Ultrathin electrospun nanofibrous membranes based on poly(γ-benzyl-L-glutamate)

Nanofibrous membranes (NfMs) composed of synthetic polymers have attracted significant attention across a diverse array of applications.^1–5 Poly(γ-benzyl-L-glutamate) (PBLG) is a synthetic polypeptide derived from glutamic acid, a member of the poly(α-amino acid) family. The properties of PBLG, including excellent biocompatibility, degradability, and intrinsic piezoelectricity, have driven extensive research focused on optimizing its potential and broadening its applicability in advanced materials science and bioengineering.^6–10 Xenofree polymers, such as PBLG, minimize risks related to immune responses and pathogen exposure while also offering enhanced reliability, quality control, and customizable properties, such as degradation rates and mechanical strength to meet specific tissue requirements.^11,12 Moreover, xenofree polymers align with ethical and regulatory guidelines, presenting lower contamination risks and extended shelf stability for safe, effective biomedical applications.¹³ The synthetic strategy of preparation of PBLG with different molecular weight allows preparing a library of specific polymers that can be tailored to specifically targeting application as nanoparticles for drug delivery.¹⁴ Porous PBLG-based scaffolds were studied for bone tissue engineering and exhibited excellent biocompatibility.¹⁵ PBLG also improved crystallization performance and wettability in PBLG/PLA fibers in carriers for a 3D model of melanoma.¹⁶ Dickinson and Hiltner investigated ways to enhance PBLG’s degradability by introducing hydrophilic groups either into the side chain or the backbone of the polymer.¹⁷ Rypacek reported that synthetic polyamino acid poly(hydroxyethyl-L-glutamine) can degrade into L-glutamic acid monomers by the action of cathepsin B, a significant enzyme in the human body.¹⁸ Fang et al. investigated the biodegradability of porous PBLG microspheres and found that the degradation rate can be tailored based on the polymer’s molecular weight, revealing an inverse linear relationship between degradation rate and molecular weight.¹⁹ Furthermore, PBLG’s secondary structure can vary depending on the type of solvent, adopting different conformations including α-helical, β-sheet, and random coil structures in solution and in the solid state.^20–23 Doty et al. found that high-molecular-weight PBLG adopts an α-helical structure when dissolved in solvents such as chloroform or dioxane, but forms a random coil conformation in dichloroacetic acid.^23,24 They concluded that specific solvent interactions with PBLG can influence its conformational stability. For example, dichloromethane, as a strong helicogenic solvent, causes PBLG helices to cluster together. However, even a small amount of TFA can disrupt these clusters, separate them into individual helices.²⁵ Minato et al. published that the addition of TFA to electrospinning solution of PBLG reduces the interactions between PBLG molecules and resulted in the production of thinner and more uniform fiber networks.²⁵ Currently, the implantation of retinal pigment epithelium (RPE) cells onto a membranous substrate that mimics Bruch’s membrane represents a promising approach in RPE cell transplantation. Bruch’s membrane, which forms the innermost layer of the choroid, has a thickness ranging from 2 to 6 μm, so it is crucial that the transplanted membrane has a comparable thickness.²⁶ A critical factor in ensuring the success of RPE cell transplantation on a solid membrane carrier is the membrane’s porosity. Given the high metabolic activity of the retina, a membrane with sufficient open porosity and larger pore sizes is essential to support RPE function and enable successful cell production and transplantation.^27–31 Recently, large studies shown that electrospun ultrathin NfMs composed of polyesters offer advantages such as high porosity, large pore size, and micrometer-scale thickness, making them effective in the field of biomedicine as an ideal support for cell therapy, particularly for the replacement of diseased eye tissue.^27–31 

