# BIOPOLYMER THIN FILMS AND COATINGS

EDITED BY : Stefan Spirk, Tiina Nypelö and Eero Kontturi PUBLISHED IN : Frontiers in Chemistry

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ISSN 1664-8714 ISBN 978-2-88963-333-3 DOI 10.3389/978-2-88963-333-3

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# BIOPOLYMER THIN FILMS AND COATINGS

Topic Editors:

Stefan Spirk, Graz University of Technology, Austria Tiina Nypelö, Chalmers University of Technology, Sweden Eero Kontturi, Aalto University, Finland

In science as well as in industry, the impetus of research on bio-based polymers has recently expanded into new terrains. The need to replace fossil-based materials with sustainable and renewable sources is one of the main drivers for the emergence and the development of new and environmentally friendly materials. While some materials applications of bio-based polymers are already very well established, for instance, in paper and textiles, others have just emerged with thin films and coatings being a recent and particular area of interest. Thin films in general are an enormous field of research both fundamentally and from an applied perspective, with uses ranging from corrosion resistance to photovoltaics and sensors. Since bio-based materials are a relatively novel source material for thin films, the research in this area is at a fresh, exciting stage at the moment.

Citation: Spirk, S., Nypelö, T., Kontturi, E., eds. (2020). Biopolymer Thin Films and Coatings. Lausanne: Frontiers Media SA. doi: 10.3389/978-2-88963-333-3

# Table of Contents

*04 Editorial: Biopolymer Thin Films and Coatings* Stefan Spirk, Tiina Nypelö and Eero Kontturi *06 Polysaccharide Thin Solid Films for Analgesic Drug Delivery and Growth of Human Skin Cells* Tina Maver, Tamilselvan Mohan, Lidija Gradišnik, Matjaž Finšgar, Karin Stana Kleinschek and Uroš Maver *19 Multilayer Density Analysis of Cellulose Thin Films* Carina Sampl, Katrin Niegelhell, David Reishofer, Roland Resel, Stefan Spirk and Ulrich Hirn *29 Design of Friction, Morphology, Wetting, and Protein Affinity by Cellulose Blend Thin Film Composition* Caterina Czibula, Gundula Teichert, Maximilian Nau, Mathias Hobisch, Chonnipa Palasingh, Markus Biesalski, Stefan Spirk, Christian Teichert and Tiina Nypelö *39 Natural Shape-Retaining Microcapsules With Shells Made of Chitosan-Coated Colloidal Lignin Particles* Tao Zou, Mika H. Sipponen and Monika Österberg *51 Spatially Resolved Crosslinking of Hydroxypropyl Cellulose Esters for the Generation of Functional Surface-Attached Organogels* Maximilian Nau, Simon Trosien, David Seelinger, Anna K. Boehm and Markus Biesalski *59 Hybrid Gibbsite Nanoplatelet/Cellulose Nanocrystal Multilayered Coatings for Oxygen Barrier Improvement* Maud Chemin, Laurent Heux, David Guérin, Laura Crowther-Alwyn and Bruno Jean *69 Ultrathin Films of Cellulose: A Materials Perspective* Eero Kontturi and Stefan Spirk *87 Aqueous Dispersions of Esterified Lignin Particles for Hydrophobic Coatings* Qi Hua, Li-Yang Liu, Muzaffer A Karaaslan and Scott Renneckar *97 Adsorption and Desorption of Organic Molecules From Thin Cellulose Films* Elias Henögl, Viktoria Haberl, Jakob Ablasser and Robert Schennach *105 Cellulose Nano-Films as Bio-Interfaces* Vikram Singh Raghuwanshi and Gil Garnier *122 Reactivity of Isocyanate-Functionalized Lignins: A Key Factor for the Preparation of Lignin-Based Polyurethanes* Mareike Zieglowski, Simon Trosien, Jochen Rohrer, Sabrina Mehlhase, Simone Weber, Kerstin Bartels, Gregor Siegert, Taina Trellenkamp, Karsten Albe and Markus Biesalski *131 Affinity of Serum Albumin and Fibrinogen to Cellulose, its Hydrophobic Derivatives and Blends* Rupert Kargl, Matej Bračič, Matic Resnik, Miran Mozetič, Wolfgang Bauer,

Karin Stana Kleinschek and Tamilselvan Mohan

## Editorial: Biopolymer Thin Films and Coatings

Stefan Spirk 1 \*, Tiina Nypelö2,3 and Eero Kontturi 4

1 Institute of Paper, Pulp and Fiber Technology, Graz University of Technology, Graz, Austria, 2 Department of Chemistry and Chemical Engineering, Chalmers University of Technology, Gothenburg, Sweden, 3 Wallenberg Wood Science Center, Gothenburg, Sweden, 4 Department of Bioproducts and Biosystems, Aalto University, Espoo, Finland

Keywords: thin films, biopolymer, cellulose, chitosan, lignin, coatings, proteins

**Editorial on the Research Topic**

#### **Biopolymer Thin Films and Coatings**

The development of thin film technology has played a decisive role in the manufacturing of hightech devices and products such as computers, mobile phones, solar cells, sensors, and displays. With polymer materials, synthetic polymers are still mostly applied while the use of bio-based materials for thin coatings and films is still at an early stage. Of these, particularly thin films of natural polysaccharides have been at the forefront of the research scene for the past 10–15 years. Although the main problems in film preparation, such as sparing solubility, have been largely overcome, many issues affecting the film performance are still not fully understood. One of the pressing issues is the influence of water on the film structure and behavior, given the special relationship that many biopolymers share with water. Other open questions include the stability and the general structural order of the films. In this special issue, we have assembled the frontiers of biopolymer ultrathin films: topical studies on water interactions, protein adsorption, and film structure among many others. The materials choices in the thin films include cellulose, cellulose derivatives, lignin, alginate, and chitosan.

Edited and reviewed by: Pellegrino Musto, Italian National Research Council (CNR), Italy

> \*Correspondence: Stefan Spirk<EMAIL_ADDRESS>
#### Specialty section:

This article was submitted to Polymer Chemistry, a section of the journal Frontiers in Chemistry

Received: 03 October 2019 Accepted: 15 October 2019 Published: 25 October 2019

#### Citation:

Spirk S, Nypelö T and Kontturi E (2019) Editorial: Biopolymer Thin Films and Coatings. Front. Chem. 7:736. doi: 10.3389/fchem.2019.00736

The special issue contains two review and 10 original research articles. The reviews yield insight into general materials perspectives (Kontturi and Spirk) as well as to the current state of ultrathin cellulose films to act as a biointerface (Raghuwanshi and Garnier). These are important emerging areas particularly with polysaccharide thin films where the research has generally been focused on their use as model systems (model surfaces). The 10 research articles, on the other hand, focus on different, mainly fundamental and materials-related aspects, such as composition, morphology and accessibility, crosslinking and interaction with biomolecules. The contribution of Kargl et al. presents affinity of proteins on cellulose, and other surfaces, and contributes to the quest for identifying the reasons behind selectivity of protein attachment on surfaces. Quantification of protein adsorption on surfaces in aqueous media is limited by the physics of the analytical techniques but also of data analysis. This contribution among others, utilizes a combination of analytical methods to cover the limitations of the individual techniques and models.

Czibula et al. investigate mixtures of two biopolymers in a film. They present film formation from cellulose derivate blends, leading to control of hydrophobic and hydrophilic domains, combined with surface roughness and structure periodicity. There is something curious about proteins adsorbing on textured surfaces that lead often to an unexpected increase in the adsorbed amount at certain compositions, as seen in previous accounts. The contribution by Czibula contributed to this topic by adding friction force morphology as a tool to screen interactions besides conventional tools such as QCM-D and SPR. Nau et al., on the other hand, focused on film formation by cross-linking hydroxypropyl cellulose that had been modified with fatty acid esters. Cross-linking and simultaneous binding on the surface was realized with a photo-induced radical cross-linker and the film properties were carefully and comprehensively monitored.

Besides macromolecular interaction, adsorption of small molecules (other than water) has been probed relatively little with cellulose thin films. Henögl et al. looked at the deposition of small molecules on cellulose surfaces with regard to their interaction capacity. They used as model compounds n-decane and deuterated methanol and found that although both molecules adsorb, the interaction of methanol with the cellulose surfaces is stronger, and supported the observation with thermodynamic data. On the other hand, the work by Sampl et al. looked more on the order and density of such cellulose thin films. The paper reveals that there are different areas in the films that feature lower density, i.e., at the air-film interface and at the film-substrate interface. Maver et al. also look into the structural feature of films, specifically hydrophilic multi-layered thin films. They further apply them as a model drug-carrier and time-dependent release system aiming for new functional and active 3D materials to be used in regenerative medicine.

Hua et al. had a more pragmatic approach to coat hydrophobic films on down-to-earth substrates like glass or Kraft pulp sheets, resulting in increased surface hydrophobicity of these materials. The lignin coating was performed using microparticles whose formation was made feasible by esterification of lignin hydroxyl groups. Likewise, Chemin et al. had their eye on utilizing thin films for genuine applications. In their work, hybrid films of cellulose nanocrystals and Gibbsite nanoplatelet films were constructed by layer-by-layer technique onto substrates in packaging applications, such as cardboard. The resulting materials exhibited improved oxygen barrier properties. Electrostatic interaction was also utilized in a different fashion by Zou et al. who used chitosan to attach as an ultrathin biofilm on negatively charged colloidal lignin nanoparticles. Such nanoparticles were especially suited for stabilizing oil-in-water Pickering emulsions for, e.g., food applications. Work of Zieglowski et al. also utilizes lignin. They provide insight in reactivity and isocyanate structure linked to Kraft lignin on polyurethane formation. Polyurethanes are an important material for coatings and use of lignin enables incorporation of the biomacromolecule into a highervalue polymer material. The rational choice of the isocyanate for prefunctionalization of lignin assists to prepare materials with better mechanical performance.

The contributions in this special issue of Biopolymer Thin Films and Coatings highlight that mastering the fundamentals of film formation and characterization already enables to reach toward utilization. The applications covered in this special issue, supported by the state-of-the-art analysis, provide a dissection of the status of the biopolymer thin film and coating field. We hope that the issue will inspires the readers to further contribute to this exciting field of fundamental and applied endeavors.

### AUTHOR CONTRIBUTIONS

All authors listed have made a substantial, direct and intellectual contribution to the work, and approved it for publication.

### ACKNOWLEDGMENTS

SS acknowledges the Christian Doppler Laboratory for Fiber Swelling and Paper Performance for financial support of the Austrian Federal Ministry of Economy, Family and Youth, and the National Foundation for Research, Technology and Development. TN acknowledges financial support of Wallenberg Wood Science Center (WWSC) and Area of Advance Materials at Chalmers University of Technology. EK acknowledges Academy of Finland (grant no. 303452) for funding.

**Conflict of Interest:** The authors declare that the research was conducted in the absence of any commercial or financial relationships that could be construed as a potential conflict of interest.

Copyright © 2019 Spirk, Nypelö and Kontturi. This is an open-access article distributed under the terms of the Creative Commons Attribution License (CC BY). The use, distribution or reproduction in other forums is permitted, provided the original author(s) and the copyright owner(s) are credited and that the original publication in this journal is cited, in accordance with accepted academic practice. No use, distribution or reproduction is permitted which does not comply with these terms.

## Polysaccharide Thin Solid Films for Analgesic Drug Delivery and Growth of Human Skin Cells

Tina Maver 1,2 \*, Tamilselvan Mohan1 \*, Lidija Gradišnik 3 , Matjaž Finšgar 4 , Karin Stana Kleinschek 1,5 and Uroš Maver 2,3

1 Laboratory for Characterization and Processing of Polymers, Faculty of Mechanical Engineering, University of Maribor, Maribor, Slovenia, 2 Department of Pharmacology, Faculty of Medicine, University of Maribor, Maribor, Slovenia, 3 Faculty of Medicine, Institute of Biomedical Sciences, University of Maribor, Maribor, Slovenia, 4 Faculty of Chemistry and Chemical Engineering, University of Maribor, Maribor, Slovenia, 5 Institute for Chemistry and Technology of Materials, Graz University of Technology, Graz, Austria

Chronic wounds not only lower the quality of patient's life significantly, but also present a huge financial burden for the healthcare systems around the world. Treatment of larger wounds often requires the use of more complex materials, which can ensure a successful renewal or replacement of damaged or destroyed tissues. Despite a range of advanced wound dressings that can facilitate wound healing, there are still no clinically used dressings for effective local pain management. Herein, alginate (ALG) and carboxymethyl cellulose (CMC), two of the most commonly used materials in the field of chronic wound care, and combination of ALG-CMC were used to create a model wound dressing system in the form of multi-layered thin solid films using the spin-assisted layer-by-layer (LBL) coating technique. The latter multi-layer system was used to incorporate and study the release kinetics of analgesic drugs such as diclofenac and lidocaine at physiological conditions. The wettability, morphology, physicochemical and surface properties of the coated films were evaluated using different surface sensitive analytical tools. The influence of in situ incorporated drug molecules on the surface properties (e.g., roughness) and on the proliferation of human skin cells (keratinocytes and skin fibroblasts) was further evaluated. The results obtained from this preliminary study should be considered as the basis for the development "real" wound dressing materials and for 3D bio-printing applications.

