluoroacetic
acid (TFA) and acetic acid were obtained from Merck. Glutaraldehyde (GA) (25% in H
2
O)
utilized for membrane crosslinking, glycine for removing unreacted GA residues, Dul-
becco modified Eagle medium (DMEM), fetal bovine serum (FBS), phosphate buffer saline
(PBS) solution, and penicillin–streptomycin used for MTT tests, were purchased from
Sigma Aldrich.
2.2. Production of Zinc-Doped Mesoporous Bioactive Glasses
Microemulsion assisted sol-gel method was employed to synthesize mesoporous
bioactive glasses nanoparticles (MBGN) based on a binary system comprising 70 mol%
SiO
2
and 30 mol% CaO. In order to incorporate additional biological functionality, Zn
2+
ions
were introduced as dopants, resulting in the synthesis of MBGN_Zn with a composition
of 70 mol% SiO
2
, 25 mol% CaO, and 5 mol% ZnO, following a previously established
protocol by the authors [
29
]. The production of nanosized bioactive glasses included
dissolving hexadecyltrimethylammonium bromide (CTAB) in deionized water and adding
ethyl acetate. Following the hydrolysis of TEOS, appropriate amounts of other precursors
including Ca(NO
3
)
2·
4H
2
O and Zn(NO
3
)
2·
6H
2
O were introduced to the mixture. After
synthesis, the resulting precipitate was washed, collected, and subjected to drying in an

Polymers 2024,16, 2066 4 of 18
oven at 60 ◦C for 24 h. Finally, the dried materials were calcinated at 700 ◦C for 2 h with a
heating rate of 2 ◦C/min.
2.3. Electrospinning and Crosslinking of Collagen Based Membranes
Collagen Type I was used to prepare collagen fibers and was obtained from the tails of
rats that were sacrificed after being used as a healthy control group in other studies. Briefly,
rat tails were soaked in a 70% ethanol solution for 30 min to eliminate any contaminants
and then immersed in a 1
×
PBS solution. After scraping the skin of the tails taken from
the PBS solution, the tendons that became prominent and rich in collagen content were
carefully separated from the tail with the help of pliers and placed in a pre-prepared 0.5 M
acetic acid solution to dissolve the collagen at +4
◦
C. The solution was filtered with the
help of multilayer gauze, dialyzed against distilled water with pH adjusted to 3 at +4
◦
C
for 3 days, then lyophilized and stored at +4 ◦C [30].
To produce the soft tissue facing layer; Type 1 collagen was dissolved in HFIP with 8%
(w/v) concentration. Selecting the appropriate solvent/solvent systems can be difficult, es-
pecially when working with natural polymers. Since the solubility of chitosan in HFIP was
low, a solvent mixture was prepared with a volume ratio of HFIP/TFA (80:20), as suggested
in the literature [
31
]. Simultaneously, chitosan was dissolved in the mixture. After the solu-
tions became homogeneous, the collagen and chitosan solutions were mixed in the ratios of
90:10, 80:20, 70:30 to optimize Col-Chi ratio aiming for homogenous nanofiber production.
Bone tissue facing layer was produced as follows: Type 1 collagen was dissolved in
HFIP at 8 wt.% [
32
]. Then, bioactive glass was added at rates of 5, 10, and 15 wt.% and
optimization studies were carried out to produce membranes consisting of homogeneous
and nano-sized fibers in which bioactive glasses were homogeneously distributed in the
fiber structure.
Through systematic optimization of each layer, compositions were identified that
exhibited the most uniform distribution of fibers and achieved the smallest fiber diameters
deemed essential to produce double-layered (DL) membranes. Notably, the compositions
Col-Chi (70:30) and Col-BG (90:10) were determined to be optimal for the fabrication of
double-layered membranes. During the double-layered membrane fabrication process, the
Col-BG layer was initially deposited onto the collector surface, followed by deposition of
the Col-Chi layer atop it (Graphical Abstract).
The membranes were produced using the Fytronix ES-9000 (Elazı˘g, Türkiye). The flow
rate during the electrospinning process was 1.0 mL/h and the distance from the needle to
the aluminum foil collector was 10–15 cm and the voltage was between 15–30 kV while
the rotating speed was 200–400 rpm. Relative humidity and temperature ranged from
20% to 30% and 20 to 25
◦
C, respectively. All electrospun nanofibrous membranes were
subjected to a vacuum oven at room temperature for one week to eliminate any possible
solvent residue. The membranes were crosslinked with 25% GA vapor in a desiccator. The
membranes were then stored in a vacuum oven at room temperature to eliminate extra
GA after being crosslinked for 24 h and subsequently rinsed with glycine to eliminate any
remaining residues of GA [33].
2.4. Characterization
2.4.1. Characterization of Bioactive Glasses and Membranes
Elemental Composition Analysis by X-ray Fluorescence Spectroscopy (XRF)
The elemental composition of bioactive glass nanoparticles was measured at am-
bient temperature by using energy dispersive X-ray fluorescence spectroscopy (EDXRF,
PANalytical Minipal, Almelo, The Netherlands).
Surface Morphology, Particle Size, and Fiber Diameter Analysis by Scanning Electron
Microscopy (SEM)
Surface morphology and particle size of the bioactive glass particles were examined
by using scanning electron microscopy (SEM, Zeiss EVO LS10, Carl Zeiss, Oberkochen,

