Iα and Iβ components being present, which is confirmed by the presence of weak absorption bands at both 744 and 710 cm−1. This can be assigned to the glucose ring deformations, compounded with glycosidic bond bending of the respective cellulose structure modifi-

**Figure 4.** FTIR absorbance spectra of the BC-AD and BC-L unmodified and plasma-modified sam-

sample is usually attributed to the 2O-H···6O-H···3O hydrogen bond group [55].

ture differently than in the air-dried samples.

The OH band around 3350 cm−1 combines the vibrations of the hydrogen bonds in cellulose. The absorption at 3235 cm−1, which is slightly stronger in the case of the BC-AD

The absorption in the 1700–1500 cm−1 range relates to adsorbed water [56]. While all the BC-AD samples show a single weak absorption at 1632 cm−1, in the BC-L samples the absorption is noticeably stronger and the absorbed range is wider, in certain cases with a dominant second absorption maximum at 1593 cm−1. This indicates there is a higher amount of adsorbed water in the BC-L samples and the water binds to the cellulose struc-

The absorption band at 1315 cm−1, which can be assigned to C-O-H bending vibrations [57], is relatively stronger in the BC-AD samples. The only absorption band where a

*3.3. Chemical Composition of BC*

cations [54].

ples.

**Table 1.** Gravimetric analysis of the BC weight loss after PM for different exposure times.

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#### *3.3. Chemical Composition of BC 3.3. Chemical Composition of BC*

FTIR spectra of the BC-AD and BC-L samples unmodified and after 60 s and 240 s PM are shown in Figure 4. Spectral interpretation and band assignment in cellulose is considered to be somewhat problematic due to the dominant role of inter- and intrachain hydrogen bond interactions that lead to numerous combination vibrations [54]. However, some general features of the material can be ascribed to the absorption bands in the present spectra. The spectra of the prepared BC correspond to cellulose I structure with both Iα and Iβ components being present, which is confirmed by the presence of weak absorption bands at both 744 and 710 cm−1 . This can be assigned to the glucose ring deformations, compounded with glycosidic bond bending of the respective cellulose structure modifications [54]. FTIR spectra of the BC-AD and BC-L samples unmodified and after 60 s and 240 s PM are shown in Figure 4. Spectral interpretation and band assignment in cellulose is considered to be somewhat problematic due to the dominant role of inter- and intra-chain hydrogen bond interactions that lead to numerous combination vibrations [54]. However, some general features of the material can be ascribed to the absorption bands in the present spectra. The spectra of the prepared BC correspond to cellulose I structure with both Iα and Iβ components being present, which is confirmed by the presence of weak absorption bands at both 744 and 710 cm−1. This can be assigned to the glucose ring deformations, compounded with glycosidic bond bending of the respective cellulose structure modifications [54].

**Figure 4.** FTIR absorbance spectra of the BC-AD and BC-L unmodified and plasma-modified sam-**Figure 4.** FTIR absorbance spectra of the BC-AD and BC-L unmodified and plasma-modified samples.

ples. The OH band around 3350 cm−1 combines the vibrations of the hydrogen bonds in cellulose. The absorption at 3235 cm−1, which is slightly stronger in the case of the BC-AD The OH band around 3350 cm−1 combines the vibrations of the hydrogen bonds in cellulose. The absorption at 3235 cm−1 , which is slightly stronger in the case of the BC-AD sample is usually attributed to the 2O-H···6O-H···3O hydrogen bond group [55].

sample is usually attributed to the 2O-H···6O-H···3O hydrogen bond group [55]. The absorption in the 1700–1500 cm−1 range relates to adsorbed water [56]. While all the BC-AD samples show a single weak absorption at 1632 cm−1, in the BC-L samples the absorption is noticeably stronger and the absorbed range is wider, in certain cases with a dominant second absorption maximum at 1593 cm−1. This indicates there is a higher amount of adsorbed water in the BC-L samples and the water binds to the cellulose struc-The absorption in the 1700–1500 cm−1 range relates to adsorbed water [56]. While all the BC-AD samples show a single weak absorption at 1632 cm−1 , in the BC-L samples the absorption is noticeably stronger and the absorbed range is wider, in certain cases with a dominant second absorption maximum at 1593 cm−1 . This indicates there is a higher amount of adsorbed water in the BC-L samples and the water binds to the cellulose structure differently than in the air-dried samples.

ture differently than in the air-dried samples. The absorption band at 1315 cm−1, which can be assigned to C-O-H bending vibrations [57], is relatively stronger in the BC-AD samples. The only absorption band where a The absorption band at 1315 cm−1 , which can be assigned to C-O-H bending vibrations [57], is relatively stronger in the BC-AD samples. The only absorption band where a consistent shift with the PM can be observed was the 1160 cm−1 which shifts about 2 cm−1 to lower wavenumber in samples after PM. Moreover, this band appeared on average about 20% weaker in those samples. These changes are in the scale of the whole spectrum rather insignificant; however, this can be expected, since PM mainly influences the very surface layer of the material and the FTIR signal is obtained over the sample depth of several micrometers. Nonetheless, the variation of this absorption band could mean the

glycosidic bond and the cellulose chains have been to some degree disrupted by the PM. The air-dried samples also show weaker maxima at 1005 and 987 cm−1 (bands related to vibrations of C-O bonds in the hydroxyl groups), while in the lyophilized samples these absorptions show merely as shoulders of the stronger bands. Especially in the case of the air-dried samples, the absorption at 1005 cm−1 shows very high variability, even within repeated measurements on a single sample. In the case of the BC-L samples with PM, a weak band was observed around 800 cm−1 . Absorptions in this region are usually assigned to deformations of the glucose ring coupled with bending of the glycosidic bond. Therefore, this could indicate the influence of PM on the glycosidic bond in the BC samples.

XPS measurements showed the changes in the surface composition, especially in the carbon and oxygen content. The samples were somewhat contaminated with Al, Si, F, S, Na, and N from the plasma vacuum chamber, and the bacteria and medium residues. The results shown in Table 2 are recalculated without the contaminating elements. The expected value for O/C from the chemical structure of cellulose is 0.83. For unmodified samples, this value is slightly lower, which is caused by the residues of media and bacteria on the surface of the material. The most significant result is the large increase in oxygen content after PM of BC-L 240 s. This could be caused by the disruption of the cellulose structure leading to the formation of highly reactive species on the surface, which react with ambient oxygen after the exposure of the modified surface to atmosphere. The increase in oxygen content after PM for BC-AD is also apparent.

**Table 2.** Carbon and oxygen atomic concentrations of BC samples measured by XPS with 0◦ detection angle.

#### *3.4. Surface Morphology of BC*

The SEM images in Figure 5 show the nanostructured morphology of unmodified BC-AD and BC-L (transparent and white part) samples. The differences between BC-AD and BC-L samples can be seen in the porosity of the material. For the L samples, the fibers are more spread out. This corresponds with the fact that the lyophilized pellicles are thicker. Illa [47], who compared lyophilization and oven drying, suggested that this phenomenon is caused by the free hydroxyl groups that can form secondary bonds. The mobility of the amorphous regions during freezing is reduced and therefore the morphology is preserved. However, during oven drying the thermal energy maintains the mobility of the amorphous chains and therefore the morphology collapses, the fibers come closer together, and those samples are thinner. Air drying at room temperature could have similar effect on the material morphology. Together with the smaller thickness of the material compared to BC-L, SEM shows that BC-AD has a more rugged surface structure with the bent fibers turned perpendicular to its surface. The differences between SEM images of BC-L transparent and BC-L white were studied as well. Those nanofibers were found to be indistinguishable in the SEM images for both samples. The plasma-modified samples did not show any differences in these areas either, so we can suppose that the thickness of the lyophilized pellicles does not affect the nanofibrillar structure and it is determined by the bacterial origin of the material.

*Nanomaterials* **2021**, *11*, x FOR PEER REVIEW 9 of 17

by the bacterial origin of the material.

by the bacterial origin of the material.

[44].

**Figure 5.** Nanostructure images of BC samples analyzed by SEM: (**a**) BC-AD, (**b**) BC-L transparent, (**c**) BC-L white. **Figure 5.** Nanostructure images of BC samples analyzed by SEM: (**a**) BC-AD, (**b**) BC-L transparent, (**c**) BC-L white. **Figure 5.** Nanostructure images of BC samples analyzed by SEM: (**a**) BC-AD, (**b**) BC-L transparent, (**c**) BC-L white.

It has been demonstrated that PM can change the material surface morphology, especially the surface roughness and porosity. This can lead to a material with different surface properties such as hydrophilicity [58]. The SEM analysis revealed ablation occurring, which is caused by surface etching during PM (Figure 6). The cellulose fibers appeared to be rugged and thinner after PM. Observed surface-terminated pores were bigger in the case of BC-AD. Those pores were wider and deeper with higher PM exposure time. The pores of BC-AD 240 s merged, forming a "brush" from the remaining parts of the fibers on surface. This very different structure is in agreement with the fact that the BC-AD 240 s sample was modified to the highest degree of the compared samples based on the gravimetric analysis. This is caused by the higher exposure time (compared to BC-AD 60 s) and by the fact that fibres in the air-dried sample are closer to each other than in the lyophilized ones. These results are in agreement with those published by Pertile et al. It has been demonstrated that PM can change the material surface morphology, especially the surface roughness and porosity. This can lead to a material with different surface properties such as hydrophilicity [58]. The SEM analysis revealed ablation occurring, which is caused by surface etching during PM (Figure 6). The cellulose fibers appeared to be rugged and thinner after PM. Observed surface-terminated pores were bigger in the case of BC-AD. Those pores were wider and deeper with higher PM exposure time. The pores of BC-AD 240 s merged, forming a "brush" from the remaining parts of the fibers on surface. This very different structure is in agreement with the fact that the BC-AD 240 s sample was modified to the highest degree of the compared samples based on the gravimetric analysis. This is caused by the higher exposure time (compared to BC-AD 60 s) and by the fact that fibres in the air-dried sample are closer to each other than in the lyophilized ones. Theseresults are in agreement with those published by Pertile et al. [44]. pecially the surface roughness and porosity. This can lead to a material with different surface properties such as hydrophilicity [58]. The SEM analysis revealed ablation occurring, which is caused by surface etching during PM (Figure 6). The cellulose fibers appeared to be rugged and thinner after PM. Observed surface-terminated pores were bigger in the case of BC-AD. Those pores were wider and deeper with higher PM exposure time. The pores of BC-AD 240 s merged, forming a "brush" from the remaining parts of the fibers on surface. This very different structure is in agreement with the fact that the BC-AD 240 s sample was modified to the highest degree of the compared samples based on the gravimetric analysis. This is caused by the higher exposure time (compared to BC-AD 60 s) and by the fact that fibres in the air-dried sample are closer to each other than in the lyophilized ones. These results are in agreement with those published by Pertile et al. [44].

It has been demonstrated that PM can change the material surface morphology, es-

did not show any differences in these areas either, so we can suppose that the thickness of the lyophilized pellicles does not affect the nanofibrillar structure and it is determined

did not show any differences in these areas either, so we can suppose that the thickness of the lyophilized pellicles does not affect the nanofibrillar structure and it is determined

**Figure 6.** BC before and after plasmatic modification as analyzed by SEM: (**a**) BC-AD, (**b**) BC-AD 60 s, (**c**) BC-AD 240 s, (**d**) BC-L, (**e**) BC-L 60 s, (**f**) BC-L 240 s. **Figure 6.** BC before and after plasmatic modification as analyzed by SEM: (**a**) BC-AD, (**b**) BC-AD 60 s, (**c**) BC-AD 240 s, (**d**) BC-L, (**e**) BC-L 60 s, (**f**) BC-L 240 s. **Figure 6.** BC before and after plasmatic modification as analyzed by SEM: (**a**) BC-AD, (**b**) BC-AD 60 s, (**c**) BC-AD 240 s, (**d**) BC-L, (**e**) BC-L 60 s, (**f**) BC-L 240 s.

#### *3.5. Gas Sorption Analysis of Porosity and Specific Surface Area*

Specific surface area of unmodified samples and samples with 240 s PM were measured by gas sorption method. These results (Table 3) are in good agreement with the SEM

observations. Firstly, compared to BC-AD, BC-L has almost five times higher porosity and specific surface area, because it maintains its structure (pellicle thickness) during the drying process. Further increase of pore volume and surface area was obtained after PM. This is due to PM fraying of the cellulose fibres. The number and size of the pores increases dramatically as well. Based on SEM images, we assumed the increase of porosity after PM for BC-AD would be greater because of the "brush" structure, which was confirmed by the BJH analysis—the porosity increased almost 13 times for BC-AD, and 6 times for BC-L. Thus, after PM the specific surface area of the BC-L and BC-AD reached comparable values. On the basis of these results, we can conclude that PM in combination with the drying process leads to the increase of pores and surface area, which results in significant changes in surface morphology which can improve cell adhesion.

**Table 3.** Specific surface area (SBET) and total pore volume (Vp) of BC samples measured by BET analysis.

#### *3.6. Contact Angle and Hydrophilicity*

The new morphology and higher oxygen content after PM can lead to changes in hydrophilicity of the material surface that can be represented by the water contact angle [59].

The contact angle measurement results (Table 4) showed that BC-AD is more hydrophobic than BC-L. The contact angles for those samples are (63.91 ± 2.69)◦ and (34.74 ± 6.8) ◦ , respectively. These differences can be attributed to the different specific surface area, which is smaller for BC-AD. For hydrophilic materials (contact angle less than 90◦ ) the higher porosity leads to a lower contact angle. This is due to the water drop being absorbed into the pores of a hydrophilic material. After PM, we observed further decrease of the contact angle that was greater in the case of BC-AD; both samples showed similar values after PM. This is consistent with much higher porosity increase for plasma-modified BC-AD compared to BC-L (Table 3). Additionally, an increase of surface oxygen content after PM represented by the O/C in XPS measurements (Table 2) leads to a more hydrophilic material. For comparison, Kurniawan et al. [45] also observed decrease of contact angle after PM with N2 and O2 plasmas while for CF4 plasma there was an increase. However, Pertile et al. [44] noticed an increase of contact angle after N2 PM. It is useful to note that each research group used different plasma discharge parameters.

**Table 4.** Values of water contact angle of BC samples measured by goniometry.

#### *3.7. In Vitro Tests of Cell Cultivation on BC Samples*

For in vitro tests, we chose the BC-AD and BC-L samples, unmodified and after 240 s PM, in order to evaluate the effect of air drying, lyophilization, and PM on the colonization of the samples with human HaCaT keratinocytes.

