Measuring Physical Properties of Electrospun Nanofiber Mats: Comparison
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Please note this is a comparison between Version 2 by Lindsay Dong and Version 1 by Andrea Ehrmann.

Electrospun nanofiber mats are nowadays often used for biotechnological and biomedical applications, such as wound healing or tissue engineering. 

  • apparent density
  • porometer
  • scanning electron microscopy (SEM)
  • specific surface area
  • fast Fourier transform (FFT)
  • Electrospun nanofiber mats

1. Introduction

Electrospinning allows for producing nanofiber mats from diverse polymers or polymer blends, including various nanoparticles, and in this way tailoring the nanofiber materials in a broad range [1,2,3][1][2][3]. Their large specific surfaces as well as other physical and chemical properties make such nanofiber mats highly suitable for biotechnological and biomedical applications, such as wound healing or tissue engineering [4,5,6][4][5][6].
Naturally, nanofiber mats for biomedical applications need special properties, especially being not cytotoxic, but depending on the exact application, they can be desired to be biodegradable or waterproof, have antibacterial of fungicide properties, etc. [7,8,9,10][7][8][9][10]. However, their morphological, mechanical and other physical properties may also be important for the planned application, although these values are often less intensively investigated than chemical and biochemical properties and often only briefly described in the methodic sections. Nevertheless, the mechanical properties are decisive for the lifetime of a nanofibrous product and the limits of its potential application, while cell adhesion depends on morphological parameters, hydrophobicity and water uptake are among the parameters controlling liquid transport, which is important for wound dressing, and porosity and water vapor/air permeability are physical parameters influencing the filtration of liquids or gases, respectively. The porosity is often mentioned as an important parameter for wound exudate transport and cell adhesion [11,12,13,14,15,16,17][11][12][13][14][15][16][17]. While the porosity describes the amount of porous volume inside the nanofiber mat volume, the pore size distribution is also often taken into account [14,15,16,18,19][14][15][16][18][19]. Other morphological parameters are the nanofiber diameters [16,20,21,22,23,24][16][20][21][22][23][24] and their orientation [25,26,27][25][26][27] as well as the surface roughness and nanofiber mat thickness [28]. Besides such structural features, the hydrophobic/hydrophilic properties of nanofiber mats [11,16][11][16] and their water uptake [17,24,29][17][24][29] are often reported. Other often-mentioned parameters are mechanical [11,15,17,18,19,20,21,23,24][11][15][17][18][19][20][21][23][24] and electrical properties [18,30][18][30] as well as water vapor and air permeability [31,32][31][32].

2. Porosity

The porosity describes the volume of voids inside a given volume of a nanofiber mat. Firstly, it must be mentioned that there can be open as well as closed pores, the latter of which are not accessible for all methods described below [33,34,35][33][34][35]. However, for typical nanofiber mats, pores can be expected to be openly accessible to any test fluid, so that for most nanofibrous membranes, no differences between the measurement principles are expected, whether they take into account closed pores or not. One of the methods that would also measure closed pores is the Archimedean principle [36]. Pati et al. used a specific gravity bottle filled with ethanol in which the nanofibrous scaffold was dipped and afterwards removed again [37]. The porosity was then calculated according to  P o r o s i t y = m 2 − m 3 − m S m 1 − m 3 × 100 % with the mass m1 of the specific gravity bottle filled with ethanol, the mass m2 of the bottle with ethanol and the scaffold, the mass m3 of the bottle after taking out the scaffold again, and mS the mass of the scaffold. Dividing both numerator and denominator by the density of ethanol, it is observable that the porosity is determined as the volume of the ethanol taken out of the bottle with the scaffold, i.e., of the ethanol that was sticking in its pores, divided by the volume of the scaffold with ethanol. Safari et al. used the same principle based on deionized water in which their nanofiber mats were immersed for 15 min, taken out, quickly dried at the sample surface and weighed, so that the porosity could be calculated as the mass of the uptaken water, divided by the sample mass [38]. Without directly using the Archimedean principle, Kahdim et al. calculated the porosity by soaking their nanofiber mats in phosphate-buffered saline (PBS) solution for 24 h, measuring the sample mass before and after PBS uptake and calculating the porosity according to this fluid uptake and the PBS density [39]. Here, it is not mentioned whether the samples were also dried on both surfaces before weighing them. Using immersion of the dried nanofiber mat in n-butanol for 2 h, Wang et al. as well as Chen et al. calculated the porosity from the densities of membrane and n-butanol as well as the measured dry and wet mass of the nanofiber mat [40,41][40][41].

3. Pore Size Distribution

While the porosity describes the overall volume of the pores in a given sample, the pore size distribution is sometimes even more important in biomedical scaffolds since it defines which pores are available for cells or can release a drug. In the easiest way, pore sizes are measured on the surface or along cross-sections of samples, typically from SEM images. Agueda et al. describe that they used ImageJ to investigate pore sizes from 3 areas per sample from SEM images taken with magnification of 2000× and 5000×, measuring 30 pores per sample [56][42]. Liu et al. similarly examined pore sizes from SEM images of their nanofiber mats, taken with magnifications of 5000× and 20,000×, averaging over 100 pore areas [57][43]. Tahami et al. also used ImageJ to measure pore sizes in SEM images, while not exactly describing the number of measurements [58][44], while Stella et al. showed histograms of the pore size distributions, which in principle allow for counting the number of measurements per sample, again taken with ImageJ from SEM images [59][45]. Only a few groups describe how they defined the pore size that they measured. Zhang et al. described measuring a reversible change in pore size by analyzing 30 pores per sample with ImageJ in their SEM images by measuring the longest diameter. Besides these 2D methods, some groups chose 3D pore measurement methods. One of them is the Barrett–Joyner–Halenda (BJH) technique, allowing analysis of pores between 1.7 nm and 300 nm [64][46]. This method is based on N2 adsorption–desorption isotherms, taken at liquid nitrogen temperature [65[47][48][49][50],66,67,68], i.e., similarly to the Brunauer–Emmett–Teller (BET) surface area measurements described in the next section. Generally, the BJH method as well as further developments are based on measuring the film formation on the mesopore walls in dependence of the condensation pressure, taking into account the so-called Kelvin-type relation describing capillary condensation, meaning that mesopores covered with an absorbed fill will instantaneously be filled [68][50]. A capillary flow porometer can also be used to investigate the pore sizes of nanofiber mats [41,70][41][51]. In this method, the sample pores are filled with a wetting liquid that is afterwards blown out of the pores by a pressurized gas or liquid [71][52], where smaller pores need a higher pressure to be emptied, i.e., the measured flow rate depends on the proportion of filled pores that block the flow so that there is zero flow at low pressure, while at a certain high pressure, all pores are emptied, and the flow rate becomes identical to the value measured for the dry sample at the same pressure [72][53]. Generally, some other methods are available, although less often reported in the recent literature, to evaluate the pore size distribution of nanofiber mats, such as mercury intrusion porosimetry [76][54]. Another method that is less well-known but often more readily available than a porometer or nitrogen absorption techniques, is thermoporometry, also known as thermoporosimetry or cryoporometry [77,78,79,80][55][56][57][58]. This calorimetric method is based on the melting or freezing point depression of the pore liquid, which can be measured with a laboratory differential scanning calorimetry (DSC) instrument by fast cooling the sample wetted with deionized water to −30 °C or lower and then slowly (e.g., with a heating rate of 0.1–1 K/min) heating it up to a temperature slightly above 0 °C [81,82][59][60].