Polymers 2024,16, 2066 5 of 18
Germany). SEM was also used to determine the fiber diameters and surface morphology of
the electrospun membranes. The samples were sputter coated with a conductive gold layer
before SEM analysis. The particle sizes of bioactive glasses were measured using ImageJ
Software Version 1.8.0 (http://imagej.nih.gov/ij/; provided in the public domain by the
National Institutes of Health, Bethesda, MD, USA) from 100 randomly chosen particles
from different SEM images. Average fiber diameters and pore sizes were determined
using ImageJ by evaluating at least 100 randomly chosen fibers/pores in SEM images.
Energy dispersive X-ray spectroscopy (EDS; Carl Zeiss, SmartEDX, Oberkochen, Germany)
analysis was also performed to determine the elemental composition of samples.
Particle Size Analysis by Dynamic Light Scattering (DLS)
The dynamic light scattering (DLS) technique (Malvern Nano ZS, Malvern Instrument
Ltd., Worcestershire, UK) was used to determine the zeta potential, particle size, and size
distribution of the bioactive glass particles. Particle size measuring was performed in water
(0.05 mg/mL) at 25
◦
C and the suspensions were sonicated for 10 min before the measurements.
Identification of Functional Groups by Fourier Transform Infrared Spectroscopy (FTIR)
The functional groups in the structure of both the bioactive glasses and the electrospun
membranes before and after SBF exposure were identified utilizing Fourier transform
infrared spectroscopy (FTIR; JASCO 6600,JASCO Ltd., Tokyo, Japan) in the wavenumber
range of 2000 to 500 cm
−1
and the number of scans was 15. Both the bioactive glasses
and membranes were placed directly onto the attenuated total reflectance (ATR) crystal
in small amounts (a few milligrams each). The crystal was cleaned thoroughly between
each measurement.
Surface Area and Pore Characteristics Analysis by Nitrogen Adsorption-Desorption Isotherms
The assessment of the specific surface area and pore characteristics of the bioactive
glasses was conducted via the analysis of N
2
-gas adsorption-desorption isotherms, em-
ploying a Brunauer–Emmett–Teller (BET) method (Micromeritics, TriStar II 3020, Norcross,
GA, USA). The bioactive glass samples were weighed (0.2 g) and placed in measurement
tubes and degassed under vacuum at 300 ◦C for 12 h prior to the adsorption experiments.
Crystallographic Analysis by X-ray Diffraction (XRD)
The X-ray diffraction (XRD) analyses were conducted to acquire the XRD patterns
of both the bioactive glass powders and the membranes before and after simulated body
fluid (SBF) exposure. These analyses were performed utilizing an XRD instrument (Rigaku,
Dmax-2200, Tokyo, Japan) operating in the 2
θ
range of 10
◦
to 60
◦
and employing Cu K
α
radiation. The experimental parameters included a step size of 0.010
◦
and a dwell time of
1◦per minute.
Thermal Analysis by Thermogravimetric-Differential Thermal Analysis (TG-DTA)
Thermogravimetric (TG) analysis of bioactive glasses and membranes were performed
with the Thermogravimetric-Differential Thermal Analyzer (TG-DTA, SIINanotechnology,
SII6000 Exstar 6300, Chiba, Japan). Throughout the heating procedure, nitrogen was passed
through an alumina sample container. The temperature was incrementally raised to 700
◦
C
at a rate of 20 ◦C per minute.
Mechanical Testing of Double-Layered Membranes
The double-layered membranes underwent mechanical testing under both dry and
wet conditions at ambient settings (20
◦
C and 50% humidity) using tensile testing machine
(Devotrans GPUG/R, ˙
Istanbul, Türkiye). The membranes were cut into 10 mm
×
30 mm
rectangular shapes, and measurements were conducted at a loading speed of 5 mm/min
with a 20 N load cell. The tensile stress–elongation curves of the specimens were plotted
using the data recorded by the machine.

Polymers 2024,16, 2066 6 of 18
2.5. In Vitro Degradation of Membranes
The degradation properties of the membranes were assessed by immersing them
in fresh phosphate-buffered saline (PBS) and SBF for four weeks. After being dried and
weighed, the membranes (10 mm
×
10 mm) were placed in falcon tubes containing 10 mL of
PBS and SBF. The membranes were then rinsed with deionized water after each incubation
period and dried before being weighed again. The biodegradability of the membranes was
calculated using the equation [34] below (Equation (1)).
Weight loss (%) = (Wi −Wd/Wi) ×100 (1)
2.6. In Vitro Bioactivity of MBGN_Zn and MBGN_Zn-Incorporated Membranes
The bioactivity of bioactive glass samples was evaluated in SBF using a technique
proposed for powder materials by Maçon et al. [
35
]. The SBF was prepared following
Kokubo’s method. In brief, MBGN_Zn powders were introduced into polyethylene falcon
tubes containing SBF at a concentration of 1.5 mg/mL. Subsequently, the bioactive glasses
were retrieved from the SBF at the end of each time interval (ranging from 1 to 4 weeks),
rinsed with distilled water and acetone to terminate any ongoing reactions, and then dried.
The membrane bioactivity was assessed using the following method: the membranes were
submerged in SBF, utilizing the equation formulated by Kokubo and Takedama for dense
materials (Equation (2)):
Vs = Sa/10 (2)
Here, Vs represents the volume of SBF in milliliters (mL), and Sa denotes the apparent
surface area of the specimen in square millimeters (mm2) [36].
2.7. In Vitro Cell Viability of Membranes
Cell viability (%) of all membranes was assessed using Saos-2 cells, employing the 3-
(4,5-dimethylthiazol-2-yl)-2,5-diphenyltetrazolium (MTT) colorimetric technique following
the ISO-10993-5 standard. Membrane extracts were initially prepared in cell medium at a
concentration of 0.2 g/mL and then diluted to concentrations of 0.1 mg/mL and 0.2 g/mL.
One milliliter of each concentration was incubated in cell medium at 37
◦
C for 24 h. The
extracts were then introduced into 96-well plates containing Saos-2 cells (1 ×104cells per
well) that had been previously incubated for 24 h at 37
◦
C with 5% CO
2
. After incubation
of the cells with the membrane extracts, 50
µ
L of MTT solution was added and incubated
for two hours at 37
◦
C with 5% CO
2
. Subsequently, the wells were rinsed three times with
Dulbecco’s phosphate-buffered saline (DPBS) to remove the MTT solution. Absorbance
values of each well were measured at 570 nm, and the cell viability (%) was calculated using
the formula provided by the ISO 10993-5 standard procedure [
37
]. The averaged findings
were statistically analyzed using one-way analysis of variance (ANOVA) and Tukey’s post
hoc test (p< 0.05).
3. Results and Discussion
3.1. Characterization of MBGN_Zn
The XRF composition of the produced MBGN, expressed in oxide form, is summarized
in Table 1. Molar percentages (%) of each oxide were listed based on XRF elementary
analysis. The deviation of the final composition from the initially calculated values has
been attributed to the washing phases of the synthesis process [
38
]. The presence of Si, Ca,
Zn, and O elements was also confirmed with EDS analyses (Figure 1b).

Polymers 2024,16, 2066 7 of 18
Table 1. Theoretical and experimental molar percentages of oxides in the structure of bioactive glass
(MBGN_Zn).
Oxide Theoretical (%) Experimental (%)
SiO270 84.62
CaO 25 11.73
ZnO 5 3.65
As shown in Figure 1a, production of MBGN_Zn particles with spherical morphology
and homogeneous size distribution was obtained via microemulsion assisted sol-gel tech-
nique. Measurements conducted using ImageJ on SEM results confirmed that the produced
bioactive glass had particle sizes ranging from 60 to 160 nm, with an average particle size of
116.26
±
19.67 nm (Figure 1c). The particle size results obtained from Zeta Sizer (Figure 1d)
were slightly larger (~140 nm) than the results obtained from SEM images due to the fact
that in DLS technique, the hydrodynamic diameter is measured, which measures electric
double-layer and the hydration shell in addition to the particle diameter [
39
]. The mi-
croemulsion assisted sol-gel technique provides significant advantages over conventional
sol-gel methods by producing nano-sized bioactive glasses with a narrower particle size
distribution. In contrast, the surfactant-free sol-gel technique typically yields glasses of
micron-sized with similar compositions [40].
Polymers2024,16,xFORPEERREVIEW8of20