Our work aims to develop ultrathin NfMs based on synthetic polypeptide PBLG for application in the eye tissue engineering. Compared to natural-based polymers, a PBLG-based carrier can offer a highly defined, xenofree alternative. Electrospun PBLG-NfMs with low thicknesses, high porosities, and favorable mechanical properties based on a novel solvent mixture and optimized parameters were examined. The key steps in the preparation, optimization, and challenges encountered during the fabrication of ultrathin PBLG-NfMs are discussed. Two solvents, 1,2-dichloroethane (DCE) and 1,3-dichloropropane (DCP), with differing properties, were tested to optimize electrospinning conditions. DCP was chosen for its lower acute toxicity compared to other halogenated solvents and for its higher boiling point relative to DCE.³² In addition, mixtures of DCE with 1% TFA and DCP with 1% TFA were prepared and tested. TFA can improve electrospinability of the both mixtures and can positively influence the final fiber structure. On the other hand, the presence of TFA traces in the final product was studied as TFA is toxic when in contact with living cells. Finally, an example of arrangement for the cell study is described.

PBLG NfMs were prepared using the electrospinning technique.³³ Polymer solutions with concentrations of 0.2, 0.3, 0.35, and 0.4 g ml⁻¹ were obtained by dissolving poly(γ-benzyl-L-glutamate) (PBLG, Mw = 201 450 g mol⁻¹, Mn = 110 380, PDI = 1.825, synthesized in our laboratory according to a previously published procedure³⁴) in two different solvents, first using 1,2-dichloroethane (DCE, 99%, M = 98.96 g mol⁻¹, bp = 83 °C, Lach-Ner, Czech Republic) and second in 1,3-dichloropropane (DCP, 99%, MW = 112.99 g mol⁻¹, bp = 122 °C, Sigma-Aldrich, USA). In some cases, 1 wt. % of trifluoroacetic acid (TFA, 99%, MW = 114.02 g mol⁻¹, Sigma-Aldrich, USA) was added to the polymer solution. The solutions were initially stirred magnetically at room temperature for 24 h until homogeneity. Afterward, the PBLG solutions were centrifuged for 30 min prior to the electrospinning procedure. The electrospinning apparatus consisted of a dispenser, with a polymer solution (KD Scientific, USA) containing a syringe fitted with a stainless steel blunt-ended needle (20G) and a DC high-voltage supply (Gamma High Voltage Research, Ormond Beach, FL 32174). During the electrospinning process, a voltage ranging from 7 to 12 kV was applied between the needle of the syringe containing the polymer solution and the collector with a gap distance from 8 to 15 cm. The polymer solution was then delivered at a flow rate ranging from 250 to 1000 μl h⁻¹. Polymeric nanofibers are produced from a liquid jet that is formed and elongated under the influence of an electric field. Formation of the Taylor cone was monitored with a video camera. Finally, the nanofibers or NfMs were collected on silicon substrates (18 × 18 mm) placed on the collector, with the process conducted at a temperature of 20–23 °C and a relative humidity of 29%–34%. Description of all samples is presented in Table I.

Scanning electron microscopy (SEM) micrographs were conducted using a Vega Plus TS 5135 (Tescan) after platinum sputter coating (4 nm) for visualization. Fiber diameter was determined using Vega software by averaging measurements of 20 randomly selected fibers across different images.

Raman spectra were recorded at room temperature with a Raman micro-spectrometer Renishaw inViaQontor. A 532 nm (2.33 eV) diode laser was used for excitation, and the scattered light was analyzed with a spectrograph featuring a holographic grating with 2400 lines mm⁻¹, achieving a spectral resolution of 1 cm⁻¹. Laser light was focused using a 100× objective lens in the pseudo-confocal mode, resulting in a spot size of 0.5 μm. To ensure accuracy and reproducibility, each Raman spectrum was acquired by averaging multiple measurements taken from different locations on the sample.

Fourier transform infrared spectra (FTIR) in region 4000–650 cm⁻¹ were recorded using a Thermo Nicolet NEXUS 870 FTIR spectrometer (MCT/A detector; 256 scans; resolution 2 cm⁻¹) equipped with the GoldenGate ATR accessory. The spectra were corrected for the carbon dioxide and humidity in the optical path.