Keywords: thin (blend) films, alginate, carboxymethyl cellulose, diclofenac, lidocaine, keratinocytes, skin fibroblasts

### INTRODUCTION

Thin films from polymeric materials present an emerging platform for various types of applications, including drug delivery (Maver et al., 2015b; Karki et al., 2016; Stana et al., 2017), sensors, microarray preparations (Mohan et al., 2012), etc. They also hold a huge scientific interest for several solid-liquid interface studies, especially for the adsorptive interaction of nanoparticles (Taajamaa et al., 2013), polyelectrolytes (Mohan et al., 2015, 2017), biomolecules (Kargl et al., 2015), etc. While significant efforts have been made to formulate drug delivery platforms from organo-soluble synthetic polymers (Vendra et al., 2011; Ponnusamy et al., 2012), the use of natural polymers

#### Edited by:

Stefan Spirk, Graz University of Technology, Austria

#### Reviewed by:

Sergiu Coseri, Institute of Macromolecular Chemistry "Petru Poni", Romania Mirjana Kostic, University of Belgrade, Serbia

#### \*Correspondence:

Tina Maver<EMAIL_ADDRESS>Tamilselvan Mohan<EMAIL_ADDRESS>
#### Specialty section:

This article was submitted to Polymer Chemistry, a section of the journal Frontiers in Chemistry

Received: 14 February 2019 Accepted: 19 March 2019 Published: 09 April 2019

#### Citation:

Maver T, Mohan T, Gradišnik L, Finšgar M, Stana Kleinschek K and Maver U (2019) Polysaccharide Thin Solid Films for Analgesic Drug Delivery and Growth of Human Skin Cells. Front. Chem. 7:217. doi: 10.3389/fchem.2019.00217

**6**

(e.g., polysaccharides) for such applications is limited and have recently gained a paramount of interest (Zelikin, 2010; Park K. et al., 2018; Park S. et al., 2018). Due to their high compatibility, bio-inertness, diverse physicochemical properties and ease of handling during manufacturing, polysaccharide thin films are excellent candidates for skin-related applications, where a controlled and targeted delivery of pain-relieving analgesic drugs are prerequisite (Miao et al., 2018). Skin is the largest organ of the integumentary system and it plays a vital protective role of the human body against pathogens. Thus, any major injury (wound) that results in the loss of its integrity, can lead to significant patient disability or even death (Singer and Clark, 1999). Such injury/wounds are often further associated with severe pain that represents a stress for the patient and is therefore a potential factor in delaying the physiological process of wound healing (McGuire et al., 2006; Guo and Dipietro, 2010). Optimizing the analgesic treatment of injured patients is thus of major importance. In order to avoid several side effects that analgesic drugs have, the local application could be a good alternative (Palao i Domenech et al., 2008; Marques et al., 2014). However, there are difficulties in quantitively controlling and prolonging the release of analgesic drug molecules such as lidocaine (LID) and diclofenac (DCF), when they are employed without any supporting matrices. Several attempts have been made to incorporate LID and DCF into nanoscale multi-layered thin films for controlling their loading capacity and release profiles (Loo et al., 2010; Jeganathan et al., 2016). LID is among the most widely used local anesthetics due to its rapid action (the effect is noticed immediately), making it suitable to alleviate the pain, caused by dressing change (Zhao et al., 2018), while DCF is a long lasting non-steroidal anti-inflammatory drug, which can provide a longer-lasting pain reduction, caused by the injury itself (Altman et al., 2015).

Since LID and DCF are both water soluble and hydrophilic, their incorporation into thin films derived from hydrophobic polymers, are still challenging. Therefore, the polysaccharides [e.g., alginate (ALG) and carboxymethyl cellulose (CMC)], which are both inherently hydrophilic and water soluble, seem suitable candidates to incorporate large amounts of both mentioned drugs in a uniform fashion. ALG is the most commonly used mucoadhesive polymer in wound healing applications, owing to its excellent water uptake capacity, non-toxicity and abundance in nature (Pawar and Edgar, 2012). It is derived from brown algae and bacteria and is classified as a linear anionic polysaccharide, composed of repeating units of β-1,4-linked D-mannuronic acid (M) and L-guluronic acid (G) in varying ratios. Depending on the type of ALG, its composition leads to the formation of tight ionically cross-linked network structures; contributing to a prolonged drug release (Paques et al., 2014). CMC, a derivative of the most abundant natural polymer cellulose, is also anionic, weakly charged and water soluble. Both were already proven to be efficient film-forming polymers and have found use in erodible and other drug formulations (Palmer et al., 2011).

Until now, the layer-by-layer (LbL) self-assembly technique has been largely used to create nanoscale multi-layered systems, which are of interest for targeted drug delivery applications (Stana et al., 2017; Park K. et al., 2018), where the simplicity and versatility of the preparation procedure lower the overall drug delivery system costs. In general, the LbL technique exploits the alternating deposition of oppositely charged substances (e.g., polyelectrolytes, polypeptides, etc.), which enables surface and bulk material decoration with a wide range of functional groups, in the final multi-layer structure (Liu et al., 2019; Zhang et al., 2019). However, the application of this technique to create multi-layered nanofilms from a single polymer or mixture of polymers with the same ionic/functional groups, and the use of the latter system in the release of therapeutic drug molecules is still only scarcely reported (Finšgar et al., 2016; Stana et al., 2017). Amongst several other fabrication techniques, spin coating is the commonly used technique for the design of multi-layered structures. Its main advantages include a high reproducibility, reliability, rapid film formation, and simultaneous solvent evaporation. On top of the mentioned, it allows to control the thickness, morphology and surface properties of the final material (Stana et al., 2017). The challenges in the fabrication of multi-layered nanofilms from water soluble and hydrophilic polymers (with similar overall properties) such as ALG, CMC, and the ALG-CMC combination by spin-assisted LbL technique was not yet explored in a systematic way. Furthermore, the potential application of these self-assembled polysaccharide multi-layers for incorporation and controlled release of LID and DCF were also not yet investigated in detail.

The motivations for this work, therefore, was to create a nanoscale multi-layered thin films, as a model drug-carrier system, in a systematic way from hydrophilic ALG and CMC. In addition, this work also aims to gain a deeper understanding of the surface and physiochemical properties of the developed multilayered system. For this purpose, aqueous solutions of ALG, CMC, and the mixture of ALG-CMC with incorporated DCF or LID drugs were used to fabricate multi-layered thin films on different solid substrates by the spin-assisted LBL coating technique. The wettability, thickness, morphology, and roughness of the multi-layered films were characterized using several surface sensitive methods. The time-dependent release of drugs from the multi-layer at body temperature and at physiological pH were also determined. The obtained release profiles were further fitted with various kinetic models to discern the release mechanism of the drugs. Finally, the influence of the multi-layer films with in situ included drugs on the proliferation of human skin cells (keratinocytes and skin fibroblasts) was evaluated. The results obtained from this preliminary study should be considered as the basis for the development of new functional and active 3D materials to be used in regenerative medicine.

### EXPERIMENTAL SECTION

### Materials and Methods

All chemicals used for the preparation of multi-layered thin films (ALG: viscosity: 0.015–0.025 Pa s, CMC: Viscosity: 2.5– 6 Pa s, DCF and LID) were purchased from Sigma-Aldrich, Germany. The tetrazolium salt MTT [3(4, 5 dimethylthiazolyl-2)-2, 5-diphenyltetrazolium bromide] for the biocompatibility testing was purchased from Sigma-Aldrich, Germany. Advanced Dulbecco's Modified Eagle's Medium (ADMEM) and Fetal Bovine Serum (FBS) were purchased from ThermoFisher, Germany. Keratinocytes, an aneuploid immortal keratinocyte cell line (HACAT) from adult human skin was kindly provided by Prof. Dr. Elsa Fabbretti (Centre for Biomedical Sciences and Engineering, University of Nova Gorica, Slovenia). Human derived skin fibroblasts (ATCC CCL-119, Detroit 551, LGC Standard) were purchased from ATCC, UK. For all experiments, ultra-pure water (18.2 M cm at 25◦C) was used, prepared from a ELGA Purelab water purification system (Veolia Water Technologies, UK).

### Sample Preparation

Three types of solutions were prepared for spin-coated. **Type I** (ALG-DCF or ALG-LID, 0.5/0.1%, w/v): 500 mg ALG was added to DCF or LID (0.1%, w/v, dissolved in water), stirred overnight with a mechanical stirrer. ALG (0.5%, w/v) was prepared in the same way without DCF or LID. **Type II**: (CMC-DCF or CMC-LID), CMC with and without DCF or LID with the same concentrations as mentioned in Type I were prepared. **Type III**. ALG-CMC-DCF or ALG-CMC-LID: ALG and CMC with DCF or LID were mixed together in a ratio of 1:1. The final concentration of the polymers and drugs in the final solution are 0.25 and 1 mg/mL, respectively. All solutions were prepared freshly before use.

### Multi-Layered Thin Film Preparation and Drug Incorporation

Silicon wafers were used as base-substrates for multi-layered thin film preparation. Prior to spin-coating, the silicon wafers were cut into pieces of 1 × 1 cm2 , soaked into a "piranha solution" [H2O2 (30 wt. %)/H2SO4 (conc.): 1:7 (v/v)], rinsed with ultrapure water and blow dried in a stream of dry nitrogen of high purity (99.999%). For creating multi-layers, the above-mentioned three types of polymer solutions with and without drugs were used. For spin-coated, 50 µL of ALG or CMC or ALG-CMC mixture with DCF or LID was deposited on the static substrate and spin-coated with a spinning speed of 4,000 revolutions per minute (rpm) and an acceleration of 2,500 rpm s−1 for 60 s. This procedure was repeated three times for each sample. In this way, three layers of the same material were created. These spincoated final three layers from ALG, CMC, and ALG-CMC are designated as ALG 3, CMC 3, and ALG-CMC 3, respectively, without incorporated drugs. The drug incorporated and spincoated three layers are designated as CMC-DCF 3 (or LID 3), ALG-DCF 3 (or LID 3), or ALG-CMC-DCF 3 (or LID 3).

### Atomic Force Microscope (AFM)

AFM was used for the topographical and surface roughness analysis of spin-coated multi-layered thin films. Prior to analysis using the Keysight 7500 AFM (Keysight Technologies, USA), the samples were attached onto round shaped metal disc sample holders. Topography images were acquired using the acoustic AC AFM mode. Silicon AFM tips (Nanosensors, Switzerland) with a nominal spring constant 12–110 N/m and a nominal resonance frequency of 210–490 kHz, were used for imaging purposes for all samples. Images of 1 × 1 µm2 were recorded, with a resolution of at least 524 × 524 pixels. All images were processed, and the corresponding roughness was calculated using freeware Gwyddion software package. For thickness determination, a straight line was created on the spin-coated sample by scratching with a sharp scalpel. Subsequently, a profile was created along the scratched straight line. This profile was then used to estimate the thickness of each spin-coated layer.

### Attenuated Total Reflection-Fourier Transform Infrared (ATR-FTIR) Analysis

The ATR-FTIR spectra of spin coated samples were measured using a Perkin Elmer FTIR System Spectrum GX Series-73565 at a wavenumber range of 4,000–650 cm−1 . A total of 32 scans were performed for all measurements with a resolution of 4 cm−1 . QCM quartz crystals (QSX301, LOT-Oriel, Germany) coated with a gold layer were used as substrates for ATR-FTIR measurements. The spin coating of polymer solutions with and without drugs were performed in the same way as with silicon wafers.

### Contact Angle Measurements

The wettability of spin-coated multi-layers was determined using a OCA15+ goniometer system (Dataphysics, Germany) with the sessile drop method. Static contact angle (SCA) measurements were carried out using ultra-pure water at ambient temperature. All measurements were carried out on at least two independent surfaces with a drop volume of 2 µL. Each SCA value was the average of at least five drops of liquid per surface.

### In vitro Release Studies

The release of DCF and LID from the multi-layered thin films was studied using an Automated Transdermal Diffusion Cells Sampling System (Logan System 912-6, Somerset, USA). For this purpose, each spin-coated wafer (1 × 1 cm2 ) was transferred to a 15 mL Franz diffusion Cell, filled with phosphate buffered saline (PBS, Sigma-Aldrich, Germany) with pH of 7.4. Its temperature was maintained at 37◦C all the time. Samples were collected over a period of 24 h at different time intervals. The absorbances were determined using a UV-Vis spectrophotometer (Cary 60 UV/VIS, Agilent, Germany), and the released amounts calculated by quantification of the absorption bands at 276 nm (DCF) and 204 nm (LID), using the Beer-Lambert law. The withdrawn sample volumes were replaced by fresh PBS. During the release profile calculations, this dilution was accounted for. The release measurements of at least three parallels of each sample were conducted and the results are reported as average value with ± standard error.

To evaluate the release profiles further, the results from the in vitro drug release test were fitted using three known models (first-order, Higuchi-model, and Korsmeyer-Peppas model). Using these models (Costa and Sousa Lobo, 2001), the release mechanisms of the respective drugs from the multi-layers were analyzed.

First-order kinetics (1) is used to describe the release of the drug where the release rate is concentration depended.

$$
\log \mathbf{Q} = \log \mathbf{Q}\_0 - \mathbf{K}\_t / 2.303 \tag{1}
$$

where Q is the amount of released drug in time "t," Q0 is the initial concentration of the drug and K is the first-order rate constant.

Higuchi-model (2) is used to express the drug release from an insoluble matrix as a square root of time dependent process based on the Fick's diffusion:

$$\mathbf{Q} = \mathbf{K} \mathbf{t}^{1/2} \tag{2}$$

where Q is the amount of drug released at time "t" and K is the Higuchi constant.

