European Journal of Pharmaceutical Sciences 66 (2015) 29–35
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European Journal of Pharmaceutical Sciences journal homepage: www.elsevier.com/locate/ejps
Nanofiber diameter as a critical parameter affecting skin cell response Jan Pelipenko, Petra Kocbek, Julijana Kristl ⇑ University of Ljubljana, Faculty of Pharmacy, Ljubljana, Slovenia
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Article history: Received 25 July 2014 Received in revised form 28 August 2014 Accepted 28 September 2014 Available online 6 October 2014 Keywords: Nanotechnology Nanofibers Electrospinning Proliferation Regenerative medicine
a b s t r a c t Electrospun polymer nanofibers have opened new opportunities in the rapidly evolving field of tissue engineering, particularly due to their topography and variability of available biomaterials. In order to better understand nanofiber influence on cell growth, the impact of their diameter was systematically examined. In this study homogenous, randomly oriented poly(vinyl alcohol) nanofibers with five different average diameters, ranging from 70 nm to 1120 nm, were produced, characterized and their impact on morphology, proliferation and mobility of keratinocytes and skin fibroblasts was evaluated. The results have shown that nanofiber diameter affects cell response and that this response is cell line specific. Nanofiber thickness affected size, morphology and actine organization of keratinocytes much more than fibroblasts. Specifically, the keratinocyte grown on nanofibers were more spherical and smaller compared to the control cells, while the fibroblasts were much less affect. They stayed almost unchanged and spread across growth surface. The cell proliferation determined based on their metabolic activity was the highest, when keratinocytes were grown on 305 nm thick nanofibers, whereas proliferation of fibroblasts grown similar nanofibers was decreased. Finally, fibroblasts exerted higher mobility than keratinocytes. Both tested cell lines on nanofiber diameters of 300 nm resulted in decreased cell mobility. These findings suggest that the control over nanofiber diameter offers promising possibility to better design the tissue scaffolds, since cells distinguish between differently sized nanofibers and respond accordingly. Ó 2014 Elsevier B.V. All rights reserved.
1. Introduction Tissues are complex structures composed of highly organized populations of individual cells. The basic principle of coordinated cellular events is cell communication with the environment; meaning interactions between cells as well as interactions of cells with the natural extracellular matrix (ECM) (Frantz et al., 2010). ECM is protein-based structure mostly composed of collagen fibers that extend in length over tens of micrometers, have thickness between 260 and 410 nm, as well as topography and structural features specific for cell response (Bettinger et al., 2009; Huang et al., 2012). When natural ECM is disturbed, either by illness or injury, regeneration process should be established to restore the tissue. Regeneration of tissues can be achieved by the combination of living cells, which provide biological func-
Abbreviations: ECM, extracellular matrix; PVA, poly(vinyl alcohol); MEM, minimum essential medium; SEM, scanning electron microscopy; PBS, phosphate buffered saline. ⇑ Corresponding author at: University of Ljubljana, Faculty of Pharmacy, Aškercˇeva cesta 7, 1000 Ljubljana, Slovenia. Tel.: +386 1 47 69 521; fax: +386 1 42 58 031. E-mail address: [email protected] (J. Kristl). http://dx.doi.org/10.1016/j.ejps.2014.09.022 0928-0987/Ó 2014 Elsevier B.V. All rights reserved.
tionality, and materials, which support the cell proliferation. Since the cell response in vivo depends on biological signals received from the surrounding environment, it is of crucial importance for particular tissue regeneration to resemble those signals (Zani and Edelman, 2010; Kim et al., 2013). Therefore, the biocompatible materials used in tissue regeneration should simulate natural behavior of treated cells. Polymer nanofibers can be used for temporal replacement of natural ECM and its functions, since they can improve regenerative process (Engel et al., 2008; Pelipenko et al., 2013a). Therefore, nanofibers are intensively explored in the field of chronic wounds, where organism is, due to the imbalance between synthesis and degradation of ECM, incapable of producing functional ECM and orchestrating the cell response leading to tissue restoration (Alves et al., 2010). Understanding interactions between cells and engineered nanostructured biomaterials is crucial for successful tissue healing. Most of these biomaterials have been studied in terms of nanofiber production abilities (Thompson et al., 2007; Yördem et al., 2008; Rošic et al., 2012; Cramariuc et al., 2013; Casasola et al., 2014) and much less has been investigated about their nanotopography and role in the cell response (Beachley and Wen, 2010; Bacakova et al., 2011).
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The cell cytoskeleton is known to have an important role in mechanosensing and mechanotransduction. Recent research has shown that filopodia first probe the environment, then long-lasting focal adhesions are established and, finally the extension of cell protrusions occurs (Albuschies and Vogel, 2013). The focal contact cell structures are mostly composed of integrins, which have size in nanometer range (Anselme et al., 2010; Huang et al., 2012). Therefore, it is assumed that they can be differently influenced by nanofibers with different diameters. The interactions between integrins and the extracellular environment (ECM or nanofibers) can result in change of cell adhesion, morphology, proliferation and even cell orientation and mobility. Nanotopography seems to have an important impact on cell behavior; therefore, it is now being explored with the aim to optimize scaffolds for tissue regeneration. The aim of our study was to investigate how to select proper nanofiber diameter to promote cell proliferation in vitro, which would probably in vivo result in healing and tissue regeneration. Firstly, electrospun poly(vinyl alcohol) (PVA) nanofibers were produced and characterized. Secondly, the impact of nanofiber diameter on the response of keratinocytes and skin fibroblasts, namely cell morphology, proliferation, and migration, was investigated in turn for further development of an effective nanofibrillar scaffolds.
2. Materials and methods 2.1. Materials PVA (MowiolÒ 20–98, Mw 125,000 g/mol) was supplied by Sigma–Aldrich (Germany), ethanol (96%) from Kefo (Slovenia). Minimum essential medium (MEM), non-essential amino acids, L-glutamin, antibiotic/antimycotic solution, fetal bovine serum, phalloidin rhodamine, trypan blue solution and Triton X-100Ò were all supplied by Sigma (Germany). Hoechst 33342 was obtained from Riedel de Haen (Germany), Cell Titer 96Ò Aqueous One Solution Cell Proliferation Assay was from Promega (Madison, WI). All other chemicals used were of analytical grade. 6, 24 and 96 well plates were from TPP (Switzerland) and custom-made inserts used in cell mobility experiments were provided by Denis Štraus, s.p. (Slovenia).