X-ray photoelectron spectroscopy (XPS) was performed using a K-Alpha⁺ spectrometer (ThermoFisher Scientific, East Grinstead, UK). Data acquisition and processing were performed with Thermo Avantage software. The NfMs were analyzed using a micro-focused, monochromated Al Kα x-ray source at an angle of incidence of 30° (measured from the surface) and an emission angle normal to the surface. The kinetic energy of the electrons was measured using a 180° hemispherical energy analyzer operated in the constant analyzer energy mode at 200 and 50 eV pass energy for the survey and high-resolution spectra, respectively. Spectral resolutions of 1.0 and 0.1 eV were used for the survey and high-resolution spectra, respectively. Surface charge compensation was applied during the measurements. All the reported XPS spectra are averages of 10 individual measurements, referenced to the C1s band of hydrocarbons at 285.0 eV. For quantification, the analyzer transmission function, Scofield sensitivity factors, and effective attenuation lengths (EALs) for photoelectrons were used. EALs were calculated using the standard TPP-2M formalism. The binding energy (BE) scale was controlled by the well-known position of the photoelectron C–C and C–H as well as C–O and C(=O)–O C 1s bands of polyethylene terephthalate and Cu 2p, Ag 3d, and Au 4f bands of metallic Cu, Ag, and Au, respectively. The BE uncertainty for the reported measurements and analysis is in the range of ±0.1 eV. The obtained XPS spectra were fitted with Voigt profiles obtained by convolving Lorentzian and Gaussian functions to determine the amounts in atomic % of individual chemical species present on the analyzed surfaces. The reported values are averages from six different positions on the sample.

Nanofibrous membrane thicknesses were measured using a KLA-Tencor P-17 profilometer (KLA-Tencor Corporation, Milpitas, CA, USA) after platinum sputter coating of 4 nm. The thickness was evaluated as the average of all the highest bands in the profile along a 1000 μm long line scan with the baseline referenced on the silicon substrate. The areal density of NfMs was calculated from the silicon wafer area and related mass of the electrospun membrane.

Mechanical testing of PBLG NfMs was carried out using a small punch device (ARES-G2, TA Instruments, USA). The samples were securely mounted on metal washers with an outer diameter of 18 mm, an inner diameter of 6.6 mm, and a thickness of 1.6 mm. A hemispherical punch with a head radius of 1.5 mm was employed for the tests. These experiments were conducted under low force conditions (<1 N) at a constant linear displacement rate of 0.016 67 mm/s (1 mm/min) until failure occurred. All the tests were performed at a controlled room temperature of 23 ± 2 °C, with both punch load and displacement continuously recorded throughout the testing process.

For tissue engineering application, the PBLG-based NfM can be prepared in the form of cultivation inserts for cell seeding. The sheets of the PBLG-based NfMs were fixed by silicon glue to the body of cultivation cup Falcon (Corning Inc., Kenneburg, USA), which was devoid of the original commercial membrane according to the procedure we published earlier.³⁰ To ensure smooth surface of the NfM even after cutting the sample from the insert, a circle frame was inserted during the electrospinning into the nanofibrous structure.²⁶ The circles (4 mm in diameter) were cut by laboratory-assembled laser microfabric station from a poly(ethylene terephthalate) foil (thickness 36 μm).