On day 1 after seeding, the HaCaT cells on BC adhered generally in higher numbers than on the control PS wells, which was, however, more pronounced in BC-L than in BC-AD (Figure 7a). This result can be attributed to a higher porosity and a larger specific surface area of BC-L, which therefore provided more space for the initial cell attachment than BC-AD. Nevertheless, the initial adhesion of cells on BC-AD was significantly improved by PM, i.e., a technique which is generally used to enhance the attractiveness of various materials for cell adhesion. The main underlying mechanism of this improvement is increase in the material hydrophilicity, manifested by a significant decrease of water drop contact angle (from approx. 64◦ to 25◦–33◦ ; Table 4). On wettable materials, the cell adhesion-mediating proteins, such as fibronectin and vitronectin, spontaneously adsorb to the materials from biological fluids (including cell culture media), are attached in an active, physiological conformation, and are well-accessible for cell adhesion receptors (e.g., integrins) on cells [10,59,60]. BC-L was sufficiently wettable even before PM (contact angle of approx. 35◦ ), and thus the PM did not further increase significantly the number of initially adhered cells, as observed on BC-AD samples. Moreover, the BC-AD samples modified with plasma were the first substrates on which the HaCaT cells started to form well-apparent and distinct islands typical for keratinocytes, which are important initial structures for creating a continuous cell layer (Figure 8). than on the control PS wells, which was, however, more pronounced in BC-L than in BC-AD (Figure 7a). This result can be attributed to a higher porosity and a larger specific surface area of BC-L, which therefore provided more space for the initial cell attachment than BC-AD. Nevertheless, the initial adhesion of cells on BC-AD was significantly improved by PM, i.e., a technique which is generally used to enhance the attractiveness of various materials for cell adhesion. The main underlying mechanism of this improvement is increase in the material hydrophilicity, manifested by a significant decrease of water drop contact angle (from approx. 64° to 25°–33°; Table 4). On wettable materials, the cell adhesion-mediating proteins, such as fibronectin and vitronectin, spontaneously adsorb to the materials from biological fluids (including cell culture media), are attached in an active, physiological conformation, and are well-accessible for cell adhesion receptors (e.g., integrins) on cells [10,59,60]. BC-L was sufficiently wettable even before PM (contact angle of approx. 35°), and thus the PM did not further increase significantly the number of initially adhered cells, as observed on BC-AD samples. Moreover, the BC-AD samples modified with plasma were the first substrates on which the HaCaT cells started to form well-apparent and distinct islands typical for keratinocytes, which are important initial structures for creating a continuous cell layer (Figure 8).

For in vitro tests, we chose the BC-AD and BC-L samples, unmodified and after 240 s PM, in order to evaluate the effect of air drying, lyophilization, and PM on the coloniza-

On day 1 after seeding, the HaCaT cells on BC adhered generally in higher numbers

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*3.7. In Vitro Tests of Cell Cultivation on BC Samples*

tion of the samples with human HaCaT keratinocytes.

**Figure 7.** Number (**a**–**c**) and viability (**d**) of human HaCaT keratinocytes on days 1, 4 and 7 after seeding on control polystyrene wells (PS) or on BC. The data were obtained by (**a**) counting cells on 18–23 microscopic images, (**b**,**c**) measuring the intensity of fluorescence of cells stained with Hoechst 33258 on 7–20 microscopic images, or (**d**) trypan blue-exclusion test from four parallel samples of each experimental group. Mean ± S.E.M. (Standard Error of Mean), One way ANOVA, Student-Newman-Keuls Method. Statistically significant differences among the experimental groups (*p* ≤ 0.05) are indicated above the columns by numbers of differing groups. **Figure 7.** Number (**a**–**c**) and viability (**d**) of human HaCaT keratinocytes on days 1, 4 and 7 after seeding on control polystyrene wells (PS) or on BC. The data were obtained by (**a**) counting cells on 18–23 microscopic images, (**b**,**c**) measuring the intensity of fluorescence of cells stained with Hoechst 33258 on 7–20 microscopic images, or (**d**) trypan blue-exclusion test from four parallel samples of each experimental group. Mean ± S.E.M. (Standard Error of Mean), One way ANOVA, Student-Newman-Keuls Method. Statistically significant differences among the experimental groups (*p* ≤ 0.05) are indicated above the columns by numbers of differing groups.

**Figure 8.** Human HaCaT keratinocytes on day 1 after seeding on control polystyrene wells (PS) or on BC. The cell nuclei were stained with Hoechst 33258. Olympus IX 51 epifluorescence microscope, DP 70 digital camera, obj. 4×, scale bar 200 µm. **Figure 8.** Human HaCaT keratinocytes on day 1 after seeding on control polystyrene wells (PS) or on BC. The cell nuclei were stained with Hoechst 33258. Olympus IX 51 epifluorescence microscope, DP 70 digital camera, obj. 4×, scale bar 200 µm.

On day 4 after seeding, however, the cell number became significantly lower on all tested BC samples in comparison with the reference PS wells (Figure 7b). In accordance with this, the cells on the images taken on day 4 covered a considerable part of the PS surface and were almost confluent, while the cells on the BC-AD and BC-L samples without PM were in an early phase of islet formation (Figure 9). This result can be explained by the fact that the flat PS surface provided a better support for the cell spreading (which is a prerequisite of the subsequent cell proliferation) than the rougher and more irregular BC surfaces. It was particularly apparent on BC-AD samples, which showed more rugged surface on SEM images (Figure 5). It is known from studies on osteoblasts that the increased material surface roughness often hampered proliferation of these cells [61,62]. This phenomenon could be even more pronounced in keratinocytes, which are epithelial cells with polarization (i.e., functional specialization) of their basal and apical cytoplasmic membrane, designated to cover surfaces of various organs, i.e., to live and grow in a 2Dlike environment. In our earlier study, the negative effect of increased surface roughness on the cell proliferation was more apparent in endothelial cells, i.e., another type of epithelial-like cells, than in osteoblasts [63]. On day 4 after seeding, however, the cell number became significantly lower on all tested BC samples in comparison with the reference PS wells (Figure 7b). In accordance with this, the cells on the images taken on day 4 covered a considerable part of the PS surface and were almost confluent, while the cells on the BC-AD and BC-L samples without PM were in an early phase of islet formation (Figure 9). This result can be explained by the fact that the flat PS surface provided a better support for the cell spreading (which is a prerequisite of the subsequent cell proliferation) than the rougher and more irregular BC surfaces. It was particularly apparent on BC-AD samples, which showed more rugged surface on SEM images (Figure 5). It is known from studies on osteoblasts that the increased material surface roughness often hampered proliferation of these cells [61,62]. This phenomenon could be even more pronounced in keratinocytes, which are epithelial cells with polarization (i.e., functional specialization) of their basal and apical cytoplasmic membrane, designated to cover surfaces of various organs, i.e., to live and grow in a 2D-like environment. In our earlier study, the negative effect of increased surface roughness on the cell proliferation was more apparent in endothelial cells, i.e., another type of epithelial-like cells, than in osteoblasts [63].

Nevertheless, the growth of HaCaT cells on BC-AD and BC-L was markedly improved by PM. On plasma-modified BC samples, the cells reached significantly higher cell numbers than on the unmodified samples (Figure 7b), and the cell islands on these samples developed into large cell colonies (Figure 9). Nevertheless, the growth of HaCaT cells on BC-AD and BC-L was markedly improved by PM. On plasma-modified BC samples, the cells reached significantly higher cell numbers than on the unmodified samples (Figure 7b), and the cell islands on these samples developed into large cell colonies (Figure 9).

As revealed by a trypan blue exclusion test performed on day 4, the cells on the tested samples generally showed a high viability, ranging from approx. 88% to 97% (Figure 7d). Surprisingly, the lowest viability values were observed in cells cultivated on lyophilized BC, especially on samples modified with plasma. These samples contained the highest amount of oxygen (Table 2), which might be associated with damage to cells by reactive oxygen species. In addition, the surfaces of lyophilized samples are highly porous and contain numerous surface irregularities and areas of variable overall thickness and hardness, which

can hamper cell spreading. It has been previously reported that keratinocytes prefer softer materials over harder ones [64,65]. Similarly, as mentioned above, the more rugged surface, which can be tolerated or even preferred by osteoblasts [66], can decrease the adhesion and proliferation of keratinocytes as well as other cells of soft tissues. *Nanomaterials* **2021**, *11*, x FOR PEER REVIEW 13 of 17

**Figure 9.** Human HaCaT keratinocytes on day 4 after seeding on control polystyrene wells (PS) or on BC. The cell nuclei were stained with Hoechst 33258. Olympus IX 51 epifluorescence microscope, DP 70 digital camera, obj. 4×, scale bar 200 µm. **Figure 9.** Human HaCaT keratinocytes on day 4 after seeding on control polystyrene wells (PS) or on BC. The cell nuclei were stained with Hoechst 33258. Olympus IX 51 epifluorescence microscope, DP 70 digital camera, obj. 4×, scale bar 200 µm.

As revealed by a trypan blue exclusion test performed on day 4, the cells on the tested samples generally showed a high viability, ranging from approx. 88% to 97% (Figure 7d). Surprisingly, the lowest viability values were observed in cells cultivated on lyophilized BC, especially on samples modified with plasma. These samples contained the highest amount of oxygen (Table 2), which might be associated with damage to cells by reactive oxygen species. In addition, the surfaces of lyophilized samples are highly porous and contain numerous surface irregularities and areas of variable overall thickness and hardness, which can hamper cell spreading. It has been previously reported that keratinocytes prefer softer materials over harder ones [64,65]. Similarly, as mentioned above, the more rugged surface, which can be tolerated or even preferred by osteoblasts [66], can decrease the adhesion and proliferation of keratinocytes as well as other cells of soft tissues. On day 7 after seeding, the cell number on the tested BC samples equalized with that On day 7 after seeding, the cell number on the tested BC samples equalized with that on reference PS; i.e., the initial increase in cell number on BC samples, observed on day 1, was lost (Figure 7c). On BC-L samples, the final cell number was even significantly lower than on the other samples. This can be a consequence of the decreased cell viability observed on day 4 (Figure 7d), due to some negative effects of BC-L samples on cell spreading and proliferation, such as their surface irregularities and possible presence of oxygen radicals. Additionally, higher porosity (Table 3) of BC-L materials can lead to higher swelling of these materials in water-containing environments, such as cell culture media, which can have rather negative effect on the cell adhesion, because it can decrease the toughness (rigidity) of the substrate material. It is known that very soft and deformable materials, such as hydrogels, cannot resist the traction forces generated by cells during their spreading and support the viable growth of cells [67,68].

on reference PS; i.e., the initial increase in cell number on BC samples, observed on day 1, was lost (Figure 7c). On BC-L samples, the final cell number was even significantly lower than on the other samples. This can be a consequence of the decreased cell viability observed on day 4 (Figure 7d), due to some negative effects of BC-L samples on cell spreading and proliferation, such as their surface irregularities and possible presence of oxygen radicals. Additionally, higher porosity (Table 3) of BC-L materials can lead to higher swelling of these materials in water-containing environments, such as cell culture media, which Nevertheless, the cell colonization of all BC samples on day 7 significantly improved in comparison with the results from the 4th day of cultivation. From the images of cells on day 7 (Figure 10) it is evident that on PS and the plasma-modified BC, the cells are fully confluent, including those on BC-L, and in some parts of these samples, they created multilayer structures. On unmodified BC samples, the cells were still growing in colonies without reaching confluence, but these colonies have markedly enlarged in comparison with day 4.

can have rather negative effect on the cell adhesion, because it can decrease the toughness (rigidity) of the substrate material. It is known that very soft and deformable materials, such as hydrogels, cannot resist the traction forces generated by cells during their spread-

confluent, including those on BC-L, and in some parts of these samples, they created multilayer structures. On unmodified BC samples, the cells were still growing in colonies without reaching confluence, but these colonies have markedly enlarged in comparison

Nevertheless, the cell colonization of all BC samples on day 7 significantly improved in comparison with the results from the 4th day of cultivation. From the images of cells

ing and support the viable growth of cells [67,68].

with day 4.

**Figure 10.** Human HaCaT keratinocytes on day 7 after seeding on control polystyrene wells (PS) or on BC. The cell nuclei were stained with Hoechst 33258. Olympus IX 51 epifluorescence microscope, DP 70 digital camera, obj. 4×, scale bar 200 µm. **Figure 10.** Human HaCaT keratinocytes on day 7 after seeding on control polystyrene wells (PS) or on BC. The cell nuclei were stained with Hoechst 33258. Olympus IX 51 epifluorescence microscope, DP 70 digital camera, obj. 4×, scale bar 200 µm.

Taken together, BC samples investigated in this study provided a good support for the adhesion, growth and viability of human HaCaT keratinocytes, comparable with standard tissue culture polystyrene, which is considered to be one of the most suitable materials for cell colonization. The effect of air drying and lyophilization on the cell colonization of BC was in general also comparable; it can be only distinguished that lyophilization had a positive effect on initial cell attachment, while the subsequent growth of cells was better on air-dried BC. However, the highest positive effect on the colonization of BC with cells has been provided by PM, as evident from the fact that the plasma-modified BC samples, either air-dried or lyophilized, were the only BC samples on which the cells were able to develop a continuous, confluent layer (an even multilayer) after one Taken together, BC samples investigated in this study provided a good support for the adhesion, growth and viability of human HaCaT keratinocytes, comparable with standard tissue culture polystyrene, which is considered to be one of the most suitable materials for cell colonization. The effect of air drying and lyophilization on the cell colonization of BC was in general also comparable; it can be only distinguished that lyophilization had a positive effect on initial cell attachment, while the subsequent growth of cells was better on air-dried BC. However, the highest positive effect on the colonization of BC with cells has been provided by PM, as evident from the fact that the plasma-modified BC samples, either air-dried or lyophilized, were the only BC samples on which the cells were able to develop a continuous, confluent layer (an even multilayer) after one week of cultivation.

#### **4. Conclusions**

week of cultivation.