2.2. Methods 2.2.1. Nanofiber preparation and characterization Aqueous PVA solution was placed in a plastic 20 ml syringe fitted with a metal needle with an inner diameter of 0.8 mm. A steady polymer solution flow rate was ensured by a syringe pump (model R-99E, Razel™ Scientific Instruments, USA) and the high voltage by a generator (model HVG-P60-R-EU, Linari Engineering s.rl. Italy). A planar stand covered with aluminum foil was used as a collector. Electrospinning parameters were as follows: polymer concentration 8–15%, w/w, relative humidity 2–60%, applied voltage 10–30 kV, and needle to collector distance 10–20 cm. In order to produce nanofibrillar supports with comparable thicknesses, the electrospinning process was performed for 10 min for each sample. After electrospinning nanofibers were thermally stabilized against dissolution in aqueous environment by dry heat at 160 °C for 30 min. The morphology of the stabilized nanofibers was determined using a scanning electron microscopy (SEM, Supra 35 VP, Carl Zeiss, Oberkochen, Germany) at an accelerating voltage of 1 kV and a secondary electron detector. Based on SEM images the average nanofiber diameter and average interfibrillar pore size were determined
by measuring 40 randomly selected nanofibers or interfibrillar pores using ImageJ 1.44p software (NIH, USA).
2.2.2. Cell culture and treatment Immortalized human keratinocytes (cell line NCTC2544, ICLC, University of Genoa, Italy) or human skin fibroblasts (cell line 149BR, Public Health England, United Kingdom) were cultured as adherent monolayers at 37 °C in a humidified atmosphere of 5% CO2 in air. They were grown in MEM supplemented with 1% (v/v) non-essential amino acids, 2 mM L-glutamin, and 100 U/ml antibiotic/antimycotic and 10% (v/v) and 15% (v/v) FBS for keratinocytes and fibroblasts, respectively. The cells were regularly subcultured when reaching 80% confluency. Their viability was tested using trypan blue exclusion assay. The experiments were performed using cell seeding density 2 104 cells/cm2 and the final cell confluency reached in the experiments did not exceed 80%.
2.2.2.1. Examination of cell morphology. The cells were grown for 3 days on nanofibrillar supports, then the cells were fixed with 4% paraformaldehyde in phosphate buffered saline (PBS, pH 7.4) for 10 min and then permeabilized with 0.1% Triton X-100Ò in PBS (pH 7.4) for 10 min. The cell nuclei were stained with the DNA intercalating dye Hoechst 33342 (5 lg/ml) for 30 min protected from light. The dye used for cell nuclei staining simultaneously stained the polymer nanofibers. Actin filaments were stained with the red-fluorescent dye phalloidin rhodamine, according to the manufacturer’s procedure. Afterwards, coverslips were fixed on the slides and analyzed under fluorescence microscope using 360/420 nm (Hoechst 33342) and 535/635 nm (Phalloidin rhodamine) excitation/emission filter sets (Olympus IX81, Tokyo, Japan). Average size of cells was determined based on analysis of 40 randomly chosen cells. Based on SEM images the average nanofiber diameter and average interfibrillar pore size were determined by measuring 40 randomly selected nanofibers or interfibrillar pores using ImageJ 1.44p software (NIH, USA).
2.2.2.2. Assessment of cell mobility. Nanofibers, electrospun on glass coverslip (/ = 15 mm), were transferred to the bottom of a 24-well plate and then custom-made insert with diameter of 1.5 mm was mounted onto nanofibrillar support, representing a model ‘‘wounded field’’ with a defined area of 1.77 mm2. Afterwards, cells (2.5 104 cells/well) were seeded and incubated at 37 °C and 5% CO2 in air for 24 h to attach on nanofibrillar support, then the insert was removed and cell mobility was evaluated after 3 and 5 days using an inverted light microscope (CKX31, Olympus, USA).
2.2.2.3. Cell proliferation. Nanofibers, electrospun on round glass cover slips (/ = 5 mm), were transferred to the bottom of a 96-well plate, then cells were seeded on them and incubated at 37 °C and 5% CO2 in air for 3 or 5 days. Their proliferation was determined by MTS assay according to the manufacturer’s procedure. The sample absorbance was measured at 490 nm using a Synergy H4 microplate reader (BioTec, USA).
2.2.3. Statistical analysis The results are expressed as means ± standard deviations. The statistical analysis was carried out using an independent samples Student’s t-test. A value of p < 0.05 was considered statistically significant.
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3. Results and discussion 3.1. Control over electrospinning parameters enabled production of various sized nanofibers Different parameters known to affect the electrospinning process, namely, polymer concentration, relative humidity, applied voltage and the distance between needle and collector, were varied (Table 1) in order to produce homogenous nanofibrillar samples differing in average fiber diameter. Although, all parameters investigated should influence on nanofiber diameter according to the literature data (Cramariuc et al., 2013; Pelipenko et al., 2013b; Rošic et al., 2013), our results have proven that the control over two parameters, namely polymer concentration and relative humidity, exerted the greatest impact on the fiber diameter and enabled preparation of nanofibers with controlled size, as demonstrated in supporting information. Randomly oriented PVA nanofibers with average diameters ranging from 69 nm to 1116 nm were produced (Fig. 1). The thinnest nanofibers were obtained using solution with low polymer concentration (8%, w/w) and high relative humidity (60%), and the thickest using solution with high polymer concentration (15%, w/w) and low relative humidity (2%). Nanofibers with diameters in between were produced using 10% (w/w) PVA
Table 1 Optimized electrospinning parameters used for production of PVA nanofibers and their corresponding diameters. Sample
A B C D E
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solution and appropriately adjusted relative humidity (Table 1). The correlation between nanofiber diameter and relative humidity was in line with our previous studies, reporting decrease in fiber diameter due to increase in relative humidity (Pelipenko et al., 2013b). In the case of high relative humidity the solvent evaporation rate is decelerated, leading to formation of thinner nanofibers. On the other hand, polymer concentration influences nanofiber diameter through the number of entanglements formed between polymer chains (Casasola et al., 2014; Rošic et al., 2013). More concentrated solution therefore resulted in thicker nanofibers, if electrospun at the same relative humidity (Table 1). All prepared nanofibers were homogenous and had a smooth surface, except nanofibers produced from the solution with the highest concentration i.e. 15% PVA (Fig. 1E), which exerted wrinkled surface. Such fibers could be beneficial for cell adhesion, since the area available for contact with cells is larger due to irregular fiber surface (Rošic et al., 2012). Besides thickness and planar arrangement of nanofibers representing surface for cell growth, also the interfibrillar pore size is important parameter for prediction of spatial cell mobility. The results showed big differences in average interfibrillar pore size in produced samples (Fig. 1). The electrospun PVA nanofiber mats composed of thicker nanofibers contained larger pore sizes compared to the nanofiber mats with thinner nanofibers. This difference had an important impact on cell response as explained in details in the following sections and should be considered when designing nanofibrillar materials for application in tissue regeneration.