Figure1.Characteristicsofbioactiveglass(MBGN_Zn);(a)SEMimage,(b)EDSresult,(c)particle
sizedistributionhistogramaccordingtoSEMimage,(d)particlesizedistributionaccordingtoDLS
analysis.
Theporediameter,specificsurfaceareaandtotalporevolumeweremeasuredtobe
5.2nm,368m
2
/g,and0.5cm
3
/g,respectively.Besidesconfirmingwiththeporediameters
between2–50nm,themesoporousnatureofthebioactiveglasseswasalsoindicatedwith
theN
2
adsorption–desorptionisothermsofsamples(Figure2b).AccordingtotheInterna-
tionalUnionofPureandAppliedChemistry(IUPAC)classification,TypeIVisotherms
signifymesoporousmaterialswithnon-uniformporesizedistribution,displayinghyste-
resisloopsduetocapillarycondensationinporesofvaryingsizes.FTIRspectraofbioac-
tiveglass(Figure2a)aftercalcinationat700°Cfor2hhaveexhibitedastrongabsorption
peakat1044cm
−1
whichisaributedto

Si–O–Siasymmetricalstretchingvibrations[41].
Additionally,aSi–O–Siwithbendingscissoringvibrationscouldbeobservedatabout800
cm
−1
whichisaributedtoringstructuresintheglassmatrix[42].TheXRDpaernof
bioactiveglassesisshowninFigure2d;nodiffractionpeakswereobservedinthegraph,
confirmingthatbioactiveglassesareamorphousinform,likeglass,andthebroadband
intherangeof15°–40°canbeaributedtotheamorphoussilicate[29,43].Thedrastic
decreaseinweightobservedintheTGAgraphofbioactiveglasssamplesbetween20–90
°C(Figure2c)hasbeenaributedtotheremovalofphysicallyadsorbedwaterandresi-
duesfromthebioactiveglassproductionprocess[44].Subsequently,thegradualreduc-
tioninweightobservedbetweentemperaturesof100–700°Cisaributedtotheremoval
ofinternalwatermoleculesduetotheslowcondensationofsilanolgroups[45,46].
Figure 1. Characteristics of bioactive glass (MBGN_Zn); (a) SEM image, (b) EDS result, (c) particle
size distribution histogram according to SEM image, (d) particle size distribution according to
DLS analysis.
The pore diameter, specific surface area and total pore volume were measured to be
5.2 nm, 368 m
2
/g, and 0.5 cm
3
/g, respectively. Besides confirming with the pore diameters
between 2–50 nm, the mesoporous nature of the bioactive glasses was also indicated with
the N
2
adsorption–desorption isotherms of samples (Figure 2b). According to the Inter-
national Union of Pure and Applied Chemistry (IUPAC) classification, Type IV isotherms
signify mesoporous materials with non-uniform pore size distribution, displaying hystere-
sis loops due to capillary condensation in pores of varying sizes. FTIR spectra of bioactive
glass (Figure 2a) after calcination at 700
◦
C for 2 h have exhibited a strong absorption

Polymers 2024,16, 2066 8 of 18
peak at 1044 cm
−1
which is attributed to Si–O–Si asymmetrical stretching vibrations [
41
].
Additionally, a Si–O–Si with bending scissoring vibrations could be observed at about
800 cm
−1
which is attributed to ring structures in the glass matrix [
42
]. The XRD pattern of
bioactive glasses is shown in Figure 2d; no diffraction peaks were observed in the graph,
confirming that bioactive glasses are amorphous in form, like glass, and the broad band
in the range of 15
◦
–40
◦
can be attributed to the amorphous silicate [
29
,
43
]. The drastic
decrease in weight observed in the TGA graph of bioactive glass samples between 20–90
◦
C
(Figure 2c) has been attributed to the removal of physically adsorbed water and residues
from the bioactive glass production process [
44
]. Subsequently, the gradual reduction
in weight observed between temperatures of 100–700
◦
C is attributed to the removal of
internal water molecules due to the slow condensation of silanol groups [45,46].
Polymers2024,16,xFORPEERREVIEW9of20
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
Figure2.Characteristicsofbioactiveglass(MBGN_Zn);(a)FTIRspectra,(b)N2ads.-desisotherm,
(c)TGAcurve,(d)XRDpaern.
3.1.1BioactivityEvaluationofMBGN_Zn
Beforeincorporatingthebioactiveglassesincollagenpolymersolutionstoprepare
thebonedefectfacinglayerofthemembranes(Col-BG),theinvitrobioactivityofglasses
wasexaminedbyimmersingthecertainamountsofbioactiveglasspowdersinSBFforup
to4weeks.Theassessmentoftheglasses’bioactivitywasconductedthroughXRD,FTIR
analyses.TheXRDpaernsofthebioactiveglassessoakedintheSBFsolutionatvarious
timesaredisplayedinFigure3a.MBGN_Znwhichcontained5%Zndisplayedbioactivity
startingfromthefirstweek.Bioactivityisdefinedbythedevelopmentofahydroxyapatite
(HA)layeronthematerial’ssurface,resultingfromionexchangebetweenthematerial
andbodilyfluids.ThepresenceofHAformationcanbeconfirmedbytheexistenceof
specificpeakcharacteristicsofHA(Figure3a)locatedat2θ=26,32,45,and56.6°corre-
spondingto(002),(211),(222),(004)planesofHA(JCPDS72–1243).Theintensityofthese
peakswasnotablyhigherbytheendofthefourthweek.Previously,authorshavenoted
theretardingeffectofZnonbioactivity,andevenobservedtheabsenceofpeaksassoci-
atedwithHA[22,47,48].Theysuggestedthatthisobservationmightvarydependingon
theprocessofSBFimmersion.WhentheFTIRanalysesfollowingSBFwereexamined,
startingfromthefirstweek,peakscorrespondingtoC–Obondsofcarbonategroupsin
theHAstructurewereobservedatwavenumbersof1500–1400cm⁻1and875–800cm⁻1.
Additionally,ashifttohigherwavenumbersinthepeaksoccurredatwavenumbersof
1100–1000cm⁻1duetophosphateformation[49].Inthisstudyweadoptedanalternative
methodforbioactivityassessment,asrecommendedforpowdersamples.Itwassug-
gestedthatthebioactivitiesofhighsurfaceareabioactivematerialsproducedviathesol-
gelmethodmaynotalignwiththetestsrecommendedbyISOforcoatinganddisc-shaped
Figure 2. Characteristics of bioactive glass (MBGN_Zn); (a) FTIR spectra, (b) N
2
ads.-des isotherm,
(c) TGA curve, (d) XRD pattern.
Bioactivity Evaluation of MBGN_Zn
Before incorporating the bioactive glasses in collagen polymer solutions to prepare
the bone defect facing layer of the membranes (Col-BG), the
in vitro
bioactivity of glasses
was examined by immersing the certain amounts of bioactive glass powders in SBF for up
to 4 weeks. The assessment of the glasses’ bioactivity was conducted through XRD, FTIR
analyses. The XRD patterns of the bioactive glasses soaked in the SBF solution at various
times are displayed in Figure 3a. MBGN_Zn which contained 5% Zn displayed bioactivity
starting from the first week. Bioactivity is defined by the development of a hydroxyapatite
(HA) layer on the material’s surface, resulting from ion exchange between the material and
bodily fluids. The presence of HA formation can be confirmed by the existence of specific
peak characteristics of HA (Figure 3a) located at 2
θ
= 26, 32, 45, and 56.6
◦
corresponding
to (002), (211), (222), (004) planes of HA (JCPDS 72–1243). The intensity of these peaks
was notably higher by the end of the fourth week. Previously, authors have noted the
retarding effect of Zn on bioactivity, and even observed the absence of peaks associated with