**4. Conclusions**  In our work, we described the differences between air drying and lyophilization with subsequent plasmatic modification of bacterial nanocellulose produced by the *Komagataeibacter sucrofermentans* bacteria strain (ATCC 700178). We found that BC-L materials maintained their structure, leading to higher porosity and specific surface area. Since the structure of BC-AD collapsed, the SEM revealed these samples to have a more rugged surface leading to almost two times higher contact angle. After plasmatic modification, the prepared samples showed a decrease of contact angle and increase of porosity and specific surface area. The O/C atomic ratio of the samples increased after PM, which suggests binding of the atmospheric oxygen to the activated surface immediately after modification. Compared to the literature, we obtained different values and trends of contact angles, morphology changes, and surface elemental composition. This is likely caused by In our work, we described the differences between air drying and lyophilization with subsequent plasmatic modification of bacterial nanocellulose produced by the *Komagataeibacter sucrofermentans* bacteria strain (ATCC 700178). We found that BC-L materials maintained their structure, leading to higher porosity and specific surface area. Since the structure of BC-AD collapsed, the SEM revealed these samples to have a more rugged surface leading to almost two times higher contact angle. After plasmatic modification, the prepared samples showed a decrease of contact angle and increase of porosity and specific surface area. The O/C atomic ratio of the samples increased after PM, which suggests binding of the atmospheric oxygen to the activated surface immediately after modification. Compared to the literature, we obtained different values and trends of contact angles, morphology changes, and surface elemental composition. This is likely caused by the different parameters of the employed plasma discharge. Furthemore, we

studied keratinocytes adhesion on our samples. We tested the initial adhesion (day 1) and subsequent growth (up to day 7) of human HaCaT keratinocytes on unmodified and plasma-modified BC-AD and BC-L. Due to its increased porosity and specific surface area, BC-L increased the initial adhesion of keratinocytes, but the subsequent cell growth was better on BC-AD. Modification with plasma for 240 s markedly accelerated the formation of typical keratinocyte islands and of continuous cell layers on both BC-AD and BC-L, which were comparable to those on standard cell culture polystyrene. Thus, the plasma-modified BC holds a great promise for skin tissue engineering and wound healing.

**Author Contributions:** Conceptualization, O.K. and L.B.; methodology, A.K. and L.S.; validation, O.K., L.B., and V.Š.; formal analysis, A.K., L.S., and O.K.; investigation, A.K., L.S., K.V., B.V., D.F., Z.K., and A.B.; resources, L.B. and V.Š.; data curation, A.K. and L.S.; writing—original draft preparation, A.K. and L.S.; writing—review and editing, O.K. and L.B.; visualization, A.K. and L.S.; supervision, O.K., L.B. and V.Š.; project administration, L.B. and V.Š.; funding acquisition, L.B. All authors have read and agreed to the published version of the manuscript.

**Funding:** This research was funded by GACR, grant number 20-01641S and OP VVV Project NAN-OTECH ITI II. No. CZ.02.1.01/0.0/0.0/18\_069/0010045.

**Conflicts of Interest:** The authors declare no conflict of interest. The funders had no role in the design of the study; in the collection, analyses, or interpretation of data; in the writing of the manuscript, or in the decision to publish the results.

#### **References**

*Article*

## **Antibacterial Multi-Layered Nanocellulose-Based Patches Loaded with Dexpanthenol for Wound Healing Applications**

**Daniela F. S. Fonseca 1 , João P. F. Carvalho 1 , Verónica Bastos 2 , Helena Oliveira 2 , Catarina Moreirinha 1 , Adelaide Almeida 2 , Armando J. D. Silvestre 1 , Carla Vilela 1,\* and Carmen S. R. Freire 1,\***

Received: 6 November 2020; Accepted: 8 December 2020; Published: 9 December 2020

**Abstract:** Antibacterial multi-layered patches composed of an oxidized bacterial cellulose (OBC) membrane loaded with dexpanthenol (DEX) and coated with several chitosan (CH) and alginate (ALG) layers were fabricated by spin-assisted layer-by-layer (LbL) assembly. Four patches with a distinct number of layers (5, 11, 17, and 21) were prepared. These nanostructured multi-layered patches reveal a thermal stability up to 200 ◦C, high mechanical performance (Young's modulus ≥4 GPa), and good moisture-uptake capacity (240–250%). Moreover, they inhibited the growth of the skin pathogen *Staphylococcus aureus* (3.2–log *CFU* mL−1 reduction) and were non-cytotoxic to human keratinocytes (HaCaT cells). The in vitro release profile of DEX was prolonged with the increasing number of layers, and the time-dependent data imply a diffusion/swelling-controlled drug release mechanism. In addition, the in vitro wound healing assay demonstrated a good cell migration capacity, headed to a complete gap closure after 24 h. These results certify the potential of these multi-layered polysaccharides-based patches toward their application in wound healing.

**Keywords:** oxidized bacterial cellulose; chitosan; alginate; layer-by-layer assembly; multi-layered patches; dexpanthenol; wound healing

#### **1. Introduction**

Natural polymers are being explored as building blocks to engineer multifunctional sustainable materials with sophisticated architectures via biomacromolecular assembly strategies. Among the prevailing methodologies to design such materials, the layer-by-layer (LbL) assembly is a simple, versatile and well-established technique with high relevance in the drug delivery domain [1]. In fact, the LbL assembly, which entails the sequential adsorption of complementary species (mostly based on opposing charges) on a substrate, is enabling the creation of drug reservoirs either in the form of planar films or capsules with a high-level of control of drug administration, targeted delivery, and lower side effects. The presence of multiple layers in a single system facilitates the design of materials with a profusion of bioactive functionalities (e.g., antioxidant activity, antibacterial activity, and wound healing capacity) that are typical to their individual precursors [1].

Polysaccharides are amid the most commonly utilized components for the design of multi-layered systems applied on drug delivery, despite the constraint of not yielding robust and free-standing films or capsules without the use of a template, either sacrificial or non-sacrificial (e.g., glass slides, mesoporous

silica, or calcium carbonate nanoparticles) [1]. In the context of planar films, bacterial cellulose (BC) membranes, viz. an exopolysaccharide produced by some non-pathogenic bacteria [2,3], can be used simultaneously as a non-sacrificial template and a drug reservoir. In fact, this robust and high purity cellulosic substrate, with a web-like entangled morphology, possesses in situ moldability that allows its direct biosynthesis in the form of membranes with personalized size and shape, revealing good in vitro biocompatibility and in vivo skin compatibility [4], alongside high water retention capacity, nanostructured porous network, high mechanical performance, and tailorable surface chemistry [5]. This set of intrinsic and unique features has been explored for the development of BC-based cutaneous drug delivery systems loaded with both hydrophilic or lipophilic active pharmaceutical ingredients (APIs) [6,7]. Nevertheless, the majority of the studies involves the simple diffusion of an API solution into the BC network followed by drying, which is a methodology that hinders the API selection and, in most instances, does not enable a controlled and targeted delivery of the APIs, particularly when they are highly water-soluble [7,8]. Herein, the use of the LbL assembly methodology, which can be performed in mild conditions in most substrates and enables the modulation of the composition, stability, and surface functionality of those substrates [1], can be a major asset for engineering BC-based membranes for wound healing applications, where a controlled and targeted drug delivery is an essential requirement.

Our interest in developing multi-layered BC derived patches by LbL assembly for cutaneous drug delivery originates from the idea of combining the peculiar properties of a freestanding exopolysaccharide robust membrane (i.e., BC) with the properties of hydrophilic ionic polysaccharides (i.e., chitosan (CH) and alginate (ALG)), and an API with wound healing potential (i.e., dexpanthenol (DEX) used in several pharmaceutical formulations in the field of dermatology and skin care). Although BC (either in its pure or oxidized forms) has already been individually blended with CH/ALG [9,10] or DEX [11], and BC nanocrystals have been used as a filler in an alginate matrix that was then covered with CH and gelatin polyelectrolytes via LbL assembly [12], there are no studies—as far as our literature search could verify—dedicated to the development of multi-layered patches composed of oxidized BC (OBC), CH, and ALG via spin-assisted LbL assembly technology, where the nanostructured porous cellulosic substrate plays the simultaneous role of a non-sacrificial template and a drug reservoir.

In this perspective, the goal of the present study is to design multi-layered patches composed of three polysaccharides, namely BC, CH, and ALG, as antibacterial cutaneous drug delivery vehicles for DEX, viz. a water-soluble API with wound healing ability. The multi-layered patches with a stratified structure were assembled via spin-assisted LbL methodology using a DEX-loaded OBC membrane as the initial negatively charged substrate, followed by the alternate adsorption of positively charged CH and negatively charged ALG polyelectrolytes, in a total of 5, 11, 17, and 21 layers. An in-depth analysis of the structure, morphology, thermal stability, mechanical properties, and moisture-uptake capacity of the multi-layered patches is presented, as well as the in vitro drug release profile at physiological pH, the in vitro antibacterial activity against *Staphylococcus aureus*, the in vitro cytotoxicity toward human HaCaT keratinocyte cells and the wound healing potential.

#### **2. Materials and Methods**

#### *2.1. Chemicals, Bacteria, and Cells*

Dexpanthenol (DEX, pharmaceutical secondary standard), glycerol (≥99.5%), phosphate buffered saline (PBS, pH 7.4), sodium hypochlorite solution (available chlorine 10–15%), sodium bromide (≥99%), sodium alginate (ALG, alginic acid sodium salt from brown algae, medium viscosity ≥2000 cP (2%, 25 ◦C)), 2,2,6,6-tetramethylpiperidine-1-oxyl (TEMPO free radical, >98%), tryptic soy broth (TSB), tryptic soy agar (TSA), 3-(4,5-dimethylthiazolyl-2)-2,5-diphenyltetrazolium bromide (MTT, 98%), and dimethyl sulfoxide (DMSO, ≥99.9%) were supplied by Sigma-Aldrich (Sintra, Portugal). Dulbecco0 s Modified Eagle0 s Medium (DMEM), fetal bovine serum (FBS), L-glutamine, penicillin/streptomycin, and fungizone were purchased from Gibco® (Life Technologies, Carlsbad, CA, USA). Type 1 ultrapure

water (resistivity of 18.2 MΩ cm (25 ◦C)) was filtered by a Simplicity® Water Purification System (Merck Millipore, Darmstadt, Germany). Other chemicals and solvents were of laboratory grade.

Chitosan (CH from shrimp, degree of deacetylation of 98%, viscosity of 2900 cP (1% solution in 1% of acetic acid)) was obtained from Mahtani Chitosan Pvt. Ltd. India. For purification, CH was dissolved in a 1% (*v*/*v*) aqueous acetic acid solution, filtered, and precipitated by neutralization with NaOH 0.5 M up to a pH of 8.5. The precipitate was rinsed with water until neutral pH, followed by freeze-drying [13].

Bacterial cellulose (BC) was biosynthesized in our laboratory by the *Gluconacetobacter sacchari* bacterial strain in the form of wet membranes [14]. *Staphylococcus aureus* (ATCC 6538) bacterium was received from American Type Culture Collection (ATCC) collection (Manassas, VA, USA). The nontumorigenic immortalized human keratinocyte HaCaT cell line was acquired from Cell Lines Services (Eppelheim, Germany).

#### *2.2. Preparation of Oxidized Bacterial Cellulose via TEMPO-Mediated Oxidation*

The oxidized bacterial cellulose (OBC) membranes were prepared via TEMPO-mediated oxidation as described elsewhere, with some modifications [15]. Shortly, 50 g of never-dried BC were suspended into deionized water containing TEMPO (0.13 g) and NaBr (1.3 g) under magnetic stirring. Then, 0.267 g of NaClO aqueous solution was slowly added, and the pH of the solution was maintained at 10.5 by adding a 0.5 M NaOH aqueous solution. The temperature was kept at 30 ◦C throughout the reaction for 30 min. The oxidation was stopped by quenching with 40 mL of absolute ethanol, followed by washing with deionized water. The OBC membranes (carboxyl content of 0.54 ± 0.09 mmol g−1 ) were stored in distilled water at 4 ◦C until further use.

#### *2.3. Preparation of Multi-Layered Patches Loaded with Dexpanthenol*

First, ALG and CH solutions (1 mg mL−1 ) were prepared in ultrapure water and 1% (*v*/*v*) acid acetic aqueous solution, respectively. Wet OBC membranes (6 × 4 cm2 ) were weighted, drained, and then soaked in 5 mL of an aqueous buffered solution (pH 7.4) of DEX (0.15% *w*/*v*) with glycerol (1% *w*/*v*). The membranes were stirred at 100 rpm and 30 ◦C for 1 h to allow the complete absorption of the solution (viz. 100% entrapment efficiency) and then left to dry at 30 ◦C for 24 h. The DEX-loaded OBC patch was coated with 250 µL of CH (zeta (ζ)-potential: 57 ± 3 mV (Zetasizer Nano ZS, Malvern Panalytical, Cambridge, United Kingdom)) and 250 µL of ALG (ζ-potential: −40 ± 3 mV) aqueous solutions by spin coating (Spin 150, APT GmbH–Automation und Produktionstechnik, Germany) at 2000 rpm during 15 s. A total of 5, 11, 17, or 21 layers of the polysaccharides were deposited, starting and finishing with CH, as compiled in Table 1.

**Table 1.** List of patches with the respective dry weight and thickness values.

a All patches contain glycerol as a plasticizer (1% *w*/*v*, 2.1 mg per cm2 of patch) that was incorporated inside the OBC; b Values are expressed as mean ± standard deviation.

#### *2.4. Characterization Techniques*

The thickness of the specimens was measured with a Mitutoyo coolant-proof digimatic micrometer MDC-25PX (Mitutoyo Corporation, Tokyo, Japan). All measurements were executed at fifteen different locations of the specimens, randomly selected, with an accuracy of 1 µm.

Attenuated total reflection-Fourier transform infrared (ATR-FTIR) spectra were computed on a Perkin-Elmer FT-IR System Spectrum BX spectrophotometer (Perkin-Elmer Inc., Waltham, MA, USA) coupled with a single horizontal Golden Gate ATR cell (Specac®, London, UK), in the wavenumber range of 600–4000 cm−1 at a resolution of 4 cm−1 over 32 scans.

Scanning electron micrographs (SEM) were recorded in a HR-FESEM SU-70 Hitachi microscope (Hitachi High-Technologies Corporation, Tokyo, Japan). The specimens were placed on an aluminum plate and formerly coated with a carbon film.

Thermogravimetric analysis (TGA) was undertaken in a SETSYS SETARAM TGA analyzer (SETARAM Instrumentation, Lyon, France). The specimens were heated from 25 ◦C to 800 ◦C at a heating rate of 10 ◦C min−1 under a nitrogen atmosphere.