The Korsmeyer-Peppas model (3) describes the release of drugs from polymeric systems, which has been successfully applied to describe the release from different modified release dosage forms:

$$\mathbf{Q}\_{\mathbf{t}}/\mathbf{Q}\_{\infty} = \mathbf{K}\mathbf{t}^{\mathbf{n}} \tag{3}$$

where Qt is the amount of drug released in time "t," Q∞ is the amount of drug released after infinite time, K is the constant, and n is the diffusional exponent. If the n value is equal 0.5, then the release mechanism is Fickian diffusion, if 0.45 < n < 0.89, then it is called non-Fickian or anomalous diffusion. In the case of "n" is between 0.89 and 1, and above 1, the drug release is the combination of both diffusion and erosion-controlled processes.

### Biocompatibility Testing

In order to test the influence of added DCF and LID on cell viability, the latter was evaluated via the reduction reaction of the tetrazolium salt MTT [3(4,5 dimethylthiazolyl-2)-2,5 diphenyltetrazolium bromide], purchased from Sigma Aldrich, Germany. This is a commonly used and reliable method to evaluate cell proliferation (Finšgar et al., 2016; Maver et al., 2018). It is based on the reduction of the yellow MTT by metabolically active cells (e.g., in part by dehydrogenase enzymes), which results in the formation of nicotinamide adenine dinucleotide and/or nicotinamide adenine dinucleotide phosphate (Maver et al., 2017). During this reaction, a purple formazan is formed intracellularly, which can be solubilized and quantified by spectrophotometric means. Thin films on silicon wafers (1 × 1 cm2 ) were exposed to the UV light for 30 min (i.e., sterilization). Further, the samples were soaked into 1.5 mL of Advanced Dulbecco's Modified Eagle's Medium (ADMEM; ThermoFisher, Germany), supplemented with 5 wt.% Fetal Bovine Serum (FBS; ThermoFisher, Germany), and incubated for 24 h at 37◦C in an atmosphere containing 5 wt.% CO2. The HaCaT and Skin Fibroblast cells (10,000 cells/well) were seeded into a 96-well microtiter plate with a final volume of 100 µL of ADMEM medium supplemented with 5 wt.% FBS. The material samples (supernatants of the starting samples) were added to the cells after 24 h of incubation at 37◦C in an atmosphere containing 5 wt.% CO2 in four parallels. As control, ADMEM medium supplemented with 5 wt.% FBS was added to the cells. After 24 h of treatment, cell viability was determined using the standard reduction of the tetrazolium salt MTT (Stergar et al., 2017; Maver et al., 2018b).

### Statistical Analysis

All numerical values are given as average values ± standard deviation. Statistical analysis was performed using SPSS Statistics 25 (IBM Corp. Armonk, NY, USA). By using the Shapiro–Wilk test it was determined that all experimental data was normally distributed, allowing the use of one-way ANOVA. P < 0.05 were considered statistically significant (these are marked with an asterisk ∗ in respective graphs).

### RESULTS AND DISCUSSION

### Surface Morphology, Layers Thickness, and Wettability

#### Surface Morphology

The surface morphology and roughness of the spin-coated multilayer films based on ALG, CMC, and the ALG-CMC mixture were characterized using atomic force microscopy (AFM). For clarity reasons, only the results obtained from the final (top) layers are shown in this section, since the same system is used for the release of DCF and LID drugs (section In vitro Release Studies). While smooth morphologies with a low surface roughness value (below 1 nm) are observed for both drug-free ALG 3 and CMC 3 coated films, the ALG-CMC mixture prepared at 1:1 ratio (top right image in **Figure 1**) did show some different surface features. Interestingly, the surface roughness of both ALG and CMC 3 samples is lower when compared to its layer thickness (see **Figure 2**). This is most likely due to tight packing and more self-organization of polymer molecules within the film structure. Surfaces with a phase separated morphology are detected for the two-component system that are composed of same charges i.e., ALG-CMC 3. Such phase separation is already noticed for the first ALG-CMC layer (**Figure S1**).

The reports on the phase separation of polymer blends from (organo-soluble) hydrophobic polymers upon spin-coating are manifold (Taajamaa et al., 2011; Strasser et al., 2016). Whereas, no studies on the formation of blend surfaces from spin-coated polysaccharides, which are hydrophilic and water-soluble, can be found to the best of our knowledge. Therefore, maximum efforts are made to explain the observed lateral phase separation in the ALG-CMC 3 multi-layered system and not to overinterpret the results at the same time. At low solute (polymer) concentration, the polymer-polymer blend in solution stays in one phase only (Doublier et al., 2000; Keating, 2012; van de Velde et al., 2015). Upon spin-coating, the increase of solute concentration will push the system into two phases with each phase enhanced in its respective component. The formation of two phases (i.e., phase separation) is often a complex phenomenon, leading to the formation of non-equilibrium morphologies, due to the rapid removal of solvent during the spin-coating process. Usually, this leads to different structures/morphology on the nano- to micrometer scale (Walheim et al., 1997). This nano- and micro-phase separation can be classified into associate and segregate, which in turn dependent on the affinity between polymers and solvent. The associate phase separation arises when both polymers are oppositely charged, and both polymers are intensified in one of the separating phases while leaving the other phases mostly with

solvent. In the case of segregate phase separation, two polymers with same charges are separated into two different phases (Fang et al., 2006). The segregate phase separation is well-known for water soluble polymers (e.g., proteins- polysaccharides) for several years (Doublier et al., 2000), but not reported for spincoated blend films. Keeping this mind, we believe that the lateral phase separation that occurred in the case of ALG-CMC blend, which stays in one phase in water, is of the segregate type, which can be explained by the transient bilayer theory (Heriot and Jones, 2005; Taajamaa et al., 2011).

According to the transient bilayer theory, the formation of the lateral phase separated structures/morphologies during spincoating, occurs because of de-wetting and interfacial instabilities between the two polymers (ALG and CMC in this study). The instability at the interface is presumably caused by the solventconcentration gradient within the film. Usually, the evaporation of the solvent is faster at the film surface than in the bulk phase, leading to a lower solvent concentration at the surface than at the polymer-substrate interface. Thus, the instability, caused by the interfacial tension at a polymer-polymer interface is a function of solvent concentration. Concretely this means that if one of the polymers out of the two polymer blend exhibits a strong interaction with the substrate, the phase separation is initiated in a surface-oriented fashion, resulting in a transient bilayer (Geoghegan et al., 1994; Walheim et al., 1997). With the evaporation of more solvent from the film, the upper layer of the film gets unstable with the formation of holes. The latter is then filled with the lower (liquid phase) layer as it is shown for ALG-CMC coated (first) layer (**Figure S1**) (Taajamaa et al., 2011). Interestingly, the phase separation is retained even after the third layer (**Figure 1**, top row and **Figure S2**). Using the phase image (**Figure S2**), the existence of two phases, with a thicker region surrounded by a thinner one, from the two-component system can be easily seen. However, we are aware of the fact that it is difficult to confirm the thicker or major phase is either ALG or CMC; which needs further investigation. From the above discussion, it is proposed that the lateral phase separation occurs upon spin-coating for the hydrophilic polysaccharides that carry similarly charged chains. The detailed understanding of these phenomenon at different ALG-CMC ratios (in water) and humidity' s is currently in progress and is out of the scope of this work. The addition of drugs (LID and DCF) leads to a change in surface morphology for both ALG 3 and CMC 3 coated layers (but not for the ALG-CMC 3 mixture). Especially, particle-like features are visible for LID incorporated ALG 3 sample, whereas a featureless and even more smooth morphology is noticed for both DCF and LID incorporated CMC 3 sample. This can be due, on one hand, an enhanced interaction between LID and functional groups in the multilayer system (as confirmed by IR) and on the other hand, potentially significant inter-diffusion of small, hydrophilic LID molecules within the entire film architecture during the film formation, which occurs during the spin-coating process. In the case of drug incorporated ALG-CMC 3 blend, the phase-separation is still visible with only minimal changes in the morphology. This could be due to the enrichment of drugs in both phases, as confirmed by ATR-FTIR and in vitro release studies, where the peaks correspond to DCF and LID and a successful release of both drugs is observed (see below).

### Layer Thickness

The thicknesses of each spin-coated layer (for respective samples) are shown in **Figure 2**. Expectedly, the layer thickness increased with the number of layers deposited for all spin-coated samples. However, there is a difference among the spin-coated materials. For ALG based samples, a layer thickness of only ∼10 nm is measured even after the spin-coated of the third layer. Whereas, a value close to 10 nm is already obtained for the first layer in CMC based samples. With the second and the third layers, the thickness value for this sample is increased from ∼22 to ∼35 nm, respectively. This can be (at least partially) explained by a higher molecular weight (700 kDa) and viscosity (2.5–6 Pa s) of the CMC solution, leading to deposition of higher polymer masses on the surface upon spin-coating. As can be seen in **Figure 2**, the thickness of the first layer of ALG-CMC blend is considerably higher when compared to the first layer of single component ALG and CMC (ALG-CMC: 17 nm > CMC: 9 nm > ALG: 5 nm). These results demonstrate that a multi-layer can be fabricated from a single or mixture of polysaccharides with the same functional groups (i.e., –COOH). In addition, it has to be pointed out that the layer thickness of drugincorporated spin-coated films (of ALG, CMC or ALG-CMC blend) did not increase considerably compared to films prepared without drugs (data not shown). This could be due to lower concentration (see **Figure 5**) and homogenous incorporation of the drugs into the spin-coated layer. This gained knowledge, in future, can exclude the use of oppositely charged components, such as carboxymethyl cellulose—chitosan (Mohan et al., 2015), hyaluronic acid- chitosan (Pérez-Álvarez et al., 2019), alginatechitosan (Stana et al., 2017; Criado-Gonzalez et al., 2019), etc., to create multi-layered system.

### Wettability

The contact angle values directly reflect surface wettability and are sensitive to chemical functionalities of the outermost layer (Stana et al., 2017). The surface wettability of spin-coated multilayers, incorporated with and without drugs, were measured

TABLE 1 | SCA(H2O) values of ALG, CMC, and ALG-CMC spin-coated multi-layers, incorporated with and without drugs.

by the static water contact angle [SCA(H2O)] measurements and the results are depicted in **Table 1** (also see in **Table S1**). It shows that the spin-coated layers of ALG 3, CMC 3 or their combination are hydrophilic [obtained SCA(H2O) values << 90◦ ] (Yuan and Lee, 2013). For all three spin-coated samples, higher SCA(H2O) values are observed compared to the SCA(H2O) values of the substrate used for spin-coated [Siwafers; SiO2 surface (10 ± 0.2◦ )] (Stana et al., 2017). The changes in the SCA(H2O) indirectly also confirm a successful deposition of the spin-coated polysaccharide materials. Compared to ALG 3 alone, slightly increased SCA(H2O) values for CMC 3 and the ALG-CMC 3 blend are observed, which can be related to the rougher and thicker surfaces of the latter samples (**Figures 1**, **2**). Another important observation based on these results is that no further (major) changes in the SCA(H2O) values resulted for the incorporation of either of the drugs. Namely, hydrophilic and highly wettable surfaces are already on their own beneficial for wound healing of specific (e.g., chronic) wounds and changes toward more hydrophobic values would potentially limit their therapeutic efficiency. Furthermore, such surfaces can even enhance the drug release efficiency by allowing penetration of body fluids and thus enabling the dissolution and/or diffusion of the drugs from the materials to the wound area (Maver et al., 2018a).

### Surface Composition

The surface composition of spin-coated multi-layers incorporated with and without drugs was analyzed using ATR-FTIR spectroscopy. **Figure 3A** shows characteristic peaks of pure ALG and CMC. For ALG, the observed peaks at 3,360, 2,903, and 1,625 cm−1 are attributed to O-H stretching, -CH stretching, the asymmetric and symmetric vibrations of -COO and C-O-C stretching, respectively. Whereas, all major peaks of CMC are observed at 3,411, 2,916, 1,583, 1,420, and 1,324 cm−1 , which are assigned to O-H and –CH stretching, and the asymmetrical and symmetrical vibrations of carboxyl groups in the CMC, respectively. Although no additional new peaks are noticed for ALG-CMC 3 coated layers, all the characteristic peaks, as shown above, of ALG and CMC are detected. These results suggest that the ALG-CMC 3 layer is composed fully of both polymers and no new bindings/interactions are involved between the functional groups.

The spectra of sodium salt of DCF and DCF loaded polymers are shown in **Figure 3B**. Characteristic DCF peaks include at 3,253 cm−1 the NH stretching of the secondary amine, at 1,571 cm−1 the –C=O stretching of the carboxyl ion, at 1,500 cm−1 the C=C ring stretching, at 1,450 cm−1 the -CH2 bending,

at 945 cm−1 the -C-O-C stretching, and at 746 cm−1 the C-Cl stretching. All these peaks are also present for all samples (ALG, CMC, and the ALG-CMC blends). **Figure 3C** shows the characteristic peaks of LID at 1,673 cm−1 for the stretching of carbonyl group, and at 1,541 cm−1 the in-plane bending of the N-H group. Also, in the case of the LID, most of these peaks are detected for all the polysaccharide based multi-layer samples. Interestingly, the -C=O peak position of CMC and ALG is altered after the incorporation of both drugs. The -C=O peak is shifted from 1,625 to 1,604 cm−1 and from 1,583 to 1,613 cm−1 for ALG and CMC, respectively. These peak shifts indicate that both drugs are involved in some sort of interactions, most likely ionic or H-bonding. Such a scenario is mostly favorable and desired for applications, especially for prolonging the release rate of the drugs. In addition, the drug molecules could also be physically incorporated, which has other advantageous that drug molecules could simply diffuse through the layers and be released into the medium or at the wound site without any hinderance from the polysaccharide in which the drugs are incorporated. It can be concluded that the ATR-FTIR results confirmed the presence, and hence a successful incorporation of both used drugs into the formed multi-layer systems regardless of their compositions. Therefore, the next step was a thorough drug release analysis of all drug incorporated samples.