In order to assess the quality of the prepared nanofibers, scanning electron microscopy (SEM) was conducted. Figures 1(a)–1(c) shows the SEM images of NF_DCE-0.2, NF_DCE-0.3, and NF_DCE-0.4 samples, prepared from PBLG solutions in DCE, with concentrations of 0.2, 0.3, to 0.4 g ml⁻¹ (denoted as C_DCE-0.2, C_DCE-0.3, and C_DCE-0.4, respectively). The nanofibers of the NF_DCE-0.2 sample prepared at the lowest concentration (C_DCE-0.2) exhibited the presence of beads. In the NF_DCE-0.3 sample prepared at the intermediate concentration (C_DCE-0.3), fibers with diameters greater than 5 μm were obtained and beads were no longer present. However, in the NF_DCE-0.4 sample prepared at the highest concentration (C_DCE-0.4), fiber production was hindered due to needle blockage during electrospinning, which was caused by the high viscosity of the polymeric solution. To enhance the electrospinning process, we repeated the experiments with the addition of 1 wt. % TFA to the PBLG solutions. TFA disrupts intermolecular interactions in the polymer, which reflects in the smooth flow of the electrospun solution and improved fiber formation during the electrospinning process. Moreover, TFA is highly volatile and accelerates solvent evaporation during electrospinning, which results in the formation of uniform nanofibers.³⁵ Other advantages of using TFA in the electrospinning process are its high dielectric constant (resulting in higher conductivity of the solution) and also miscibility with a wide range of organic solvents.^36,37 Figures 1(d)–1(f) shows the SEM images of NF_DCE-TFA-0.2, NF_DCE-TFA-0.3, and NF_DCE-TFA-0.4 samples prepared from TFA-enhanced solutions, with concentration of 0.2, 0.3, and 0.4 g ml⁻¹ (denoted as C_{DCE- TFA-0.2}, C_DCE-TFA-0.3, and C_DCE-TFA-0.4, respectively).

It can be observed that the issue of bead formation persists in the NF_DCE-TFA-0.2 sample prepared with the lowest concentration (C_DCE-TFA-0.2). For the NF_DCE-TFA-0.3 and NF_{DCE-TFA 0.4} samples prepared from higher concentrations (C_DCE-TFA-0.3 and C_DCE-TFA-0.4, respectively), maintaining a stable Taylor cone for a sufficient duration to enable membrane preparation proved challenging.

This difficulty remained despite modifications to various electrospinning parameters, including the needle-to-collector gap, voltage, and flow rate. In addition, SEM images indicate that an increase in concentration is accompanied by a significant rise in fiber diameters. The same experiments as previously performed with the DCE solvent were then repeated using a different solvent, DCP. During the electrospinning process with DCP, the stability time of the Taylor cone was observed, increasing from 10 to 60 s compared to the previous experiments. In addition, the DCP solution allowed the electrospinning to be performed at lower flow rates, around 600 μl h⁻¹, compared to the higher flow rates of 1000 μl h⁻¹ used with DCE solutions. Figures 2(a)–2(c) shows the SEM images of NF_DCP-0.2, NF_DCP-0.3, and NF_DCP-0.4 samples, prepared using the DCP solvent with concentrations of 0.2, 0.3, to 0.4 g ml⁻¹ (denoted as C_DCP-0.2, C_DCP-0.3, and C_DCP-0.4 respectively).

SEM images show that the lowest C_DCP-0.2 concentration yields nanofibers with beads, while higher concentrations of C_DCP-0.3 and C_DCP-0.4 produce bead-free fibers. The fibers exhibited good morphological quality compared to those prepared with DCE solvents. Furthermore, fiber diameters were reduced to ∼2 μm. After the addition of TFA, the stability of the Taylor cone improved from 60 to 600 s, and the flow rate was adjusted to low values up to 300 μl h⁻¹, which led to a decrease in fiber diameters to as low as 0.6 μm. Figures 2(d)–2(f) show the SEM images of NF_DCP-TFA-0.2, NF_DCP-TFA-0.3, and NF_DCP-TFA-0.4 samples prepared from these TFA-enhanced solutions, with concentration of 0.2 and 0.3–0.4 g ml⁻¹ (denoted as C_{DCP- TFA-0.2}, C_DCP-TFA-0.3, and C_DCP-TFA-0.4 respectively). Based on SEM analysis, it was found that when concentration (C_{DCP- TFA-0.2}) was used, the formed nanofibers in the NF_DCP-TFA-0.2 sample showed a slight presence of beads compared to the NF_DCP-TFA-0.3 sample, and small-diameter compared to the NF_DCP-TFA-0.4 sample. During tests with C_DCP-TFA-0.3 concentration, increasing the flow rate from 300 to 1000 μl h⁻¹ led to an increase in mean diameter values from 0.6 to 1.1 μm. To ensure the complete elimination of beads, an optimized solution of PBLG in DCP + 1 wt. % TFA with concentration of 0.35 g ml⁻¹, denoted C_0.35, was selected and new optimized parameters were established, as presented in Table II.