Tensile mechanical tests were conducted on a uniaxial Instron 5966 testing apparatus (Instron Corp., Rockville, MD, USA) in traction mode at a deformation rate of 10 mm min−1 using a 500 N static load cell and gauge length of 30 mm. The specimens were rectangular strips (5 × 1 cm2 ) dried at 40 ◦C. All measurements were held on seven replicates.

The moisture-uptake capacity was assessed by laying the dry specimens (2×2 cm2 ) in a conditioned cabinet with 100% relative humidity at 25 ◦C for 22 h. After that period, the weight (*Ww*) of the moist specimens was measured, and the moisture-uptake was determined according to the equation:

$$\text{Moisture} - \text{uptake} \left( \% \right) = \left( W\_w - W\_0 \right) \times W\_0^{-1} \times 100 \tag{1}$$

where *W*0 is the initial weight of the dry specimens [16].

#### *2.5. In Vitro Drug Release Assays*

DEX-loaded patches (2 × 2 cm2 ) were immersed in a vessel containing 30 mL of 0.01 M PBS (pH 7.4) and the dissolution tests were then carried out at 32 ◦C and 50 rpm during 90 h. At regular time intervals, aliquots of 3 mL of each solution were withdrawn, and the same volume of fresh PBS was added to maintain a constant volume. The DEX concentration in each aliquot was determined by UV–Vis spectroscopy (Shimadzu UV-1800 UV–Vis spectrophotometer, Shimadzu Corp., Kyoto, Japan) at 206 nm with the linear calibration curve: *y* = 0.135*x* + 0.0167 (R2 = 0.9978), in the range of 10–100 µm mL−1 . The DEX content at each time was plotted as a cumulative percentage release concentration calculated using the formula:

$$\mathcal{C}\_{\text{cumulative}} = \mathcal{C}\_{n} + \left[ (\mathfrak{Z} \times \mathcal{C}\_{n-1}) / \mathfrak{Z}0 \right] \tag{2}$$

where *Cn* and *Cn*−1 are the concentrations of DEX at time *n* and *n* − 1, respectively. Nine replicates were performed for each specimen [8].

#### *2.6. In Vitro Antibacterial Activity*

The antibacterial activity of the OBC-based membranes was examined against *S. aureus*. The bacterial pre-inoculum cultures were grown for 24 h in TSB growth medium at 37 ◦C under horizontal shaking at 120 rpm until reaching a concentration of 108 to 109 colony forming units per mL (*CFU* mL−1 ). Each film specimen (4 × 6 cm2 ) was placed in contact with 5 mL of bacterial suspension via a ten-fold dilution of the overnight grown culture in PBS (pH 7.4). A bacterial cell suspension was tested as the control while OBC membranes, neat and DEX-loaded, were tested as blank references. All specimens were incubated at 37 ◦C under horizontal shaking at 120 rpm. At 24 h contact time, aliquots (500 µL) of each specimen and controls were collected and the bacterial concentration

(*CFU* mL−1 ) was determined by plating serial dilutions on TSA medium. The plates were incubated at 37 ◦C for 48 h. The *CFU* were determined on the most appropriate dilution on the agar plates. Two independent experiments were carried out and, for each, two replicates were plated. The bacteria log reduction of the specimens was calculated as follows:

$$\log \text{CFL mL}^{-1} \\ \text{reduction} = \log \text{CFL}\_{\text{control}} - \log \text{CFL}\_{\text{patch}} \tag{3}$$

#### *2.7. In Vitro Cytotoxicity Assay*

The cytotoxicity of the patches was evaluated in human keratinocytes cell line (HaCaT cells) by the MTT assay [17]. Briefly, cells were grown in complete DMEM medium supplemented with 10% FBS, 2 mM L-glutamine, 10,000 U mL−1 penicillin/streptomycin, and 250 µg mL−1 fungizone at 37 ◦C in 5% CO2 humidified atmosphere. Cells were observed daily under an inverted phase-contrast Eclipse TS100 microscope (Nikon, Tokyo, Japan). The tests were performed in triplicates of OBC, OBC/DEX, and OBC/DEX/CH/ALG\_21. Specimens of 1 × 1 cm2 were prepared, sterilized by ultraviolet radiation, and then incubated with 1.0 mL of complete DMEM medium at 37 ◦C with 5% CO2 for 24 h to prepare the specimen extract.

Meanwhile, one 96-well plate was prepared with 5 × 4 wells filled up with 6000 cells/well and the cells were then incubated in complete culture medium for 24 h for adhesion. After that time, cell culture medium (in the 96-well plates) was substituted by 100 µL of each of the extracts obtained from the incubated specimens and cells were then additionally incubated for either 24 h or 48 h. As a negative control, HaCaT cells were incubated with complete DMEM medium and treated identically as described for the specimens.

At the end of the incubation time, 50 µL of MTT (at a concentration of 1 g L−1 ) was added to each well and incubated for 4 h at 37 ◦C in a 5% CO2 humidified atmosphere. After that, the culture medium with MTT was removed and replaced by 150 µL of DMSO and the plate was positioned in a shaker for 2 h in the dark to totally dissolve the formazan crystals. The absorbance of the specimens was measured with a BioTek Synergy HT plate reader (Synergy HT Multi-Mode, BioTeK, Winooski, VT, USA) at 570 nm with blank corrections. The cell viability was calculated with respect to the control cells:

$$\text{Cell viability} \left( \% \right) = \left[ \left( \text{Abs}\_{\text{sample}} - \text{Abs}\_{\text{DMSO}} \right) / \left( \text{Abs}\_{\text{control}} - \text{Abs}\_{\text{DMSO}} \right) \right] \times 100 \tag{4}$$

where the *Abssample* is the absorbance of the specimen; *AbsDMSO* is the absorbance of the *DMSO* solvent; and *Abscontrol* is the absorbance of the control.

#### *2.8. In Vitro Wound Healing (Scratch) Assay*

The effect of the multi-layered patches on the migration capability of HaCaT keratinocyte cells was evaluated using the scratch assay. Cells were seeded in a 6-well plate at 5 × 105 cells/well and the linear wound was generated with a sterile 200 µL plastic pipette tip. Specimens of 1 × 1 cm2 were prepared, sterilized by ultraviolet radiation, and then incubated with 1.0 mL of complete DMEM medium at 37 ◦C, with 5% CO2 for 24 h to prepare the specimen extract. The tests were performed in triplicates of OBC/DEX, OBC/DEX/CH/ALG\_11, and OBC/DEX/CH/ALG\_21. As a negative control, HaCaT cells were treated identically as described for the specimens. Each specimen extract was applied to the scratch and the cells were then further incubated for 9 and 24 h at 37 ◦C, with 5% CO2. Optical micrographs (Eclipse TS100 microscope, Nikon, Tokyo, Japan) of the scratched area of each condition were taken and compared to the corresponding 0 h time point.

#### *2.9. Statistical Analysis*

The statistical analysis was conducted with OriginPro 9.0.0 (OriginLab Corporation, Northampton, MA, USA), where the statistical significance was assessed by the analysis of variance (ANOVA) and Tukey's test with the statistical significance established at *p* < 0.05.

#### **3. Results and Discussion** cellulose membranes with negatively charged groups at the surface (Figure 1a). The water-soluble radical reagent TEMPO is a catalytic oxidant that jointly with NaBr and NaClO selectively converts

**3. Results and Discussion**

Polysaccharides-based multi-layered patches composed of OBC, CH, and ALG were fabricated by spin-assisted LbL coating assembly technology, as illustrated in Figure 1. This assembly methodology was selected due to its simplicity and the stratified structure it conveys to the materials [1]. Therefore, the first step to prepare the multi-layered membranes comprised the functionalization of BC via the environmentally friendly TEMPO-mediated oxidation to obtain oxidized bacterial cellulose membranes with negatively charged groups at the surface (Figure 1a). The water-soluble radical reagent TEMPO is a catalytic oxidant that jointly with NaBr and NaClO selectively converts the cellulose C6 hydroxyl groups into carboxylic moieties in aqueous media, without compromising the morphological integrity of the fibers [18]. the cellulose C6 hydroxyl groups into carboxylic moieties in aqueous media, without compromising the morphological integrity of the fibers [18]. The OBC membrane was then loaded with DEX (i.e., OBC/DEX membrane, Figure 1b), which is a water-soluble active pharmaceutical ingredient (API) used in several pharmaceutical formulations in the field of dermatology and skin care due to its capacity to act as a skin moisturizer, skin barrier restorer, and facilitator of wound healing [19]. All membranes contain 0.32 mg of DEX per cm2 of OBC, selected based on equivalent commercial formulations [19]. Further to this, glycerol (1% *w*/*v*, 2.1 mg per cm2 of membrane) was incorporated in the membrane as a plasticizer to upsurge their malleability and conformability to the skin surface [20].

of BC via the environmentally friendly TEMPO-mediated oxidation to obtain oxidized bacterial

*Nanomaterials* **2020**, *10*, 2469 6 of 16

Polysaccharides-based multi-layered patches composed of OBC, CH, and ALG were fabricated by spin-assisted LbL coating assembly technology, as illustrated in Figure 1. This assembly methodology was selected due to its simplicity and the stratified structure it conveys to the materials

**Figure 1.** Schematic representation of (**a**) TEMPO-mediated oxidation of BC yielding OBC, (**b**) loading of OBC with DEX, (**c**) spin coating LbL assembly of the polysaccharides multi-layered patch with 5 layers, and (**d**) digital photographs of the OBC/DEX/CH/ALG patches with 5, 11, 17, and 21 layers. **Figure 1.** Schematic representation of (**a**) TEMPO-mediated oxidation of BC yielding OBC, (**b**) loading of OBC with DEX, (**c**) spin coating LbL assembly of the polysaccharides multi-layered patch with 5 layers, and (**d**) digital photographs of the OBC/DEX/CH/ALG patches with 5, 11, 17, and 21 layers.

Afterward, four multi-layered patches with a distinct number of layers (5, 11, 17, and 21) of the positively charged CH (ζ-potential: 57 ± 3 mV) and the negatively charged ALG (ζ-potential: −40 ± 3 mV) polyelectrolytes were prepared (Figure 1c). CH and ALG polysaccharides were selected as the The OBC membrane was then loaded with DEX (i.e., OBC/DEX membrane, Figure 1b), which is a water-soluble active pharmaceutical ingredient (API) used in several pharmaceutical formulations in the field of dermatology and skin care due to its capacity to act as a skin moisturizer, skin barrier restorer, and facilitator of wound healing [19]. All membranes contain 0.32 mg of DEX per cm2 of OBC, selected based on equivalent commercial formulations [19]. Further to this, glycerol (1% *w*/*v*, 2.1 mg per cm2 of membrane) was incorporated in the membrane as a plasticizer to upsurge their malleability and conformability to the skin surface [20].

Afterward, four multi-layered patches with a distinct number of layers (5, 11, 17, and 21) of the positively charged CH (ζ-potential: 57 ± 3 mV) and the negatively charged ALG (ζ-potential: −40 ± 3 mV) polyelectrolytes were prepared (Figure 1c). CH and ALG polysaccharides were selected as the oppositely charged polyelectrolytes because of their efficient film-forming ability and vast applicability in the LbL assembly technology [1]. Furthermore, CH was chosen as the final layer in the four multi-layered membranes due to its known antibacterial activity toward a wide variety of microorganisms [13,21], largely linked to the presence of amino groups [22]. This bioactive property is particularly pertinent in the context of skin regeneration to avoid the growth of pathogenic microorganisms that will contribute to microbial wound infections [23,24].

All multi-layered patches are whitish and homogeneous, as displayed in the photographs of Figure 1d, confirming the even distribution of DEX and of the polyelectrolytes (i.e., CH and ALG). Predictably, the weight and thickness of the multi-layered patches increased with the increasing number of layers as listed Table 1. The nanostructured patches were characterized in terms of structure by ATR-FTIR spectroscopy, morphology by SEM, thermal stability by TGA, mechanical properties by tensile tests and moisture-uptake capacity. Additionally, the in vitro drug release profile, in vitro antibacterial activity, in vitro cytotoxicity, and in vitro wound healing potential were also evaluated.

#### *3.1. Structural and Morphological Characterization*

The modification of BC via TEMPO-mediated oxidation to obtain OBC with negatively charged groups was confirmed by ATR-FTIR spectroscopy through the presence of the typical absorption bands of the BC backbone at 1029 cm−1 (C–O stretching), 1316 cm−1 (O–H in plane bending), 2897 cm−1 (C–H stretching), and 3340 cm−1 (O–H stretching) [25], and the emergence of the absorption band at 1590–1640 cm−1 assigned to the antisymmetric vibration of ionized carboxyl groups (νas(COO−), Figure 2a) [26]. Moreover, the SEM micrograph of the OBC membrane surface (Figure 2c) shows the characteristic three-dimensional nanofibrillar structure of BC [5,14]. This means that the oxidation of BC did not affect the morphology (Figure 2c) nor the visual appearance of the membranes.

The incorporation of DEX via diffusion into the three-dimensional porous network of OBC was also validated by ATR-FTIR spectroscopy (Figure 2b) through the presence of the typical absorption bands of each component (Figure 2a), whilst the ones allocated to DEX, namely at 3308 cm−1 (O–H and N–H stretching), 1636 cm−1 (C=O stretching, amide I band), and 1535 cm−1 (amide II band) [26,27] are not completely observable given the small amount of API present in the patch (0.32 mg of DEX per cm2 of OBC). In the SEM micrograph of the surface of the OBC/DEX patch (Figure 2c), the nanofibrillar structure of OBC is still visible, suggesting that all DEX was absorbed into the three-dimensional porous network of OBC and none remains at the surface of the patch, confirming the above vibrational spectroscopy data.

The ATR-FTIR spectra of the multi-layered patches (Figure 2b) portrayed similarities with those of their precursors (Figure 2a), mainly with OBC, but also with chitosan at 3342 cm−1 (O–H stretching), 1638 cm−1 (C=O stretching and N−H bending), 1592 cm−1 (−NH2 bending), 1380 cm−1 (−CH2 bending), 1148 cm−1 (C−O−C antisymmetric stretching), 1078 and 1032 cm−1 (C−O stretching) [28], and ALG at 3250 cm−1 (O–H stretching), and 1596 cm−1 (COO− antisymmetric stretching) and 1406 cm−1 (COO− symmetric stretching) [29]. Moreover, the absorption bands of glycerol at 1100 cm−1 and 1030 cm−1 (C–O stretching vibrations of alcohols) and 3250 cm−1 (O–H stretching), [7] are fully superimposed with the vibrations of the polysaccharides, viz. OBC, CH, and ALG.