### In vitro Release Studies

Controlled and local release of pain relieving, and antiinflammatory drugs presents an important step toward a more successful chronic wound healing. Such drugs can not only diminish the pain from the wound itself, but also reduce the pain induced stress, which is known to hinder the healing efficiency, especially in the long term (Maver et al., 2015a). The used drugs herein (LID and DCF), come from different pharmacodynamic groups (LID is a local anesthetic and DCF a non-steroidal anti-inflammatory drug), and hence lower the sensation of pain through different mechanisms, improving the overall therapeutic potential of the developed multi-layered materials even further (Maver et al., 2017). Since they are both water soluble, DCF and LID can be easily mixed and homogeneously incorporated within the herein used hydrophilic host materials (ALG and CMC). Altogether, such systems provide an excellent platform for rapid design of novel multifunctional wound dressing materials as was shown before (Maver et al., 2019). In this study, novel polysaccharide drug loaded multi-layer thin film systems were fabricated that serve as a representative model of "real" dressing materials, enabling their optimization already during development. Since it is important to ensure an efficient pharmacotherapy i.e., controlled drug delivery for the newly developed systems, a lot of emphasis has been put on the evaluation of the results obtained from the in vitro drug release studies.

The DCF and LID release profiles from the different polysaccharide multi-layers are shown in **Figure 4**. The release of both drugs is performed at physiological pH with PBS buffer and at a simulated body temperature (37◦C) using the "Franzdiffusion" cell-based drug release system, which is the best currently available system to simulate the release from wound dressings. The released mass of DCF as a function of time is depicted in **Figure 4A**. Immediately it can be observed that there are notable differences in the released mass of DCF from the three systems. As it seems, the DCF release is favored from the "pure" ALG multi-layers, which is not really surprising considering that DCF is in its essence an "acid." Whereas, CMC can exhibit multiple carboxylic groups per one sugar unit, this is not the case with ALG, which has only one such group in its structure. Therefore, increased repulsive forces could be present in the case of CMC multi-layer systems, which probably led already in the preparation steps to smaller incorporated DCF amounts, hence finally contributed to a smaller overall DCF release. In addition, as can be seen from the calculated first derivatives of the release data (**Figure 4B**), the profiles of all systems show that multiple mechanisms might be involved in the release of DCF as the time progresses (breaks in the curves of first derivatives). This corresponds to the fact that release of hydrophilic drugs from hydrophilic multi-layer thin films present not just a "simple" diffusion-controlled mechanism but is often further accompanied by a combination of swelling and erosion of polymer chains upon contact with a buffer solution. As soon as the drug becomes soluble and polymer chains becomes mobile, the diffusion of drugs through the multi-layered films occurs with a different rate. A careful evaluation of the release profiles shows a fast (burst) release (∼45%) within the first 30 min,

which most likely corresponds to the release and dissolution of "trapped" or unbound drug molecules within the polymer structure. The latter is followed by a slower and prolonged release (∼50%) for the next 6 h (360 min), which is related to the combination of swelling and partial erosion (mentioned above). The last release stage is the so called "plateau," which present a diffusion-controlled release of the remainder of the drug from the remaining (mostly intact) part of the multilayer system. Another important observation based on the obtained release results is that regardless of the composition, all three different samples seems to follow the same combined release mechanism (confirmed by the calculation of the first derivatives shown in **Figure 4B**). This has another implication for the potential clinical use of such materials. Namely, by simply choosing another polymer (or the combination thereof), the physician can also control the overall used/delivered drug dose to the wound. Further studies would be necessary to show how we can further control the released drug amount by changing the incorporated drug amount, but this is out of the scope of this study. Developing the thought about the potential clinical application of such materials further, it can be said that such a release profile is essentially as desired in release of analgesic drugs in chronic wound treatment. Whereas, the initial "large" released amount of the analgesic drug is essential to provide a strong pain-relieving action as soon as possible, the sustained release is important to lower the pain sensation until the dressing is changed. The sustained drug release further improves the quality of life of the patient in long-term during treatment.

Similar deductions about the release mechanism can be made for the system with incorporated drug LID, although there is a different order of the systems in regard of the total release LID amount. Namely, here the CMC based system show the highest release amount, which is most likely again connected with the polymer and drug molecular structure. LID presents an amide, which has another secondary amine in its structure. The latter is capable to form interactions (e.g., ionic bonds) with carboxylic groups, which might have contributed to a higher incorporated dose of the drug during the preparation step (i.e., spin-coating). Since CMC has a higher amount of such groups, more of the drug could have been interacting with the polymer chains in the multi-layer film structure. This data fits with the ATR-IR results where the shift of -C=O peaks of polymers are observed as the result of ionic interactions. Two more conclusions can be made based on the results shown in **Figure 4**. Firstly, the multi-layer systems with incorporated LID had a higher overall incorporated amount of the drug. And secondly, regardless of the drug type, the combined polymer system (ALG-CMC) has shown the most sustained release. To ease further discussions of the latter, the overall released mass of DCF and LID from three types of multi-layers are compared and the results are shown in **Figure 5**. The total released mass of DCF is higher for ALG 3 with a thickness of ca.11 nm than the other two multi-layers (CMC 3: 36 nm, ALG-CMC 3: 33 nm). It is suggested that the DCF as a small molecule with a molecular weight of 296.2 g/mol is expected to diffuse faster in and out of the thin multi-layer films. From this release result, it can be stated that even the thinnest multi-layer films can be used to incorporate and release

drugs in therapeutic doses in a controlled manner similar to that of three times thicker CMC 3 or ALG-CMC 3 films. Among other two multi-layers, a slightly lower mass is found for ALG-CMC 3. A reason can be that the diffusion of DCF through the thicker ALG-CMC 3 and hierarchically structured film from the two polymer components is prevented, and thus the overall released mass is lower. However, a different trend is observed for LID incorporated multi-layers. While ALG 3 and ALG-CMC 3 showed almost the same amount of the released mass, CMC 3 multi-layers showed a slightly higher mass release, which is the opposite to the results observed for DCF incorporated CMC 3 multi-layer. For LID incorporated layers (**Figure 4C**), although the release profile is similar to DCF, the overall released mass is 10 times higher when compared to DCF. This can be related to differences in the solubility of both drugs. It is suggested that upon contact with water, compared to DCF (solubility in water: 0.005 mg/mL), the hydrochloride form of LID (used in this study, solubility in water: 0.6 mg/mL) incorporated within the hydrophilic ALG-CMC 3 matrix dissolves and diffuses quicker through the film, and thus a larger amount of drug is released over time. Overall, the data show that it is possible to carefully tune the release rate of drug using one particular system, as it is demonstrated by several others (Vasilev et al., 2011; Wang et al., 2016; Maver et al., 2018a; Speer et al., 2019). Using thinner and thicker multi-layer coatings, the release rate can be extended even further (desired in treatment of chronic wounds is for multiple days).

To further evaluate the kinetics and mechanisms of drug (DIC and LID) release from the multi-layers, the in vitro release data were fitted by linear regression analysis according to different kinetic models. Although different kinetic models were considered, only the models (First-order, Higuchi-model, and Korsmeyer-Peppas model, see **Table 2**) that gave reasonable fits for the release data were selected [based on the square of the correlation coefficient (r 2 ) value]. The kinetic model that resulted in the highest r 2 value was considered as the best fit for the release data. Among the tested three kinetic models, the Korsmeyer–Peppas model had the best fit with the highest r 2 value for all multi-layers. Slightly lower r 2 values were calculated

for the Higuchi-model (reflected in **Table 2** and **Figure 6**). Since the first-order model resulted in much lower r 2 values, it was not considered for further interpretation and discussion of the results. Considering the best fit (highest r 2 value in **Table 2**), as well as considering the calculated first derivative results based on the release data (shown in **Figures 4B,D**), it can be assumed that more than one mechanism is involved in the release of the drug from all three multi-layers (Keny et al., 2009). For the fitting of the results, the release data from 0 to 360 min were used. The best fit was obtained with Korsmeyer-Peppas model, which is among the most common models to describe the drug release kinetics from polymeric systems, when the release mechanism is unknown, or more than one release phenomena is involved. The release component "n" value from the Korsmeyer– Peppas Equation (3) was shown to be related (at least partially) to different release mechanisms and can therefore be used for an estimation of the latter. From **Table 2** it can be seen that the "n" value is found to vary with type of multi-layers used, whereas the absolute value is in all cases above 0.89 for both drugs. This calculation leads to the conclusion that the release of both drugs from all system is an anomalous non-Fickian diffusion transport (super case-II transport mechanism). This means that the drug molecules are released through a diffusion process in a highly hydrated polysaccharide matrix that involved in the dissolution or relaxation of polymer chains (might be either swelling, erosion, or both) (Nayak and Pal, 2013). Since the Higuchi model came close to the best fit (r 2 : 0.96–0.97), let us consider what it would mean, if the latter model would hold. Considering that the Higuchi model presents the derivation of an equation that allowed for the quantification of drug release from thin ointment films, containing finely dispersed drug (applicable in perfect sink conditions), this model could also potentially be used to explain the release mechanism in this study. For example, the obtained r 2 values suggest that the release of drugs from the multi-layered system is controlled by diffusion, which is totally opposite to non-Fickian diffusion approach. Therefore, according to the best fit and highest r 2 value, we believe that the release of both drugs from all three multi-layered system follows Korsmeyer–Peppas model and no other kinetic models.

### Biocompatibility Testing

In vitro biocompatibility testing with human skin cells is an important indication for developed material safety and (depending on the type of material and purpose) to some extent

FIGURE 6 | Drug release data of DCF and LID loaded polymers fitted to various kinetic models.

FIGURE 7 | (A) HaCaT cells' viability determination using the MTT assay for all samples and the control; ADMEM + FBS (red line in graph), (B) Skin Fibroblast cells' viability determination using the MTT assay for all samples and the control; ADMEM + FBS (red line in graph). Error bars represent standard deviation of the replicate measurements. Statistical significance is defined as \*P < 0.05 (ANOVA test) for all samples compared to the control.

efficiency for wound application. In comparison to most available studies, where animal-derived or cancer-transformed cells are used, the results obtained using human cells can serve as a much better approximation of the actual influence of the developed materials in clinical use (Stana et al., 2017). The viability of keratinocytes (HaCaT) and skin fibroblasts (SF) with the drug incorporated multi-layers was tested to confirm the potential of prepared materials for wound care applications. Since extracts of the developed materials were tested against the skin cells, pure cell growth media (ADMEM + FBS) was used as the control sample. From **Figure 7**, which show the obtained results of this testing, it is immediately apparent that all samples with one exception among each of the developed drug filled multi-layer systems, outperformed the control sample. The exception is LID incorporated ALG3 and DCF incorporated ALG-CMC3 multilayer; however, no significant deviation was measured even in these samples, as well as no differences in the cell morphology was detected for neither of these samples, indicating that no toxicity whatsoever is present. Important conclusions, which can be made here; no toxic degradation products are formed in neither of the samples (namely, these would significantly affect the cell growth in a negative sense), and the tested multi-layers promoted the growth of cells, especially the skin fibroblasts (**Figure 7B**). After 48 h, the viability of skin fibroblasts significantly increased in almost all cases, whereas the best results were obtained for the drug-free and DCF included ALG3 multi-layer. Unexpectedly, the incorporation of LID did partially lower the viability of the cells compared to the drug free materials, although the values still remained above the control. Similar results were acquired for skin fibroblasts exposed to CMC 3 with and without included drugs. In the case of ALG-CMC 3, the viability decreased a little after addition of DCF, but nevertheless it remained above the one measured for the control sample. Similar results, only with somewhat lower values, were obtained in the case of HaCaT cells.

Altogether, the obtained results prove that regardless of the polymer-drug combinations used in this study, especially when considering the different mechanisms of action of the LID and DCF drugs, they did not exhibit any toxic effect on neither of the most abundant human skin cells (keratinocytes and fibroblasts). Therefore, we can conclude that tested materials with and without included drugs are biocompatible. The measured higher viabilities in most sample cases compared to the control, require further studies to evaluate the physiological mechanism behind, whereas at the moment, the obtained results indicate that the developed materials are very suitable for use in wound care.

### CONCLUSIONS

A simple and rapid method to fabricate nanoscale multilayered system from water soluble polysaccharides using the combination of spin-coated and LBL techniques is demonstrated. The potential application of multi-layered thin films created from ALG and CMC for the effective incorporation and controlled release of two analgesic drug molecules (DIC and LID) at a physiological condition is investigated in detail. Upon spincoating, the phase separation occurred for the ALG-CMC mixture, which could not be seen for either ALG or CMC. Such phase separated features are not influenced by the incorporation of either DIC or LID. The presence of latter can be confirmed by ATR-FTIR measurements; however, no substantial changes in the surface wettability are observed before and after the incorporation of drug molecules. Although the film thickness is increased for all three system with a number of deposition steps, a thicker film (highly adsorbed mass) is formed for both CMC and ALG-CMC. A rapid "burst" release followed by a prolonged release kinetics are observed for both drugs regardless

### REFERENCES

of the multilayered system, while the overall released mass is lower for ALG-CMC compared to either ALG or CMC coated surfaces alone. The release properties of both drugs are suited for Korsmeyer–Peppas model, which showed a best fit and the highest "r 2 value" when compared to other two models. The release behavior is predicted to be non-Fickian based on the exponent "n" value which is above 0.89 in all cases. Although the influence of incorporated drugs in the multi-layered thin films is not significant in the growth of both tested skin cells, a pounced growth of skin fibroblast cells is detected already for drug-free multi-layered films. Overall, the knowledge gained through this study is highly beneficial not only to develop a highly multifunctional materials but also for applications related to targeted drug-delivery and tissue engineering, in general.