4. Specific Surface Area

Among the very special properties of nanofiber mats is their large specific surface area. While this value is often mentioned as a reason why nanofiber mats are especially useful for a certain application, its value is scarcely measured. The most common measurement technique is based on the aforementioned BET adsorption–desorption isotherms of N2 gas on the sample surface [68,88,89,90,91,92,93,94][50][61][62][63][64][65][66][67]. Generally, a wider hysteresis loop in the adsorption–desorption curve indicates a more mesoporous structure of the nanofiber mat [68][50]. To evaluate these curves, it is necessary to differentiate between the different adsorption isotherms. Many nanofiber mats reported in the literature belong to type IV [95][68], while type I or a pressure-dependent change between these types are also found [96,97][69][70]. Differentiation between the different types of adsorption isotherms is possible by fitting a range of possible equations to the measured curves, while a first idea of the type can already be gained by looking at the slopes of the measured adsorption–desorption curves [98][71].

5. Nanofiber Diameter

The diameters of nanofibers in an electrospun membrane are usually obtained from SEM images and either given as average with standard deviation or as distribution, sometimes as distribution boxplots [101][72], but mostly as a histogram. In the latter case, typically 100 or more fiber diameters per sample are measured to prepare a histogram [39,41,48,49,90][39][41][63][73][74]. In most cases, the diameters are measured manually by ImageJ [14[14][42][43][50][75],43,56,57,68], while a few groups mention other software [41,44,90][41][63][76] or do not mention the software used [70][51].

6. Nanofiber Orientation

Oriented nanofibers can be produced, e.g., by a fast rotating collector cylinder. Similarly to the nanofiber diameter distribution, the orientation of the nanofibers in an electrospun membrane is also usually determined from SEM images. The fiber orientations can be measured manually in ImageJ [108][77] or other software [109][78]. An interesting possibility to automatically detect fiber orientations is given by the ImageJ plugin OrientationJ. Another possibility to evaluate fiber orientation automatically in ImageJ is given by the inbuilt fast Fourier transform (FFT) function as well as the Oval Profile plugin to receive a radial direction intensity plot [110,111,112][79][80][81]. The latter can also be given as a polar plot [113][82], which is often more intuitively understandable. These automatic orientation examinations have the advantage of taking into account all fiber parts, while manual calculations naturally have to be limited to certain parts of the fibers and are thus susceptible to subjective decisions of the evaluator. On the other hand, automatic calculations of the fiber orientations are highly error-prone if the fibers are too thin, i.e., only a few pixels per diameter, which will lead to favoring 0°, ±45° and ±90° orientations [110][79]. Thus, the choice of the images will potentially influence the results and has to be done with care.

7. Surface Roughness

The surface roughness of electrospun nanofiber mats can influence their hydrophobicity to a certain extent. When researchers mention measuring the roughness related to electrospun nanofiber mats, sometimes the roughness of the whole membrane is meant, while in other cases the roughness of single nanofibers is addressed. Correspondingly, different measurement methods are necessary to detect these different orders of magnitude of roughness. Havlícek et al., e.g., show roughness measurements based on confocal laser scanning microscope (CLSM) images [61][83]. As preparation, they coated the investigated samples with a thin gold layer to enable better visibility of the relatively transparent nanofibers. In this way, 3D maps of the nanofibrous surfaces were prepared. Field emission SEM (FE-SEM) images were used by Shahverdi et al., who investigated nanofiber mat surfaces with Fiji software (National Institute of Health, 9000 Rockville Pike, Bethesda, MD 20892, USA), leading to relatively noisy 3D images for most samples on which a qualitative comparison of the fiber roughness was performed [114][84]. El-Morsy et al. used Gwyddion (http://gwyddion.net/ (accessed on 25 March 2023)) to evaluate FE-SEM images, showing average roughness Ra of around 100 nm for different fiber material compositions, again with high noise [115][85]. Other studies show similarly noisy 3D maps, created by Gwyddion or other software from SEM images [116][86], although the noise could be reduced by using SEM images with higher magnification [117][87].  Besides the aforementioned methods based on surface images, taken by SEM or AFM, it is also possible to use a laser surface profilometer. In this way, Kichi et al. reported roughness Ra in the range of 3–6 µm, i.e., apparently taking into account a larger area of the nanofiber mat, as could be expected due to the optical measurement and the correspondingly limited resolution [123][88]. Even a mechanical stylus-based profilometer was used to measure the roughness of nanofiber mats, finding Ra values around 160–260 nm, however, for a lateral resolution of approx. 60 µm [124][89].

8. Nanofiber Mat Thickness

While the macroscopic thickness of a nanofiber mat seems to be simply measurable at first glance, there are, nevertheless, diverse methods with their advantages and disadvantages, sometimes influencing the result by the measurement. One of the problematic methods is using a micrometer caliper since its pressure limitation is usually not sufficient to avoid compression of fine nanofiber mats, similarly to microscopic textile fabrics. Liu et al. tried to compensate for this effect by folding the membranes twice before measuring, i.e., by measuring four layers instead of only one [57][43], while other groups did not comment on this problem [125][90]. A typical textile thickness measurement instrument, which has a larger measurement area and causes less pressure on the investigated sample, was applied by Pakolpakcil et al., who used a digital thickness gauge for nonwovens and measured at 10 points on the nanofiber mat [126][91]. To fully avoid this influence of the measurement on the measured value, some studies used optical methods to investigate the thickness of an electrospun membrane. Ryu et al. applied light transmittance measurements to investigate the sample thickness [127][92]. For this, they prepared nanofiber mats with different electrospinning times between 15 min and 75 min, measured their light transmittance and the thickness, the latter by cross-sectional microscopic images, and used the Beer–Lambert law correlating both values. This enabled a real-time thickness measurement during electrospinning.

9. Hydrophobicity/Hydrophilicity

Hydrophobic or hydrophilic properties of nanofiber mats can be significantly influenced by surface functionalization, e.g., by plasma treatment [132][93]. The hydrophobic or hydrophilic properties of nanofiber mats are mostly determined by contact angle measurements, mostly applying the sessile-drop method in which a small droplet (e.g., with 5 µL volume [49[74][94],133], sometimes less [43][75]) is placed on the sample, and a microscope with camera is used to take photographs from the side, often at defined times, enabling fitting the contact angles on the photographs. There are also contact measurement instruments that are commercially available [38,42,44,46,48,108,134][38][73][76][77][95][96][97]. For custom-made setups, evaluation of the contact angles is possible with ImageJ, either manually or with a plugin such as DropSnake [57][43].