PVA conc. (w/w, %)
Tipcollector distance (cm)
Applied voltage (kV)
Relative humidity (%)
Nanofiber’s diameter ± SD (nm)
3.2. Effect of nanofibers on cell morphology
8 10
15
15
60
69 ± 19 161 ± 42 305 ± 48 667 ± 83 1116 ± 354
Cells can change their morphology according to the properties of the substrate they are growing on. Keratinocytes cultured on nanofibers showed changed cytoskeletal organization compared to the control cells growing on a glass coverslip (Fig. 2). The control cells were well spreaded over the growth surface and tightly connected to each other. Those at the edges, formed focal contacts,
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Fig. 1. SEM images of nanofibers tested: (A) 69 ± 19 nm, (B) 161 ± 42 nm, (C) 305 ± 48, (D) 667 ± 83 nm and (E) 1116 ± 354 nm and corresponding average sizes of interfibrillar pores.
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marked with arrows in Fig. 2. The actin filaments in cells grown on nanofibers could not be clearly seen, contrary to the control cells with well-organized actin filaments. Not only the internal structure but also the average size of cells was affected by fiber diameter as follows: 20.9 ± 2.7 lm on 69 nm nanofibers, 19.0 ± 3.3 lm on 161 nm nanofibers, 14.5 ± 2.6 lm on 305 nm nanofibers, 15.9 ± 2.7 lm on 667 nm nanofibers, 25.9 ± 6.1 lm on 1116 nm nanofibers, and 45.3 ± 7.7 lm on flat glass coverslip. The cells grown on thinnest nanofibers were more spherical and significantly smaller compared to the control cells, whereas the morphology of keratinocytes grown on thicker nanofibers was governed by spatial nanofiber arrangement. In the case of thicker nanofibers, the interfibrillar pores were bigger (Fig. 1) and therefore enabled partial cell infiltration into the nanofibrillar support, which consequently affected the cell morphology (Guimarães et al., 2010). More rounded cell morphology on randomly structured nanofibers compared to the cell morphology on glass coverslip has been reported previously (Pelipenko et al., 2013a). Contrary to keratinocytes, the morphology and actin organization of fibroblasts was much less affected by nanofiber diameter (Fig. 2). The cells were spread across the growth surface with average size of 80 lm along their length. The ability of fibroblasts to spread unhindered across nanosized obstacles has been reported also by Dalby and coworkers, who explored fibroblast mobility on nanocolumns (160 nm high, 100 nm wide) (Dalby et al., 2004).
3.3. Effect of nanofibers on cell mobility
Fig. 2. Keratinocytes and skin fibroblasts grown on nanofibers with different diameters and control cells grown on a glass coverslip. Actine filaments were stained using phalloidin red, whereas cell nuclei and nanofibers using Hoechst 33342. Due to the weak fluorescence of nanofibers and interference with other fluorophore, nanofibers are not well seen in the images of merged fluorescent signals; therefore, they are presented also separately. The arrows indicate focal contacts between cells and underlying glass coverslip.
The term cell mobility describes the mobility as a result of cell proliferation and spreading of daughter cells across the growth substrate and not the movement of a single cell across the growth substrate (Bertoncelj et al., 2014). The cell mobility was evaluated based on the comparison of the size of the ‘‘wounded field’’ at the investigated time point and its size at the beginning of the experiment (Fig. 3, t = 0 d). However, the effect of the cell size was not eliminated and could have also influenced the speed of ‘‘wounded field’’ closure; therefore, it was treated as a contribution to the overall cell mobility. In both tested cell lines, the highest cell mobility was observed on nanofibrillar supports composed of thin nanofibers (69 and 161 nm) (Fig. 3). Further increase in size of nanofibers resulted in decreased cell mobility. When nanofibers were thicker, interfibrillar pores were larger (Fig. 1) and cells could have penetrated partially into the nanofibrillar support, resulting in reduced overall cell mobility. In comparison to the control cells grown on a glass coverslip, cell mobility on nanofibers was reduced, which could be explained by physical entrapment of cells in the nanofibrillar network (Guimarães et al., 2010; Pelipenko et al., 2013a) and improved focal cell adhesion (Toh et al., 2006; Albuschies and Vogel, 2013). Another reason for lower cell mobility on nanofibers in comparison to the glass coverslip could be stiffness of the nanofibrillar support (Wang et al., 2012), which explains also the differences in cell mobility on various sized nanofibers. In one of our previous studies, we demonstrated that nanofiber stiffness strongly depends on nanofiber diameter (Jankovic´ et al., 2013). When cells are cultured on a soft substrates, such as polyacrylamide gels, cell–cell interactions are favoured over cell interactions with supporting substrate (Alves et al., 2010), resulting in lower cell mobility. However, when the supporting growth substrate becomes stiffer, interactions between cell and supporting substrate prevail and increased cell mobility is observed (Toh et al., 2006). The results of cell mobility are in line
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Fig. 3. The effect of nanofiber diameter on cell mobility. Cells grown on a flat glass coverslip were used as a control. The arrows indicate the area with increased cell density around the ‘‘wounded field’’.