Polymers 2024,16, 2066 9 of 18
HA [
22
,
47
,
48
]. They suggested that this observation might vary depending on the process
of SBF immersion. When the FTIR analyses following SBF were examined, starting from
the first week, peaks corresponding to C–O bonds of carbonate groups in the HA structure
were observed at wavenumbers of 1500–1400 cm
−1
and 875–800 cm
−1
. Additionally, a shift
to higher wavenumbers in the peaks occurred at wavenumbers of 1100–1000 cm
−1
due to
phosphate formation [
49
]. In this study we adopted an alternative method for bioactivity
assessment, as recommended for powder samples. It was suggested that the bioactivities
of high surface area bioactive materials produced via the sol-gel method may not align
with the tests recommended by ISO for coating and disc-shaped materials [
35
]. Through
our study, we have demonstrated that mesoporous bioactive glasses containing Zn exhibit
bioactivity as early as the first week of immersion.
Polymers2024,16,xFORPEERREVIEW10of20
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
materials[35].Throughourstudy,wehavedemonstratedthatmesoporousbioactive
glassescontainingZnexhibitbioactivityasearlyasthefirstweekofimmersion.
Figure3.XRDpaerns(a)andFTIRspectra(b)ofbioactiveglass(MBGN_Zn)beforeandafterSBF
incubationatthedifferenttimeperiods(1week,2weeks,3weeks,4weeks).
3.2.CharacterizationMembranes
3.2.1.SEMAnalysesofElectrospunMembranes
Theelectrospinningtechniqueisamethodthatenablestheproductionofmembranes
possessingasimilarporesizeandfiberstructuretotheECM.Researchhasdemonstrated
thatthediameterofthemembranefibercanaffectcellularactivitiessuchasadhesion,
proliferation,andmigration,aributedtoalargersurfaceareaenablingbeercell-surface
interaction[50].Thefindingsofthisstudyrevealedthattheaveragefiberdiameterof
membraneswas324±60nmfor100Col(Figure4a),265±141nmfor70:30Col-Chi(Figure
4b),and349±124nmfor90:10Col-BG(Figure4c)membranes.Thelargerfiberdiameters
inthemembranescontainingbioactiveglasseshavepreviouslybeenaributedtothein-
creasedviscosityofthepolymericsuspensionduetothepresenceoffillerparticles[34].
AccordingtoRadetal.,theincreaseinfiberdiameterobservedupontheadditionofbio-
activeglassisaributedtotheagglomerationofbioactiveglassparticleswithinthefibers
[51].Conversely,thesmallerdiametersofchitosan-containingmembranesmaybelinked
tointeractionsbetweencollagenandchitosan,renderingthemmiscibleatamolecular
level.Thepresenceofultra-thinfibersinthe40nmrangecouldresultfromtheseinterac-
tions,andanotherfactorthathasbeenproposedistheformationoforganicsaltsdueto
theTFAsolventaddedtodissolvechitosan[52].Organicsaltshavebeenreportedtoin-
creasethechargedensity,potentiallyinfluencingthefiberdiameterwhenaddedtoelec-
trospinningsolutionsinsmallamounts[53].Thefiberdiametershaveincreasedforall
compositionsaftercrosslinking.Theaveragefiberdiameterofmembraneswas614±251
nmfor100Col(Figure4d),463±175nmforCol-Chi(Figure4e),and702±149nmforCol-
BG(Figure4f)membranes.Theincreaseinfiberdiameterofcollagenmembranesfollow-
ingcrosslinkingwithGAcanbeascribedtotheimprovedstructuralintegrityanddenser
arrangementofcollagenfibers,resultinginthemergingoffibers[54].
Figure 3. XRD patterns (a) and FTIR spectra (b) of bioactive glass (MBGN_Zn) before and after SBF
incubation at the different time periods (1 week, 2 weeks, 3 weeks, 4 weeks).
3.2. Characterization Membranes
3.2.1. SEM Analyses of Electrospun Membranes
The electrospinning technique is a method that enables the production of membranes
possessing a similar pore size and fiber structure to the ECM. Research has demonstrated
that the diameter of the membrane fiber can affect cellular activities such as adhesion,
proliferation, and migration, attributed to a larger surface area enabling better cell-surface
interaction [
50
]. The findings of this study revealed that the average fiber diameter of mem-
branes was 324
±
60 nm for 100 Col (Figure 4a), 265
±
141 nm for 70:30 Col-Chi (Figure 4b),
and 349
±
124 nm for 90:10 Col-BG (Figure 4c) membranes. The larger fiber diameters in the
membranes containing bioactive glasses have previously been attributed to the increased
viscosity of the polymeric suspension due to the presence of filler particles [
34
]. Accord-
ing to Rad et al., the increase in fiber diameter observed upon the addition of bioactive
glass is attributed to the agglomeration of bioactive glass particles within the fibers [
51
].
Conversely, the smaller diameters of chitosan-containing membranes may be linked to
interactions between collagen and chitosan, rendering them miscible at a molecular level.
The presence of ultra-thin fibers in the 40 nm range could result from these interactions,
and another factor that has been proposed is the formation of organic salts due to the TFA
solvent added to dissolve chitosan [
52
]. Organic salts have been reported to increase the
charge density, potentially influencing the fiber diameter when added to electrospinning
solutions in small amounts [
53
]. The fiber diameters have increased for all compositions
after crosslinking. The average fiber diameter of membranes was 614
±
251 nm for 100 Col
(Figure 4d), 463
±
175 nm for Col-Chi (Figure 4e), and 702
±
149 nm for Col-BG (Figure 4f)
membranes. The increase in fiber diameter of collagen membranes following crosslinking
with GA can be ascribed to the improved structural integrity and denser arrangement of
collagen fibers, resulting in the merging of fibers [54].

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Figure4.SEMimagesof(a)100Col,(b)Col-Chi,(c)Col-BGbeforecrosslinking,and(d)100Col,(e)
Col-Chi,(f)Col-BGaftercrosslinking.
Inhibitingthepassageofmicroorganismstothedefectareaislargelydependenton
theporesizeofthemembrane.Randomlyorientednonwovenmembranesfacilitatethe
creationofultrasmallporeformations[55].Itwasobservedthattheporesizesofthepro-
ducedmembranesarelimitedtoafewmicrons.Theporesizesweremeasuredas0.972±
0.337µm,0.625±0.216µm,and2.263±0.734µmrespectivelyfor100Col,Col-Chi,and
Col-BGsamples(Figure5).Thenumberandsizeofporesweremeasuredtobelargerfor
Col-BGmembranes.Lawetal.explainedthelargerporesizesbylargerfiberdiameters
[27].Nelsonetal.alsoreportedapositivecorrelationbetweenfiberdiameterandthepore
sizeofelectrospunmembranes[56].
Figure5.Poresizedistributionhistogramsof(a)100Col,(b)Col-Chi,and(c)Col-BGcrosslinked
membranes(measurementsconductedusingImageJonSEMimages).