10. Water Uptake

The water uptake of an electrospun nanofiber mat can be defined in different ways—by the uptake in the pores around the fibers, or by the uptake inside the fibers, causing swelling, which is especially the case for electrospun hydrogels [139][98]. The water uptake of the material itself can be tested in the bulk form, e.g., by measuring the water uptake of a film of the examined material [38]. It is calculated by W a t e r   u p t a k e = m 1 − m 0 m 0 × 100 % with the masses m0 of the dry sample and m1 of the sample after immersion in water for a defined time. Often, distilled or deionized water is used, and immersion times are usually around 1–2 days [38,42,140][38][95][99]. The water uptake may vary upon adding fillers, such as nanoclays [138][100]. While typical values of water uptake are around several percent for many materials, it can also be in the range of 200–600% for very hydrophilic, porous scaffolds [38,43,49,134,141][38][74][75][97][101]. For hydrogels, even increasing values of several thousand percent during a few minutes were measured [45,142,143][102][103][104]. 

11. Mechanical Properties

The mechanical properties of electrospun nanofiber mats depend on the fiber material and orientation, but also on the crystallinity of the fibers. They are mostly investigated by tensile tests [56,57][42][43], often with test speeds of 1–10 mm/min [41[41][63][95],42,90], sometimes even 20–30 mm/min [46[72][96],101], depending on the sample size and elongation at break. A few papers report measuring stress–strain curves with a constant force ramp rate, e.g., 0.3 N/min [88][61]. Safari et al. examined the difference between dry and wet state and found a significantly higher elongation at break and lower tensile strength in the wet state of Poly(N-vinylcaprolactam)/poly(vinyl acetate) copolymer nanofiber mats [38]. Zadeh et al. found that the percentage of carbon nanotubes in the polyurethane nanofiber mats influenced Young’s modulus of the investigated samples [49][74]. A very special tensile test, based on single electrospun fibers, was reported by Munawar and Schubert [144][105]. Working with well-aligned fibers, they rolled a fiber bundle for the tensile testing, clamped it in a single-fiber tensile tester, and afterwards cut the tested area and weighed the tested part of the fiber bundle to enable calculation of Young’s modulus. In this way, they could measure along the fiber axes instead of taking the mechanical properties of a whole nanofiber mat, averaging over arbitrary fiber orientation. Besides tensile and compressive tests, some groups also reported bursting tests of nanofiber mats. Jalalah et al. applied the standard bursting strength test according to ISO 13938-2:1999 and found a linear correlation between nanofiber mat thickness and bursting strength [89][62]. Nejad et al. found a significant increase in the bursting strength of their nanofiber mats by adding PCL to poly(ethylene terephthalate) (PET). It was, however, calculated from the tensile strength tests [53][106]. The latter also tested suture retention according to ISO 7198:2016 by tensile tests with a well-defined suture thread that was inserted 2 mm from the top edge of the electrospun strip, so that the tensile test led to pulling the suture through the graft [53][106].

12. Electrical Conductivity

The conductivity of nanofibers depends on their material, thickness, crystallinity, etc. Conductive nanofiber mats can stimulate cell attachment, proliferation and differentiation [145][107]. This is why the conductivity of electrospun membranes is often measured [9]. On the other hand, soft and compressible textile fabrics generally pose a challenge to measurements of their conductivity since the contact between the measuring instrument and conductive parts of the sample may be prohibited by non-conductive fibers, and the fibrous structure reduces the contact area if contact pins are used, as is usual for multimeters [146][108]. Generally, samples can be measured with the two-electrode methods (as in common multimeters), the four-electrode method, which is capable of eliminating the contact resistance, and methods with even more electrodes [147][109]. The four-wire measurement (also known as four-terminal sensing or four-point probe) uses two outer current-introducing and two inner voltage-sensing electrodes, in this way becoming independent from the contact resistances. The van der Pauw method works similarly: while the four contacts are not aligned but positioned along the sample perimeter, the van der Pauw method measures the average sample resistivity, whereas the linear four-point probe method measures the resistivity along the electrode orientation [148][110]. Due to the expected high contact resistance, textile fabrics should normally be measured with the linear four-point probe or the van der Pauw method, depending on the desired information and the sample geometry.

13. Water Vapor Permeability

The water vapor permeability of an electrospun membrane is correlated with its porosity and is especially important for wound dressing applications, where too high water vapor permeability results in fast hydration and thus scars, while too low values let exudates accumulate and thus increase the risk of infection [150,151][111][112]. Quantitatively, the water vapor transmission should be between 76 and 9360 g/(m2 day) to improve wound healing [152,153][113][114]. Gu et al. reduced this optimum window to 2000–2500 g/(m2 day) [154][115]. Mostly, the water vapor transmittance is measured by gravimetry, where the sample is fixed on the opening, with defined diameter (e.g., 1.23 cm), of a round bottle that is filled with a defined volume of distilled water (e.g., 5 mL) and placed in an oven at typically 37 °C for 24 h [155][116]. The water vapor transmittance (WVTR) is calculated as WVTR = − Δ W A T with the mass change −ΔW of the water in the container, the exposed area A and the measurement time T [42][95]. Usually, the same measurement is performed with an open container as the reference [155][116].

14. Air Permeability

Air permeability is one of the parameters often measured for macroscopic textiles, but less often for electrospun nanofiber mats, potentially because it can partly be estimated from water vapor transmission tests [153][114]. Nevertheless, some studies report measuring the air permeability of nanofibrous scaffolds directly, usually giving the transmitted air volume per area and time, i.e., in the unit cm³/(cm2 s) or cm/s. Pakolpakcil et al. used a commercial air permeability tester at a fixed pressure of 100 Pa and a test area of 20 cm2, resulting in values of around 10–12 cm/s [126][91]. Using the same parameters, Sun et al. found values of around 1–3 cm/s for their nanofiber mats electrospun from polyamide and multi-wall carbon nanotubes [165][117].

15. Thermal Properties

The thermal conductivity of an electrospun nanofiber mat is often correlated with its electrical conductivity; however, in many cases, only the thermal conductivity is measured. Depending on the planned application, sometimes a high thermal conductivity is sought, while often the high porosity of a nanofiber mat, combined with the low thermal conductivity of most polymers, is used to prepare heat-blocking nanofibrous membranes instead. Thermal conductivity can be evaluated, e.g., by a diffusivity measurement instrument at a defined temperature, often not far above the room temperature [174][118]. Measurements at high temperatures, however, are also possible using the hot disk method, e.g., using infrared thermography on the upper side of the sample, which is placed on the hot disk [175,176,177][119][120][121].