with the size dependant mechanical behavior of nanofibers, i.e. the decrease in mobility (compared to the growth on the glass coverslip) on thicker nanofibers was more prominent than the decrease in cell mobility on thinner nanofibers. Furthermore, the area with locally increased cell density, visible as a darker ring around the ‘‘wounded field’’, was observed and was the most prominent in the case of keratinocytes cultured 5 days on 667 nm nanofibers (Fig. 3), indicating increased cell proliferation, which could be attributed to hindered contact inhibition at the edge of the model ‘‘wounded field’’ (Gray et al., 2008). The higher mobility of fibroblasts, in comparison to keratinocytes, can be explained by the fact that fibroblasts are larger than keratinocytes (80 lm compared to 40 lm, when cultured on glass coverslip) and, therefore, spread over larger surface area. Additionally, fibroblasts are exposed to the nanofibrillar structures of ECM in their natural environment and are in the case of injury forced to migrate toward wounded area and produce elements of new ECM. On the other hand, the main function of keratinocytes is to form densely packed cell layers, namely a barrier epithelium over newly formed cell layers and ECM in dermis (Reinhart-King et al., 2008). This phenomenon was observed also in vitro through the manner ‘‘wounded field’’ was closing. In the case of keratinocytes, the ‘‘wounded field’’ was closing due to the even cell proliferation from the edge of the ‘‘wounded field’’. In the case of fibroblasts, the irregular pattern of cell penetration in ‘‘wounded field’’ caused difficulties in experimental determination of ‘‘wounded field’’ size over the time, indicated also by high standard deviations of the measurements (Fig. 3). To sum up, cells cultured on thinner nanofibers exerted higher mobility than those cultured on thicker nanofibers. However, the
mobility was still much lower compared to the cell mobility on a glass coverslip. 3.4. Effect of nanofibers on cell proliferation Keratinocyte and fibroblast proliferation evaluated indirectly via cell metabolic activity was affected by nanofiber diameter as shown in Fig. 4. In the case of keratinocytes, the highest metabolic activity was observed for cells growing on 305 nm thick nanofibers at both tested time points, however, it was not significantly different from the metabolic activity of keratinocytes cultured on 667 nm thick nanofibers. The lowest metabolic activity i.e. significantly lower than metabolic activity of cells grown on all other investigated nanofibers, was observed for cells grown on 69 nm nanofibers. The obtained results correlate well with changes observed in average cell size. Faster cell proliferation resulted in smaller cells. The smallest cells (15.5 ± 2.6 lm) were observed in sample grown on 305 nm nanofibers, where the highest metabolic activity was also determined. Increase in fiber diameter up to 1116 ± 12.2 nm resulted in less stimulated cell proliferation, due to decreased resemblance between nanofibers and elements of natural ECM, which are mostly 260–410 nm in size (Bettinger et al., 2009). Fibers thicker than 1000 nm do not stimulate cell proliferation as efficient as thinner ones as reported previously (Leong et al., 2009). In the case of fibroblasts cultured on 69 and 161 nm thick nanofibers the proliferation was significantly decreased at both investigated time points, while on thicker nanofibers it was comparable to the growth of control cells. Higher degree of cell differentiation and slower cell metabolism are two key characteristics,
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(Wang et al., 2012; Yeh et al., 2012). However this is not in line with our results, indicating the highest keratinocyte proliferation on 305 nm thick nanofibers. Furthermore, the cell proliferation is expected to be decreased with increase in nanofiber diameter, because nanofibers exert size-dependant mechanical properties i.e. increased nanofiber stiffness with decrease in their diameter (Jankovic´ et al., 2013). This conclusion holds true only for keratinocytes grown on investigated nanofibers smaller than 1000 nm. Additionally, it was shown that the cell response does not depend only on stiffness of growth surface, but is also dependent on surface morphology i.e. dimensions of underlying nanofibers (Alves et al., 2010), confirmed also in our study. Cells are complex entities with a remarkable, inherent capacity to sense, integrate, and respond to environmental stimuli, such as nanotopography of growth surface. The current in vitro research was performed separately on keratinocytes and skin fibroblasts and has revealed some important results, which represent a good starting point for the future more complex in vitro studies using keratinocyte and fibroblast co-cultures, which better resemble in vivo conditions. Interplay and orchestrated activity between both cell types is important for wound healing in vivo, therefore, results obtained on individual cell lines indicate the direction of the cell response on nanofibers, however, prior more precise conclusions about the effects of nanofibers on wound healing in vivo can be made, the cell response in more complex environment should be systematically investigated. 4. Conclusions
Fig. 4. The effect of nanofiber diameter on keratinocyte and skin fibroblast metabolic activity determined 3 and 5 days after cell attachment. The cell metabolic activity is expressed relative to metabolic activity of cells grown on a flat glass coverslip.
which distinguish fibroblasts from keratinocytes and can explain their less prominent short term response due to the changed growth surface (Bertoncelj et al., 2014). Our results showed cell line and nanofiber diameter dependant cell proliferation on nanofibrillar supports, however, the stimulated cell proliferation caused by nanofibers is mainly reported in the literature (Fagotto and Gumbiner, 1996). According to Wang et al. and Yeh et al., the highest cell proliferation is expected on glass coverslip, since it represents the stiffest growth surface
The effect of nanofiber diameter on skin cell growth was systematically investigated over a broad range of fiber diameters, prepared by electrospinning of PVA solutions with adjusted polymer concentration and precise regulation of relative humidity during production process. It has been shown that nanofiber diameter affects cell response, which is also cell line specific. Keratinocytes are more sensitive to nanofiber size than fibroblasts, since their morphology was changed due to growth on nanofibers, whereas morphological changes of fibroblasts were less obvious. Furthermore, the proliferation of keratinocytes was a function of nanofiber diameter, with a maximum reached, when cells were growing on 305 nm thick nanofibers. Proliferation of fibroblasts was reduced, when cells were grown on thin nanofibers, and was comparable to the growth on the glass coverslip, when fibers were thicker. Finally, it was proven that the cell mobility is decreased if cells are grown on thicker nanofibers. Base on the results, it can be concluded that nanofiber diameter is an important parameter, influencing cell response. Therefore, it should be precisely considered when nanofibrillar materials intended for application in biomedical field, such as wound dressing or tissue scaffolds, are being designed. Based on the obtained in vitro results combination of thin and thick nanofibers could be recommended for skin tissue regeneration, since thicker nanofibers (300–700 nm) stimulate cell proliferation, while cell mobility, resulting in wound closure, is higher on thinner nanofibers. Acknowledgments The authors gratefully acknowledge the financial support provided by the Slovenian Research Agency for Programme Number P1-0189, Projects J1-4236 and J1-6746, and Grant Number 100011-310213. Appendix A. Supplementary material Supplementary data associated with this article can be found, in the online version, at http://dx.doi.org/10.1016/j.ejps.2014.09.022.