Figure 4. SEM images of (a) 100Col, (b) Col-Chi, (c) Col-BG before crosslinking, and (d) 100Col,
(e) Col-Chi, (f) Col-BG after crosslinking.
Inhibiting the passage of microorganisms to the defect area is largely dependent on
the pore size of the membrane. Randomly oriented nonwoven membranes facilitate the
creation of ultrasmall pore formations [
55
]. It was observed that the pore sizes of the
produced membranes are limited to a few microns. The pore sizes were measured as
0.972
±
0.337
µ
m, 0.625
±
0.216
µ
m, and 2.263
±
0.734
µ
m respectively for 100 Col, Col-
Chi, and Col-BG samples (Figure 5). The number and size of pores were measured to be
larger for Col-BG membranes. Law et al. explained the larger pore sizes by larger fiber
diameters [
27
]. Nelson et al. also reported a positive correlation between fiber diameter
and the pore size of electrospun membranes [56].
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Figure4.SEMimagesof(a)100Col,(b)Col-Chi,(c)Col-BGbeforecrosslinking,and(d)100Col,(e)
Col-Chi,(f)Col-BGaftercrosslinking.
Inhibitingthepassageofmicroorganismstothedefectareaislargelydependenton
theporesizeofthemembrane.Randomlyorientednonwovenmembranesfacilitatethe
creationofultrasmallporeformations[55].Itwasobservedthattheporesizesofthepro-
ducedmembranesarelimitedtoafewmicrons.Theporesizesweremeasuredas0.972±
0.337µm,0.625±0.216µm,and2.263±0.734µmrespectivelyfor100Col,Col-Chi,and
Col-BGsamples(Figure5).Thenumberandsizeofporesweremeasuredtobelargerfor
Col-BGmembranes.Lawetal.explainedthelargerporesizesbylargerfiberdiameters
[27].Nelsonetal.alsoreportedapositivecorrelationbetweenfiberdiameterandthepore
sizeofelectrospunmembranes[56].
Figure5.Poresizedistributionhistogramsof(a)100Col,(b)Col-Chi,and(c)Col-BGcrosslinked
membranes(measurementsconductedusingImageJonSEMimages).

Figure 5. Pore size distribution histograms of (a) 100 Col, (b) Col-Chi, and (c) Col-BG crosslinked
membranes (measurements conducted using ImageJ on SEM images).
3.2.2. Bioactivity Studies of Membranes
The bioactivity of membranes was assessed by immersing them in SBF for 4 weeks.
Peaks related to the (211) plane of HA at 2
θ
= 32
◦
could be observed starting from the first
week of immersion, while peaks related to (222) at around 2
θ
= 45
◦
were visible after 3 weeks
of immersion (Figure 6a) [
49
,
57
]. Confirmation of the formation of HA on the surface of

Polymers 2024,16, 2066 11 of 18
membranes during immersion in SBF was obtained from the FTIR spectra shown in Figure 6b.
After 4 weeks of immersion, a peak at 600 and 570 cm
−1
corresponding to the P–O bending
vibrations associated with the PO
43−
group in the crystalline layer of HA was observed.
Additionally, phosphate group (P–O stretching) was indicated by a peak in the region between
1100 and 1000 cm
−1
. The peaks in the region between 1500 and 1400 cm
−1
were attributed to
the C–O stretching, belonging to the CO32−groups in the carbonated HA layer [58].
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3.2.2.BioactivityStudiesofMembranes
ThebioactivityofmembraneswasassessedbyimmersingtheminSBFfor4weeks.
Peaksrelatedtothe(211)planeofHAat2θ=32°couldbeobservedstartingfromthefirst
weekofimmersion,whilepeaksrelatedto(222)ataround2θ=45°werevisibleafter3
weeksofimmersion(Figure6a)[49,57].ConfirmationoftheformationofHAonthesur-
faceofmembranesduringimmersioninSBFwasobtainedfromtheFTIRspectrashown
inFigure6b.After4weeksofimmersion,apeakat600and570cm−1correspondingtothe
P–ObendingvibrationsassociatedwiththePO43−groupinthecrystallinelayerofHAwas
observed.Additionally,phosphategroup(P–Ostretching)wasindicatedbyapeakinthe
regionbetween1100and1000cm−1.Thepeaksintheregionbetween1500and1400cm−1
wereaributedtotheC–Ostretching,belongingtotheCO32−groupsinthecarbonated
HAlayer[58].
Figure6.XRDpaerns(*indicatesHA,JCPDS;72–1243)(a)andFTIRspectra(b)ofmembranes
beforeandafterSBFincubationatthedifferenttimeperiods(1week,2weeks,3weeks,4weeks).
TheconfirmationofHAformationwasfurthersupportedbySEManalyses(Figure
7).SEMimagesofthemembranesafterimmersioninSBFfor1,2,3,and4weeksshowed
theprogressionofcalciumphosphateprecipitates.Precipitatesbecamevisibleattheend
ofoneweek,andtheentiresurfacewascoveredbytheendofthefourthweek.
Figure 6. XRD patterns (* indicates HA, JCPDS; 72–1243) (a) and FTIR spectra (b) of membranes
before and after SBF incubation at the different time periods (1 week, 2 weeks, 3 weeks, 4 weeks).
The confirmation of HA formation was further supported by SEM analyses (Figure 7).
SEM images of the membranes after immersion in SBF for 1, 2, 3, and 4 weeks showed the
progression of calcium phosphate precipitates. Precipitates became visible at the end of
one week, and the entire surface was covered by the end of the fourth week.
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Figure7.SEMimagesofCol-BGmembranes;attheendof(a)1week,(b)2weeks,(c)3weeks,(d)4
weeksinSBF.
Overall,theSEMimages,XRDresults,andFTIRspectraprovidestrongevidencefor
theformationofHAonthemembranesurfacesduringimmersioninSBF,confirmingtheir
bioactivityandpotentialforbone-bondingapplications.
3.2.3.ThermalAnalysisofMembranes
Themasslossofcollagenoccursintwostages:first,thedesorptionofphysically
boundwaterwithinthecollagenstructure,andsecond,thethermaldegradationofpoly-
mericchains,duringwhichthepeptidebondswithinthecollagenmoleculesbeginto
breakdown.Chitosandegradesinthreestages.Inadditiontothetwostagesobservedfor
collagen-lossofstructurallyboundwaterandthermaldegradationofpolymericchains-
chitosanhasanextrainitialstep.Thisadditionalstep,occurringatlowertemperatures,is
duetothelossofabsorbedorweaklyboundwaterandistypicallyrecognizedasthefirst
droponthethermogravimetricanalysis(TGA)curve(Figure8)[54].

Figure8.TGAcurvesofmembranesattheconstantheatingrate(20°Cperminute).
Figure 7. SEM images of Col-BG membranes; at the end of (a) 1 week, (b) 2 weeks, (c) 3 weeks,
(d) 4 weeks in SBF.