References

  1. Teo, W.E.; Ramakrishna, S. A review on electrospinning design and nanofibre assemblies. Nanotechnology 2006, 17, R89.
  2. Ramakrishna, S.; Fujihara, K.; Teo, W.E.; Yong, T.; Ma, Z.; Ramaseshan, R. Electrospun nanofibers: Solving global issues. Mater. Today 2006, 9, 40–50.
  3. Grothe, T.; Wehlage, D.; Böhm, T.; Remche, A.; Ehrmann, A. Needleless electrospinning of PAN nanofibre mats. Tekstilec 2017, 60, 290–295.
  4. Klinkhammer, K.; Seiler, N.; Grafahrend, D.; Gerardo-Nava, J.; Mey, J.; Brook, G.A.; Möller, M.; Dalton, P.D.; Klee, D. Deposition of electrospun fibers on reactive substrates for In Vitro investigations. Tissue Eng. Part C 2009, 15, 77–85.
  5. Mamun, A. Review of possible applications of nanofibrous mats for wound dressings. Tekstilec 2019, 62, 89–100.
  6. Gao, S.T.; Tang, G.S.; Hua, D.W.; Xiong, R.H.; Han, J.Q.; Jiang, S.H.; Zhang, Q.L.; Huang, C.B. Stimuli-responsive bio-based polymeric systems and their applications. J. Mater. Chem. B 2019, 7, 709–729.
  7. Rasouli, R.; Barhoum, A.; Bechelany, M.; Dufresne, A. Nanofibers for biomedical and healthcare applications. Macromol. Biosci. 2019, 19, 1800256.
  8. Wehlage, D.; Blattner, H.; Mamun, A.; Kutzli, I.; Diestelhorst, E.; Rattenholl, A.; Gudermann, F.; Lütkemeyer, D.; Ehrmann, A. Cell growth on electrospun nanofiber mats from polyacrylonitrile (PAN) blends. AIMS Bioeng. 2020, 7, 43–54.
  9. Tanzli, E.; Ehrmann, A. Electrospun nanofibrous membranes for tissue engineering and cell growth. Appl. Sci. 2021, 11, 6929.
  10. Zahedi, P.; Khatibi, A.; Fallah-Darrehchi, M. ANtimicrobial electrospun membranes. In Electrospun and Nanofibrous Membranes; Elsevier: Amsterdam, The Netherlands, 2023; pp. 501–519.
  11. Jafari, S.; Hosseini Salekdeh, S.S.; Solouk, A.; Yousefzadeh, M. Electrospun polyethylene terephthalate (PET) nanofibrous conduit for biomedical application. Polym. Adv. Technol. 2020, 31, 284–296.
  12. More, N.; Ranglani, D.; Kharche, S.; Kapusetti, G. Electrospun mat of thermal-treatment-induced nanocomposite hydrogel of polyvinyl alcohol and cerium oxide for biomedical applications. J. Appl. Polym. Sci. 2020, 137, 49426.
  13. Ramos, C.; Lanno, G.-M.; Laidmäe, I.; Meos, A.; Härmas, R.; Kogermann, K. High humidity electrospinning of porous fibers for tuning the release of drug delivery systems. Int. J. Polym. Mater. Polym. Biomater. 2021, 70, 880–892.
  14. Chen, S.X.; John, J.V.; McCarthy, A.; Xie, J.W. New forms of electrospun nanofiber materials for biomedical applications. J. Mater. Chem. B 2020, 8, 3733–3746.
  15. Zou, S.Z.; Wang, X.R.; Fan, S.N.; Yao, X.; Zhang, Y.P.; Shao, H.L. Electrospun regenerated Antheraea pernyi silk fibroin scaffolds with improved pore size, mechanical properties and cytocompatibility using mesh collectors. J. Mater. Chem. B 2021, 9, 5514–5527.
  16. Selvaras, T.; Alshamrani, S.A.; Gopal, R.; Jaganathan, S.K.; Sivalingam, S.; Kadiman, S.; Saidin, S. Biodegradable and antithrombogenic chitosan/elastin blended polyurethane electrospun membrane for vascular tissue integration. J. Biomed. Mater. Res. B 2023, 111, 1171–1181.
  17. Entekhabi, E.; Nazarpak, M.H.; Shafieian, M.; Mohammadi, H.; Firouzi, M.; Hassannejad, Z. Fabrication and in vitro evaluation of 3D composite scaffold based on collagen/hyaluronic acid sponge and electrospun polycaprolactone nanofibers for peripheral nerve regeneration. J. Biomed. Mater. Res. A 2021, 109, 300–312.
  18. Yardimci, A.I. Comparative Study of the Structural, Mechanical and Electrochemical Properties of Polyacrylonitrile (PAN)-Based Polypyrrole (PPy) and Polyvinylidene Fluoride (PVDF) Electrospun Nanofibers. J. Macromol. Sci. B 2022, 61, 1103–1115.
  19. Zhang, H.N.; Zhang, T.T.; Qiu, Q.H.; Qin, X.H. Quaternary ammonium salt–modified polyacrylonitrile/polycaprolactone electrospun nanofibers with enhanced antibacterial properties. Text. Res. J. 2021, 91, 2194–2203.
  20. Jirofti, N.; Golandi, M.; Movaffagh, J.; Ahmadi, F.S.; Kalalinia, F. Improvement of the Wound-Healing Process by Curcumin-Loaded Chitosan/Collagen Blend Electrospun Nanofibers: In Vitro and In Vivo Studies. ACS Biomater. Sci. Eng. 2021, 7, 3886–3897.
  21. Akbarzadeh, M.; Pezeshki-Modaress, M.; Zandi, M. Biphasic, tough composite core/shell PCL/PVA-GEL nanofibers for biomedical application. J. Appl. Polym. Sci. 2020, 137, 48713.
  22. Islam, M.A.; Begum, H.A.; Shahid, M.A.; Ali, Y. Antibacterial electrospun nanofibers from poly (vinyl alcohol) and Mikania micrantha with augmented moisture properties: Formation and evaluation. J. Text. Inst. 2021, 112, 1602–1610.
  23. Stella, S.M.; Vijayalakshi, U. Influence of chemically modified Luffa on the preparation of nanofiber and its biological evaluation for biomedical applications. J. Biomed. Mater. Res. A 2019, 107, 610–620.
  24. Du, Z.W.; Jia, S.W.; Xiong, P.; Cai, Z.J. Preparation of protein nanoparticle-coated poly(hydroxybutyrate) electrospun nanofiber based scaffold for biomedical applications. Int. J. Polym. Mater. Polym. Biomater. 2022, 71, 677–691.
  25. Malik, S.; Hussain, T.; Nazir, A.; Khenoussi, N.; Cheema, S.A. Oriented electrospun nanofibers on stand-alone multi-segmented cylindrical collectors. J. Text. Inst. 2021, 112, 955–964.
  26. Munawar, M.A.; Schubert, D.W. Highly Oriented Electrospun Conductive Nanofibers of Biodegradable Polymers-Revealing the Electrical Percolation Thresholds. ACS Appl. Polym. Mater. 2021, 3, 2889–2901.
  27. Han, Y.S.; Hong, H.J.; Park, S.M.; Kim, D.S. Metal–Electrolyte Solution Dual-Mode Electrospinning Process for In Situ Fabrication of Electrospun Bilayer Membrane. Adv. Mater. Interfaces 2020, 7, 2000571.