Regarding the morphology (Figure 2c), the SEM micrographs of the surface of the multi-layered membranes did not disclose the three-dimensional fibrillar structure of OBC because it was completely covered by the various CH and ALG layers. Furthermore, the micrographs also depicted the formation of micron-size particles credited to the polyelectrolytes, which underlines the homogeneous and total coverage of the negatively charged OBC substrate. Indeed, in the case of the patch with 21 layers, the micrograph resembles the formation of a particulate film.

**Figure 2.** (**a**,**b**) ATR-FTIR spectra (vibrational modes: *ν* = stretching, *δ* = bending) of BC, OBC, CH, ALG, DEX (**a**), OBC/DEX, and OBC/DEX/CH/ALG-based multi-layered patches with 5, 11, 17, and 21 layers (**b**), and (**c**) SEM surface micrographs (×30.0 k magnification) of OBC, OBC/DEX, and **Figure 2.** (**a**,**b**) ATR-FTIR spectra (vibrational modes: ν = stretching, δ = bending) of BC, OBC, CH, ALG, DEX (**a**), OBC/DEX, and OBC/DEX/CH/ALG-based multi-layered patches with 5, 11, 17, and 21 layers (**b**), and (**c**) SEM surface micrographs (×30.0 k magnification) of OBC, OBC/DEX, and OBC/DEX/CH/ALG-based multi-layered patches with 5, 11, 17, and 21 layers.

#### OBC/DEX/CH/ALG-based multi-layered patches with 5, 11, 17, and 21 layers. *3.2. Thermal Stability and Mechanical Properties*

*3.2. Thermal Stability and Mechanical Properties*  The thermal stability of the multi-layered patches was measured by thermogravimetric analysis under a nitrogen atmosphere. According to Figure 3, the data did not vary appreciably with the increasing number of polyelectrolyte layers (i.e., 5, 11, 17, and 21 layers), with all membranes exhibiting a double weight-loss degradation profile with thermal stability up to about 200 °C (Figure 3b). Apart from the initial volatilization of water at ca. 100 °C, the small first step with a maximum rate of decomposition temperature at ca. 240 °C is allocated to the volatilization of glycerol [30] and degradation of DEX [27], while the second step with a maximum rate of decomposition temperature in the range 320—330 °C is related to the polysaccharide skeleton degradation, namely OBC [31], CH [13], and ALG [32]. In this second-step, the maximum degradation temperature slightly decreases The thermal stability of the multi-layered patches was measured by thermogravimetric analysis under a nitrogen atmosphere. According to Figure 3, the data did not vary appreciably with the increasing number of polyelectrolyte layers (i.e., 5, 11, 17, and 21 layers), with all membranes exhibiting a double weight-loss degradation profile with thermal stability up to about 200 ◦C (Figure 3b). Apart from the initial volatilization of water at ca. 100 ◦C, the small first step with a maximum rate of decomposition temperature at ca. 240 ◦C is allocated to the volatilization of glycerol [30] and degradation of DEX [27], while the second step with a maximum rate of decomposition temperature in the range 320–330 ◦C is related to the polysaccharide skeleton degradation, namely OBC [31], CH [13], and ALG [32]. In this second-step, the maximum degradation temperature slightly decreases with the increasing number of layers, most likely due to the marginally lower thermal stability of the

polyelectrolytes [13,32]. Furthermore, the final residue of the multi-layered patches at 800 ◦C varied between 25% for OBC/DEX/CH/ALG\_5 and 35% for OBC/DEX/CH/ALG\_21, which is mainly attributed to the increase of CH and ALG in the membranes. However, it is worth remarking that all multi-layered OBC/DEX/CH/ALG-based membranes are thermally stable up to 200 ◦C, and therefore, can undergo the sterilization procedures (e.g., autoclaving at ca. 120 ◦C) required for biomedical applications. between 25% for OBC/DEX/CH/ALG\_5 and 35% for OBC/DEX/CH/ALG\_21, which is mainly attributed to the increase of CH and ALG in the membranes. However, it is worth remarking that all multi-layered OBC/DEX/CH/ALG-based membranes are thermally stable up to 200 °C, and therefore, can undergo the sterilization procedures (e.g., autoclaving at ca. 120 °C) required for biomedical applications.

polyelectrolytes [13,32]. Furthermore, the final residue of the multi-layered patches at 800 °C varied

**Figure 3.** (**a**) Thermograms and (**b**) derivative curves of the multi-layered patches: OBC/DEX/CH/ALG\_5, OBC/DEX/CH/ALG\_11, OBC/DEX/CH/ALG\_17, and OBC/DEX/CH/ALG\_21. **Figure 3.** (**a**) Thermograms and (**b**) derivative curves of the multi-layered patches: OBC/DEX/CH/ALG\_5, OBC/DEX/CH/ALG\_11, OBC/DEX/CH/ALG\_17, and OBC/DEX/CH/ALG\_21.

The mechanical properties of the multi-layered polysaccharides-based patches were also analyzed, and the data obtained from the stress-strain curves are condensed in Table 2. The negatively charged OBC substrate exhibits an elongation at break of 3.4 ± 0.6%, tensile strength of 150 ± 25 MPa and Young's modulus of 8.2 ± 1.4 GPa. Furthermore, the inclusion of DEX (dissolved in a buffered solution with glycerol (1% *w*/*v*)) into the OBC porous structure promoted a decrease in both Young's modulus and tensile strength to 3.5 ± 1.2 GPa and 97 ± 15 MPa, respectively, complemented by an increase in the elongation at break to 4.9 ± 0.6%. This behavior is comparable to the trend obtained for pure BC membranes after the incorporation of APIs such as diclofenac [33] and lidocaine [34] into the BC three-dimensional structure. Furthermore, Tanrıverdi et al. also reported a decrease in the tensile strength of synthetic nanofiber mats, namely poly(lactic-*co*-glycolic acid), poly(ethylene oxide), and poly(ɛ-caprolactone), loaded with DEX [35]. The mechanical properties of the multi-layered polysaccharides-based patches were also analyzed, and the data obtained from the stress-strain curves are condensed in Table 2. The negatively charged OBC substrate exhibits an elongation at break of 3.4 ± 0.6%, tensile strength of 150 ± 25 MPa and Young's modulus of 8.2 ± 1.4 GPa. Furthermore, the inclusion of DEX (dissolved in a buffered solution with glycerol (1% *w*/*v*)) into the OBC porous structure promoted a decrease in both Young's modulus and tensile strength to 3.5 ± 1.2 GPa and 97 ± 15 MPa, respectively, complemented by an increase in the elongation at break to 4.9 ± 0.6%. This behavior is comparable to the trend obtained for pure BC membranes after the incorporation of APIs such as diclofenac [33] and lidocaine [34] into the BC three-dimensional structure. Furthermore, Tanrıverdi et al. also reported a decrease in the tensile strength of synthetic nanofiber mats, namely poly(lactic-*co*-glycolic acid), poly(ethylene oxide), and poly(ε-caprolactone), loaded with DEX [35].

After the LbL assembly of the oppositely charged polyelectrolytes (i.e., CH and ALG) on the negatively charged OBC substrate, the mechanical performance of the multi-layered membranes **Table 2.** Young's modulus, tensile strength, and elongation at break of the OBC, OBC/DEX, and the multi-layered patches.

**Table 2.** Young's modulus, tensile strength, and elongation at break of the OBC, OBC/DEX, and the a All patches contain glycerol as a plasticizer.

multi-layered patches. **Patch a Young's Modulus/GPa Tensile Strength/MPa Elongation at Break/%**  OBC 8.2 ± 1.4 150 ± 25 3.4 ± 0.6 OBC/DEX 3.5 ± 1.2 97 ± 15 4.9 ± 0.6 OBC/DEX/CH/ALG\_5 4.2 ± 0.8 142 ± 39 4.3 ± 0.7 OBC/DEX/CH/ALG\_11 4.4 ± 0.8 154 ± 52 4.0 ± 1.1 OBC/DEX/CH/ALG\_17 9.8 ± 2.7 166 ± 45 2.0 ± 0.4 OBC/DEX/CH/ALG\_21 10.9 ± 4.0 188 ± 57 2.2 ± 0.4 After the LbL assembly of the oppositely charged polyelectrolytes (i.e., CH and ALG) on the negatively charged OBC substrate, the mechanical performance of the multi-layered membranes increased with the increasing number of layers. The Young's modulus increased from 4.2 ± 0.8 GPa for OBC/DEX/CH/ALG\_5 to 10.9 ± 4.0 GPa for OBC/DEX/CH/ALG\_21, whereas the tensile strength increased from 142 ± 39 MPa for OBC/DEX/CH/ALG\_5 to 188 ± 57 MPa for OBC/DEX/CH/ALG\_21. On the contrary, the elongation at break decreased from 4.3 ± 0.7% for OBC/DEX/CH/ALG\_5 to 2.2 ± 0.4% for OBC/DEX/CH/ALG\_21, meaning that the membranes became less elastic as the number of layers increased from 5 to 21. Nonetheless, these hydrophilic multi-layered patches are satisfactorily malleable to conform to the surface of the skin.

a All patches contain glycerol as a plasticizer.

Worth noting is the fact that these OBC/DEX/CH/ALG multi-layered patches exhibited a higher tensile strength than the similar non-multi-layered films based on hydrogen peroxide oxidized BC (HOBC) or periodic acid oxidized BC (POBC) containing CH and ALG, which were prepared by blending the homogenized HOBC or POBC pulp with a CH and ALG gel solution, and studied for application as wound dressings [10]. blending the homogenized HOBC or POBC pulp with a CH and ALG gel solution, and studied for application as wound dressings [10]. *3.3. Moisture-Uptake Capacity*  The moisture-uptake capacity of the multi-layered patches was determined to anticipate their

(HOBC) or periodic acid oxidized BC (POBC) containing CH and ALG, which were prepared by

*Nanomaterials* **2020**, *10*, 2469 10 of 16

Worth noting is the fact that these OBC/DEX/CH/ALG multi-layered patches exhibited a higher

#### *3.3. Moisture-Uptake Capacity* interaction with moisture and exudates when applied to wound sites. Hence, the patches were left in a chamber with 100% relative humidity at 25 °C for 22 h. According to Figure 4, all

The moisture-uptake capacity of the multi-layered patches was determined to anticipate their interaction with moisture and exudates when applied to wound sites. Hence, the patches were left in a chamber with 100% relative humidity at 25 ◦C for 22 h. According to Figure 4, all OBC/DEX/CH/ALG patches absorbed moisture over time and were able to reach values of 240–250% after 22 h of exposure. Furthermore, the time-dependent data did not vary appreciably with the increasing number of layers (i.e., 5, 11, 17, and 21 layers), meaning that these patches present an analogous moisture-uptake capacity. This behavior is obviously credited to the hydrophilic nature of the precursors of the OBC/DEX/CH/ALG-based membranes, as in fact shown for the non-multi-layered films composed of OBC, CH, and ALG [10]. OBC/DEX/CH/ALG patches absorbed moisture over time and were able to reach values of 240–250% after 22 h of exposure. Furthermore, the time-dependent data did not vary appreciably with the increasing number of layers (i.e., 5, 11, 17, and 21 layers), meaning that these patches present an analogous moisture-uptake capacity. This behavior is obviously credited to the hydrophilic nature of the precursors of the OBC/DEX/CH/ALG-based membranes, as in fact shown for the non-multilayered films composed of OBC, CH, and ALG [10]. Therefore, any of these multi-layered polysaccharide-based patches can promote (if necessary) a moist environment at the skin interface and remove the surplus of exudates or other body fluids when applied to the skin surface.

**Figure 4.** (**a**) Moisture-uptake capacity and (**b**) DEX cumulative release profile of OBC, OBC-loaded with DEX, and multi-layered polysaccharides-based patches. **Figure 4.** (**a**) Moisture-uptake capacity and (**b**) DEX cumulative release profile of OBC, OBC-loaded with DEX, and multi-layered polysaccharides-based patches.

*3.4. In Vitro Drug Release Profile*  The in vitro release profile of the multi-layered polysaccharides-based patches was tested in PBS Therefore, any of these multi-layered polysaccharide-based patches can promote (if necessary) a moist environment at the skin interface and remove the surplus of exudates or other body fluids when applied to the skin surface.

(pH 7.4) at 32 °C that simulates the usual pH and temperature of the skin [36]. Figure 4b shows the

#### cumulative release profiles of DEX from the OBC/DEX/CH/ALG-based membranes, which were *3.4. In Vitro Drug Release Profile*

compared with the in vitro release of DEX from a membrane composed solely of OBC and DEX. In general, the release profile of the patches is quite standard, with an initial burst followed by a plateau where the DEX release achieves the highest value. While the OBC/DEX and OBC/DEX/CH/ALG\_5 patches reached a maximum cumulative release of ca*.* 95% after 16 h, the The in vitro release profile of the multi-layered polysaccharides-based patches was tested in PBS (pH 7.4) at 32 ◦C that simulates the usual pH and temperature of the skin [36]. Figure 4b shows the cumulative release profiles of DEX from the OBC/DEX/CH/ALG-based membranes, which were compared with the in vitro release of DEX from a membrane composed solely of OBC and DEX.

OBC/DEX/CH/ALG\_11 achieved ca. 87% after 16 h and OBC/DEX/CH/ALG\_17 attained ca*.* 85% after 28 h. In the case of the OBC/DEX/CH/ALG\_21 patch, it only reached a maximum cumulative release of ca. 65% after 90 h. This is a direct indication that the multi-layered systems retarded the DEX release from the OBC membrane, particularly for 21 layers. Moreover, this slow-release profile is suited for wounds that require longer therapeutic treatments combined with a local skin moisturizer with antibacterial properties and wound healing ability. When compared with non-multi-layered BC-based membranes loaded with water-soluble APIs [7,8], these OBC/DEX/CH/ALG multi-layered patches presented a controlled release profile that only reached high cumulative release values after several hours. In general, the release profile of the patches is quite standard, with an initial burst followed by a plateau where the DEX release achieves the highest value. While the OBC/DEX and OBC/DEX/CH/ALG\_5 patches reached a maximum cumulative release of ca. 95% after 16 h, the OBC/DEX/CH/ALG\_11 achieved ca. 87% after 16 h and OBC/DEX/CH/ALG\_17 attained ca. 85% after 28 h. In the case of the OBC/DEX/CH/ALG\_21 patch, it only reached a maximum cumulative release of ca. 65% after 90 h. This is a direct indication that the multi-layered systems retarded the DEX release from the OBC membrane, particularly for 21 layers. Moreover, this slow-release profile is suited for wounds that require longer therapeutic treatments combined with a local skin moisturizer with antibacterial properties and wound healing ability.