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Huang, C., Fu, X., Liu, J., Qi, Y., Li, S., Wang, H., 2012. The involvement of integrin beta1 signaling in the migration and myofibroblastic differentiation of skin fibroblasts on anisotropic collagen-containing nanofibers. Biomaterials 33, 1791–1800. Jankovic´, B., Pelipenko, J., Škarabot, M., Muševicˇ, I., Kristl, J., 2013. The design trend in tissue-engineering scaffolds based on nanomechanical properties of individual electrospun nanofibers. Int. J. Pharm. 455, 338–347. Kim, H.N., Jiao, Alex, Hwang, Nathaniel S., Kim, Min Sung, Kang, Do Hyun, Kim, Deok-Ho, Suh, Kahp-Yang, 2013. Nanotopography-guided tissue engineering and regenerative medicine. Adv. Drug Deliv. Rev. 65, 536–558. Leong, M.F., Chian, K.S., Mhaisalkar, P.S., Ong, W.F., Ratner, B.D., 2009. Effect of electrospun poly(D,L-lactide) fibrous scaffold with nanoporous surface on attachment of porcine esophageal epithelial cells and protein adsorption. J. Biomed. Mater. Res. A 89, 1040–1048. Pelipenko, J., Kocbek, P., Govedarica, B., Rošic, R., Baumgartner, S., Kristl, J., 2013a. The topography of electrospun nanofibers and its impact on the growth and mobility of keratinocytes. Eur. J. Pharm. Biopharm. 84, 401–411. Pelipenko, J., Kristl, J., Jankovic´, B., Baumgartner, S., Kocbek, P., 2013b. The impact of relative humidity during electrospinning on the morphology and mechanical properties of nanofibers. Int. J. Pharm. 456, 125–134. Reinhart-King, C.A., Dembo, M., Hammer, D.A., 2008. Cell–cell mechanical communication through compliant substrates. Biophys. J. 95, 6044–6051. Rošic, R., Pelipenko, J., Kocbek, P., Baumgartner, S., Bešter-Rogacˇ, M., Kristl, J., 2012. The role of rheology of polymer solutions in predicting nanofiber formation by electrospinning. Eur. Polym. J. 48, 1374–1384. Rošic, R., Pelipenko, J., Kristl, J., Kocbek, P., Bešter-Rogacˇ, M., Baumgartner, S., 2013. Physical characteristics of poly(vinyl alcohol) solutions in relation to electrospun nanofiber formation. Eur. Polym. J. 49, 290–298. Thompson, C.J., Chase, G.G., Yarin, A.L., Reneker, D.H., 2007. Effects of parameters on nanofiber diameter determined from electrospinning model. Polymer 48, 6913– 6922. Toh, Y.C., Ng, S., Khong, Y.M., Zhang, X., Zhu, Y., Lin, P.-C., et al., 2006. Cellular responses to a nanofibrous environment. NanoToday 1, 34–43. Wang, Y., Wang, G., Luo, X., Qiu, J., Tang, C., 2012. Substrate stiffness regulates the proliferation, mobility, and differentiation of epidermal cells. Burns 38, 414– 420. Yeh, Y.T., Hur, S.S., Chang, J., Wang, K.C., Chiu, J.J., Li, Y.S., Chien, S., 2012. Matrix stiffness regulates endothelial cell proliferation through septin 9. PLoS ONE 7 (10), e46889. http://dx.doi.org/10.1371/journal.pone.0046889. Yördem, O.S., Papila, M., Mencelog˘lu, Y.Z., 2008. Effects of electrospinning parameters on polyacrylonitrile nanofiber diameter: an investigation by response surface methodology. Mater. Des. 29, 34–44. Zani, B.G., Edelman, E.R., 2010. Cellular bridges: routes for intercellular communication and cell mobility. Commun. Integr. Biol. 3, 215–220.
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European Journal of Pharmaceutical Sciences journal homepage: www.elsevier.com/locate/ejps
Nanofiber diameter as a critical parameter affecting skin cell response Jan Pelipenko, Petra Kocbek, Julijana Kristl ⇑ University of Ljubljana, Faculty of Pharmacy, Ljubljana, Slovenia
a r t i c l e
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Article history: Received 25 July 2014 Received in revised form 28 August 2014 Accepted 28 September 2014 Available online 6 October 2014 Keywords: Nanotechnology Nanofibers Electrospinning Proliferation Regenerative medicine
a b s t r a c t Electrospun polymer nanofibers have opened new opportunities in the rapidly evolving field of tissue engineering, particularly due to their topography and variability of available biomaterials. In order to better understand nanofiber influence on cell growth, the impact of their diameter was systematically examined. In this study homogenous, randomly oriented poly(vinyl alcohol) nanofibers with five different average diameters, ranging from 70 nm to 1120 nm, were produced, characterized and their impact on morphology, proliferation and mobility of keratinocytes and skin fibroblasts was evaluated. The results have shown that nanofiber diameter affects cell response and that this response is cell line specific. Nanofiber thickness affected size, morphology and actine organization of keratinocytes much more than fibroblasts. Specifically, the keratinocyte grown on nanofibers were more spherical and smaller compared to the control cells, while the fibroblasts were much less affect. They stayed almost unchanged and spread across growth surface. The cell proliferation determined based on their metabolic activity was the highest, when keratinocytes were grown on 305 nm thick nanofibers, whereas proliferation of fibroblasts grown similar nanofibers was decreased. Finally, fibroblasts exerted higher mobility than keratinocytes. Both tested cell lines on nanofiber diameters of 300 nm resulted in decreased cell mobility. These findings suggest that the control over nanofiber diameter offers promising possibility to better design the tissue scaffolds, since cells distinguish between differently sized nanofibers and respond accordingly. Ó 2014 Elsevier B.V. All rights reserved.
1. Introduction Tissues are complex structures composed of highly organized populations of individual cells. The basic principle of coordinated cellular events is cell communication with the environment; meaning interactions between cells as well as interactions of cells with the natural extracellular matrix (ECM) (Frantz et al., 2010). ECM is protein-based structure mostly composed of collagen fibers that extend in length over tens of micrometers, have thickness between 260 and 410 nm, as well as topography and structural features specific for cell response (Bettinger et al., 2009; Huang et al., 2012). When natural ECM is disturbed, either by illness or injury, regeneration process should be established to restore the tissue. Regeneration of tissues can be achieved by the combination of living cells, which provide biological func-
Abbreviations: ECM, extracellular matrix; PVA, poly(vinyl alcohol); MEM, minimum essential medium; SEM, scanning electron microscopy; PBS, phosphate buffered saline. ⇑ Corresponding author at: University of Ljubljana, Faculty of Pharmacy, Aškercˇeva cesta 7, 1000 Ljubljana, Slovenia. Tel.: +386 1 47 69 521; fax: +386 1 42 58 031. E-mail address: [email protected] (J. Kristl). http://dx.doi.org/10.1016/j.ejps.2014.09.022 0928-0987/Ó 2014 Elsevier B.V. All rights reserved.
tionality, and materials, which support the cell proliferation. Since the cell response in vivo depends on biological signals received from the surrounding environment, it is of crucial importance for particular tissue regeneration to resemble those signals (Zani and Edelman, 2010; Kim et al., 2013). Therefore, the biocompatible materials used in tissue regeneration should simulate natural behavior of treated cells. Polymer nanofibers can be used for temporal replacement of natural ECM and its functions, since they can improve regenerative process (Engel et al., 2008; Pelipenko et al., 2013a). Therefore, nanofibers are intensively explored in the field of chronic wounds, where organism is, due to the imbalance between synthesis and degradation of ECM, incapable of producing functional ECM and orchestrating the cell response leading to tissue restoration (Alves et al., 2010). Understanding interactions between cells and engineered nanostructured biomaterials is crucial for successful tissue healing. Most of these biomaterials have been studied in terms of nanofiber production abilities (Thompson et al., 2007; Yördem et al., 2008; Rošic et al., 2012; Cramariuc et al., 2013; Casasola et al., 2014) and much less has been investigated about their nanotopography and role in the cell response (Beachley and Wen, 2010; Bacakova et al., 2011).