  28. Maurya, A.K.; Weidenbacher, L.; Spano, F.; Fortunato, G.; Rossi, R.M.; Frenz, M.; Dommann, A.; Neels, A.; Sadeghpour, A. Structural insights into semicrystalline states of electrospun nanofibers: A multiscale analytical approach. Nanoscale 2019, 15, 7176–7187.
  29. Shao, Z.G.; Chen, J.Y.; Ke, L.-J.; Wang, Q.F.; Wang, X.; Li, W.W.; Zheng, G.F. Directional Transportation in a Self-Pumping Dressing Based on a Melt Electrospinning Hydrophobic Mesh. ACS Biomater. Sci. Eng. 2021, 7, 5918–5926.
  30. Sengupta, P.; Ghosh, A.; Bose, N.; Mukherjee, S.; Chowdhury, A.R.; Datta, P. A comparative assessment of poly(vinylidene fluoride)/conducting polymer electrospun nanofiber membranes for biomedical applications. J. Appl. Polym. Sci. 2020, 137, 49115.
  31. Gwon, G.; Choi, H.J.; Bae, J.H.; Binti Zulkifli, N.A.; Jeong, W.; Yoo, S.S.; Hyun, D.C.; Lee, S.W. An All-Nanofiber-Based Substrate-Less, Extremely Conformal, and Breathable Organic Field Effect Transistor for Biomedical Applications. Adv. Funct. Mater. 2022, 32, 2204645.
  32. Shi, S.; Si, Y.F.; Han, Y.T.; Wu, T.; Irfan Iqbal, M.; Fei, B.; Li, R.K.Y.; Hu, J.L.; Qu, J.P. Non-Toxic Crosslinking of Electrospun Gelatin Nanofibers for Tissue Engineering and Biomedicine—A Review. Adv. Mater. 2022, 34, 2107938.
  33. Kim, H.K.; Chung, H.J.; Park, T.G. Biodegradable polymeric microspheres with “open/closed” pores for sustained release of human growth hormone. J. Control. Release 2006, 112, 167–174.
  34. Kim, Y.J.; Kang, J.H.; Shen, B.W.; Wang, Y.Q.; He, Y.; Lee, M.S. Open–closed switching of synthetic tubular pores. Nat. Commun. 2015, 6, 8650.
  35. Liu, H.; Zhao, X.P. Thermal Conductivity Analysis of High Porosity Structures with Open and Closed Pores. Int. J. Heat Mass Transf. 2022, 183A, 122089.
  36. Yang, J.; Shi, G.X.; Bei, J.Z.; Wang, S.G.; Cao, Y.L.; Shang, Q.X.; Yang, G.H.; Wang, W.J. Fabrication and surface modification of macroporous poly(L-lactic acid) and poly(L-lactic-co-glycolic acid) (70/30) cell scaffolds for human skin fibroblast cell culture. J. Biomed. Mater. Res. 2002, 62, 438–446.
  37. Pati, F.; Adhikari, B.; Dhara, S. Development of chitosan-tripolyphosphate non-woven fibrous scaffolds for tissue engineering application. J. Mater. Sci. Mater. Med. 2012, 23, 1085–1096.
  38. Safari, S.; Ehsani, M.; Zandi, M. Stimuli-responsive electrospun nanofibers based on PNVCL-PVAc copolymer in biomedical applications. Prog. Biomater. 2021, 10, 245–258.
  39. Kahdim, Q.S.; Abdelmoula, N.; Al-Karagoly, H.; Albukhaty, S.; Al-Saaidi, J. Fabrication of a Polycaprolactone/Chitosan Nano fibrous Scaffold Loaded with Nigella sativa Extract for Biomedical Applications. BioTech 2023, 12, 19.
  40. Wang, N.; Yang, Y.J.; Al-Deyab, S.S.; El-Newehy, M.; Yu, J.Y.; Ding, B. Ultra-light 3D nanofibre-nets binary structured nylon 6–polyacrylonitrile membranes for efficient filtration of fine particulate matter. J. Mater. Chem. A 2015, 3, 23946–23954.
  41. Chen, Y.J.; Mensah, A.; Wang, Q.Q.; Li, D.W.; Qiu, Y.Y.; Wie, Q.F. Hierarchical porous nanofibers containing thymol/beta-cyclodextrin: Physico-chemical characterization and potential biomedical applications. Mater. Sci. Eng. C 2020, 115, 111155.
  42. Agueda, J.R.S.; Madrid, J.; Mondragon, J.M.; Lim, J.; Tan, A.; Wang, I.; Duguran, N.; Bondoc, A. Synthesis and Characterization of Electrospun Polyvinylidene Fluoride-based (PVDF) Scaffolds for Renal Bioengineering. J. Phys. Conf. Ser. 2021, 2071, 012005.
  43. Liu, W.Y.; Walker, G.; Price, S.; Yang, X.D.; Li, J.; Bunt, C. Electrospun Membranes as a Porous Barrier for Molecular Transport: Membrane Characterization and Release Assessment. Pharmaceutics 2021, 13, 916.
  44. Tahami, S.R.; Nemati, N.H.; Keshvari, H.; Khorasani, M.T. Effect of Electrical Potential on the Morphology of Polyvinyl Alcohol/ Sodium Alginate Electrospun Nanofibers, Containing Herbal Extracts of Calendula Officinalis for Using in Biomedical Applications. J. Mod. Process. Manuf. Prod. 2020, 9, 43–46.
  45. Stella, S.M.; Sridhar, T.M.; Ramprasath, R.; Gimbun, J.; Vijayalakshmi, U. Physio-Chemical and Biological Characterization of Novel HPC (Hydroxypropylcellulose):HAP (Hydroxyapatite):PLA (Poly Lactic Acid) Electrospun Nanofibers as Implantable Material for Bone Regenerative Application. Polymers 2023, 15, 155.
  46. McLaren, R.L.; Laycock, C.J.; Brousseau, E.; Owen, G.R. Examining slit pore widths within plasma-exfoliated graphitic material utilising Barrett–Joyner–Halenda analysis. New J. Chem. 2021, 45, 12071–12080.
  47. Yang, H.M.; Song, X.L.; Zhang, X.C.; Ao, W.Q.; Qiu, G.H. Synthesis of vanadium-doped SnO2 nanoparticles by chemical co-precipitation method. Mater. Lett. 2003, 57, 3124–3127.
  48. Sing, K.S.W.; Williams, R.T. Physisorption Hysteresis Loops and the Characterization of Nanoporous Materials. Absorpt. Sci. Technol. 2004, 22, 773–782.
  49. Kim, C.-Y.; Lee, J.-K.; Kim, B.-I. Synthesis and pore analysis of aerogel–glass fiber composites by ambient drying method. Colloids Surf. A Physicochem. Eng. Asp. 2008, 313–314, 179–182.
  50. Bazzi, M.; Shabani, I.; Mohandesi, J.A. Enhanced mechanical properties and electrical conductivity of Chitosan/Polyvinyl Alcohol electrospun nanofibers by incorporation of graphene nanoplatelets. J. Mech. Behav. Biomed. Mater. 2022, 125, 104975.
  51. Zhang, S.Y.; Yan, D.; Zhao, L.F.; Lin, J.Y. Composite fibrous membrane comprising PLA and PCL fibers for biomedical application. Compos. Commun. 2022, 34, 101268.