When compared with non-multi-layered BC-based membranes loaded with water-soluble APIs [7,8], these OBC/DEX/CH/ALG multi-layered patches presented a controlled release profile that only reached high cumulative release values after several hours.

The in vitro drug release curves (Figure 4b) were further adjusted to kinetic models to identify the diffusional release mechanism of DEX from the multi-layered OBC/DEX/CH/ALG-based membranes. The data were better fitted to the Korsmeyer–Peppas kinetic model [37,38], *Mt*/*M*∞ = *ktn* , with *Mt* as the DEX content released at time *t*, *M*∞ is the DEX content released at infinite time, *k* is the kinetic constant, and *n* is the diffusion constant denoting the release mechanism [37,39]. Based on this kinetic model, only the cumulative release values of *Mt*/*M*∞ < 60% are fitted, and therefore, a release exponent (*n*) of 0.45 was obtained for OBC/DEX, 0.68 for OBC/DEX/CH/ALG\_5, 0.69 for OBC/DEX/CH/ALG\_11, 0.68 for OBC/DEX/CH/ALG\_17, and 0.70 for OBC/DEX/CH/ALG\_21 (Table 3). These curve fitting parameters are representative of Fickian diffusion (i.e., diffusion-controlled drug release mechanism, in the case of OBC/DEX, and non-Fickian transport (0.5 < *n* < 1.0) (i.e., diffusion and swelling controlled drug release mechanism), for the case of the multi-layered polysaccharide-based membranes [37–39].

**Table 3.** Fitting parameters of the Korsmeyer–Peppas kinetic model and the corresponding drug release mechanism.

#### *3.5. In Vitro Antibacterial Activity*

The antibacterial activity of the multi-layered polysaccharide-based patches was assayed against *S. aureus* (a Gram-positive bacterium) for 24 h, as shown in Figure 5a. This bacterium has been extensively studied because it is the culprit of several skin infections, particularly in patients with skin injuries [23]. As expected, neither the control nor the OBC membrane hindered the growth of the Gram-positive pathogenic microorganism. This observation is consistent with the literature, since OBC does not inhibit the growth of *S. aureus*, as previously reported [40]. Nevertheless, when DEX is loaded into the OBC membrane, the bacterial growth of *S. aureus* was reduced by 0.8–log *CFU* mL−1 after 24 h. Although DEX is not an antibacterial agent [41], it seems that its combination with OBC conveyed a small growth inhibition to the OBC/DEX membrane.

Regarding the multi-layered polysaccharide-based patches, they are responsible for ca. 3.2–log *CFU* mL−1 reduction after 24 h (Figure 5a). Since the general tendency of the four membranes was the same, it is fair to claim that all OBC/DEX/CH/ALG patches are antibacterial agents due to the presence of chitosan. In fact, several studies have shown that the marine polysaccharide derived from chitin (i.e., chitosan) presents antibacterial activity toward a wide variety of microorganisms [21] including *S. aureus* [13], largely linked to the presence of amino groups [22,24]. This bioactive property is of paramount importance for skin regeneration to avert the growth of pathogenic microorganisms causing microbial wound infections [23,24].

24 h and 94 ± 9% after 48 h. This behavior was predictable because DEX is a non-cytotoxic drug used

With regard to the multi-layered patch with the higher number of layers (i.e., OBC/DEX/CH/ALG\_21), it was also deemed as a non-cytotoxic material because the cell viability was 95 ± 5% after 24 h and 97 ± 6% after 48 h (Figure 5b). Furthermore, the optical micrographs of the HaCaT cells (Figure 5c) support the cell viability data by evidencing that neither the cell morphology nor the cell count varied in relation to the control after 24 h of cell incubation with the OBC, OBC/DEX, and OBC/DEX/CH/ALG\_21 membranes. Actually, all values were high above the 70% threshold of

in several pharmaceutical formulations in the field of dermatology and skin care [19].

**Figure 5.** (**a**) Antibacterial activity (24 h of exposure) of OBC, OBC-loaded with DEX, and multilayered patches (the symbol \* represents the means with a significant difference (*p* < 0.05) from the control and OBC); (**b**) cell viability of HaCaT cells after 24 h and 48 h of exposure to negative control, OBC, OBC/DEX, and OBC/DEX/CH/ALG\_21, and (**c**) the corresponding optical micrographs of HaCaT cells after 24 h of exposure. **Figure 5.** (**a**) Antibacterial activity (24 h of exposure) of OBC, OBC-loaded with DEX, and multi-layered patches (the symbol \* represents the means with a significant difference (*p* < 0.05) from the control and OBC); (**b**) cell viability of HaCaT cells after 24 h and 48 h of exposure to negative control, OBC, OBC/DEX, and OBC/DEX/CH/ALG\_21, and (**c**) the corresponding optical micrographs of HaCaT cells after 24 h of exposure.

#### *3.6. In Vitro Cytotoxicity Assays*

*3.7. In Vitro Wound Healing Activity*  Having proved that the multi-layered polysaccharide-based patches are robust, thermally stable, and present a controlled drug release profile, antibacterial activity against *S. aureus* and noncytotoxicity toward HaCaT cells, the next step focused on the wound healing potential of these patches as delivery vehicles for DEX. Therefore, the wound healing activity of the multi-layered polysaccharide-based patches with 11 and 21 layers (OBC/DEX/CH/ALG\_11 and OBC/DEX/CH/ALG\_21) as well as of the OBC/DEX patch was assessed using the in vitro wound healing scratch assay [47]. Therefore, the influence of the patches on the cell migration was studied for HaCaT keratinocyte cells after 9 and 24 h of exposure. Predictably, and as shown in Figure 6, the cells cultivated in a medium containing only culture medium (i.e., control) presented a good The cytotoxic behavior of the multi-layered patch with 21 layers (OBC/DEX/CH/ALG\_21) as well as that of OBC and OBC/DEX was assessed in human HaCaT keratinocyte cells via the indirect MTT assay. This cell line was elected because it has been intensely used as a model for epidermal cells [42–45]. Figure 5b shows that the HaCaT cells metabolic activity was normal for the negative control (100% cell viability) as well as for the OBC membrane (ca. 98%) for both exposure times (24 and 48 h). Therefore, the OBC membrane is non-cytotoxic toward HaCaT cells, which is coherent with the results achieved in the literature for other cell lines such as NIH3T3 fibroblast cells [31]. Furthermore, the cell viability of the OBC patch loaded with DEX was almost unaltered with values of 97 ± 7% after 24 h and 94 ± 9% after 48 h. This behavior was predictable because DEX is a non-cytotoxic drug used in several pharmaceutical formulations in the field of dermatology and skin care [19].

migration capacity with the complete occlusion of the scratch after 24 h. Overall, and in the case of the patches, the HaCaT keratinocyte cells situated near the boundary of the scratched region moved toward the vacant region as time progressed, achieving a full closure after 24 h. Furthermore, the cell density at the back of the scratch site is also growing with time, which points to the proliferation of the HaCaT keratinocyte cells. However, in terms of temporal progression, the patches containing DEX, namely OBC/DEX, OBC/DEX/CH/ALG\_11 and With regard to the multi-layered patch with the higher number of layers (i.e., OBC/DEX/ CH/ALG\_21), it was also deemed as a non-cytotoxic material because the cell viability was 95 ± 5% after 24 h and 97 ± 6% after 48 h (Figure 5b). Furthermore, the optical micrographs of the HaCaT cells (Figure 5c) support the cell viability data by evidencing that neither the cell morphology nor the cell count varied in relation to the control after 24 h of cell incubation with the OBC, OBC/DEX, and OBC/DEX/CH/ALG\_21 membranes. Actually, all values were high above the 70% threshold of cell viability [46], which reflects the potential in vivo performance of these safe and compatible multi-layered patches for application in the biomedical field.

#### *3.7. In Vitro Wound Healing Activity*

Having proved that the multi-layered polysaccharide-based patches are robust, thermally stable, and present a controlled drug release profile, antibacterial activity against *S. aureus* and non-cytotoxicity toward HaCaT cells, the next step focused on the wound healing potential of these patches as delivery vehicles for DEX. Therefore, the wound healing activity of the

multi-layered polysaccharide-based patches with 11 and 21 layers (OBC/DEX/CH/ALG\_11 and OBC/DEX/CH/ALG\_21) as well as of the OBC/DEX patch was assessed using the in vitro wound healing scratch assay [47]. Therefore, the influence of the patches on the cell migration was studied for HaCaT keratinocyte cells after 9 and 24 h of exposure. Predictably, and as shown in Figure 6, the cells cultivated in a medium containing only culture medium (i.e., control) presented a good migration capacity with the complete occlusion of the scratch after 24 h. natural compounds known to expedite the wound healing process, and by Silva et al. [16] for multifunctional nanofibrous patches composed of nanocellulose and lysozyme nanofibers. Given that wound closure is a signal of wound healing ability [49], these antibacterial and noncytotoxic multi-layered polysaccharide-based patches have the potential to foster wound healing and, simultaneously, hinder skin infections. Moreover, the drug release profile can be tuned to the specific type of wound and required treatment time by simply adjusting the number of polysaccharide layers.

were documented by Sharma et al. [48] for curcumin conjugated with hyaluronic acid, which are two

*Nanomaterials* **2020**, *10*, 2469 13 of 16

OBC/DEX/CH/ALG\_21, demonstrated higher coverage of the wound-like gap area after 9 h, induced by cell growth and migration, when compared to the positive control. Despite the analogous behavior of the HaCaT keratinocyte cells after 24 h of exposure to the three patches (Figure 6), the optical micrographs after 9 h show a slightly slower cell growth and migration for the patches with 11 and 21 layers. In fact, the behavior of the different patches after 9 h of exposure (Figure 6) seems to be in line with the in vitro drug release profile of DEX, where the OBC/DEX patch is the one releasing the higher amount of DEX in the shorter period of time and the multi-layered OBC/DEX/CH/ALG\_21 membrane with a controlled drug release profile is the one releasing the lower content of DEX after

**Figure 6.** Optical micrographs of the scratch assay of the HaCaT cells after 9 and 24 h of exposure to control, OBC/DEX, OBC/DEX/CH/ALG\_11, and OBC/DEX/CH/ALG\_21 patches. **Figure 6.** Optical micrographs of the scratch assay of the HaCaT cells after 9 and 24 h of exposure to control, OBC/DEX, OBC/DEX/CH/ALG\_11, and OBC/DEX/CH/ALG\_21 patches.

**4. Conclusions**  Functional multi-layered patches composed of OBC, CH, ALG, and DEX were prepared for wound healing applications. Four multi-layered patches with different number of layers, viz. 5, 11, 17, and 21, were assembled by the spin assisted LbL coating technique of oppositely charged CH and ALG polyelectrolytes onto the OBC cellulosic substrate loaded with DEX. These patches with a stratified structure had thermal stability up to almost 200 °C, mechanical properties with a Young's modulus higher than 4 GPa, and good moisture-uptake capacity (240–250%). Additionally, they inhibited the growth of *Staphylococcus aureus* with 3.2–log *CFU* mL–1 reduction and are non-cytotoxic to human keratinocytes (HaCaT cells) with a cell viability of about 97% after 48 h. The in vitro DEX release profile from the patches is time-dependent and is prolonged with the increasing number of layers, and the in vitro wound healing assay showed a good migration capacity with the full occlusion of the wound scratch after 24 h. These data validate the potentiality of these antibacterial multilayered polysaccharides-based patches as vehicles for the topical delivery of DEX, which is a promoter of wound healing. Overall, and in the case of the patches, the HaCaT keratinocyte cells situated near the boundary of the scratched region moved toward the vacant region as time progressed, achieving a full closure after 24 h. Furthermore, the cell density at the back of the scratch site is also growing with time, which points to the proliferation of the HaCaT keratinocyte cells. However, in terms of temporal progression,the patches containing DEX, namely OBC/DEX, OBC/DEX/CH/ALG\_11 and OBC/DEX/CH/ALG\_21, demonstrated higher coverage of the wound-like gap area after 9 h, induced by cell growth and migration, when compared to the positive control. Despite the analogous behavior of the HaCaT keratinocyte cells after 24 h of exposure to the three patches (Figure 6), the optical micrographs after 9 h show a slightly slower cell growth and migration for the patches with 11 and 21 layers. In fact, the behavior of the different patches after 9 h of exposure (Figure 6) seems to be in line with the in vitro drug release profile of DEX, where the OBC/DEX patch is the one releasing the higher amount of DEX in the shorter period of time and the multi-layered OBC/DEX/CH/ALG\_21 membrane with a controlled drug release profile is the one releasing the lower content of DEX after 24 h, as depicted in Figure 4b. Similar observations in terms of wound closure as a function of time were documented by Sharma et al. [48] for curcumin conjugated with hyaluronic acid, which are two natural compounds known to expedite the wound healing process, and by Silva et al. [16] for multifunctional nanofibrous patches composed of nanocellulose and lysozyme nanofibers.

Given that wound closure is a signal of wound healing ability [49], these antibacterial and non-cytotoxic multi-layered polysaccharide-based patches have the potential to foster wound healing and, simultaneously, hinder skin infections. Moreover, the drug release profile can be tuned to the specific type of wound and required treatment time by simply adjusting the number of polysaccharide layers.

#### **4. Conclusions**

Functional multi-layered patches composed of OBC, CH, ALG, and DEX were prepared for wound healing applications. Four multi-layered patches with different number of layers, viz. 5, 11, 17, and 21, were assembled by the spin assisted LbL coating technique of oppositely charged CH and ALG polyelectrolytes onto the OBC cellulosic substrate loaded with DEX. These patches with a stratified structure had thermal stability up to almost 200 ◦C, mechanical properties with a Young's modulus higher than 4 GPa, and good moisture-uptake capacity (240–250%). Additionally, they inhibited the growth of *Staphylococcus aureus* with 3.2–log *CFU* mL−1 reduction and are non-cytotoxic to human keratinocytes (HaCaT cells) with a cell viability of about 97% after 48 h. The in vitro DEX release profile from the patches is time-dependent and is prolonged with the increasing number of layers, and the in vitro wound healing assay showed a good migration capacity with the full occlusion of the wound scratch after 24 h. These data validate the potentiality of these antibacterial multi-layered polysaccharides-based patches as vehicles for the topical delivery of DEX, which is a promoter of wound healing.