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The cell cytoskeleton is known to have an important role in mechanosensing and mechanotransduction. Recent research has shown that filopodia first probe the environment, then long-lasting focal adhesions are established and, finally the extension of cell protrusions occurs (Albuschies and Vogel, 2013). The focal contact cell structures are mostly composed of integrins, which have size in nanometer range (Anselme et al., 2010; Huang et al., 2012). Therefore, it is assumed that they can be differently influenced by nanofibers with different diameters. The interactions between integrins and the extracellular environment (ECM or nanofibers) can result in change of cell adhesion, morphology, proliferation and even cell orientation and mobility. Nanotopography seems to have an important impact on cell behavior; therefore, it is now being explored with the aim to optimize scaffolds for tissue regeneration. The aim of our study was to investigate how to select proper nanofiber diameter to promote cell proliferation in vitro, which would probably in vivo result in healing and tissue regeneration. Firstly, electrospun poly(vinyl alcohol) (PVA) nanofibers were produced and characterized. Secondly, the impact of nanofiber diameter on the response of keratinocytes and skin fibroblasts, namely cell morphology, proliferation, and migration, was investigated in turn for further development of an effective nanofibrillar scaffolds.
2. Materials and methods 2.1. Materials PVA (MowiolÒ 20–98, Mw 125,000 g/mol) was supplied by Sigma–Aldrich (Germany), ethanol (96%) from Kefo (Slovenia). Minimum essential medium (MEM), non-essential amino acids, L-glutamin, antibiotic/antimycotic solution, fetal bovine serum, phalloidin rhodamine, trypan blue solution and Triton X-100Ò were all supplied by Sigma (Germany). Hoechst 33342 was obtained from Riedel de Haen (Germany), Cell Titer 96Ò Aqueous One Solution Cell Proliferation Assay was from Promega (Madison, WI). All other chemicals used were of analytical grade. 6, 24 and 96 well plates were from TPP (Switzerland) and custom-made inserts used in cell mobility experiments were provided by Denis Štraus, s.p. (Slovenia).
2.2. Methods 2.2.1. Nanofiber preparation and characterization Aqueous PVA solution was placed in a plastic 20 ml syringe fitted with a metal needle with an inner diameter of 0.8 mm. A steady polymer solution flow rate was ensured by a syringe pump (model R-99E, Razel™ Scientific Instruments, USA) and the high voltage by a generator (model HVG-P60-R-EU, Linari Engineering s.rl. Italy). A planar stand covered with aluminum foil was used as a collector. Electrospinning parameters were as follows: polymer concentration 8–15%, w/w, relative humidity 2–60%, applied voltage 10–30 kV, and needle to collector distance 10–20 cm. In order to produce nanofibrillar supports with comparable thicknesses, the electrospinning process was performed for 10 min for each sample. After electrospinning nanofibers were thermally stabilized against dissolution in aqueous environment by dry heat at 160 °C for 30 min. The morphology of the stabilized nanofibers was determined using a scanning electron microscopy (SEM, Supra 35 VP, Carl Zeiss, Oberkochen, Germany) at an accelerating voltage of 1 kV and a secondary electron detector. Based on SEM images the average nanofiber diameter and average interfibrillar pore size were determined
by measuring 40 randomly selected nanofibers or interfibrillar pores using ImageJ 1.44p software (NIH, USA).
2.2.2. Cell culture and treatment Immortalized human keratinocytes (cell line NCTC2544, ICLC, University of Genoa, Italy) or human skin fibroblasts (cell line 149BR, Public Health England, United Kingdom) were cultured as adherent monolayers at 37 °C in a humidified atmosphere of 5% CO2 in air. They were grown in MEM supplemented with 1% (v/v) non-essential amino acids, 2 mM L-glutamin, and 100 U/ml antibiotic/antimycotic and 10% (v/v) and 15% (v/v) FBS for keratinocytes and fibroblasts, respectively. The cells were regularly subcultured when reaching 80% confluency. Their viability was tested using trypan blue exclusion assay. The experiments were performed using cell seeding density 2 104 cells/cm2 and the final cell confluency reached in the experiments did not exceed 80%.
2.2.2.1. Examination of cell morphology. The cells were grown for 3 days on nanofibrillar supports, then the cells were fixed with 4% paraformaldehyde in phosphate buffered saline (PBS, pH 7.4) for 10 min and then permeabilized with 0.1% Triton X-100Ò in PBS (pH 7.4) for 10 min. The cell nuclei were stained with the DNA intercalating dye Hoechst 33342 (5 lg/ml) for 30 min protected from light. The dye used for cell nuclei staining simultaneously stained the polymer nanofibers. Actin filaments were stained with the red-fluorescent dye phalloidin rhodamine, according to the manufacturer’s procedure. Afterwards, coverslips were fixed on the slides and analyzed under fluorescence microscope using 360/420 nm (Hoechst 33342) and 535/635 nm (Phalloidin rhodamine) excitation/emission filter sets (Olympus IX81, Tokyo, Japan). Average size of cells was determined based on analysis of 40 randomly chosen cells. Based on SEM images the average nanofiber diameter and average interfibrillar pore size were determined by measuring 40 randomly selected nanofibers or interfibrillar pores using ImageJ 1.44p software (NIH, USA).
2.2.2.2. Assessment of cell mobility. Nanofibers, electrospun on glass coverslip (/ = 15 mm), were transferred to the bottom of a 24-well plate and then custom-made insert with diameter of 1.5 mm was mounted onto nanofibrillar support, representing a model ‘‘wounded field’’ with a defined area of 1.77 mm2. Afterwards, cells (2.5 104 cells/well) were seeded and incubated at 37 °C and 5% CO2 in air for 24 h to attach on nanofibrillar support, then the insert was removed and cell mobility was evaluated after 3 and 5 days using an inverted light microscope (CKX31, Olympus, USA).
2.2.2.3. Cell proliferation. Nanofibers, electrospun on round glass cover slips (/ = 5 mm), were transferred to the bottom of a 96-well plate, then cells were seeded on them and incubated at 37 °C and 5% CO2 in air for 3 or 5 days. Their proliferation was determined by MTS assay according to the manufacturer’s procedure. The sample absorbance was measured at 490 nm using a Synergy H4 microplate reader (BioTec, USA).
2.2.3. Statistical analysis The results are expressed as means ± standard deviations. The statistical analysis was carried out using an independent samples Student’s t-test. A value of p < 0.05 was considered statistically significant.