  52. Peinador, R.I.; Calvo, J.I.; Aim, R.B. Comparison of Capillary Flow Porometry (CFP) and Liquid Extrusion Porometry (LEP) Techniques for the Characterization of Porous and Face Mask Membranes. Appl. Sci. 2020, 10, 5703.
  53. Jena, A.; Gupta, K. Pore Volume of Nanofiber Nonwovens. Int. Nonwovens J. 2005, 2.
  54. He, X.; Wang, Y.-n.; Zhou, J.F.; Wang, H.B.; Ding, W.; Shi, B. Suitability of Pore Measurement Methods for Characterizing the Hierarchical Pore Structure of Leather. J. Am. Leather Chem. Assoc. 2019, 114, 41–47.
  55. Liu, Y.; Lannutti, J.J. Characterization of electrospun porosities: Current techniques. In Proceedings of the Nanofiber, Applications and Related Technologies NART 2021, Istanbul, Turkey, 8–10 September 2021; pp. 54–63.
  56. Appell, M.; Jackson, M.A. Applications of Nanoporous Materials in Agriculture. Adv. Appl. Nanotechnol. Agric. 2013, 1143, 167–176.
  57. Orsolini, P.; Michen, B.; Huch, A.; Tingaut, P.; Caseri, W.R.; Zimmermann, T. Characterization of Pores in Dense Nanopapers and Nanofibrillated Cellulose Membranes: A Critical Assessment of Established Methods. ACS Appl. Mater. Interfaces 2015, 7, 25884–25897.
  58. Raja, I.S.; Fathima, N.N. Gelatin–Cerium Oxide Nanocomposite for Enhanced Excisional Wound Healing. ACS Appl. Bio Mater. 2018, 1, 487–495.
  59. Hao, J.J.; Lu, C.X.; Zhou, P.C.; Li, D.H. Pore structure development of polyacrylonitrile nascent fibers in water stretching process. Thermochim. Acta 2013, 569, 42–47.
  60. Kanungo, I.; Fathima, N.N.; Rao, J.R.; Nair, B.U. Influence of PCL on the material properties of collagen based biocomposites and in vitro evaluation of drug release. Mater. Sci. Eng. C 2013, 33, 4651–4659.
  61. Balasubramaniam, B.; Kumar, S.A.; Singh, K.A.; Bhunia, S.; Verma, K.; Tian, L.M.; Gupta, R.K.; Gaharwar, A.K. Electrically Conductive MoS2 Reinforced Polyacrylonitrile Nanofibers for Biomedical Applications. Adv. NanoBiomed Res. 2022, 2, 2100105.
  62. Jalalah, M.; Ahmad, A.; Saleem, A.; Bilal Qadir, M.; Khaliq, Z.; Khan, M.Q.; Nazir, A.; Faisal, M.; Alsaiari, M.; Irfan, M.; et al. Electrospun Nanofiber/Textile Supported Composite Membranes with Improved Mechanical Performance for Biomedical Applications. Membranes 2022, 12, 1158.
  63. Chen, Y.J.; Qiu, Y.Y.; Chen, W.B.F.; Wei, Q.F. Electrospun thymol-loaded porous cellulose acetate fibers with potential biomedical applications. Mater. Sci. Eng. C 2020, 109, 110536.
  64. Chen, S.; Shen, L.L.; Huang, D.; Du, J.; Fan, X.X.; Wie, A.L.; Chen, W.Y. Facile synthesis, microstructure, formation mechanism, in vitro biocompatibility, and drug delivery property of novel dendritic TiO2 nanofibers with ultrahigh surface area. Mater. Sci. Eng. C 2020, 115, 111100.
  65. Cheng, H.; Li, X.N.; Li, T.H.; Qin, D.F.; Tang, T.F.; Li, Y.P.; Wang, G.X. Electrospun Nanofibers with High Specific Surface Area to Prepare Modified Electrodes for Electrochemiluminescence Detection of Azithromycin. J. Nanomater. 2021, 2021, 9961663.
  66. Li, W.Y.; Chao, S.; Li, Y.M.; Bai, F.Q.; Teng, Y.K.; Li, X.; Li, L.J.; Wang, C. Dual-layered composite nanofiber membrane with Cu-BTC-modified electrospun nanofibers and biopolymeric nanofibers for the removal of uremic toxins and its application in hemodialysis. J. Membr. Sci. 2022, 642, 119964.
  67. Arabpour, Z.; Baradaran-Rafii, A.; Bakhshaiesh, N.L.; Ai, J.; Ebrahimi-Barough, S.; Malekabadi, H.E.; Nazeri, N.; Vaez, A.; Salehi, M.; Sefat, F.; et al. Design and characterization of biodegradable multi layered electrospun nanofibers for corneal tissue engineering applications. J. Biomed. Mater. Res. 2019, 107, 2340–2349.
  68. Lim, S.K.; Hwang, S.-H.; Chang, D.I.; Kim, S.H. Preparation of mesoporous In2O3 nanofibers by electrospinning and their application as a CO gas sensor. Sens. Actuators B Chem. 2010, 149, 28–33.
  69. Prajapati, Y.N.; Verma, N. Adsorptive desulfurization of diesel oil using nickel nanoparticle-doped activated carbon beads with/without carbon nanofibers: Effects of adsorbate size and adsorbent texture. Fuel 2017, 189, 186–194.
  70. Othman, F.E.C.; Yusof, N.; Petru, M.; Md Nordin, N.A.H.; Hamid, M.F.; Ismail, A.F.; Rushdan, A.I.; Hassan, S.A. Polyethy leneimine-impregnated activated carbon nanofiber composited graphene-derived rice husk char for efficient post-combustion CO2 capture. Nanotechnol. Rev. 2022, 11, 926–944.
  71. Al-Ghouti, M.A.; Da’ana, D.A. Guidelines for the use and interpretation of adsorption isotherm models: A review. J. Hazard. Mater. 2020, 393, 122383.
  72. Chaiarwut, S.; Ekabutr, P.; Chuysinuan, P.; Chanamuangkon, T.; Supaphol, P. Surface immobilization of PCL electrospun nanofibers with pexiganan for wound dressing. J. Polym. Res. 2021, 28, 344.
  73. Khataei, S.; Al-Musawi, M.H.; Asadi, K.; Ramezani, S.; Abbasian, M.; Ghorbani, M. Effect of molecular weight and content of polyvinylpyrrolidone on cell proliferation, loading capacity and properties of electrospun green tea essential oil-incorporated polyamide-6/polyvinylpyrrolidone nanofibers. J. Drug Deliv. Sci. Technol. 2023, 82, 104310.
  74. Zadeh, Z.E.; Solouk, A.; Shafieian, M.; Nazarpak, M.H. Electrospun polyurethane/carbon nanotube composites with different amounts of carbon nanotubes and almost the same fiber diameter for biomedical applications. Mater. Sci. Eng. C 2021, 118, 111403.
  75. Ghaee, A.; Bagheri-Khoulenjani, S.; Afshar, H.A.; Bogheiri, H. Biomimetic nanocomposite scaffolds based on surface modified PCL-nanofibers containing curcumin embedded in chitosan/gelatin for skin regeneration. Comp. B Eng. 2019, 177, 107339.