In order to produce small-diameter fibers ranging from 300 to 500 nm, which are ideal for tissue engineering applications, a low flow rate of 250 μl h⁻¹ was used. Figure 3 shows SEM observations made on the optimized sample NfM prepared with the concentration C_0.35.

Further SEM micrographs (Fig. 3) of NfM sample revealed the absence of beads and structural defects, along with significant uniformity in the size and shape of the nanofibers when using DCP with 1 wt. % of TFA solution with the optimized parameters. Detailed observations of the fiber diameter distribution^38,39 showed that the average diameter of the nanofibers was 0.53 ± 0.01 μm. These findings confirm the effectiveness of the optimized concentration and electrospinning parameters.

In order to elucidate the conformation of PBLG, Raman spectroscopy analysis was performed not only in NfMs, but also in PBLG with DCE or DCP solutions without and with TFA at room temperature. The Raman spectra of the PBLG solutions are presented in Fig. 4. The results of these analyses highlighted distinctive spectral features of PBLG, such as amide II band at 1555 cm⁻¹. Two ring stretching bands or two aromatic vibrations (Aro₁ and Aro₂) are associated with phenyl groups at 1587 and 1608 cm⁻¹ respectively. The amide I band at 1653 cm⁻¹ primarily arises from the vibrations of the C=O (carbonyl) bond in the amide groups. The ester stretching vibrations band at 1732 cm⁻¹ arises from the vibrations of the C=O bond in the ester groups. The amide A band is associated with the N–H stretching vibrations at 3292 cm⁻¹.

By focusing on the frequency of the amide I band in the Raman spectra of samples prepared from DCE and DCP, it was observed that the band typically seen around 1630 cm⁻¹, which is known to be sensitive to different conformations, is absent. This indicates that the β-sheet conformation is not present. Conversely, the amide I band remained constant at 1652 cm⁻¹, indicating that our PBLG samples primarily adopt an α-helix conformation at room temperature. These observations confirm the sensitivity of PBLG to environmental conditions, particularly the choice of solvent, in determining its structural conformation. This indicates that there is a weak interaction between PBLG and the solvents used.

On the other hand, the comparison between normalized Raman spectra showed that the addition of 1 wt. % TFA does not affect the positions of the Raman bands, indicating the preservation of the α-helix conformation. For the Raman spectra of the PBLG in DCP and PBLG in DCP + 1 wt. % TFA solutions, it is also observed that there is a slight variation in the intensity ratios amide I/Aro₁ from 1.62 to 1.85 and amide A/Aro₁ from 0.95 to 1.15, and it can also be seen that the amide A band is the most sensitive, showing an increase in both intensity and integrated intensity (area) following the addition of TFA. This variation of bands profiles could be attributed to the disruption of aggregates of α-helices in response to the presence of the TFA.

The NfM samples were dried for 24 h at 27 °C under vacuum before performing the Raman measurements. The Raman spectra of the membranes shown in Fig. 5 illustrate the lack of change in the intensity ratios and areas of the Raman bands. This lack of change confirms that the effects of TFA on PBLG in the solid state are preserved. On the other hand, the amide I band shifts to lower frequencies from 1653 cm⁻¹ (in the liquid state) to 1651 cm⁻¹ (in the solid state), while maintaining the same full width at half maximum (FWHM) value of 12 cm⁻¹. When the PBLG solution transforms into a membrane, there is indeed a change in the state of matter, from a liquid phase to a solid phase. Solidification leads to a more compact, ordered, and much dense structure. The molecules in the nanofibers are closer together and better organized than in the solution. In Raman spectroscopy, vibrations can be affected by the compactness and organization of the material. In a more compact material, intermolecular interactions are stronger, which can modulate the observed vibration frequencies.