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3. Results and discussion 3.1. Control over electrospinning parameters enabled production of various sized nanofibers Different parameters known to affect the electrospinning process, namely, polymer concentration, relative humidity, applied voltage and the distance between needle and collector, were varied (Table 1) in order to produce homogenous nanofibrillar samples differing in average fiber diameter. Although, all parameters investigated should influence on nanofiber diameter according to the literature data (Cramariuc et al., 2013; Pelipenko et al., 2013b; Rošic et al., 2013), our results have proven that the control over two parameters, namely polymer concentration and relative humidity, exerted the greatest impact on the fiber diameter and enabled preparation of nanofibers with controlled size, as demonstrated in supporting information. Randomly oriented PVA nanofibers with average diameters ranging from 69 nm to 1116 nm were produced (Fig. 1). The thinnest nanofibers were obtained using solution with low polymer concentration (8%, w/w) and high relative humidity (60%), and the thickest using solution with high polymer concentration (15%, w/w) and low relative humidity (2%). Nanofibers with diameters in between were produced using 10% (w/w) PVA
Table 1 Optimized electrospinning parameters used for production of PVA nanofibers and their corresponding diameters. Sample
A B C D E
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solution and appropriately adjusted relative humidity (Table 1). The correlation between nanofiber diameter and relative humidity was in line with our previous studies, reporting decrease in fiber diameter due to increase in relative humidity (Pelipenko et al., 2013b). In the case of high relative humidity the solvent evaporation rate is decelerated, leading to formation of thinner nanofibers. On the other hand, polymer concentration influences nanofiber diameter through the number of entanglements formed between polymer chains (Casasola et al., 2014; Rošic et al., 2013). More concentrated solution therefore resulted in thicker nanofibers, if electrospun at the same relative humidity (Table 1). All prepared nanofibers were homogenous and had a smooth surface, except nanofibers produced from the solution with the highest concentration i.e. 15% PVA (Fig. 1E), which exerted wrinkled surface. Such fibers could be beneficial for cell adhesion, since the area available for contact with cells is larger due to irregular fiber surface (Rošic et al., 2012). Besides thickness and planar arrangement of nanofibers representing surface for cell growth, also the interfibrillar pore size is important parameter for prediction of spatial cell mobility. The results showed big differences in average interfibrillar pore size in produced samples (Fig. 1). The electrospun PVA nanofiber mats composed of thicker nanofibers contained larger pore sizes compared to the nanofiber mats with thinner nanofibers. This difference had an important impact on cell response as explained in details in the following sections and should be considered when designing nanofibrillar materials for application in tissue regeneration.
PVA conc. (w/w, %)
Tipcollector distance (cm)
Applied voltage (kV)
Relative humidity (%)
Nanofiber’s diameter ± SD (nm)
3.2. Effect of nanofibers on cell morphology
8 10
15
15
60
69 ± 19 161 ± 42 305 ± 48 667 ± 83 1116 ± 354
Cells can change their morphology according to the properties of the substrate they are growing on. Keratinocytes cultured on nanofibers showed changed cytoskeletal organization compared to the control cells growing on a glass coverslip (Fig. 2). The control cells were well spreaded over the growth surface and tightly connected to each other. Those at the edges, formed focal contacts,
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Fig. 1. SEM images of nanofibers tested: (A) 69 ± 19 nm, (B) 161 ± 42 nm, (C) 305 ± 48, (D) 667 ± 83 nm and (E) 1116 ± 354 nm and corresponding average sizes of interfibrillar pores.
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marked with arrows in Fig. 2. The actin filaments in cells grown on nanofibers could not be clearly seen, contrary to the control cells with well-organized actin filaments. Not only the internal structure but also the average size of cells was affected by fiber diameter as follows: 20.9 ± 2.7 lm on 69 nm nanofibers, 19.0 ± 3.3 lm on 161 nm nanofibers, 14.5 ± 2.6 lm on 305 nm nanofibers, 15.9 ± 2.7 lm on 667 nm nanofibers, 25.9 ± 6.1 lm on 1116 nm nanofibers, and 45.3 ± 7.7 lm on flat glass coverslip. The cells grown on thinnest nanofibers were more spherical and significantly smaller compared to the control cells, whereas the morphology of keratinocytes grown on thicker nanofibers was governed by spatial nanofiber arrangement. In the case of thicker nanofibers, the interfibrillar pores were bigger (Fig. 1) and therefore enabled partial cell infiltration into the nanofibrillar support, which consequently affected the cell morphology (Guimarães et al., 2010). More rounded cell morphology on randomly structured nanofibers compared to the cell morphology on glass coverslip has been reported previously (Pelipenko et al., 2013a). Contrary to keratinocytes, the morphology and actin organization of fibroblasts was much less affected by nanofiber diameter (Fig. 2). The cells were spread across the growth surface with average size of 80 lm along their length. The ability of fibroblasts to spread unhindered across nanosized obstacles has been reported also by Dalby and coworkers, who explored fibroblast mobility on nanocolumns (160 nm high, 100 nm wide) (Dalby et al., 2004).
3.3. Effect of nanofibers on cell mobility
Fig. 2. Keratinocytes and skin fibroblasts grown on nanofibers with different diameters and control cells grown on a glass coverslip. Actine filaments were stained using phalloidin red, whereas cell nuclei and nanofibers using Hoechst 33342. Due to the weak fluorescence of nanofibers and interference with other fluorophore, nanofibers are not well seen in the images of merged fluorescent signals; therefore, they are presented also separately. The arrows indicate focal contacts between cells and underlying glass coverslip.
The term cell mobility describes the mobility as a result of cell proliferation and spreading of daughter cells across the growth substrate and not the movement of a single cell across the growth substrate (Bertoncelj et al., 2014). The cell mobility was evaluated based on the comparison of the size of the ‘‘wounded field’’ at the investigated time point and its size at the beginning of the experiment (Fig. 3, t = 0 d). However, the effect of the cell size was not eliminated and could have also influenced the speed of ‘‘wounded field’’ closure; therefore, it was treated as a contribution to the overall cell mobility. In both tested cell lines, the highest cell mobility was observed on nanofibrillar supports composed of thin nanofibers (69 and 161 nm) (Fig. 3). Further increase in size of nanofibers resulted in decreased cell mobility. When nanofibers were thicker, interfibrillar pores were larger (Fig. 1) and cells could have penetrated partially into the nanofibrillar support, resulting in reduced overall cell mobility. In comparison to the control cells grown on a glass coverslip, cell mobility on nanofibers was reduced, which could be explained by physical entrapment of cells in the nanofibrillar network (Guimarães et al., 2010; Pelipenko et al., 2013a) and improved focal cell adhesion (Toh et al., 2006; Albuschies and Vogel, 2013). Another reason for lower cell mobility on nanofibers in comparison to the glass coverslip could be stiffness of the nanofibrillar support (Wang et al., 2012), which explains also the differences in cell mobility on various sized nanofibers. In one of our previous studies, we demonstrated that nanofiber stiffness strongly depends on nanofiber diameter (Jankovic´ et al., 2013). When cells are cultured on a soft substrates, such as polyacrylamide gels, cell–cell interactions are favoured over cell interactions with supporting substrate (Alves et al., 2010), resulting in lower cell mobility. However, when the supporting growth substrate becomes stiffer, interactions between cell and supporting substrate prevail and increased cell mobility is observed (Toh et al., 2006). The results of cell mobility are in line
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Fig. 3. The effect of nanofiber diameter on cell mobility. Cells grown on a flat glass coverslip were used as a control. The arrows indicate the area with increased cell density around the ‘‘wounded field’’.