Polymers 2024,16, 2066 12 of 18
Overall, the SEM images, XRD results, and FTIR spectra provide strong evidence for
the formation of HA on the membrane surfaces during immersion in SBF, confirming their
bioactivity and potential for bone-bonding applications.
3.2.3. Thermal Analysis of Membranes
The mass loss of collagen occurs in two stages: first, the desorption of physically bound
water within the collagen structure, and second, the thermal degradation of polymeric
chains, during which the peptide bonds within the collagen molecules begin to break down.
Chitosan degrades in three stages. In addition to the two stages observed for collagen -loss
of structurally bound water and thermal degradation of polymeric chains-chitosan has an
extra initial step. This additional step, occurring at lower temperatures, is due to the loss
of absorbed or weakly bound water and is typically recognized as the first drop on the
thermogravimetric analysis (TGA) curve (Figure 8) [54].
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
Figure7.SEMimagesofCol-BGmembranes;attheendof(a)1week,(b)2weeks,(c)3weeks,(d)4
weeksinSBF.
Overall,theSEMimages,XRDresults,andFTIRspectraprovidestrongevidencefor
theformationofHAonthemembranesurfacesduringimmersioninSBF,confirmingtheir
bioactivityandpotentialforbone-bondingapplications.
3.2.3.ThermalAnalysisofMembranes
Themasslossofcollagenoccursintwostages:first,thedesorptionofphysically
boundwaterwithinthecollagenstructure,andsecond,thethermaldegradationofpoly-
mericchains,duringwhichthepeptidebondswithinthecollagenmoleculesbeginto
breakdown.Chitosandegradesinthreestages.Inadditiontothetwostagesobservedfor
collagen-lossofstructurallyboundwaterandthermaldegradationofpolymericchains-
chitosanhasanextrainitialstep.Thisadditionalstep,occurringatlowertemperatures,is
duetothelossofabsorbedorweaklyboundwaterandistypicallyrecognizedasthefirst
droponthethermogravimetricanalysis(TGA)curve(Figure8)[54].

Figure8.TGAcurvesofmembranesattheconstantheatingrate(20°Cperminute).
Figure 8. TGA curves of membranes at the constant heating rate (20 ◦C per minute).
As observed in the graph, the TGA curves of membranes containing chitosan are
slightly different from those that do not contain chitosan. The observed variation has been
attributed to the modified thermal stability of the composite material as a result of the
interactions between hydroxyl (–OH), amino (–NH
2
), and carbonyl (–C=O) groups found in
collagen and chitosan [
59
]. Another factor that can be implicated in the various curves is the
application of acidic solvent TFA which used to increase the solubility of the chitosan [
60
].
It is known that the low pH causes the chitosan’s amino groups to protonate. This allows
NH
3+
in the chitosan structure and –COO
−
in the aspartic and glutamic residues in the
collagen structure to interact electrostatically, leading to a decrease in the thermal stability
of Col-Chi membranes [59].
3.2.4. Mechanical Properties of Membranes
Nonwoven electrospun collagen-based membranes are highly susceptible to degrada-
tion even with minimal exposure to water, and their fiber structure can be easily degraded
by moisture [
32
]. Therefore, crosslinking of collagen-based electrospun membranes before
use is important to ensure the barrier properties of the membrane and to obtain more
controlled degradation behavior. Compared to synthetic polymers, collagen typically ex-
hibits a lower tensile strength due to its natural biomaterial composition. The mechanical
properties of the double-layered electrospun membranes were evaluated through tensile
testing under both dry (DL24-Dry and DL48-Dry) and wet conditions (DL24-Wet). Prior to
testing, the membranes were immersed in SBF for 2 min to observe their behavior under
wet conditions. Additionally, double-layered membranes crosslinked for 48 h (DL48-Dry)
were also tested to assess the effect of crosslinking duration on mechanical properties.
Figure 9illustrates the stress/elongation curves for the double-layered samples. The hy-
drated crosslinked collagen membranes displayed significantly higher elongation and

Polymers 2024,16, 2066 13 of 18
lower stress compared to the dry samples. This outcome was expected, as previously
mentioned in the literature, due to the weakening of non-covalent interactions by water
molecules [61].
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Asobservedinthegraph,theTGAcurvesofmembranescontainingchitosanare
slightlydifferentfromthosethatdonotcontainchitosan.Theobservedvariationhasbeen
aributedtothemodifiedthermalstabilityofthecompositematerialasaresultofthe
interactionsbetweenhydroxyl(–OH),amino(–NH
2
),andcarbonyl(–C=O)groupsfound
incollagenandchitosan[59].Anotherfactorthatcanbeimplicatedinthevariouscurves
istheapplicationofacidicsolventTFAwhichusedtoincreasethesolubilityofthechitosan
[60].ItisknownthatthelowpHcausesthechitosan’saminogroupstoprotonate.This
allowsNH
3+
inthechitosanstructureand–COO
−
intheasparticandglutamicresiduesin
thecollagenstructuretointeractelectrostatically,leadingtoadecreaseinthethermalsta-
bilityofCol-Chimembranes[59].
3.2.4.MechanicalPropertiesofMembranes
Nonwovenelectrospuncollagen-basedmembranesarehighlysusceptibletodegra-
dationevenwithminimalexposuretowater,andtheirfiberstructurecanbeeasilyde-
gradedbymoisture[32].Therefore,crosslinkingofcollagen-basedelectrospunmem-
branesbeforeuseisimportanttoensurethebarrierpropertiesofthemembraneandto
obtainmorecontrolleddegradationbehavior.Comparedtosyntheticpolymers,collagen
typicallyexhibitsalowertensilestrengthduetoitsnaturalbiomaterialcomposition.The
mechanicalpropertiesofthedouble-layeredelectrospunmembraneswereevaluated
throughtensiletestingunderbothdry(DL24-DryandDL48-Dry)andwetconditions
(DL24-Wet).Priortotesting,themembraneswereimmersedinSBFfor2mintoobserve
theirbehaviorunderwetconditions.Additionally,double-layeredmembranescross-
linkedfor48h(DL48-Dry)werealsotestedtoassesstheeffectofcrosslinkingdurationon
mechanicalproperties.Figure9illustratesthestress/elongationcurvesforthedouble-lay-
eredsamples.Thehydratedcrosslinkedcollagenmembranesdisplayedsignificantly
higherelongationandlowerstresscomparedtothedrysamples.Thisoutcomewasex-
pected,aspreviouslymentionedintheliterature,duetotheweakeningofnon-covalent
interactionsbywatermolecules[61].