### DATA AVAILABILITY

All datasets generated for this study are included in the manuscript and/or the **Supplementary Files**.

### AUTHOR CONTRIBUTIONS

TiM performed all experiments and contributed substantially to the writing of this manuscript. TaM contributed substantially to the writing, handled submission and revising of the manuscript. LG performed experimental work related to cell testing. MF, KS, and UM contributed to writing and revising of the manuscript.

### ACKNOWLEDGMENTS

The authors acknowledge the financial support from the Slovenian Research Agency for Research (grant numbers: P2- 0118, P3-0036, P2-0032, J1-9169, I0-0029, and Z2-8168).

### SUPPLEMENTARY MATERIAL

The Supplementary Material for this article can be found online at: https://www.frontiersin.org/articles/10.3389/fchem. 2019.00217/full#supplementary-material

influence on human primary endothelial cells. Biomacromolecules 18, 413–421. doi: 10.1021/acs.biomac.6b01504

**Conflict of Interest Statement:** The authors declare that the research was conducted in the absence of any commercial or financial relationships that could be construed as a potential conflict of interest.

Copyright © 2019 Maver, Mohan, Gradišnik, Finšgar, Stana Kleinschek and Maver. This is an open-access article distributed under the terms of the Creative Commons Attribution License (CC BY). The use, distribution or reproduction in other forums is permitted, provided the original author(s) and the copyright owner(s) are credited and that the original publication in this journal is cited, in accordance with accepted academic practice. No use, distribution or reproduction is permitted which does not comply with these terms.

## Multilayer Density Analysis of Cellulose Thin Films

Carina Sampl 1,2†, Katrin Niegelhell 1,2†, David Reishofer 1†, Roland Resel 3†, Stefan Spirk 1,2 \* † and Ulrich Hirn1,2†

*1 Institute for Paper, Pulp and Fibre Technology, Graz University of Technology, Graz, Austria, 2 CD-Laboratory for Fibre Swelling and Paper Performance, Graz University of Technology, Graz, Austria, 3 Institute of Solid State Physics, Graz University of Technology, Graz, Austria*

An approach for the multilayer density analysis of polysaccharide thin films at the example of cellulose is presented. In detail, a model was developed for the evaluation of the density in different layers across the thickness direction of the film. The cellulose thin film was split into a so called "roughness layer" present at the surface and a "bulk layer" attached to the substrate surface. For this approach, a combination of multi-parameter surface plasmon resonance spectroscopy (SPR) and atomic force microscopy (AFM) was employed to detect changes in the properties, such as cellulose content and density, thickness and refractive index, of the surface near layer and the bulk layer. The surface region of the films featured a much lower density than the bulk. Further, these results correlate to X-ray reflectivity studies, indicating a similar layered structure with reduced density at the surface near regions. The proposed method provides an approach to analyse density variations in thin films which can be used to study material properties and swelling behavior in different layers of the films. Limitations and challenges of the multilayer model evaluation method of cellulose thin films were discussed. This particularly involves the selection of the starting values for iteration of the layer thickness of the top layer, which was overcome by incorporation of AFM data in this study.

#### Edited by:

*Giuseppe Mensitieri, University of Naples Federico II, Italy*

### Reviewed by:

*Ram Gupta, Pittsburg State University, United States Marino Lavorgna, Institute of Polymers, Composite and Biomaterials, Italian National Research Counci, Italy*

> \*Correspondence: *Stefan Spirk<EMAIL_ADDRESS>
*†Members of NAWI Graz and the European Polysaccharide Network of Excellence (EPNOE)*

#### Specialty section:

*This article was submitted to Polymer Chemistry, a section of the journal Frontiers in Chemistry*

Received: *22 January 2019* Accepted: *27 March 2019* Published: *16 April 2019*

#### Citation:

*Sampl C, Niegelhell K, Reishofer D, Resel R, Spirk S and Hirn U (2019) Multilayer Density Analysis of Cellulose Thin Films. Front. Chem. 7:251. doi: 10.3389/fchem.2019.00251* Keywords: multilayer analysis, surface plasmon resonance, atomic force microscopy, cellulose thin film, X-ray reflectivity

### INTRODUCTION

The past decades have seen tremendous efforts to explore and to understand processes occurring at surfaces of materials, which was accompanied by the development of new surface sensitive techniques and methodologies. These techniques are either capable of determining the surface properties such as morphology, topography, chemical structure, and composition, or they are utilized to monitor dynamic phenomena taking place at interfaces, such as adsorption, modification, or wetting, to name a few (Vickerman and Gilmore, 2009; Zaera, 2011).

Atomic force microscopy (AFM) (Binnig et al., 1986), for instance, is an imaging technique for the investigation of surfaces on an atomic scale by scanning the surface using a cantilever with a sharp tip while it is maintained at a constant force or a constant height above the sample. Nowadays, the observation of surfaces in real-time using fast scanning devices (one frame per second and faster) allows for visualizing dynamic biological processes at interfaces. For example, enzymatic degradation of biopolymers such as cellulose can be monitored and videos of the degradation process can be acquired (Giessibl, 2003; Jalili and Laxminarayana, 2004; Niegelhell et al., 2016).

Surface plasmon resonance (SPR) spectroscopy is a surface sensitive technique that is able to monitor processes in real-time as well. In contrast to atomic force microscopy, it detects changes in the chemical environment, i.e., the refractive index n, near a metal surface (∼100 nm) (Raether, 1977). The technique is based on the resonance of surface plasmons, which originates from the oscillation of charge densities on the metal surface caused by freely moving electron gas (Ritchie, 1957). Acquisition of SPR spectra is accomplished by focusing a p-polarized light source onto a metal surface (e.g., a glass slide coated with a thin gold layer) and recording the intensity of the reflected light by a detection system (e.g., photodiode array detectors). State of the art multi-parameter SPR devices, acquire spectra in dependence of the incidence angle of the light, which allows for the investigation of processes in real-time and in varying ambient media (i.e., gas or liquid) at different wavelengths (i.e., 670 and 785 nm) in a single experiment. The resulting curves are evaluated by a multilayer fitting procedure, whereby thickness (d) and refractive index (n) of the examined layers are derived (Geddes et al., 1994). In a standard SPR spectrometer, where the measurement is performed with only a single wavelength, determination of both -layer thickness and refractive index- is not possible without assuming or knowing one of the values. Multiparameter SPR (MP-SPR) systems measuring at two or more wavelengths overcome this problem (Peterlinz and Georgiadis, 1996; Liang et al., 2010).

When observing phenomena with SPR spectroscopy in realtime, the change in refractive index related to the whole sample is detected. In most cases, this information is sufficient because it allows for monitoring adsorption processes at the interfaces, in terms of deposited mass and adsorption kinetics. Many examples do exist which investigate biomolecule deposition on cellulose thin films mainly with the aim to establish sensor systems or antifouling surfaces (Orelma et al., 2011, 2012; Niegelhell et al., 2016, 2017, 2018; Strasser et al., 2016; Mohan et al., 2017; Weißl et al., 2018). Since water incorporation into the film changes its refractive index, swelling has been investigated by SPR as well for the whole film (Kontturi et al., 2013). However, when it comes to more complex questions, i.e., whether the swelling of such a thin film is different on the "surface" than in the "bulk," other approaches with more complex fitting procedures are required. Up to now, the fitting procedures used in SPR data evaluation have only been used to describe the cellulose film as a single layer. When splitting the film into more than one layer, more information is required for the fitting procedure.

Herein, the investigation of such a multilayer analysis approach to evaluate density variations of different layers in thin films, at the example of cellulose, is presented. A model describing the cellulose thin film as a multilayer system was developed. The implementation of AFM data into the multilayer fitting model was of outmost importance in order to obtain starting values for the fitting procedure concerning thickness of bottom and top layer. This combination of analysis methods and the presented multilayer density analysis approach yielded spatial resolution along the z-axis of the examined films. Compositions of the entire films as well as differences between the surface regions and the bulk were evaluated. The approach was compared to X-ray reflectivity (XRR) measurements of cellulose thin films, which were also analyzed to obtain thickness and density data for a multilayer cellulose film system.

### THEORETICAL BACKGROUND

### Determination of Thickness and Refractive Index of Cellulose Thin Films: The 2-Wavelength Method (2-λ-Method)

The simulation of a sample measured at a single wavelength provides a refractive index (n)—thickness (d) continuum (n decreases when d increases) without a unique solution, since the surface plasmon wave vector ksp is dependent on both entities.

$$k\_{\text{SP}} \propto \ n \* d \tag{1}$$

In order to determine a unique solution, measurements at two wavelengths resulting in two different sets of n - d continua are required. Since the refractive index is dependent on the wavelength, the n - d continuum measured at one wavelength can be shifted to the other wavelength (Peterlinz and Georgiadis, 1996). The unique solution is found at the intersection of these continuum solutions.

$$k\_{\text{SP1}} = \; n\_{\text{\&1}} \ast d \; and \; \tag{2}$$

$$k\_{\text{SP2}} = \ n\_{\text{\&2}} \* d \tag{3}$$

where

$$n\_{\lambda 2} = \left( n\_{\lambda 1} + \frac{dn}{d\lambda} \left( \lambda\_2 - \lambda\_1 \right) \right) and \tag{4}$$

$$k\_{\rm SP1} = \; n\_{\lambda 1} \* d \; and \tag{5}$$

$$k\_{\rm SP2} = \left( n\_{\lambda 1} + \frac{dn}{d\lambda} \left( \lambda\_2 - \lambda\_1 \right) \right) \ast d \tag{6}$$

The dependence of refractive index on the wavelength, i.e., chromatic dispersion (dn/dλ), is approximated to be linear for relatively small wavelength changes (a few hundred nm).

For cellulose, the dn/dλ at 670 and 785 nm is −0.0338 and −0.0204 µm−1 , respectively (Kasarova et al., 2007). The average, −0.0271 µm−1 , was used in the calculations as a chromatic dispersion value of pure cellulose. The corresponding value for air is −0.00000856 µm−1 (Ciddor, 1996). The n - d curves of the cellulose thin films obtained by the two wavelengths were plotted in the same graph. The n - d curve of the measurement performed at 785 nm was shifted by the dn/dλ value of cellulose and by the dn/dλ value of the ambient medium (air). The intersection points of the shifted curves (air and cellulose) measured at 785 nm with the curve measured at 670 nm results in two unique solutions. An average (nfilm) of the n values obtained by the intersection points was determined. The proportion of cellulose of the volume of the film, a corresponding to the obtained nfilm value, was calculated according to Equation (7):

$$n\_{film} = \ a \cdot n\_{cellulose} + \ (1 - a) \cdot n\_{medium} \tag{7}$$

where ncellulose is 1.466 (Kasarova et al., 2007) and nmedium is 1.00028 for air (Ciddor, 1996).

Based on a, the value of dn/dλ was corrected by Equation (8) (Kontturi et al., 2013):

$$dn/d\lambda\_{\text{film}} = |a \cdot dn/d\lambda\_{\text{cellulose}} + (1 - a) \cdot dn/d\lambda\_{\text{medium}} \tag{8}$$

Then, the intersection point of the n - d curve measured at 670 nm and the n - d curve shifted from 785 to 670 nm with the new dn/dλ results in the unique solution (n and d) of the cellulose thin film (for a detailed example see **Figure 4** in section results and discussion). The composition a of the film was determined by Equation (7). The densities of the films were calculated from the composition and the density values of pure cellulose and air (see **Supplementary Material**) similar to Equation (7).

### Multilayer Analysis of Cellulose Thin Films

The procedure to obtain thickness d and the density (via refractive index n) of a thin film has been described in detail in the last section (Theoretical Background). These MP-SPR measurements are the basis for the multilayer analysis of thin film samples (**Figure 1**). The investigated film is split into two layers, namely the roughness layer (RL) and the bulk layer (BL). Thickness and density are determined for each layer.

The roughness layer RL describes the layer that contains the boundary between the film and the ambient medium i.e., the roughness of the film. The RL is composed of cellulose and air, the material fraction (mf) refers to the percentage of material within the volume of the RL.

Thin film data required to make use of the model is attained by MP-SPR spectroscopy. This data does not contain information on surface roughness, i.e., the roughness layer RL is treated as a flat layer with thickness dt and density ρ t m. The roughness layer thickness (RLTAFM) as well as its material fraction (mfAFM RL ) were acquired from AFM topography experiments. AFM data was evaluated as illustrated in **Figure 2** and implemented into simulations. A detailed description of the AFM evaluations is shown in the experimental section and in the **Supplementary Material**.

### Multilayer Data Analysis for 2-Layer Thin Films Analysis

We are using the modeling software Winspall to analyze film thickness and refractive index (density) from SPR measurements. The key approach of performing multilayer analysis is modeling the thickness for two layers of material instead of one layer of material in Winspall, in combination with the 2-λ method for determining n and d of the respective layers. Starting from a 1 layer analysis and using the AFM measurement results as starting values for the roughness layer thickness as starting values for the iteration a sequence of models is built to successively refine and confirm the results for refractive index (density) and film thickness in top- and bottom layer of the thin film. A step-by step description for this modeling procedure is shown in **Figure 3**. Finally, equations 9 to 12 are used to calculate the final results.

$$
\rho\_{RL}^{AFM} = \frac{m f^{AFM}}{100} \,\,\, \dot{\rho}\_m^t \,\, \,\, \,\, \rho\_m^t \tag{9}
$$

$$RTL^{\text{SPR}} = \frac{d\_l \cdot 100}{\text{cellulose content [\%]}} \tag{10}$$

$$
\rho\_{RL}^{\text{SPR}} = \frac{m f^{\text{SPR}}[\%]}{100} \cdot \rho\_m^t \tag{11}
$$

$$mf^{\text{SPR}} = \frac{d\_t \cdot 100}{RLT^{AFM}} \,. \tag{12}$$

Here ρ AFM/SPR RL is the density of the roughness layer and mfAFM/SPR is the material fraction calculated from AFM or MP-SPR data. RLT is the roughness layer thickness, dt is the thickness of the top layer and ρ t m is the density of the top layer from MP-SPR multilayer evaluation.