In order to ensure that the solvent was evaporated properly, the samples were subjected to annealing for two hours under vacuum at a temperature of 70 °C. Analysis of the spectra before and after annealing revealed a decrease in intensity ratios (amide I/Aro₁) from 1.87 to 1.79 and (amide A/Aro₁) from 1.08 to 0.96. This decrease was particularly prominent in the amide A band, which also showed a decrease in integrated intensity following annealing (Fig. 6).

It is very important here to note that there is a reversal of behavior contrasts with what was observed in the spectrum of the PBLG in DCP solution after the addition of TFA. Consequently, this reduction was attributed to intensity ratio and area to the evaporation of TFA following annealing. X-ray photoelectron spectroscopy (XPS) characterizations were carried out to confirm our hypothesis. In addition, the characteristic signature of the α-helix conformation was observed, indicating that the PBLG nanofibers exhibited remarkable thermal stability at the chosen annealing temperature, which did not affect the quality of the nanofibers.

Fourier-transform infrared spectroscopy (FTIR) analyses were also conducted to thoroughly evaluate the structural properties and interactions within samples.^40,41 Regarding the conformation of the secondary structure of PBLG, the presence of the amide I band around 1650 cm⁻¹ in the FTIR spectra confirms the predominance of the α-helical conformation (Fig. 7). These results are consistent with observations from Raman spectroscopy, which does not reveal a band around 1630 cm⁻¹, typical of β-sheet structures. Furthermore, the impact of adding TFA is clearly manifested in the FTIR spectra, where variations in absorption intensity indicate changes in molecular interactions. In addition, Fig. 8 shows the decreased intensity of amide I and amide A after annealing, as previously observed in Raman spectra.

XPS analysis was employed to investigate the chemical composition of the NfM particularly concerning the presence of TFA. Figure 9 shows the changes in composition before and after thermal treatment for two hours under vacuum at a temperature of 70 °C.

The high resolution XPS spectra taken in the C 1s region, show C–C and C–H (from the aliphatic and phenyl groups) at 285.0 eV, C–O and C–N contributions at 286.5 eV, C(=O)–NH amide group at 288.2 eV, and C(=O)–O ester contribution at 289.0 eV. The spectra taken in the N 1s region show one peak at 400.2 eV, originating from the amide NH–C(=O). In the O 1s region, the spectra of the membranes can be deconvoluted with two contributions characteristic for the O=C and C–O moieties at 532.1 and 533.5 eV. The presence of amide and ester groups was further verified in the high-resolution spectra taken in the N 1s and O 1s regions. The band at 291.6 eV is a shake-up satellite arising from π → π* transition in the phenyl rings. Its appearance shields the observation of the possible traces CF₃ groups from TFA. Nevertheless, the F 1s spectrum of the non-thermally annealed structures points to the presence of TFA before the treatment. The low observed amounts of TFA in the XPS spectra can also be related to the fact that the XPS measurements are performed under high vacuum conditions. However, the thermal annealing leads to complete disappearance of the fluorine signals (Table III) and preservation of the chemical structure of the nanofibers.

In order to determine the thickness of the nanofibrous membrane, profilometric measurements were conducted. Figure 10 shows the variation of the average value of thickness for the NfM samples prepared with the times of 30, 90, and 125 s.

The average values of the thickness (D_avr) are presented in Table IV. The average value of the membrane thickness increases with the preparation time.

where d_s is the density of the porous structure (g cm⁻³) and d is the density of the structure material = 1.19 g cm⁻³.

Table IV shows that as the surface density increases (corresponding to an increase in thickness), there is a slight decrease in membrane porosity. This inverse relationship between porosity and thickness is due to increased fiber compaction, which reduces the available interstitial space as the thickness of the membrane increases.

The mechanical properties of NfMs are inherently linked to their specific structure and architecture, tailored to meet the demands of various applications.^42–44 Uniaxial stress–strain testing is a widely used technique for assessing the mechanical properties of NfMs.^26,45 However, due to the membrane’s thickness being less than 10 μm, handling larger samples for this type of test proved impractical.²⁶ Therefore, we opted for a small punch test, which effectively evaluates the mechanical properties of miniature specimens in a biaxial configuration.^46,47 Figure 11 shows the punch load-displacement curves for the NfM₃₀, NfM₉₀, and NfM₁₂₅ samples.