  76. Esmaeili, E.; Eslami-Arshaghi, T.; Hosseinzadeh, S.; Elahirad, E.; Jamalpoor, Z.; Hatamie, S.; Soleimani, M. The biomedical potential of cellulose acetate/polyurethane nanofibrous mats containing reduced graphene oxide/silver nanocomposites and curcumin: Antimicrobial performance and cutaneous wound healing. Int. J. Biol. Macromol. 2020, 152, 418–427.
  77. Li, Y.; Shen, Q.; Shen, J.; Ding, X.B.; Liu, T.; He, J.H.; Zhu, C.Y.; Zhao, D.; Zhu, J.D. Multifunctional Fibroblasts Enhanced via Thermal and Freeze-Drying Post-treatments of Aligned Electrospun Nanofiber Membranes. Adv. Fiber Mater. 2021, 3, 26–37.
  78. Cai, Z.J.; Xiong, P.; He, S.Q.; Zhu, C. Improved piezoelectric performances of highly orientated poly(β-hydroxybutyrate) electrospun nanofiber membrane scaffold blended with multiwalled carbon nanotubes. Mater. Lett. 2019, 240, 213–216.
  79. Hellert, C.; Wortmann, M.; Frese, N.; Grötsch, G.; Cornelißen, C.; Ehrmann, A. Adhesion of Electrospun Poly(acrylonitrile) Nanofibers on Conductive and Isolating Foil Substrates. Coatings 2021, 11, 249.
  80. Storck, J.L.; Grothe, T.; Mamun, A.; Sabantina, L.; Klöcker, M.; Blachowicz, T.; Ehrmann, A. Orientation of electrospun magnetic nanofibers near conductive areas. Materials 2020, 13, 47.
  81. Bazrafshan, Z.; Stylios, G.K. Custom-built electrostatics and supplementary bonding in the design of reinforced Collagen-g-P (methyl methacrylate-co-ethyl acrylate)/nylon 66 core-shell fibers. J. Mech. Behav. Biomed. Mater. 2018, 87, 19–29.
  82. He, H.J.; Wang, Y.M.; Farkas, B.; Nagy, Z.K.; Molnar, K. Analysis and prediction of the diameter and orientation of AC electrospun nanofibers by response surface methodology. Mater. Des. 2020, 194, 108902.
  83. Havlícek, K.; Svobodová, L.; Bakalova, T.; Lederer, T. Influence of electrospinning methods on characteristics of polyvinyl butyral and polyurethane nanofibres essential for biological applications. Mater. Des. 2020, 194, 108898.
  84. Shahverdi, F.; Barati, A.; Salehi, E.; Arjomandzadegan, M. Biaxial electrospun nanofibers based on chitosan-poly (vinyl alcohol) and poly (ε-caprolactone) modified with CeAlO3 nanoparticles as potential wound dressing materials. Int. J. Biol. Macromol. 2022, 221, 736–750.
  85. El-Morsy, M.A.; Afifi, M.; Ahmed, M.K.; Awwad, N.S.; Ibrahium, H.A.; Alqahtani, M.S. Electrospun nanofibrous scaffolds of polycaprolactone containing binary ions of Pd/vanadate doped hydroxyapatite for biomedical applications. J. Drug Deliv. Sci. Technol. 2022, 70, 103153.
  86. El-Naggar, M.E.; Shalaby, E.S.; Abd-Al-Aleem, A.H.; Abu-Saied, M.A.; Youssef, A.M. Synthesis of environmentally benign antimicrobial dressing nanofibers based on polycaprolactone blended with gold nanoparticles and spearmint oil nanoemulsion. J. Mater. Res. Technol. 2021, 15, 3447–3460.
  87. Teaima, M.H.; Abdelnaby, F.A.; Fadel, M.; El-Nabarawi, M.A.; Shoueir, K.R. Synthesis of Biocompatible and Environmentally Nanofibrous Mats Loaded with Moxifloxacin as a Model Drug for Biomedical Applications. Pharmaceutics 2020, 12, 1029.
  88. Kichi, M.K.; Torkaman, R.; Mohammadi, H.; Toutounchi, A.; Kharaziha, M.; Alihosseini, F. Electrochemical and in vitro bioactivity behavior of poly (ε-caprolactone) (PCL)-gelatin-forsterite nano coating on titanium for biomedical application. Mater. Today Commun. 2020, 24, 101326.
  89. Drobota, M.; Gradinaru, L.M.; Vlad, S.; Bargan, A.; Butnaru, M.; Angheloiu, M.; Afori, M. Preparation and Characterization of Electrospun Collagen Based Composites for Biomedical Applications. Materials 2020, 13, 3961.
  90. Lasenko, I.; Sanchaniya, J.V.; Kanukuntla, S.P.; Ladani, Y.; Viluma-Gudmona, A.; Kononova, O.; Lusis, V.; Tipans, I.; Selga, T. The Mechanical Properties of Nanocomposites Reinforced with PA6 Electrospun Nanofibers. Polymers 2023, 15, 673.
  91. Pakolpakcil, A.; Draczynski, Z.; Szulc, J.; Stawski, D.; Tarzynska, N.; Bednarowicz, A.; Sikorski, D.; Hernandez, C.; Sztajnowski, S.; Krucinska, I.; et al. An In Vitro Study of Antibacterial Properties of Electrospun Hypericum perforatum Oil-Loaded Poly(lactic Acid) Nonwovens for Potential Biomedical Applications. Appl. Sci. 2021, 11, 8219.
  92. Ryu, H.I.; Koo, M.S.; Kim, S.J.; Kim, S.K.; Park, Y.-A.; Park, S.M. Uniform-thickness electrospun nanofiber mat production system based on real-time thickness measurement. Sci. Rep. 2020, 10, 20847.
  93. Mozaffari, A.; Gashti, M.P. Air Plasma Functionalization of Electrospun Nanofibers for Skin Tissue Engineering. Biomedicines 2022, 10, 617.
  94. Chen, S.X.; John, J.V.; McCarthy, A.; Carlson, M.A.; Li, X.W.; Xie, J.W. Fast transformation of 2D nanofiber membranes into pre-molded 3D scaffolds with biomimetic and oriented porous structure for biomedical applications. Appl. Phys. Rev. 2020, 7, 021406.
  95. Salehi, M.; Niyakan, M.; Ehterami, A.; Haghi-Daredeh, S.; Nazarnezhad, S.; Abbaszadeh-Goudarzi, G.; Vaez, A.; Hashemi, S.F.; Rezaei, N.; Mousavi, S.R. Porous electrospun poly(ε-caprolactone)/gelatin nanofibrous mat containing cinnamon for wound healing application: In vitro and in vivo study. Biomed. Eng. Lett. 2020, 10, 149–161.
  96. Ahmed, M.K.; Mansour, S.F.; Al-Wafi, R.; Abdel-Fattah, E. Nanofibers scaffolds of co-doped Bi/Sr-hydroxyapatite encapsulated into polycaprolactone for biomedical applications. J. Mater. Res. Technol. 2021, 13, 2297–2309.
  97. Movahedi, M.; Salehi, A.O.M.; Hajipour, F.P.; Etemad, S. Casein release and characterization of electrospun nanofibres for cartilage tissue engineering. Bull. Mater. Sci. 2022, 45, 76.