The nine steps of the 2-layer analysis as shown in **Figure 3** are described below. Notation is explained in the caption of **Figure 3**. For a detailed example over all 9 steps of the analysis of a cellulose thin film, see **Table S1**.

**STEP 1:** Starting point is a 1-layer analysis of the entire film according to the 2-λ method. The result is thickness d SPR and density ρ SPR (refractive index n SPR, respectively) for the entire cellulose film.

**STEP 2:** The film is split into two layers –the top and bottom layer- denoted as d 0 t and d0 b . Whereas, the starting d 0 t values for evaluation are derived from AFM data with d 0 t = RLTAFM/2. Starting d 0 b value is set to d SPR—d 0 t . The refractive index of the entire cellulose thin film n SPR is chosen as starting refractive index value for both, top (n 0 t ) and bottom layer (n 0 b ).

**STEP 3:** Iteration for the corresponding refractive indices; Result: refractive indices n 0 t and n 0 b .

**STEP 4:** Evaluation of the bottom layer by 2-λ method yielding d 1 b and n 1 b , while keeping the values for the top layer (d 0 t , n 0 t ) constant.

**STEP 5:** The roughness layer thickness obtained from AFM imaging (RLTAFM) is implemented into the model as new thickness value for the top layer (d 1 t ).

**STEP 6 and 7:** The corresponding refractive index (n 1 t ) is iterated in the simulation program and d 2 t and n 2 t of the top layer is evaluated by the 2-λ method, while the bottom layer values (d 1 b , n 1 b ) are kept constant.

**STEP 8:** Evaluation of the bottom layer by the 2-λ method while keeping the new values for the top layer (d 2 t , n2 t ) constant, in order to validate the simulations.

**STEP 9:** To calculate the roughness layer's thickness (RLT), the density (ρ SPR/AFM RL ) and the material fraction (mfSPR/AFM) data from AFM and SPR as iterated above are calculated using Equations (9–12).

### EXPERIMENTAL

### Materials

Trimethylsilyl cellulose (TMSC, DSSi = 2.8, Avicel, Mw = 185.000 g·mol−1 , Mn = 30.400 g·mol−1 , PDI = 6.1 determined by SEC in chloroform) was purchased from TITK (Rudolfstadt, Germany). Chloroform (CHCl3, 99.3%), hydrochloric acid (HCl, 37 %), hydrogen peroxide (H2O2, 30% in water) and sulfuric acid (H2SO4, 95%) were purchased from Sigma Aldrich and used as received. MilliQ water (resistivity = 18 M·cm) from a Millipore water purification system (Millipore, USA) was used for all experiments. A total of three films was measured with SPR and AFM and the 2-layer analysis was performed.

layer (*BL, db,* ρ *b m*).

### Substrate Cleaning and Film Preparation

SPR sensor slides—glass sensors with chromium adhesion layer (∼5 nm) and gold coating (∼50 nm) (CEN102Au)—were obtained from Cenibra, (Bramsche, Germany). In order to remove adventitious carbon, SPR sensors were cleaned before use by treatment with piranha solution (freshly prepared from sulfuric acid and hydrogen peroxide in a 3:1 (v/v) ratio) over a period of 10 min. Afterwards the SPR slides were extensively rinsed with MilliQ water and dried in a stream of nitrogen. The silicon wafer substrates (native oxide layer, 1.4 × 1.4 cm²) for the XRR measurements were cleaned with "piranha" acid [H2SO4:H2O2 = 7:3 (v/v)] for 30 min and neutralized afterwards with distilled water.

Cellulose thin films were prepared by spin coating trimethylsilyl cellulose (0.75 wt% in CHCl3) onto the gold slides. One hundred microliters of TMSC-solution were deposited on the substrate and then rotated for 60 s at a spinning speed of 4,000 rpm and an acceleration of 2,500 rpm·s −1 . TMSC was converted to cellulose by treatment with hydrochloric acid (HCl) vapor. The TMSC films were placed into a petri dish (diameter 5 cm) containing 3 ml of 10% HCl. The dish was covered with its cap and the films were exposed to the HCl vapors for 15 min. The regeneration was verified by water contact angle and ATR-IR measurements as reported elsewhere (Kontturi et al., 2003; Woods et al., 2011; Mohan et al., 2012a,b).

### Multi-Parameter Surface Plasmon Resonance (MP-SPR) Spectroscopy

Two-wavelength MP-SPR spectroscopy experiments were performed with an MP-SPR NaviTM 210A Vasa instrument (by BioNavis Ltd., Tampere, Finland) equipped with two light source pairs providing 670 and 785 nm in each of the two measurement channels. All measurements were performed using a full angular scan (39−78◦ , scan speed: 8◦ ·s −1 ) in three parallels. SPR data was processed with BioNavis Dataviewer software. The full angular scans were simulated with the optical fitting software Winspall 3.01 (which is freely available from the Max-Planck Institute for Polymer Research (Mainz, Germany), http://www2. mpip-mainz.mpg.de/groups/knoll/software, 12.6.2013). The multilayer fitting approach is based on the Fresnel equations and the recursion formalism. The SPR signal of the pure sensor surface was simulated first in order to obtain the background

for the subsequent evaluation of the cellulose thin film. The two wavelengths cross point analyses were performed both using Microsoft Office Excel 2010 and Origin 8.6.

### Atomic Force Microscopy (AFM)

AFM imaging was performed in atomic force microscopy tapping mode with a Veeco multimode scanning probe microscope (Bruker, USA). The images were scanned using silicon cantilevers (NCH-VS1-W, Nanoworld, Switzerland) with a resonance frequency of 320 kHz and a force constant of 42 N·m−1 . All images were processed using Nanoscope software package (V7.30r1sr3, Veeco).

The roughness layer thickness RLTAFM is evaluated from the topography images measured by AFM, compare **Figure 2**. The roughness layer is the region of the cellulose film where both, material and ambient medium (water or air) are found, i.e., the region containing the boundary between cellulose film and surrounding. The thickness of this region depends on the roughness of the film. We have defined the roughness layer as the z-directional layer containing 95% of the surface roughness. For calculation, the histogram of the topography distribution is evaluated. From each edge of the histogram, 2.5% of the topography values are clipped off, the topography range comprising the remaining

95% of height values is defined as the roughness layer thickness. We are not using 100% of the values because the topography distribution is close to normally distributed which means there are always some far outliers which would bloat the roughness layer beyond the true surface interaction range.

### X-Ray Reflectivity XRR

X-ray reflectivity measurements were performed using a PANalytical Empyrean goniometer system with radiation produced by a copper sealed tube (λ = 0.154178 nm). The primary side of the reflectometer was equipped with a 20 mm beam mask, a multilayer mirror, a 1/32◦ slit, and an automatic beam attenuator. On the secondary side, a receiving slit of 0.1 mm and a Soller slit of 0.02 rad were used in front the PANalytical PIXEL3D point detector. The sample stage was a domed DHS 900 from Anton Paar (Resel et al., 2003), equipped with a SHT15 humidity sensor to monitor the relative humidity and the temperature during the measurements. The relative humidity (RH) was controlled using a S-503 humidity generator from Michell instruments. For each humidity step an equilibration time of 30 min was accomplished. XRR measurements were performed in the 2θ region 0.030–9.999◦ with a step size of 0.006◦ . The evaluation of the data was performed with the X'Pert Reflectivity (Panalytical, C6H10O5 for cellulose was used) software package providing information on the electron density, layer thickness, and the roughness of the films by applying (Parratt, 1954) formalism and the disturbance term of Nevot–Croce(Nevot and Croce, 1980). A three layer model was required to fit the experimental data of the cellulose film (50% relative humidity, same as in laboratory where the SPR spectrometer is located), resulting in a very thin layer at the Si-cellulose interface (d = 0.6 nm, ρ = 0.8 g cm−3 ), a bulk layer (d = 43.9 nm, ρ = 1.39 g cm−3 ) and a surface layer (d = 4.2 nm, ρ = 1.09 g cm−3 ). Total film thickness was 48.7 nm, with a density of 1.41 g cm−3 at 50% relative humidity. For the sake of comparison, data obtained at other humidity levels (0, 25, and 70% was acquired. The difference in layer thickness compared to the gold substrate stems from the different wettability of the silicon wafer (which is required for XRR) leading to higher thicknesses.

### RESULTS AND DISCUSSION

### Characterization and Preparation of Cellulose Thin Films

Cellulose thin films were prepared by spin coating of trimethylsilyl cellulose (TMSC), dissolved in chloroform (CHCl3), and subsequent conversion to cellulose by a regeneration step completed with HCl vapors, cleaving off the silyl-groups by formation of TMSCl (Kontturi et al., 2003; Kontturi and Lankinen, 2010). The completed conversion to cellulose was proven by ATR-IR spectroscopy, proving the disappearance of bands related to the TMS group (υSiC, δSiOC at 1,250 and 852 cm−1 ) while the typical OH stretching vibrations (υOH) for cellulose at 3,200–3,600 cm−1 appeared (Ehmann et al., 2015). The AFM images display smooth, homogeneous surface topography (**Figure 4**). Evaluation of thickness from AFM-imaging results in a RLT (roughness layer thickness) of 6.4 nm and a cellulose material fraction (mfAFM) of 47.9 % in that film.

### 2-Layer Analysis of Thickness and Density in Thin Films

Thickness and refractive indices of three individual cellulose thin films were first determined for the entire film using the 2-λ method (**Figure 5**) as described above. In principle, cellulose thin films can be prepared with high reproducibility and only slight variations in film thickness. For example, the thickness evaluation of three thin films applying the 2-λ method led to just minor deviations (d SPR of 36.4 ± 0.5 nm). The refractive indices at 785 and 670 nm are 1.408 ± 0.028 and 1.410 ± 0.024, respectively, which can be used to determine the cellulose content a in the whole film (Equation 7, 88.8 ± 0.1%). Based on that result, the film was subjected to multilayer density analysis as outlined in the previous sections (**Figure 3**). The first result was that the

FIGURE 5 | 2-λ method: (A) Continuous solutions (*n* – *d* curves) obtained at 670 and 785 nm at the example of a dry cellulose thin film (film 1 in Table S3) spin coated from CHCl3. (B) The curve measured at 785 nm was shifted by the *dn/d*λ of air (−0.00000856 µm−1 ) and cellulose (−0.0271 µm−1 ) in order to determine the composition of the film. The new *dn/d*λ value for the film consisting of air and cellulose was re-estimated from the composition (average of refractive indices at both intersection points) with equations 7–8. (C) The *n* – *d* curve recorded at 785 nm was shifted by the resulting *dn/d*λ to obtain the unique solution (*d* = 36.9 nm, *n* = 1.406, *a* = 89 % cellulose) for the investigated thin film.

refractive indices of the top and bottom layer did not yield the same result. We found much smaller values for the top layer

*RLTSPR, mfSPR and* ρ *SPR*/*AFM RL were calculated according to Equations (9-*−*12). Note that for XRR, films featured a different thickness due to the use of a different substrate and the values that have been given in the table were obtained at 50% relative humidity.*

(nt = 1.237 ± 0.034 at 670 nm; 1.242 ± 0.030 at 785 nm) than for the bottom layer (nb = 1.421 ± 0.012 for 670 and 1.420 ± 0.011 for 785 nm). As a consequence, the cellulose content in the bottom layer (a SPR b = 91.8 ± 0.1 %) is much larger than in the top layer (a SPR t = 52.2 ± 0.1 %). The thickness of the entire films and the sum of the top and bottom layer thicknesses obtained by our multilayer analysis are consistent (one layer analysis: d SPR = 36.4 ± 0.5 vs. two layer analysis 36.5 ± 0.6 nm (bottom db = 33.7 ± 0.7 and top layer dt = 2.8 ± 0.2 nm). Further, the density of the whole film (ρ SPR = 1.34 ± 0.09 g·cm−3 ) is very well-represented by the individual densities of the top and bottom layers (bottom ρ b m = 1.39 ± 0.04 g·cm−3 and top ρ t m = 0.76 ± 0.10 g·cm−3 ). The results from the multilayer density analysis using data obtained from AFM and MP-SPR spectroscopy experiments are depicted in **Table 1.** Evaluation results of the three cellulose thins films are shown in **Table S2**. Further, each individual thin film is evaluated in a multiple manner and compared to the other films, in order to demonstrate stability of the fitting approach and indicate deviations (see **Table S3**).

### Comparison of AFM, SPR, and XRR Data

The resulting data from AFM, MP-SPR spectroscopy, and XRR was compared to validate the employed strategy for multilayer analysis. The material fraction (mf, composed of cellulose and air) and RLT were acquired by AFM. The cellulose content, i.e., the density of the material (ρ t m), and the thickness of the layer consisting of material, i.e., the top layer (dt), were obtained from MP-SPR spectroscopy. For each technique, values which correspond to the same parameter determined by the other technique (ρ AFM RL , ρ SPR RL , RLTSPR, and mfSPR) were calculated from the measured results (mfAFM, RLTAFM, dt and ρ t m), according to Equations (9–12). The outcome of these calculations is summarized in **Table 1**. While there are some minor deviations, the results are very consistent for AFM and 2-layer MP-SPR approach. The deviations can to some extent be attributed to the different measurement areas of the applied techniques and small variations in the cellulose thin films. For instance, AFM covers an area of 10 × 10 µm2 , whereas the area of the MP-SPR spectroscopy lasers (Ø = 0.6 mm) is ∼0.28 mm2 .