Figure9.Stress-elongationgraphofmembranes.
Itisanticipatedthatthewatercontentofthehydrated,crosslinkedcollagenmem-
braneshadenhancethemobilityofpolymerchains,leadingtotheobservedhighelonga-
tioninthehydratedsamples[62].Asthecrosslinkingtimeincreased,themembranes’
abilitytowithstandstressdecreased,possiblyduetotheformationofamorerigid
Figure 9. Stress-elongation graph of membranes.
It is anticipated that the water content of the hydrated, crosslinked collagen mem-
branes had enhance the mobility of polymer chains, leading to the observed high elongation
in the hydrated samples [
62
]. As the crosslinking time increased, the membranes’ ability to
withstand stress decreased, possibly due to the formation of a more rigid structure. It has
been previously reported that an increase in crosslinking density does not always lead to
an increase in mechanical properties [
63
]. In a study conducted by Guo et al., mechanical
analysis of membranes produced via electrospinning revealed that collagen membranes
had a tensile strength of 2.13
±
0.17 MPa, which increased to 6.16
±
1.43 MPa upon the
addition of chitosan. These results highlight the significant enhancement of mechanical
properties facilitated by chitosan [64].
3.2.5. Biodegradation of Membranes
The natural degradation capability of electrospun membranes is a critical property,
influencing their biocompatibility and potential for tissue regeneration. Maintaining struc-
tural integrity is key for these membranes to exhibit barrier properties during the healing
process [
65
]. When the degradation behavior of the membranes in SBF and PBS was com-
pared, it was observed that in SBF (Figure 10a), the degradation rates for all samples (Col,
Col-Chi, Col-BG, and DL) generally increased over time. Col-BG consistently exhibited the
highest degradation rate among all samples, surpassing Col and Col-Chi. The increased
degradation rate observed in the Col-BG samples in both PBS and SBF after a 4-week incu-
bation period could be attributed to the presence of bioactive glasses with a high surface
area in the membrane structure. This led to greater liquid adsorption, and the gradual
dissolution of the glass within the membrane structure may have also contributed to the
higher weight loss observed in these samples [
34
]. DL showed the lowest degradation rate
among the samples, although it still increased steadily over the 4-week period. This could
be attributed to the formation of a crosslinked layer primarily at the surface, resulting in a
lower degree of crosslinking at the deeper layers of the membrane. Consequently, these
deeper layers may have become more prone to degradation once the better crosslinked
surface layer was removed [66].

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structure.Ithasbeenpreviouslyreportedthatanincreaseincrosslinkingdensitydoesnot
alwaysleadtoanincreaseinmechanicalproperties[63].InastudyconductedbyGuoet
al.,mechanicalanalysisofmembranesproducedviaelectrospinningrevealedthatcolla-
genmembraneshadatensilestrengthof2.13±0.17MPa,whichincreasedto6.16±1.43
MPaupontheadditionofchitosan.Theseresultshighlightthesignificantenhancementof
mechanicalpropertiesfacilitatedbychitosan[64].
3.2.5.BiodegradationofMembranes
Thenaturaldegradationcapabilityofelectrospunmembranesisacriticalproperty,
influencingtheirbiocompatibilityandpotentialfortissueregeneration.Maintaining
structuralintegrityiskeyforthesemembranestoexhibitbarrierpropertiesduringthe
healingprocess[65].WhenthedegradationbehaviorofthemembranesinSBFandPBS
wascompared,itwasobservedthatinSBF(Figure10a),thedegradationratesforallsam-
ples(Col,Col-Chi,Col-BG,andDL)generallyincreasedovertime.Col-BGconsistently
exhibitedthehighestdegradationrateamongallsamples,surpassingColandCol-Chi.
TheincreaseddegradationrateobservedintheCol-BGsamplesinbothPBSandSBFafter
a4-weekincubationperiodcouldbeaributedtothepresenceofbioactiveglasseswitha
highsurfaceareainthemembranestructure.Thisledtogreaterliquidadsorption,and
thegradualdissolutionoftheglasswithinthemembranestructuremayhavealsocontrib-
utedtothehigherweightlossobservedinthesesamples[34].DLshowedthelowestdeg-
radationrateamongthesamples,althoughitstillincreasedsteadilyoverthe4-weekpe-
riod.Thiscouldbeaributedtotheformationofacrosslinkedlayerprimarilyatthesur-
face,resultinginalowerdegreeofcrosslinkingatthedeeperlayersofthemembrane.
Consequently,thesedeeperlayersmayhavebecomemorepronetodegradationoncethe
beercrosslinkedsurfacelayerwasremoved[66].

Figure10.Degradation(%)ofmembranesovertime(1,2,3,and4week(s))in(a)SBFand(b)PBS
(dataweregivenasmeanandstandarddeviationsandindicatedwitherrorbars,pvalues<0.05(n
=3)).
Incontrast,inPBS(Figure10b),thedegradationratesweregenerallylowercompared
toSBFforallsamples.Col-BGstilldemonstratedthehighestdegradationrate,followed
byCol-Chi,Col,andDL.However,therateofincreaseindegradationovertimeappeared
tobeslowerinPBScomparedtoSBFforallsamples.Overall,thedegradationbehaviorin
SBFwasmorepronouncedandacceleratedcomparedtoPBS,withCol-BGconsistently
showingthehighestdegradationrateacrossbothenvironments.ThedegradationinSBF
provedtobemorecomplex,likelyduetomineralaccumulationonthemembranesurface
[67].Awell-designedGBRbarriershouldgraduallydegradeovertime.Degradable
Figure 10. Degradation (%) of membranes over time (1, 2, 3, and 4 week(s)) in (a) SBF and (b) PBS
(data were given as mean and standard deviations and indicated with error bars, pvalues < 0.05
(n= 3)).
In contrast, in PBS (Figure 10b), the degradation rates were generally lower compared
to SBF for all samples. Col-BG still demonstrated the highest degradation rate, followed by
Col-Chi, Col, and DL. However, the rate of increase in degradation over time appeared to
be slower in PBS compared to SBF for all samples. Overall, the degradation behavior in SBF
was more pronounced and accelerated compared to PBS, with Col-BG consistently showing
the highest degradation rate across both environments. The degradation in SBF proved
to be more complex, likely due to mineral accumulation on the membrane surface [
67
].
A well-designed GBR barrier should gradually degrade over time. Degradable membranes
offer the advantage of naturally decomposing without requiring surgical removal, but there
are challenges in controlling the degradation process of these collagen membranes [68].
3.2.6. Cell Viability Evaluation of Membranes
The MTT assay is a colorimetric method commonly used to assess cell viability and pro-
liferation. The metabolic activity of cells is measured by converting MTT, a yellow tetrazolium
salt, into purple formazan crystals through mitochondrial enzyme activity. For optimal tissue
regeneration, the electrospun membranes should facilitate the adhesion and proliferation of
target cells. The least toxicity is a desirable property of any material in contact with the living
tissues. Collagen–chitosan complexes have been demonstrated to be non-cytotoxic when
tested against various cell lines using different techniques [
64
,
69
,
70
]. It was observed that all
sample groups exhibited cell viability above 70% at various concentrations studied, indicating
that the produced membranes do not have a cytotoxic effect and demonstrate biocompatible
properties (Figure 11) [
37
]. Statistically significant differences were found between different
concentrations within all sample groups (p< 0.05). Moreover, significant differences were
noted among Col-Chi, Col-BG, and DL samples at the same concentration. Upon analyzing
the results at identical concentrations, it was noted that membranes doped with bioactive
glass displayed higher cell viability (%) than other samples. Particularly, samples containing
double-layered membranes and bioactive glass containing membranes exhibited the highest
cell viability (%). While cell viability values generally increased with concentration, this
rise was more noticeable in membranes containing bioactive glass. Consequently, it can be
inferred that the contribution of bioactive glass becomes more pronounced at higher concen-
trations. The results showed that the samples containing chitosan exhibited the lowest cell
viability. Collagen has been reported to influence chitosan’s cytocompatibility, primarily by
widening its pore aperture, thus enhancing chitosan’s water retention capacity. Consequently,
this may facilitate cell adhesion and infiltration into the pores, promoting the formation of
three-dimensional growths [
69
]. In a study conducted by Wang et al., it was demonstrated
that a chitosan–collagen composite film enhances osteoblast functions and mineralization in
MC3T3-E1 cells by increasing Erk1/2 phosphorylation, thereby enhancing Runx2 transcrip-