**Author Contributions:** Conceptualization, C.V. and C.S.R.F.; Investigation, D.F.S.F., J.P.F.C., V.B., C.M. and C.V.; Resources, H.O., A.A., A.J.D.S. and C.S.R.F.; Writing—original draft preparation C.V.; Writing—review and editing, D.F.S.F., J.P.F.C., V.B., H.O., C.M., A.A., A.J.D.S., C.V. and C.S.R.F.; Supervision, C.V. and C.S.R.F.; Funding acquisition, H.O., A.A., A.J.D.S. and C.S.R.F. All authors have read and agreed to the published version of the manuscript.

**Funding:** This work was developed within the scope of the projects CICECO-Aveiro Institute of Materials (UIDB/50011/2020 & UIDP/50011/2020) and CESAM (UIDB/50017/2020), financed by national funds through the Portuguese Foundation for Science and Technology (FCT)/MCTES. FCT is also acknowledged for the doctoral grant to D.F.S.F. (PD/BD/115621/2016), and the research contracts under Scientific Employment Stimulus to H.O. (CEECIND/04050/2017) and C.V. (CEECIND/00263/2018). The AgroForWealth project (CENTRO-01-0145-FEDER-000001) is acknowledged for the research grant to J.P.F.C., while the research contract of V.B. (CDL-CTTRI-161-ARH/2018) was funded by the FCT project POCI-01-0145-FEDER-031794.

**Conflicts of Interest:** The authors declare no conflict of interest.

#### **References**

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© 2020 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 (http://creativecommons.org/licenses/by/4.0/).

## *Article* **The Effect of High Lignin Content on Oxidative Nanofibrillation of Wood Cell Wall**

**Simon Jonasson 1 , Anne Bünder 2 , Linn Berglund 1 , Magnus Hertzberg 3 , Totte Niittylä 2 and Kristiina Oksman 1,4,\***

**Abstract:** Wood from field-grown poplars with different genotypes and varying lignin content (17.4 wt % to 30.0 wt %) were subjected to one-pot 2,2,6,6-Tetramethylpiperidin-1-yl)oxyl catalyzed oxidation and high-pressure homogenization in order to investigate nanofibrillation following simultaneous delignification and cellulose oxidation. When comparing low and high lignin wood it was found that the high lignin wood was more easily fibrillated as indicated by a higher nanofibril yield (68% and 45%) and suspension viscosity (27 and 15 mPa·s). The nanofibrils were monodisperse with diameter ranging between 1.2 and 2.0 nm as measured using atomic force microscopy. Slightly less cellulose oxidation (0.44 and 0.68 mmol·g −1 ) together with a reduced process yield (36% and 44%) was also found which showed that the removal of a larger amount of lignin increased the efficiency of the homogenization step despite slightly reduced oxidation of the nanofibril surfaces. The surface area of oxidized high lignin wood was also higher than low lignin wood (114 m2 ·g −1 and 76 m2 ·g −1 ) which implicates porosity as a factor that can influence cellulose nanofibril isolation from wood in a beneficial manner.

**Keywords:** cellulose nanofibrils; wood; lignin; TEMPO-oxidation

#### **1. Introduction**

The valorization of wood and other lignocelluloses generally include a variety of costly mechanical-chemical refining steps in order to overcome the inherent recalcitrance that comes from lignified cell walls and complex inaccessible crystalline cellulose microfibril architectures [1]. Effectively nanofibrillating the cell wall is an attractive prospect where cellulose nanofibrils (CNFs) are attainable as a final product or as an intermediate for further treatments that utilizes the increased accessibility [2]. For instance, through improved saccharification of nanofibrils compared to initial wood fibers [3]. An important factor that has made CNF isolation more favorable is the implementation of various pretreatments in addition to traditional pulping processes. These swell the cell wall structure and reduces its structural integrity, thereby making it more susceptible to complete nano-fibrillation. The choice of pretreatment, cellulosic feedstock and mechanical disintegration method varies across literature and is a subject of intense research activity [4]. A common process in this context is that of 2,2,6,6-Tetramethylpiperidin-1-yl)oxyl (TEMPO) catalyzed oxidation. This pretreatment mediates nanofibrillation through selective carboxylation of the cellulose C6-hydroxyl groups through N-oxyl radical generation [5]. The treatment has been studied in great depth in relation to delignified pulps from a variety of sources [6]. It has also been demonstrated that feedstock containing lignin, such as thermomechanical pulp [7], hemp [8] and wood [9], can be simultaneously delignified and carboxylated using the

**Citation:** Jonasson, S.; Bünder, A.; Berglund, L.; Hertzberg, M.; Niittylä, T.; Oksman, K. The Effect of High Lignin Content on Oxidative Nanofibrillation of Wood Cell Wall. *Nanomaterials* **2021**, *11*, 1179. https:// doi.org/10.3390/nano11051179

Academic Editor: Carla Vilela

Received: 23 March 2021 Accepted: 21 April 2021 Published: 29 April 2021

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treatment. The relation between these procedures and nanofibrillation is complex where CNFs with residual lignin are attainable despite substantial carboxylation [10,11]. It has also been reported that the presence of the TEMPO-catalyst significantly increases carboxylation whilst keeping the delignifying efficiency constant [7]. The understanding of how these phenomena occurs in relation to native wood structures with varying lignin is currently unexplored. Available literature revolves around already processed feedstocks or focuses on pure mechanical fibrillation [12]. A natural step would thus be to combine TEMPOoxidation with variation in lignin content of unprocessed feedstocks. The associated novelty includes highlighting TEMPO-oxidation as a metric of fibrillation suitability for raw wood with variation in lignin content, something which could be applicable to commercial endeavors where TEMPO-oxidation is an attractive prospect in obtaining CNFs. Direct TEMPO-oxidation of wood has only recently been the subject of few dedicated studies [9,13]. These studies were furthermore showcasing one type of wood whereas the study presented herein aims to explore an additional factor of wood characteristics. This can increase the knowledge and feasibility of direct oxidation of wood by elucidating how wood properties, in this case native lignin content, influences the process.

In this study, we investigated field-grown poplar trees with large (17.4–30.0 wt %) systematic differences in lignin content. The goal was to analyze how lignin content associated wood recalcitrance is manifested in relation to nanofibrillation after pretreatments that simultaneously delignifies and carboxylates the wood fibers. Increased understanding of how oxidative treatments interplay with variation in lignin content of the feedstock is beneficial to a wide range of CNF isolation endeavors through elucidation of the link between wood and final CNFs.

#### **2. Materials and Methods**

#### *2.1. Materials*

Six different wood samples with specific chemical composition (see Table 1) were supplied by SweTree Technologies AB, Umeå Sweden. The samples were selected out of a library of trees (see Supplementary Material) based on the largest variety of total lignin content as determined using pyrolysis GS/MS [14]. The variation was validated using principal component analysis (see Supplementary Material). Three wood samples with similar lignin content were chosen as center points, approximately corresponding to lignin content in between the lowest and the highest samples. This was done in order to be able to evaluate trends between lignin content and characterization if present.

**Table 1.** Composition of the wood feedstock used in this study shown in wt %.

1 as determined using Test Methods T 203 C TAPPI-standard [15] with three technical replicates on the basis of dried chlorite holocellulose.

High-purity sodium chlorite (77.5–82.5%), standard hydrochloric acid solution (0.5 N), standard sodium hydroxide solution (0.1 N), sodium hydroxide beads (≥97%, ACS) were purchased from VWR, Solna, Sweden. TEMPO (99%), sodium hypochlorite (NaClO, 6–14% active chlorine) and Congo red (analytical standard) were purchased from Sigma-Aldrich, Stockholm, Sweden AB. All chemicals were used as received.

#### *2.2. Oxidation and Homogenisation*

Wood powder of the grinded poplar trees were sieved through a 300–500 µm mesh prior to sampling (2 g) and soaking in (100 mL) distilled water for 24 h prior to further

treatment. Primary oxidant sodium chlorite 5.0 g·g −1 wood and TEMPO (17.5 mg·g −1 wood) was added together with a phosphate buffer (0.1 M, 100 mL, pH = 6.8) to the vessel (250 mL) containing the wet powder. Each vessel with respective suspension was closed and submerged into a SBS40 shaking water bath (Cole-Parmer Stuart, Staffordshire, United Kingdom) at 60 ◦C for 1 h at 200 rpm in order to dissolve NaClO2 and TEMPO. Shaking water bath was used to eliminate experimental error from variation in heating rate and stirring and to make sure each tree sample was subjected to identical processing conditions. The reaction was initiated through addition of sodium hypochlorite (2 mL·g −1 wood) and then left for 48 h. The methodology was adapted from [16] with changes to the amount of primary oxidant and feedstock characteristics. The fully delignified and carboxylated wood was filtered thoroughly and then homogenized one pass without recirculation using an APV-2000 high-pressure homogenizer (SPX Flow Inc, Silkesborg, Denmark) with an average flow rate of 4 mL·s −1 at a pressure of 1000 bar. The concentration of respective sample was adjusted prior to and after homogenization to eliminate unwanted effects from variation in solid content.

#### *2.3. Viscosity of CNF Suspensions*

The viscosity of the oxidized and homogenized suspensions was measured at the same concentrations (0.183 ± 0.01 wt %) using a Vibro Viscometer (SV-10, A&D Company Limited, Tokyo, Japan) with a tuning fork vibration method at a vibrational frequency of 30 Hz. The measurements were repeated three times at room temperature for each sample.

#### *2.4. Carboxylate Content*

Carboxylate content was analyzed through the electric conductivity titration method adapted from [17]. 150 mL of respective CNF (cellulose nanofibril) suspensions (≈0.2 wt %) were protonated through addition of 0.1 M hydrochloric acid and 0.01 M sodium chloride. The suspensions were titrated with fresh 0.01 M sodium hydroxide through 0.5 mL increments until pH 10 was reached. The amount of carboxylate groups (mmol·g −1 ) induced from TEMPO-oxidation was calculated through Equation (1) below where c is the concentration of the sodium hydroxide, V2 and V1 is the volume of added sodium hydroxide at the end and start, respectively, m is the mass of the cellulosic material in the sample, calculated through subtraction of added acid, base and salt mass from the oven dried suspension. The titration was repeated three times.

$$\text{Carboxylate groups} \left(\frac{\text{mmol}}{\text{g}}\right) = \frac{\text{C}(\text{V}\_2 - \text{V}\_1)}{\text{m}} \tag{1}$$

#### *2.5. Atomic Force Microscopy*

Tapping-mode atomic force microscopy (AFM) Veeco MultiMode scanning probe, Santa Barbara, CA, USA, was used to confirm the presence of nanofibrils and analyze the morphology. Antimony-doped silicon cantilevers (TESPA-V2, Bruker, Camarillo, CA, USA) with a spring constant of 42 N·m−1 and a nominal tip radius of 8 nm were used for the analysis. Samples were prepared by depositing a small droplet of the CNF suspension (0.001 wt %) on a freshly cleaved mica plate and letting it air dry for ≥ 5 h. Around 100 fully individualized nanofibrils, total from four micrographs were analyzed for each sample and presented as a mean.

#### *2.6. Process and Nanofibril Yield*

Process yield was calculated as the washed gravimetric yield after the chemical treatment relative dry wood mass. The fraction of the process yield that comprised individual, colloidally stable nanofibrils were further quantified through centrifugation of the suspensions at 12,000× *g* (Avanti J25i, Beckman Coulter Inc. Brea, CA, USA) for 20 min at an approximate consistency of 0.2 wt %. The suspensions were decanted and the solids retained in the sediment were dried for 24 h at 95 ◦C. This was repeated three times and the

nanofibril-fraction was then calculated according to the Equation (2), where mp and ms is the dry precipitate and supernatant mass in the sample, respectively. The nanofibril-fraction (Φ) was presented as fraction of the process yield.

$$\Phi = 1 - \frac{\mathbf{m\_p}}{\mathbf{m\_p} + \mathbf{m\_s}} \tag{2}$$

#### *2.7. Thermogravimetric Analysis*

Dried CNF suspensions were subjected to thermogravimetric analysis TGA-Q500, TA Instruments, (New Castle, DE, USA) to analyze the relative amount of residue after pyrolysis of the oxidized wood samples and to compare the thermal stability of the CNFs. A heating rate of 10 ◦C·min−1 from room temperature to 900 ◦C was used in a nitrogen atmosphere.

#### *2.8. Klason Lignin and Cellulose Content*

Cellulose content of the wood holocellulose and dried CNF suspensions were calculated based on extraction using 17.5 M NaOH according to TAPPI-standard [15], where the oven dried gravimetric yield after filtration was used to estimate α-cellulose content. Klason lignin for the dry CNF suspensions was analyzed according to standard [18] through hydrolysis using sulfuric acid

#### *2.9. Network Manufacturing*

CNF suspensions were degassed for 30 min in a vacuum oven prior to vacuum filtration on hardened filter paper (Whatman, Grade 52, GE Healthcare, Machelen, Belgium, Pore size: 7 µm). The wet networks were carefully peeled from the filter paper and dried to around 14 % solid content prior. The stable networks were placed between two blotting papers and pressed (2 kN·m−2 ) for 10 h. The dry networks were further compression molded using Fontijne Grotnes LPC-300 (Vlaardingen, Netherlands), between two mylar films (Lohmann Technologies, Knowl Hill, UK) at a pressure of 0.32 MPa and at a temperature of 120 ◦C.

#### *2.10. X-ray Diffraction*

An X-ray diffractometer (PANalytical, Almelo, Netherlands) equipped with a PIXcel3d detector was used to measure the degree of crystallinity of the dried nanofibrils. Cu-Kα radiation (λ = 0.154 nm) was used during analysis with operation voltage of 45 kV and a current of 40 mA. The crystallinity index (CI) was calculated according to the peak height method after baseline correction [19]. This was done according to Equation (3), where I200 is the intensity of the crystalline (200) peak at 2θ = 22.4 and Iam is the intensity minimum between nearby crystalline peaks at 2θ = 18.

$$\text{CI} \left( \% \right) = \frac{\text{I}\_{200} - \text{I}\_{\text{am}}}{\text{I}\_{200}} \times 100 \tag{3}$$

#### *2.11. Mechanical Testing*

The dry networks were cut into rectangular samples (40 mm × 5 mm) using a mechanical punch. The samples were then stored at 50% RH for at least two days prior to mechanical characterization. Mechanical testing was performed using a Shimadzu AG-X universal testing machine (Kyoto, Japan) with a 500 N load cell. Testing was performed at a crosshead speed of 10%·min−1 , with the strain being measured using a video extensometer (high-speed camera, HPV-X2). The gauge length was set to 20 mm for each measurement. Seven specimens were analyzed for each network batch. The tensile strength was reported as the maximum strength at break. Young's modulus was calculated from the slope of the stress–strain curve at early strain (0.1–0.5 %).