X-Ray reflectivity was employed to further validate the approach. However, the results are not directly comparable in terms of absolute layer thicknesses since gold substrates (used in SPR spectroscopy) cannot be used for that technique. Instead, films have been deposited on silicon wafers which have a different surface free energy, leading to different cellulose film thickness. Further, the film thickness is a function of humidity level; by increasing humidity levels, layer thickness increases (from 0 to 70% r.h. thickness increase by 14%, see **Table S4**, for full data and **Figure 6A** for the XRR curves including fitting). At the same humidity levels as in the labs where the AFM and the SPR spectrometer are located (50% r.h.), we observed a surface layer with reduced density (thickness: 4.2 nm, ρ: 1.05 g· cm−3 ) as well as a bulk layer with a higher density (thickness: 43.9 nm, ρ: 1.39 g cm−3 ). The share of the top layer on the overall layer thickness is 7.7 and 9.1% for cellulose thin films on gold and silicon wafers, respectively. Given the different experimental setup and the difference in the substrates, the XRR data is in excellent agreement with the multilayer density approach using AFM/SPR. The densities of the prepared films are in the range of molecular dynamics simulations on amorphous cellulose (1.34–1.39 g cm−3 ) (Mazeau and Heux, 2003; Chen et al., 2004) but significantly lower than those of reported bulk amorphous substrates (1.48 g cm−3 ) (Mark, 1982). It should be noted that XRR revealed a thin layer (0.6 nm) at the cellulose-Si interface with rather low density (0.8 g cm−3 ). This is a known phenomenon for many polymer films but could not be revealed by SPR spectroscopy. The density profile of the layered system obtained by XRR is depicted in **Figure 6B**.

In summary both, MP-SPR and XRR yield a similar layer structure of the cellulose films, with a lower material density in a few nanometers thick top layer of the film and a higher density in the bulk of the film.

### Potential and Limitations of the Method

Finally, the limitations and requirements of the multilayer density analysis approach need to be elucidated. The crucial point in the presented multilayer approach is to have appropriate starting values for the thickness of the bottom and the top layer (dt 0 and db 0 ) for the data analysis in order to obtain physically meaningful results. Unique solutions for given RLT/2 values were only obtained when the starting values for the thickness were in between d SPR and the value obtained by iteration of the simulation program (d 0 t ). A randomly chosen starting value for the RLT does not lead to a result corresponding to the real RLT of the thin film. Furthermore, some randomly chosen starting values of the RLT did not even yield intersection points of the n - d curves. However, by using AFM data providing starting values for RLT, physically meaningful results can be obtained, which was confirmed by XRR. The importance of implementing AFM data as starting values for the fitting process is shown in more detail in **Table S1**. Comparability of fitting results and deviations within multiple evaluations of the individual thin films is described in **Table S3** (**Supplementary Material**). The main advantage of SPR is that it can be used with a wide range of liquids while by XRR this is a rather challenging task. The multilayer analysis of thin films to explore swelling processes in different

liquids is a potentially powerful application and will be applied in future reports.

### CONCLUSION

We demonstrated a multilayer density analysis of thin films at the example of cellulose in air. We showed that the combination of AFM and SPR data allows for analyzing the density variation and thickness in surface near regions of cellulose thin films. The key finding, a layer with reduced density and a thickness of a few nanometers at the top of the cellulose film, was confirmed by XRR measurements on similar films. We believe that this approach has the potential to reveal the density in multilayer structures of a wide range of materials (biopolymer films, porous thin films), e.g., for density analysis in thin film structures or for water uptake during swelling of cellulose with different types of liquids. Still, the approach features some limitations and further improvements need to be accomplished. A major shortcoming is that the starting values used for the splitting of the layer are very important to get meaningful results in the fitting procedure, i.e., an educated guess is required, which was provided by AFM in this

study. A further limitation is the used modeling software which seems to have minor problems with numerical stability (see **Tables S2, S3**), when inappropriate starting values were used. Further improvements of the approach in future studies will focus on more robustness toward the starting values used for the fitting procedures. A promising approach is to employ 4 or 6- different wavelengths in MP-SPR measurements and to use this additional information together with adequate modeling software for an improved multilayer analysis of thin films.

### AUTHOR CONTRIBUTIONS

All authors listed have made a substantial, direct and intellectual contribution to the work, and approved it for publication.

### FUNDING

Federal Ministry of Economy, Family and Youth and the National Foundation for Research, Technology and Development, Austria.

### REFERENCES

### ACKNOWLEDGMENTS

The financial support of the Federal Ministry of Economy, Family and Youth and the National Foundation for Research, Technology and Development, Austria, is gratefully acknowledged. We also thank the industrial partners Mondi, Océ, Kelheim, and SIG Combibloc for their support. Martin Steer, Jakob Häglsperger, and Johannes Rögner are gratefully acknowledged for providing inspiration for this work.

### SUPPLEMENTARY MATERIAL

The Supplementary Material for this article can be found online at: https://www.frontiersin.org/articles/10.3389/fchem. 2019.00251/full#supplementary-material

Supplementary Material | Supporting information is available including AFM topography evaluation, background MP-SPR curves,additional simulation experiments for multilayer analysis, results of multilayer analysis and comparison of multiple evaluations.

surface plasmon resonance spectroscopy. Opt. Commun. 130, 260–266. doi: 10.1016/0030-4018(96)00238-6

cellulose xanthate: properties and characterization. Cellulose 25, 711–721. doi: 10.1007/s10570-017-1576-3

**Conflict of Interest Statement:** The authors declare that the research was conducted in the absence of any commercial or financial relationships that could be construed as a potential conflict of interest.

Copyright © 2019 Sampl, Niegelhell, Reishofer, Resel, Spirk and Hirn. This is an open-access article distributed under the terms of the Creative Commons Attribution License (CC BY). The use, distribution or reproduction in other forums is permitted, provided the original author(s) and the copyright owner(s) are credited and that the original publication in this journal is cited, in accordance with accepted academic practice. No use, distribution or reproduction is permitted which does not comply with these terms.

## Design of Friction, Morphology, Wetting, and Protein Affinity by Cellulose Blend Thin Film Composition

Caterina Czibula1,2†, Gundula Teichert 3†, Maximilian Nau4 , Mathias Hobisch3 , Chonnipa Palasingh5 , Markus Biesalski 4 , Stefan Spirk 2,3, Christian Teichert 1,2 \* and Tiina Nypelö5,6 \*

#### Edited by:

Giuseppe Mensitieri, University of Naples Federico II, Italy

#### Reviewed by:

Artur J. M. Valente, University of Coimbra, Portugal Ram Gupta, Pittsburg State University, United States

#### \*Correspondence:

Christian Teichert<EMAIL_ADDRESS>Tiina Nypelö<EMAIL_ADDRESS>
†These authors have contributed equally to this work

#### Specialty section:

This article was submitted to Polymer Chemistry, a section of the journal Frontiers in Chemistry

Received: 28 January 2019 Accepted: 26 March 2019 Published: 03 May 2019

#### Citation:

Czibula C, Teichert G, Nau M, Hobisch M, Palasingh C, Biesalski M, Spirk S, Teichert C and Nypelö T (2019) Design of Friction, Morphology, Wetting, and Protein Affinity by Cellulose Blend Thin Film Composition. Front. Chem. 7:239. doi: 10.3389/fchem.2019.00239 1 Institute of Physics, Montanuniversität Leoben, Leoben, Austria, 2 Christian Doppler Laboratory for Fiber Swelling and Paper Performance, Graz University of Technology, Graz, Austria, 3 Institute of Paper, Pulp and Fiber Technology, Graz University of Technology, Graz, Austria, 4 Macromolecular Chemistry and Paper Chemistry, Technical University Darmstadt, Darmstadt, Germany, 5 Division of Applied Chemistry, Department of Chemistry and Chemical Engineering, Chalmers University of Technology, Gothenburg, Sweden, 6 Wallenberg Wood Science Center, Gothenburg, Sweden

Cellulose derivate phase separation in thin films was applied to generate patterned films with distinct surface morphology. Patterned polymer thin films are utilized in electronics, optics, and biotechnology but films based on bio-polymers are scarce. Film formation, roughness, wetting, and patterning are often investigated when it comes to characterization of the films. Frictional properties, on the other hand, have not been studied extensively. We extend the fundamental understanding of spin coated complex cellulose blend films via revealing their surface friction using Friction Force Microscopy (FFM). Two cellulose derivatives were transformed into two-phase blend films with one phase comprising trimethyl silyl cellulose (TMSC) regenerated to cellulose with hydroxyl groups exposed to the film surface. Adjusting the volume fraction of the spin coating solution resulted in variation of the surface fraction with the other, hydroxypropylcellulose stearate (HPCE) phase. The film morphology confirmed lateral and vertical separation and was translated into effective surface fraction. Phase separation as well as regeneration contributed to the surface morphology resulting in roughness variation of the blend films from 1.1 to 19.8 nm depending on the film composition. Friction analysis was successfully established, and then revealed that the friction coefficient of the films could be tuned and the blend films exhibited lowered friction force coefficient compared to the single-component films. Protein affinity of the films was investigated with bovine serum albumin (BSA) and depended mainly on the surface free energy (SFE) while no direct correlation with roughness or friction was found. BSA adsorption on film formed with 1:1 spinning solution volume ratio was an outlier and exhibited unexpected minimum in adsorption.

Keywords: blend films, spinodal decomposition, cellulose, friction, protein adsorption, adhesion

### INTRODUCTION

Spinodal decomposition of polymer blends can generate thin films with multi-phase surface composition, and often, complex morphology (Heriot and Jones, 2005). Lateral separation of the phases on a substrate results in patterns where islands of one phase reside within the other phase. Vertical separation can lead to heterogeneous distribution in the film's z-direction induced by surface energy differences of the components and can influence the surface morphology (Karim et al., 1998).

An intriguing area of research is the exploration of blend thin films obtained with spin coating. During the spin coating step, macroscopically homogeneous solutions of two polymers phase separate into the domains (Dalnoki-Veress et al., 1997). The ratio of the two polymers, molecular weight, solvent and spin coating conditions define the feature sizes and shapes (Xue et al., 2012). Synthetic polymer phase separation is exploited in coatings and organic electronics (Halls et al., 1995), while for biopolymer blend thin films their utilization and fundamentals, are still in their origins.

Cellulose synthetized by nature is notorious for insolubility having only few one-component solvents such as Nmethylmorpholine-N-oxide (Medronho et al., 2012). In industrial fiber production, cellulose insolubility has been circumvented by derivatization followed by conversion to cellulose commonly referred to as regeneration. Similar approaches have been applied in thin film manufacturing involving derivatization, dissolution, film formation, and regeneration of the derivatized film to cellulose (Schaub et al., 1993; Kontturi et al., 2003a,b). Spin coating, and propagation of repeating submicron patterns, requires volatile and good solvents for the blend components. Trimethylsilyl cellulose (TMSC) dissolved in toluene or chloroform has been used for single-component cellulose films as well as for blend films (Kontturi et al., 2009, 2010; Nyfors et al., 2009; Niegelhell et al., 2016, 2017; Strasser et al., 2016). Regeneration takes place upon exposure to hydrochloric acid vapor (Schaub et al., 1993).

Cellulose in products is often an inert component intended to protect or seal, such as package products, or to carry functionalities as in paper-based diagnostics (Pelton, 2009). As a substrate it can be used to accommodate follow-up chemistries, for example, to bind peptides to further tune it for specific protein or antibody affinity and sensing of biological molecules (Orelma et al., 2012a,b; Zhang et al., 2013). In addition to detection, protein adsorption is a way to design medical materials or to control fouling of surfaces. The inhibition of protein deposition, on the other hand, is an asset when bacterial growth and biofilm formation are required to be blocked.

Biofilm formation on surfaces takes place in two phases where the first one includes reversible physical attachment of bacteria and the second irreversible, cellular phase. Surface roughness, wetting and surface configuration have been identified as key parameters for bacterial adhesion (An and Friedman, 1998). The effect of hydrophobicity is directly in connection with the properties of the bacteria while with respect to roughness there is direct evidence that increase in roughness and hence, in the surface area—promotes bacterial adhesion. Roughness is different from surface configuration that refers to patterning on the surface. The periodicity and size of surface patterns has been found to be a parameter to inhibit bacterial adhesion. While bacteria preferentially adhere to irregularities that conform to their size since this maximizes bacteria-surface area (Katsikogianni and Missirlis, 2004), there is evidence that specific surface pattern design can prevent the attachment. A pattern mimicking a skin of shark was able to significantly reduce biofilm formation (Chung et al., 2007). The key parameters of the biofilm inhibiting films are non-random patterns with a hierarchy where the size is optimized to the size of a specific bacteria (Schumacher et al., 2007a,b). Here, we apply spinodal separation to generate periodical cellulose blend film hierarchies and evaluate the morphology and configuration and discuss their relation to antifouling surfaces.