Polymers 2024,16, 2066 15 of 18
tional activity. Moreover, the film induced the overexpression of osteoblastic marker genes
such as Runx2 and Type I collagen in these cells [71].
Polymers2024,16,xFORPEERREVIEW16of20


membranesoffertheadvantageofnaturallydecomposingwithoutrequiringsurgicalre-
moval,buttherearechallengesincontrollingthedegradationprocessofthesecollagen
membranes[68].
3.2.6.CellViabilityEvaluationofMembranes
TheMTTassayisacolorimetricmethodcommonlyusedtoassesscellviabilityand
proliferation.ThemetabolicactivityofcellsismeasuredbyconvertingMTT,ayellowte-
trazoliumsalt,intopurpleformazancrystalsthroughmitochondrialenzymeactivity.For
optimaltissueregeneration,theelectrospunmembranesshouldfacilitatetheadhesion
andproliferationoftargetcells.Theleasttoxicityisadesirablepropertyofanymaterial
incontactwiththelivingtissues.Collagen–chitosancomplexeshavebeendemonstrated
tobenon-cytotoxicwhentestedagainstvariouscelllinesusingdifferenttechniques
[64,69,70].Itwasobservedthatallsamplegroupsexhibitedcellviabilityabove70%at
variousconcentrationsstudied,indicatingthattheproducedmembranesdonothavea
cytotoxiceffectanddemonstratebiocompatibleproperties(Figure11)[37].Statistically
significantdifferenceswerefoundbetweendifferentconcentrationswithinallsample
groups(p<0.05).Moreover,significantdifferenceswerenotedamongCol-Chi,Col-BG,
andDLsamplesatthesameconcentration.Uponanalyzingtheresultsatidenticalcon-
centrations,itwasnotedthatmembranesdopedwithbioactiveglassdisplayedhighercell
viability(%)thanothersamples.Particularly,samplescontainingdouble-layeredmem-
branesandbioactiveglasscontainingmembranesexhibitedthehighestcellviability(%).
Whilecellviabilityvaluesgenerallyincreasedwithconcentration,thisrisewasmoreno-
ticeableinmembranescontainingbioactiveglass.Consequently,itcanbeinferredthatthe
contributionofbioactiveglassbecomesmorepronouncedathigherconcentrations.The
resultsshowedthatthesamplescontainingchitosanexhibitedthelowestcellviability.
Collagenhasbeenreportedtoinfluencechitosan’scytocompatibility,primarilybywid-
eningitsporeaperture,thusenhancingchitosan’swaterretentioncapacity.Consequently,
thismayfacilitatecelladhesionandinfiltrationintothepores,promotingtheformation
ofthree-dimensionalgrowths[69].InastudyconductedbyWangetal.,itwasdemon-
stratedthatachitosan–collagencompositefilmenhancesosteoblastfunctionsandminer-
alizationinMC3T3-E1cellsbyincreasingErk1/2phosphorylation,therebyenhancing
Runx2transcriptionalactivity.Moreover,thefilminducedtheoverexpressionofosteo-
blasticmarkergenessuchasRunx2andTypeIcollageninthesecells[71].

Figure11.Thecellviability(%)valuesofmembranesatdifferentextractconcentrationsafter24hof
incubation(dataweregivenasmean±standarddeviations(n=6)—differentsymbols(*,**,***)
indicatedwitherrorbarsonthebarshowastatisticallysignificantdifference,pvalues<0.05).
Figure 11. The cell viability (%) values of membranes at different extract concentrations after 24 h
of incubation (data were given as mean
±
standard deviations (n= 6)—different symbols (*, **, ***)
indicated with error bars on the bar show a statistically significant difference, pvalues < 0.05).
4. Conclusions
This study highlights the significant role of tailored material design in enhancing the
bone regeneration process, specifically through the application of guided bone regeneration
techniques. We have developed a highly effective approach to improve tissue regeneration
by designing a double-layered membrane that addresses the distinct needs of both soft
and hard tissues. The bone tissue facing layer, composed of collagen and zinc-doped
bioactive glass, may promote the growth of bone tissue, while the soft tissue layer, made of
collagen and chitosan, may meet the needs of soft tissue. We employed electrospinning
to create nanofibers that imitate the extracellular matrix, proving that the membranes
displayed desirable bioactivity, biodegradability, and biocompatibility. The results indicate
that the double-layered membranes made from polymer-bioactive glass composite, which
were created in this research, have great potential for use in tissue engineering. These
membranes may provide an advanced solution to the difficulties faced in bone regeneration
in dental applications.
Author Contributions: Conceptualization, D.A. and A.C.Ö.; formal analysis, D.A. and A.C.Ö.;
investigation, D.A.; methodology, D.A. and C.Ö.; resources, K.T., Y.M.S. and S.Y.; supervision, K.T.,
Y.M.S. and S.Y.; validation, D.A.; visualization, D.A., A.C.Ö. and C.Ö.; writing—original draft, D.A.,
A.C.Ö. and C.Ö.; writing—review and editing, D.A., A.C.Ö. and S.Y. All authors have read and
agreed to the published version of the manuscript.
Funding: This research received funding from Yıldız Technical University Scientific Research Projects
Coordination Unit under project number FBA-2021-4533.
Institutional Review Board Statement: Not applicable.
Data Availability Statement: The data presented in this study are available on request from the
corresponding author.
Acknowledgments: The author(s) extend sincere appreciation to Istanbul Arel University Polymer
Technologies and Composite Application and Research Center for providing access to the electro-
spinning device and facilitating characterization analysis. Ali Can ÖZARSLAN also thanks the
financial support from the TUBITAK under BIDEB/2250 - Performance-Based Scholarships Pro-
gram for PhD. Cem ÖZEL and Dilan ALTAN also thanks the financial support from the TUBITAK
under the BIDEB/2211 National PhD Scholarship Program and BIDEB/2250—Performance-Based

Polymers 2024,16, 2066 16 of 18
Scholarships Program for PhD. Dilan ALTAN also expresses gratitude to Yildiz Technical University
for collaborating with the Turkish Council of Higher Education (YÖK) in providing the 100/2000
Doctoral Scholarship. This work is dedicated to the memory of Kadriye Tuzlakoglu, whose invaluable
guidance and support continue to inspire. Though Kadriye Tuzlakoglu is no longer with us, her
impact on our academic and personal journey will be forever remembered and cherished. Graphical
abstract was created with BioRender, ©biorender.com.
Conflicts of Interest: The authors declare that they have no known competing financial interests or
personal relationships that could have appeared to influence the work reported in this paper.
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