with the size dependant mechanical behavior of nanofibers, i.e. the decrease in mobility (compared to the growth on the glass coverslip) on thicker nanofibers was more prominent than the decrease in cell mobility on thinner nanofibers. Furthermore, the area with locally increased cell density, visible as a darker ring around the ‘‘wounded field’’, was observed and was the most prominent in the case of keratinocytes cultured 5 days on 667 nm nanofibers (Fig. 3), indicating increased cell proliferation, which could be attributed to hindered contact inhibition at the edge of the model ‘‘wounded field’’ (Gray et al., 2008). The higher mobility of fibroblasts, in comparison to keratinocytes, can be explained by the fact that fibroblasts are larger than keratinocytes (80 lm compared to 40 lm, when cultured on glass coverslip) and, therefore, spread over larger surface area. Additionally, fibroblasts are exposed to the nanofibrillar structures of ECM in their natural environment and are in the case of injury forced to migrate toward wounded area and produce elements of new ECM. On the other hand, the main function of keratinocytes is to form densely packed cell layers, namely a barrier epithelium over newly formed cell layers and ECM in dermis (Reinhart-King et al., 2008). This phenomenon was observed also in vitro through the manner ‘‘wounded field’’ was closing. In the case of keratinocytes, the ‘‘wounded field’’ was closing due to the even cell proliferation from the edge of the ‘‘wounded field’’. In the case of fibroblasts, the irregular pattern of cell penetration in ‘‘wounded field’’ caused difficulties in experimental determination of ‘‘wounded field’’ size over the time, indicated also by high standard deviations of the measurements (Fig. 3). To sum up, cells cultured on thinner nanofibers exerted higher mobility than those cultured on thicker nanofibers. However, the
mobility was still much lower compared to the cell mobility on a glass coverslip. 3.4. Effect of nanofibers on cell proliferation Keratinocyte and fibroblast proliferation evaluated indirectly via cell metabolic activity was affected by nanofiber diameter as shown in Fig. 4. In the case of keratinocytes, the highest metabolic activity was observed for cells growing on 305 nm thick nanofibers at both tested time points, however, it was not significantly different from the metabolic activity of keratinocytes cultured on 667 nm thick nanofibers. The lowest metabolic activity i.e. significantly lower than metabolic activity of cells grown on all other investigated nanofibers, was observed for cells grown on 69 nm nanofibers. The obtained results correlate well with changes observed in average cell size. Faster cell proliferation resulted in smaller cells. The smallest cells (15.5 ± 2.6 lm) were observed in sample grown on 305 nm nanofibers, where the highest metabolic activity was also determined. Increase in fiber diameter up to 1116 ± 12.2 nm resulted in less stimulated cell proliferation, due to decreased resemblance between nanofibers and elements of natural ECM, which are mostly 260–410 nm in size (Bettinger et al., 2009). Fibers thicker than 1000 nm do not stimulate cell proliferation as efficient as thinner ones as reported previously (Leong et al., 2009). In the case of fibroblasts cultured on 69 and 161 nm thick nanofibers the proliferation was significantly decreased at both investigated time points, while on thicker nanofibers it was comparable to the growth of control cells. Higher degree of cell differentiation and slower cell metabolism are two key characteristics,
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(Wang et al., 2012; Yeh et al., 2012). However this is not in line with our results, indicating the highest keratinocyte proliferation on 305 nm thick nanofibers. Furthermore, the cell proliferation is expected to be decreased with increase in nanofiber diameter, because nanofibers exert size-dependant mechanical properties i.e. increased nanofiber stiffness with decrease in their diameter (Jankovic´ et al., 2013). This conclusion holds true only for keratinocytes grown on investigated nanofibers smaller than 1000 nm. Additionally, it was shown that the cell response does not depend only on stiffness of growth surface, but is also dependent on surface morphology i.e. dimensions of underlying nanofibers (Alves et al., 2010), confirmed also in our study. Cells are complex entities with a remarkable, inherent capacity to sense, integrate, and respond to environmental stimuli, such as nanotopography of growth surface. The current in vitro research was performed separately on keratinocytes and skin fibroblasts and has revealed some important results, which represent a good starting point for the future more complex in vitro studies using keratinocyte and fibroblast co-cultures, which better resemble in vivo conditions. Interplay and orchestrated activity between both cell types is important for wound healing in vivo, therefore, results obtained on individual cell lines indicate the direction of the cell response on nanofibers, however, prior more precise conclusions about the effects of nanofibers on wound healing in vivo can be made, the cell response in more complex environment should be systematically investigated. 4. Conclusions
Fig. 4. The effect of nanofiber diameter on keratinocyte and skin fibroblast metabolic activity determined 3 and 5 days after cell attachment. The cell metabolic activity is expressed relative to metabolic activity of cells grown on a flat glass coverslip.
which distinguish fibroblasts from keratinocytes and can explain their less prominent short term response due to the changed growth surface (Bertoncelj et al., 2014). Our results showed cell line and nanofiber diameter dependant cell proliferation on nanofibrillar supports, however, the stimulated cell proliferation caused by nanofibers is mainly reported in the literature (Fagotto and Gumbiner, 1996). According to Wang et al. and Yeh et al., the highest cell proliferation is expected on glass coverslip, since it represents the stiffest growth surface
The effect of nanofiber diameter on skin cell growth was systematically investigated over a broad range of fiber diameters, prepared by electrospinning of PVA solutions with adjusted polymer concentration and precise regulation of relative humidity during production process. It has been shown that nanofiber diameter affects cell response, which is also cell line specific. Keratinocytes are more sensitive to nanofiber size than fibroblasts, since their morphology was changed due to growth on nanofibers, whereas morphological changes of fibroblasts were less obvious. Furthermore, the proliferation of keratinocytes was a function of nanofiber diameter, with a maximum reached, when cells were growing on 305 nm thick nanofibers. Proliferation of fibroblasts was reduced, when cells were grown on thin nanofibers, and was comparable to the growth on the glass coverslip, when fibers were thicker. Finally, it was proven that the cell mobility is decreased if cells are grown on thicker nanofibers. Base on the results, it can be concluded that nanofiber diameter is an important parameter, influencing cell response. Therefore, it should be precisely considered when nanofibrillar materials intended for application in biomedical field, such as wound dressing or tissue scaffolds, are being designed. Based on the obtained in vitro results combination of thin and thick nanofibers could be recommended for skin tissue regeneration, since thicker nanofibers (300–700 nm) stimulate cell proliferation, while cell mobility, resulting in wound closure, is higher on thinner nanofibers. Acknowledgments The authors gratefully acknowledge the financial support provided by the Slovenian Research Agency for Programme Number P1-0189, Projects J1-4236 and J1-6746, and Grant Number 100011-310213. Appendix A. Supplementary material Supplementary data associated with this article can be found, in the online version, at http://dx.doi.org/10.1016/j.ejps.2014.09.022.
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