#### *2.12. Sample Porosity and Moisture Content*

Density of the networks were measured by calculation based on the specimen volume and weight. Porosity (P) was derived using Equation (4), where ρs and ρc is the density of the specimen and cellulose (1.5 g·cm−3 ), respectively [20] Moisture content of the specimen was calculated from difference in weight after 24 h in oven at 105 ◦C

$$P = 1 - \frac{\rho\_s}{\rho\_c} \tag{4}$$

#### *2.13. Wood Porosity Analysis*

Intact wood samples at the extreme values of initial lignin content (17.4 and 30.0%) were oxidized in an identical manner as described in Section 2.2. The oxidized samples were carefully washed by submerging the samples into 20 L of distilled water. The fragile delignified and carboxylated samples were left to soak for four hours and the process was repeated three times with fresh distilled water. Brunauer–Emmett–Teller (BET) specific surface area analysis of the samples were performed by first freezing the wet samples at −20 ◦C for 10 h prior to freeze-drying for 72 h using Alpha 1-2 LD plus (Martin Christ Gefriertrocknungsanlagen GmbH, Osterode am Harz, Germany). The samples were degassed overnight at 120 ◦C followed by N2 physiosorption using a Micromeritics Gemini VII surface area analyzer (Norcross, GA, USA). A total of 30–60 mg of oxidized wood sample was analyzed for a respective sample.

Surface area of the oxidized wood was also estimated in the wet state by analyzing congo red absorption of the cellulosic fibers as described by [12]. Briefly, the absorbed dye was quantified using a UV-vis spectrophotometer (GENESYS, 10 UV, Thermo Scientific, Schwerte, Germany) at the absorption maxima 500 nm and then translated to surface area (SSA) using Equations (5) and (6), where E is the solution concentration of congo red at equilibrium (mg·mL−1 ), Amax is the absorbed amount of congo red on the sample (mg·g −1 ), Kad is the equilibrium constant, N is Avogadro's constant, SA is the theoretical surface area of a congo red molecule (1.73 nm2 ) and MW is the molecular weight of congo red (697 g·mole−1 ).

$$\frac{\mathbf{E}}{\mathbf{A}} = \frac{1}{\mathbf{K}\_{\text{ad}}\mathbf{A}\_{\text{max}}} + \frac{\mathbf{E}}{\mathbf{A}\_{\text{max}}} \tag{5}$$

$$\text{SSA} = \frac{\text{A}\_{\text{max}} \times \text{N} \times \text{SA}}{\text{MW} \times 10^{21}} \tag{6}$$

Samples cut in the wet state were freeze-dried and analyzed using a scanning electron microscope JEOL (JSM-IT300, Tokyo, Japan) at an acceleration voltage of 5 kV. A 15 nm layer of platinum was sputtered on the samples prior to analysis.

#### **3. Results and Discussion**

#### *3.1. Fibrillation Efficiency and Nanofibril Characteristics*

Suspensions of oxidized and homogenized wood samples showed a difference in viscosity between 9 and 27 mPa·s (shown in Figure 1a) is indicative of a qualitatively higher degree of fibrillation [21]. A similar behavior is reflected in the fraction of the yield that comprises fine colloidally stable nanofibrils, (shown in Figure 1b). A total of 45–68 wt % of the yield was recovered upon fractionating. A higher recovery of nanofibrils from wood with higher lignin content can be seen which is supported by the increased viscosity. The process yield ranged between 36 and 44 wt % of dry initial wood and decreased with more initial lignin content of the wood.

This can be understood from the complete dissolution of the lignin after being subjected to the catalytic conditions in this study. Klason lignin difference was also negligible after the treatment where all samples had an estimated residual lignin content of <1 wt %, indicating that a full dissolution of lignin took place following oxidation. A similar enhanced delignification has been observed after treatment of thermomechanical pulp using the related TEMPO/NaBr/NaClO-system [7]. It is further noteworthy that the nanofibril

stars.

study [28].

CNFs.

yield relative initial dry wood is similar for high lignin containing wood despite lower relative cellulose content. This is indicative of effects associated with higher initial lignin content and is the subject of discussion in Section 3.3. process yield ranged between 36 and 44 wt % of dry initial wood and decreased with more initial lignin content of the wood.

*Nanomaterials* **2021**, *11*, 1179 6 of 14

**Figure 1.** Viscosity (**a**), process yield (**b**) and nanofibril yield (b, red) of CNF suspensions obtained from wood with varying initial lignin content. Data represent mean ± SD of six different CNF samples (*n* = 3 replicates per sample). Data points for respective samples indicated with black and red **Figure 1.** Viscosity (**a**), process yield (**b**) and nanofibril yield (b, red) of CNF suspensions obtained from wood with varying initial lignin content. Data represent mean ± SD of six different CNF samples (*n* = 3 replicates per sample). Data points for respective samples indicated with black and red stars.

This can be understood from the complete dissolution of the lignin after being subjected to the catalytic conditions in this study. Klason lignin difference was also negligible after the treatment where all samples had an estimated residual lignin content of <1 wt %, indicating that a full dissolution of lignin took place following oxidation. A similar enhanced delignification has been observed after treatment of thermomechanical pulp using Carboxylate content of the final suspensions is shown in Figure 2 together with AFM micrographs of the CNFs that comprises the suspensions. Carboxylate content ranged between 0.44 and 0.68 mmol·g −1 of cellulose and was higher for low lignin wood compared to high. The range of the height of the individualized CNFs was measured between 1.2 and 2.0 nm and are thus thinner than most models suggest for elementary fibrils of wood cellulose. Significant differences in height were only apparent when comparing low lignin wood (sample #1) to the rest. *Nanomaterials* **2021**, *11*, 1179 7 of 14 Figure 2. The correlation between the size distribution as obtained from AFM and other metrics of fibrillation is furthermore more accurate for highly fibrillated samples as in this

the related TEMPO/NaBr/NaClO-system [7]. It is further noteworthy that the nanofibril

crofibrils are the subjects of a variable oxidation throughout this study. This can be explained by looking at the oxidation process where lignin hydroxyl groups are efficiently oxidized and depolymerized by TEMPO-systems through cleaving of C-C and ether **Figure 2.** AFM micrographs (**a**) of the CNFs with corresponding height (**b**) and carboxylate content (**c**). Error bars in (**b**) derived from standard deviation of the size distribution. Data represent mean ± SD of six different CNF samples with 50–100 measurements per sample. Data points for **Figure 2.** AFM micrographs (**a**) of the CNFs with corresponding height (**b**) and carboxylate content (**c**). Error bars in (**b**) derived from standard deviation of the size distribution. Data represent mean ± SD of six different CNF samples with 50–100 measurements per sample. Data points for respective samples indicated with black stars.

respective samples indicated with black stars.

of irregular shapes highlighted (**right**).

2014), which implies that the smaller individual elementary fibrils comprising these mi-

bonds [22]. Lignin is present in both the thin middle lamella and the cell wall layers of wood [23] and can expectedly be preferentially oxidized during the process. This would decrease the effectiveness of the cellulose oxidation since lignin is more accessible than From the AFM analysis there were also particles present across the samples, especially apparent in CNFs made from high lignin wood. A larger scale AFM micrograph of The relation between carboxylate content and initial lignin content indicates a decrease in carboxylation with more initial lignin of the feedstock although the correlation cannot be extrapolated to a larger range of carboxylation due to a rather narrow interval that the

into smaller nanofibrils and even individual polymer chains [25]. This fragmentation phenomenon has been reported by multiple authors [24–27]. The increased carboxylation taking place for the cellulose derived from low lignin wood would according to this hypothesis be related to the decreased height of the CNFs from low lignin wood as shown in

**Figure 3.** AFM micrograph of CNF (**left**) made from high lignin wood with particles and clusters

This is hypothesized to be residues from processing. Even though extensive filtration took place during the isolation process it is likely that trapping of non-cellulosic components occurs inside the cell wall during oxidation and liberated together with the CNFs during homogenization. This is further supported by an increase in char after pyrolysis of the samples as shown in Figure 4. A mass fraction between 0.16 and 0.38 was remaining after pyrolysis of nanofibrils with more residues remaining for CNFs made from high lignin wood. The thermal stability of all CNFs was also similar as shown in Figure 4 and this study [28].

carboxylation ranges within. Interestingly, at 0.37 mmol·g −1 it has been calculated that all the surfaces of native cellulose microfibrils have undergone oxidation (Sjöstedt, 2014), which implies that the smaller individual elementary fibrils comprising these microfibrils are the subjects of a variable oxidation throughout this study. This can be explained by looking at the oxidation process where lignin hydroxyl groups are efficiently oxidized and depolymerized by TEMPO-systems through cleaving of C-C and ether bonds [22]. Lignin is present in both the thin middle lamella and the cell wall layers of wood [23] and can expectedly be preferentially oxidized during the process. This would decrease the effectiveness of the cellulose oxidation since lignin is more accessible than cellulose and the process is defined by a specified amount of chlorite (5 g·g −1 wood). This has been reported previously [24] and explained through splitting of the elementary fibril into smaller nanofibrils and even individual polymer chains [25]. This fragmentation phenomenon has been reported by multiple authors [24–27]. The increased carboxylation taking place for the cellulose derived from low lignin wood would according to this hypothesis be related to the decreased height of the CNFs from low lignin wood as shown in Figure 2. The correlation between the size distribution as obtained from AFM and other metrics of fibrillation is furthermore more accurate for highly fibrillated samples as in this study [28]. **Figure 2.** AFM micrographs (**a**) of the CNFs with corresponding height (**b**) and carboxylate content (**c**). Error bars in (**b**) derived from standard deviation of the size distribution. Data represent mean ± SD of six different CNF samples with 50–100 measurements per sample. Data points for respective samples indicated with black stars. From the AFM analysis there were also particles present across the samples, especially apparent in CNFs made from high lignin wood. A larger scale AFM micrograph of

*Nanomaterials* **2021**, *11*, 1179 7 of 14

Figure 2. The correlation between the size distribution as obtained from AFM and other metrics of fibrillation is furthermore more accurate for highly fibrillated samples as in this

From the AFM analysis there were also particles present across the samples, especially apparent in CNFs made from high lignin wood. A larger scale AFM micrograph of that sample is shown in Figure 3, where particles and clusters are scattered among the CNFs. that sample is shown in Figure 3, where particles and clusters are scattered among the CNFs.

**Figure 3.** AFM micrograph of CNF (**left**) made from high lignin wood with particles and clusters of irregular shapes highlighted (**right**). **Figure 3.** AFM micrograph of CNF (**left**) made from high lignin wood with particles and clusters of irregular shapes highlighted (**right**).

This is hypothesized to be residues from processing. Even though extensive filtration took place during the isolation process it is likely that trapping of non-cellulosic components occurs inside the cell wall during oxidation and liberated together with the CNFs during homogenization. This is further supported by an increase in char after pyrolysis of the samples as shown in Figure 4. A mass fraction between 0.16 and 0.38 was remaining after pyrolysis of nanofibrils with more residues remaining for CNFs made from high lignin wood. The thermal stability of all CNFs was also similar as shown in Figure 4 and this This is hypothesized to be residues from processing. Even though extensive filtration took place during the isolation process it is likely that trapping of non-cellulosic components occurs inside the cell wall during oxidation and liberated together with the CNFs during homogenization. This is further supported by an increase in char after pyrolysis of the samples as shown in Figure 4. A mass fraction between 0.16 and 0.38 was remaining after pyrolysis of nanofibrils with more residues remaining for CNFs made from high lignin wood. The thermal stability of all CNFs was also similar as shown in Figure 4 and this is in agreement with what commonly is reported for CNFs derived from the TEMPO/NaClO/NaClO2-treatment.

Pyrolysis of lignin generally leads to a higher char formation compared to cellulose and hemicellulose [29]. In the absence of lignin, however, as shown for these samples, the residues have to originate from another source. Higher amount of char formation from the pyrolysis of deprotonated CNFs have been reported, where a difference up to 16 wt % was reported solely from the deprotonation of the carboxylate groups [30]. This effect together

with the presence of residues from the oxidation explains this phenomenon and is further elaborated upon in Section 3.3 with porosity analysis of the TEMPO-oxidized wood. Thermal degradation occurs earlier compared to chlorite bleached wood, but later compared to cellulose that has undergone the harsher TEMPO/NaBr/NaClO-treatment [27]. The lack of variation amongst the samples is also supported by the carboxylate content (Figure 2), where the differences are relatively small in comparison to the whole range of carboxylate content that is possible to obtain after TEMPO-oxidation of all available hydroxyl groups on the nanofibril surfaces. *Nanomaterials* **2021**, *11*, 1179 8 of 14 is in agreement with what commonly is reported for CNFs derived from the TEMPO/NaClO/NaClO2-treatment.

**Figure 4.** Thermal degradation curves showing mass loss (**a**), derivative weight (**b**) mass loss curve with zoom-in (200–400 °C) (**c**) and corresponding mass fraction of residues remaining at 900 °C (**d**) **Figure 4.** Thermal degradation curves showing mass loss (**a**), derivative weight (**b**) mass loss curve with zoom-in (200–400 ◦C) (**c**) and corresponding mass fraction of residues remaining at 900 ◦C (**d**) Data points for respective samples indicated with black stars.

#### Data points for respective samples indicated with black stars. *3.2. Dry Network Characteristics*

wide range of CNFs [32].

Pyrolysis of lignin generally leads to a higher char formation compared to cellulose and hemicellulose [29]. In the absence of lignin, however, as shown for these samples, the residues have to originate from another source. Higher amount of char formation from the pyrolysis of deprotonated CNFs have been reported, where a difference up to 16 wt % was reported solely from the deprotonation of the carboxylate groups [30]. This effect together with the presence of residues from the oxidation explains this phenomenon and X-ray diffractograms of dried networks and corresponding degree of crystallinity is presented in Figure 5. The degree of crystallinity of the networks was similar at a range between 77 and 82% with a slight decrease from low to high lignin wood. The native cellulose Iβ crystal structure was retained after processing for all samples, as indicated by the diffractogram peak location [31]. Both the absolute value of crystallinity and tendencies for a decrease in crystallinity as a function of fibrillation agree with previous analysis of a wide range of CNFs [32].