  98. Gashti, M.P.; Dehdast, S.A.; Berenjian, A.; Shabani, M.; Zarinabadi, E.; Fard, G.C. PDDA/Honey Antibacterial Nanofiber Composites for Diabetic Wound-Healing: Preparation, Characterization, and In Vivo Studies. Gels 2023, 9, 173.
  99. Merin, D.D.; Jose, R.A.; Arulananth, T.S.; Sundarraj, A.A.; Inbamalar, T.M.; Meharie, M.G. Nanoclay-Incorporated Polycaprolactone Matrix via Electrospinning Techniques-Enriched Spectroscopic Responses. J. Nanomater. 2023, 2023, 1194158.
  100. Liu, Y.Y.; Chen, X.Q.; Xin, J.H. Hydrophobic duck feathers and their simulation on textile substrates for water repellent treatment. Bioinspiration Biomim. 2008, 3, 046007.
  101. Serbezeanu, D.; Vlad-Bubulac, T.; Rusu, D.; Gradisteanu Pircalabioru, G.; Samoila, I.; Dinescu, S.; Aflori, M. Functional Polyimide-Based Electrospun Fibers for Biomedical Application. Materials 2019, 12, 3201.
  102. Chen, J.; Zhang, T.H.; Hua, W.K.; Li, P.Y.; Wang, X.F. 3D Porous poly(lactic acid)/regenerated cellulose composite scaffolds based on electrospun nanofibers for biomineralization. Colloids Surf. A Physicochem. Eng. Asp. 2020, 585, 124048.
  103. Sazegar, M.; Bazgir, S.; Katbab, A.A. Preparation and characterization of water-absorbing gas-assisted electrospun nanofibers based on poly(vinyl alcohol)/chitosan. Mater. Today Commun. 2020, 25, 101489.
  104. Abdolbaghian, H.; Bazgir, S. Fabrication and characterization of gas-assisted core-shell hydrogel nanofibers as a drug release system with antibacterial activity. Eur. Polym. J. 2022, 174, 111302.
  105. Munawar, M.A.; Schubert, D.W. Revealing Electrical and Mechanical Performances of Highly Oriented Electrospun Conductive Nanofibers of Biopolymers with Tunable Diameter. Int. J. Mol. Sci. 2021, 22, 10295.
  106. Nejad, M.R.; Yousefzadeh, M.; Solouk, A. Electrospun PET/PCL small diameter nanofibrous conduit for biomedical application. Mater. Sci. Eng. C 2020, 110, 110692.
  107. Pedrotty, D.M.; Koh, J.; Davis, B.H.; Taylor, D.A.; Wolf, P.; Niklason, L.E. Engineering skeletal myoblasts: Roles of threedimensional culture and electrical stimulation. Am. J. Physiol. Heart Circ. Physiol. 2005, 288, H1620–H1626.
  108. Schwarz-Pfeiffer, A.; Obermann, M.; Weber, M.O.; Ehrmann, A. Smarten up garments through knitting. IOP Conf. Ser. Mater. Sci. Eng. 2016, 141, 012008.
  109. Tyurin, I.N.; Getmantseva, V.V.; Andreeva, E.G. Van der Pauw Method for Measuring the Electrical Conductivity of Smart Textiles. Fibre Chem. 2019, 51, 139–146.
  110. Blachowicz, T.; Ehrmann, G.; Ehrmann, A. Recent Developments in Additive Manufacturing of Conductive Polymer Composites. Macromol. Mater. Eng. 2023; 2200692, early view.
  111. Archana, D.; Dutta, J.; Dutta, P.K. Evaluation of chitosan nano dressing for wound healing: Characterization, in vitro and in vivo studies. Int. J. Biol. Macromol. 2013, 57, 193–203.
  112. Mi, F.-L.; Shyu, S.-S.; Wu, Y.-B.; Lee, S.-T.; Shyong, J.-Y.; Huang, R.-N. Fabrication and characterization of a sponge-like asymmetric chitosan membrane as a wound dressing. Biomaterials 2001, 22, 165–173.
  113. Chen, X.; Wang, X.; Wang, S.; Zhang, X.; Yu, J.; Wang, C. Mussel-inspired polydopamine-assisted bromelain immobilization onto electrospun fibrous membrane for potential application as wound dressing. Mater. Sci. Eng. C. Mater. Biol. Appl. 2020, 110, 110624.
  114. Zhong, G.F.; Qiu, M.Y.; Zhang, J.B.; Jiang, F.C.; Yue, X.; Huang, C.; Zhao, S.Y.; Zeng, R.; Zhang, C.; Qu, Y. Fabrication and characterization of electrospinning nanofibers embedded with Bletilla striata polysaccharide and Rosmarinic acid to promote wound healing. Int. J. Biol. Macromol. 2023, 234, 123693.
  115. Gu, S.-Y.; Wang, Z.-M.; Ren, J.; Zhang, C.-Y. Electrospinning of gelatin and gelatin/poly(l-lactide) blend and its characteristics for wound dressing. Mater. Sci. Eng. C 2009, 29, 1822–1828.
  116. Naseri-Nosar, M.; Farzamfar, S.; Sahrapeyma, H.; Ghorbani, S.; Bastami, F.; Vaez, A.; Salehi, M. Cerium oxide nanoparticle-containing poly (ε-caprolactone)/gelatin electrospun film as a potential wound dressing material: In vitro and in vivo evaluation. Mater. Sci. Eng. C 2017, 81, 366–372.
  117. Sun, N.; Wang, G.-G.; Zhao, H.-X.; Cai, Y.-W.; Li, J.-Z.; Li, G.-Z.; Zhang, X.-N.; Wang, B.-L.; Han, J.-C.; Wang, Y.H.; et al. Waterproof, breathable and washable triboelectric nanogenerator based on electrospun nanofiber films for wearable electronics. Nano Energy 2021, 90, 106639.
  118. Lv, X.H.; Tang, Y.; Tian, Q.F.; Wang, Y.P.; Ding, T. Ultra-stretchable membrane with high electrical and thermal conductivity via electrospinning and in-situ nanosilver deposition. Compos. Sci. Technol. 2020, 200, 108414.
  119. Zhang, X.S.; Wang, B.; Wu, N.; Han, C.; Wu, C.Z.; Wang, Y.D. Flexible and thermal-stable SiZrOC nanofiber membranes with low thermal conductivity at high-temperature. J. Europ. Ceram. Soc. 2020, 40, 1877–1885.
  120. Li, Z.J.; Cheng, B.; Ju, J.G.; Kang, W.M.; Liu, Y. Development of a novel multi-scale structured superhydrophobic nanofiber membrane with enhanced thermal efficiency and high flux for membrane distillation. Desalination 2021, 501, 114834.
  121. Han, Z.Y.; Cheng, Z.Q.; Chen, Y.; Liang, Z.W.; Li, H.F.; Ma, Y.J.; Feng, X. Fabrication of highly pressure-sensitive, hydrophobic, and flexible 3D carbon nanofiber networks by electrospinning for human physiological signal monitoring. Nanoscale 2019, 11, 5942–5950.
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