A key parameter derived from the slope of the linear portion of these curves is stiffness, which indicates the membrane’s capacity to resist deformation under applied forces. The data show a clear relationship between membrane thickness and stiffness, as thicknesses increased from 1.7 to 4.5 μm. In particular, the slope (a) of the tangent to the linear segment of the curves increased significantly, moving from 0.016 to 0.13. This corresponds to a notable enhancement in average stiffness, with values rising from 16 ± 2 N m⁻¹ for NfM₃₀, to 51 ± 2 N m⁻¹ for NfM₉₀, and reaching 130 ± 2 N m⁻¹ for NfM₁₂₅. The measured stiffness values of ultrathin PBLG-based nanofibrous membranes are in accordance with polylactide-based electrospun membranes of the similar thicknesses from our previous work.²⁶ 

During biological testing (cell staining), it is often necessary to cut the sample with cultivated cells from the cultivation insert. However, ultrathin NfM collapses when wet after cutting and it is really difficult to manipulate.

By including a circle frame in the electrospun structure, the sample with cultivated cells can be easily cut by biopsy punch around the frame from the insert and transferred to the Petri dish. Such a setting will allow not only easy manipulation but also can help protect cultivated cells from possible sample bending or rolling. In Fig. 12(a), the design of cultivation inserts with PBLG-based ultrathin NfMs of different thicknesses is shown. Even the membrane with the lowest thickness 1.7 μm was suitable for the circle frame incorporation. The detail of the cultivation inserts with 3.8 μm thick NfM is shown in Fig. 12(b) and the detail of the structure of the same sample observed by SEM (nanofibers and part of the circle frame) in Fig. 12(c). Such an arrangement of ultrathin NfMs could find application in specific branches of tissue engineering, e.g., in regeneration of the eye.

In summary, our study focused on the preparation and characterization of electrospun PBLG NfMs. By carefully selecting solvents and optimizing parameters, we successfully produced high-quality, bead-free membranes, as evidenced by SEM observations. Raman spectroscopy provided valuable insights into the conformational dynamics of PBLG in both solution and solid state, revealing the predominance of an α-helical conformation. The addition of 1 wt. % TFA improved the electrospinning process with minimal impact on the Raman and FTIR band profiles, confirming the preservation of the α-helical conformation. Furthermore, annealing at 70 °C resulted in the evaporation of TFA, as confirmed by XPS, while maintaining the α-helical conformation. The small punch test demonstrated a strong relationship between membrane thickness and mechanical stiffness, suggesting that thicker membranes exhibit higher toughness. Finally, application of NfM equipped with a supporting frame for cell cultivation arrangement was presented. The comprehensive analysis described in this study deepens our understanding of PBLG nanofiber-based membranes. Future studies such as biological tests are required to further enhance these chemical and physical properties, establishing a critical foundation for assessing their suitability and performance in tissue engineering applications.

This study was supported by the European Regional Development Fund (Grant No. CZ.02.01.01/00/22_008/0004562 Project “Excellence in Regenerative Medicine”).

The authors have no conflicts to disclose.

M. Souibgui: Conceptualization (equal); Data curation (equal); Investigation (equal); Methodology (equal); Validation (equal); Writing – original draft (equal). Z. Morávková: Data curation (equal); Formal analysis (equal); Writing – original draft (equal). O. Pop Georgievski: Data curation (equal); Formal analysis (equal); Writing – original draft (equal). J. Hodan: Data curation (equal); Formal analysis (equal). M. A. Thottappali: Formal analysis (equal). V. Cimrová: Data curation (equal); Writing – original draft (equal). J. Dvořáková: Formal analysis (equal); Writing – original draft (equal). V. Proks: Writing – original draft (equal). H. Studenovska: Conceptualization (equal); Investigation (equal); Methodology (equal); Supervision (equal); Validation (equal); Writing – original draft (equal).

The data that support the findings of this study are available from the corresponding author upon reasonable request.