Atomic force microscopy (AFM) is a non-destructive analytical tool applying low forces and is therefore suited for nanoscale characterization of soft polymer and biological surfaces. Apart from morphology studies, phase contrast in tapping mode imaging (Tamayo and Garcia, 1996), chemical contrast with functionalized AFM tips (Frisbie et al., 1994), and mechanical contrasts (Chyasnavichyus et al., 2015; Kocun et al., 2017) are established investigation routines. Phase and adhesion contrast measurements have been explored also on cellulosic films to achieve chemical contrast with functionalized probes (Ganser et al., 2016). Friction behavior of blend films can be studied on the nanoscale by friction force microscopy (FFM) (Mate et al., 1987; Marti et al., 1990; Meyer and Amer, 1990). In FFM, the AFM tip scans in contact mode normal to the cantilever's long axis, and the resulting cantilever torsion is related to the friction coefficient. Friction contrast for polymers by FFM has been demonstrated for phaseseparated thin organic films (Overney et al., 1992, 1994) where differences in the friction signal between hydrocarbon and fluorocarbon containing domains were found. More recently, FFM was employed to correlate friction to viscoelastic relaxation (Hammerschmidt et al., 1996; Sondhauß et al., 2015) and to characterize photoreactive organic surface patterns of spin casted thin films (Hlawacek et al., 2009; Shen et al., 2014). Polyisoprene and polystyrene blend coatings were recently reported to show an unexpected tribological synergy (Emerson et al., 2017). By varying the composition of the blend films, it was possible to tune the tribological properties and achieve friction coefficients which are much lower than for the pure films.

A blend of TMSC (regenerable to cellulose) and hydroxypropylcellulose stearate (HPCE) resulted in micropatterned films with varying aliphatic surface concentration. HPCE, a cellulose derivative, exhibits long alkyl side chains which may act as brushes and in combination with cellulose impact, wetting, adhesion, and protein adsorption. The surface morphology, roughness and lateral correlation length, of the films was quantitatively studied using AFM. The friction behavior was analyzed by FFM and compared to the adhesive properties of the surfaces of the pure and blend films obtained from AFM force spectroscopy. Finally, the comprehensive information obtained by surface characterization was employed to determine the influence of adhesive and tribological surface properties of the cellulose-HPCE blend films on bovine serum albumin (BSA) adsorption investigated by surface plasmon resonance spectroscopy (SPR) and Quartz Crystal Microbalance with Dissipation monitoring (QCM-D).

### MATERIALS AND METHODS

### Materials

Hydroxypropylcellulose stearate (HPCE, Mw 134,700 g mol−1 , Mn 107,400 g mol−1 , polydispersity index 1.25, degree of substitution (DS) 3.0) was synthesized according to a literature protocol (Nau et al., 2018). Trimethylsilyl cellulose (TMSC, From Avicel, Mw 185,000 g mol−1 , Mn 30,400 g mol−1 , polydispersity index 6.1, DS 2.8) was purchased from TITK (Thuringian Institute of Textile and Plastics Research, Germany). The structures are shown in **Figure 1**. Chloroform (99.3%), disodium phosphate heptahydrate (Na2HPO4·7H2O), sodium dihydrogen phosphate monohydrate (NaH2PO4·H2O), hydrochloric acid (37%), and bovine serum albumin (BSA) were purchased from Sigma-Aldrich and were used as received. Silicon wafers used as film substrates were cut 1 cm × 2 cm. Surface plasmon resonance (SPR) gold sensor slides (CEN102 AU) were purchased from Cenibra (Germany). Milli-Q water (resistivity = 18.2 −1 cm−1 at 25◦C) from a Millipore water purification system (Millipore, U.S.A.) was used for contact angle, SPR, and QCM-D investigations.

### Substrate Cleaning and Film Preparation

The film substrates were cleaned by immersing them in an insitu produced peroxymonosulfuric acid containing H2O2 (30

wt%)/ H2SO4 (1:3 v/v) for 10 min for SPR spectroscopy slides or 30 min for silicon wafers, respectively. After rinsing with deionized water, the wafers were dried with nitrogen gas, rinsed and stored in deionized water. TMSC and HPCE were dissolved in chloroform in a concentration of 0.75 wt%, using a water bath heated to 30◦C, and 120 h on a magnetic stirrer. Right before use the solutions were filtered through 0.45µm PVDF filters (Chromafil) and mixed in volumetric ratios labeled further on as TMSC:HPCE 1:0, 1:3, 1:1; 3:1, and 0:1. A volume of 100 µl was used for spin coating and operated for 60 s with an acceleration of 2,500 rpm s−1 and a speed of 4,000 rpm.

The conversion of TMSC into cellulose was implemented in a polystyrene petri dish (5 cm in diameter) containing 3 ml of 10 wt% HCl. The substrates were exposed to HCl vapor for 12 min. The regeneration of cellulose from TMSC was verified by contact angle and ATR-IR measurements (Alpha FT-IR spectrometer, Bruker, U.S.A.) using an attenuated total reflection attachment and obtaining spectra between 4,000 and 400 cm−1 with 48 scans and a resolution of 4 cm−1 . The data was analyzed with OPUS 4.0 software.

### Profilometry

Thickness of the thin films was determined by scratching the films with a scalpel and measuring the profile of a scan length of 1,000µm and a duration of 3 s using a DETAK 150 Stylus Profiler from Veeco (Bruker, USA) on a hydraulic balanced stone table with a diamond stylus with a radius of 12.5µm and a force of 3 mg. Three films of each sample were measured at 6 locations before and after regeneration. Film thickness and film roughness was calculated from the resulting profile using Software Vision 64.

### Contact Angle (CA) and Surface Free Energy (SFE) Determination

Static contact angle measurements were performed with a Drop Shape Analysis System DSA100 (Krüss GmbH, Germany) with a T1E CCD video camera (25 frames per second) and the DSA1 v 1.90 software. All measurements were performed at least three times on minimum two manufactured films with Milli-Q water and diiodomethane using a droplet size of 3 µL and a dispense rate of 400 µL min−1 . Static CAs were calculated with the Young-Laplace equation, and the SFE was determined with the Owen-Wendt-Rabel-Kaelble (OWRK) method. Surface tension of 50.80 and 72.80 mN m−1 for diidomethane and water were used, respectively.

### Adsorption Experiments

A phosphate buffer containing 8.1 mM disodium phosphate, 1.9 mM sodium phosphate and 100 mM sodium chloride at pH 7.4 was used to carry out the adsorption experiments of BSA on the films. Surface Plasmon Resonance Spectroscopy (SPR) was performed with a MP-SPR Navi 200 from Bionavis Ltd (Finland), using 785 nm laser in both measurement channels. The attached autosampler MP-SPR Navi 210A was set to 20 µl min−1 flow rate. The equilibration of the thin films was observed by measuring the spectra with full angular scan (39–78◦ ) and scan speed of 8◦ s −1 at 24.5◦ and plot the SPR-angle over time. A concentration of 1.0 mg ml−1 of BSA was dissolved in the buffer and exposed to the thin films for 10 min. Adsorbed mass (Ŵ) was calculated with the de Feijter equation,

$$
\Gamma = \frac{\Delta \Theta^\* \kappa^\* d\_{\mathcal{P}}}{d n/d c} \tag{1}
$$

using the refractive index increment (dn/dc) 0.182 cm3 g −1 . The 12 is the angular response of the surface plasmon resonance. For thin layers (<100 nm), k × dp can be considered constant and can be obtained by calibration of the instrument by determination of the decay wavelength ld. Here it was 1.09 × 10−7 cm/◦ (at 670 nm) and 1.9 × 10−7 cm/◦ (at 785 nm) in aqueous systems.

Quartz Crystal Microbalance and Dissipation (QCM-D) instrument (model E4) from Q-Sense (Sweden) was used with gold sensors purchased from QuartzPro (Sweden). The attached peristaltic pump was set to 0.1 ml min−1 . Adsorption was performed in the same conditions as the SPR analyses. The data was analyzed using Johannsmann modeling (Johannsmann et al., 1992; Naderi and Claesson, 2006).

### Atomic Force Microscopy

Most AFM and all FFM measurements were acquired using an Asylum Research MFP-3D AFM (USA). The instrument is equipped with a closed-loop planar x-y-scanner with a scanning range of 85µm × 85µm and a z-range of 15µm. The tapping mode AFM images were recorded with standard silicon probes (Olympus AC160TS, cantilever spring constant ∼30 N m−1 , tip radius ∼10 nm). The measurements were obtained in ambient conditions at 50 ± 8% relative humidity and a temperature of 22 ± 1 ◦C. Topography images of three independent positions were recorded for each sample. All the data was processed in the open-source software Gwyddion (Necas and Klapetek, 2012). For the 5µm × 5µm images, a roughness analysis (Teichert, 2002) was performed by calculating the 1D heightheight correlation function:

$$C\left(\mathbf{x}\right) = \left\langle \left[ z\left(\mathbf{x}\_0 + \mathbf{x}\right) - \left< z \right> \right] \left[ z\left(\mathbf{x}\_0 - \left< z \right>\right) \right] \right\rangle \tag{2}$$

of each scan line and then averaging over all lines. The resulting values were fitted with the function:

$$C(\mathfrak{x}) = \sigma^2 \, e^{-\left(\frac{|\mathfrak{x}|}{\mathfrak{f}}\right)^{2\alpha}} \,\tag{3}$$

TABLE 1 | RMS roughness and lateral correlation length of the roughness of the blend films prior (TMSC:HPCE) and after (cellulose:HPCE) regeneration.

The data is obtained from analysis of the 5µm × 5µm topography images.

The parameters σ, ξ, and α are used to characterize the surface roughness (Teichert, 2002). The σ denotes the root mean square (RMS) roughness, i.e., the standard deviation of the height values, which is a common measure for the vertical roughness. The lateral correlation length ξ describes the lateral fluctuation of the height values and α is the so-called Hurst parameter or roughness exponent. It determines the shape of C(x) and quantifies the jaggedness of the surface.

For FFM, which is recorded in contact mode, NT-MDT CSG10/Au probes with a tip radius of about 30 nm and a low cantilever spring constant of 0.1 N m−1 were employed. Images with frame size of 5µm × 5µm were obtained with a constant scan speed of 2.5µm s−1 . A vertical force of about 10 nN was applied during the measurements. For acquisition of an FFM image, standard contact mode AFM scan including the lateral trace and retrace channel were recorded. The raw lateral signals were converted to friction images by subtracting the lateral retrace from the lateral trace signal and dividing it by two for each image (Kalihari et al., 2010; Shen et al., 2014). This eliminates topography artifacts and a possible offset.

The quantitative interpretation in terms of friction coefficient is not straightforward, and literature includes several calibration methods (Klapetek, 2018). Lateral force sensitivity calibration was done here according to the wedge calibration method of Varenberg et al. (2003). Quantitative friction images were obtained by multiplying the resulting friction image data with the lateral force sensitivity using the Gwyddion software.

AFM force spectroscopy measurements to investigate the adhesion properties of the film surfaces were performed with a scan rate of 2 Hz and a force distance of 0.5µm. For these measurements, HSC60 probes from Team Nanotec (Germany) were used which have a cantilever spring constant of about 50 N/m and a tip radius of 60 nm. Here, 32 × 32 px² maps were obtained on 5µm × 5µm topography scans.

A Veeco Multimode Quadrax MM AFM (Bruker, USA) in tapping mode using standard silicon probes (NCH-VS1-W, NanoWorld AG) was used for recording film topography after they were rinsed with chloroform.

### Surface Morphology and Area Determination

To quantify the observed surface features for the individual blend films, ten individual cross-sections of the features were obtained from the topography images for each film with the Gwyddion software to determine the height and width of the features. The values are given as mean ± standard deviation.

The AFM topography images were used for calculating effective surface fractions of the blend films. These are referred to as "effective surface fraction" or "surface fraction" later in the manuscript. The film component fraction derived from spinning solution is referred to as "volume fraction" or "volume ratio." Masked surfaces were evaluated by the surface area estimate method in Gwyddion, computed by simple triangulation that considers heights and spatial relations in the surface. For this purpose, additional points were added in-between four neighboring points using the mean values of these pixels. Four triangles are formed, and the surface area is approximated by summing their areas. Masking was done by threshold in z-value and adjusting with a pen tool. Single noise pixel was removed by grain filtering function. For films that showed a reliable phase contrast in the tapping mode topography analysis, the masks were determined from the phase information.

### RESULTS AND DISCUSSION

Spin coated TMSC and HPCE films formed into smooth one-component film morphologies—indicating a uniform film formation—while the blend films containing both derivatives resulted in films with spinodal decomposition and are presented via three composition ratios, 3:1, 1:1, and 1:3 (**Figure 2A**). These TMSC:HPCE blend film surfaces consisted of domains which either formed cavities (TMSC:HPCE 3:1) or protrusions in lower

sub-micrometer range. Protrusions with height 35 ± 10 nm and width of 780 ± 130 nm were determined for TMSC:HPCE 1:1 and of height of 55 ± 10 nm and width of 340 ± 50 nm for TMSC:HPCE 1:3. The protrusions increased in height with increasing HPCE content but decreased in lateral size.

The regeneration of TMSC into cellulose is accompanied by the cleavage of the silyl groups and simultaneous formation of hydrogen bonds of the created hydroxyl groups. This led to densification and reduction in film thickness (**Figure S1**), similar to reported elsewhere (Wolfberger et al., 2015), of the regenerated parts and consequently, more complex morphology (**Figure 2B**). HPCE is unaffected by the treatment (compare **Figures 2A,B**). After conversion to cellulose, the blend film cellulose fraction mass was reduced which translated into an inverted morphology compared to the TMSC:HPCE blend films. The cellulose:HPCE 3:1 surface was characterized by protrusions with arbitrary size and a height of 40 ± 10 nm, whereas the cellulose:HPCE 1:1 and 1:3 films contained cavities which decrease in lateral size. The 1:1 sample showed random-shaped surface features with a lateral width of 660 ± 80 nm, but the more spherical-shaped features on the 1:3 blend film surface had a width of only 250 ± 25 nm. An overview of this data is presented in **Table 1**.