Production of Biocompatible Protein Functionalized Cellulose

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Bio-interactions and Biocompatibility

Production of Biocompatible Protein Functionalized Cellulose Membranes by a Top-down Approach Franck Quero, Abraham Quintro, Nicole Orellana, Genesis Opazo, Andreas Mautner, Alonso Jaque, Fabiola Valdebenito, Marcos Flores, and C.A. Acevedo ACS Biomater. Sci. Eng., Just Accepted Manuscript • DOI: 10.1021/acsbiomaterials.9b01015 • Publication Date (Web): 30 Aug 2019 Downloaded from pubs.acs.org on September 3, 2019

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ACS Biomaterials Science & Engineering

Production of Biocompatible Protein Functionalized Cellulose Membranes by a Topdown Approach Franck Quero1,2*, Abraham Quintro1, Nicole Orellana3, Genesis Opazo1, Andreas Mautner4, Alonso Jaque1, Fabiola Valdebenito1, Marcos Flores5 and Cristian Acevedo3,6

1. Laboratorio de Nanocelulosa y Biomateriales, Departamento de Ingeniería Química, Biotecnología y Materiales, Facultad de Ciencias Físicas y Matemáticas, Universidad de Chile, Avenida Beauchef 851, Santiago, Chile.

2. Millennium Nucleus on Smart Soft Mechanical Metamaterials, Avenida Beauchef 851, Santiago, Chile.

3. Centro de Biotecnología "Dr. Daniel Alkalay Lowitt", Universidad Técnica Federico Santa María, Avenida España 1680 Valparaíso, Chile.

4. Polymer and Composite Engineering (PaCE) Group, Institute of Materials Chemistry and Research, Faculty of Chemistry, University of Vienna, Vienna, Austria.

5. Laboratorio de Superficies y Nanomateriales, Departamento de Física, Facultad de Ciencias Físicas y Matemáticas, Universidad de Chile, Avenida Beauchef 850, Santiago,

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Chile. 6. Departamento de Física, Universidad Técnica Federico Santa María, Avenida España 1680 Valparaíso, Chile.

(*)Corresponding

Author. F. Quero, Email: [email protected].

KEYWORDS: Cellulose fibrils; membranes; top-down approach; proteins; tunicate; biocompatibility.

ABSTRACT Protein functionalized cellulose fibrils were isolated from the tunic of Pyura chilensis and subsequently used to produce protein functionalized cellulose membranes. Bleached cellulose membranes were also obtained and used as reference material. FTIR and Raman spectroscopy demonstrated that the membranes are mostly constituted of cellulose along with the presence of residual proteins and pigments. Protein functionalized cellulose membranes were found to possess 3.1% of protein at their surface as measured by X-ray photoelectron spectroscopy.

Powder

X-ray

diffraction,

scanning

electron

microscopy

and

thermogravimetric analysis were used to identify and semi-quantify the amount of residual sand grains present within the structure of the membranes. The presence of residual proteins was found not to significantly affect the tensile mechanical properties of the membranes. Streaming ζ-potential was used to assess surface charges of the membranes. Below pH 4, non-bleached cellulose membranes possessed highly negative surfaces charges, and also significantly less negative surface charges at physiological pH when compared to bleached


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cellulose membranes. No significant difference was found with respect to growth kinetics of myoblasts at the surface of the membranes for cell culturing times of 48 and 72 h. After 48 h of culture, protein functionalized cellulose-based membranes that possess 3.1% of proteins at their surface (H1) were, however, found to promote higher cell density, cell spreading and more orientated shape cell morphology when compared to the other cellulose-based membranes (H3 and B) evaluated in the present study.

INTRODUCTION In recent years, natural materials including chitin, chitosan, starch, cellulose and gelatin have shown great promises for their use in regenerative medicine and tissue engineering applications.1-2 Some of the advantages of natural materials over most synthetic materials include their renewable origin, biodegradability, non-toxicity and biocompatibility to name only a few.3-4 Nanocellulose is a relatively new class of nanomaterial. It is typically obtained from plant cellulose using top-down strategies through various physical, chemical and biological methods.5-8 Nanocellulose has shown great potential for use in biomedicine owing to its biocompatibility, either in the form of cellulose nanofibers (CNFs) or cellulose nanocrystals.3,9 Bacterial cellulose is another class of nanocellulose that can be obtained from bacteria under specific conditions.10 This method for obtaining nanocellulose is referred to as bottom-up approach. Bacterial cellulose has been intensively investigated for regenerative medicine and tissue engineering applications and has shown great potential owing to its high purity compared to nanocellulose obtained from other sources.3,11 Sea animals referred to as tunicates are another source of cellulose.12 Tunicate cellulose has been evaluated for tissue engineering applications, mostly in the form of cellulose nanocrystals.13-14 These were found



to be particularly relevant to direct the morphology of muscle cells14 and promote skeletal muscle myogenesis.13 Nanocellulose can be used as a building block to fabricate cellulose-based structure assemblies,15 including flat and dense cellulose-based membranes. Up to date, cellulose nanopaper membranes have been mostly evaluated for filtration and separation applications, including the removal of virus, copper and nitrates from aqueous media, to name a few examples.16-18 Nanocellulose has also been evaluated for biomedical and electrobiocompatible device application including wearable bioelectronics19-20, biosensors21-22 and neural interfaces,23 where biocompatibility is needed.24-25 Dense films and membranes, which can be considered as two dimensional (2D) structures, are also relevant materials for tissue engineering applications.26 When developing scaffold materials to be used as substrates for cell cultures, it is advisable to work initially with 2D structures to study single variables.26 2D structures are also relevant for skin tissue engineering and are useful materials to complement other scaffold structures used for musculoskeletal repair or cardiovascular applications.27-28 Up to date, cellulose nanocrystals have been mostly investigated to fabricate dense cellulose-based membranes for tissue engineering applications26 and literature in this domain utilizing cellulose fibrils or nanofibers is scarce. One of the drawbacks of nanocellulose, however, is that its surface lacks bioactivity.3, 24 One strategy that has been proposed to render cellulose-based membranes bioactive and to provide more chemical functionality to cellulose (including -COOH and NH2 moieties) is to link proteins, peptides or amino acids to its surface.29-36 The cellulose source that has been mostly used so far in this regard is bacterial cellulose, where the modification of its surface by protein, peptides and amino acids was found to provide significant benefits for cell cultures,29-30,32-34 including the modification of cell morphology.37 In these works, linking


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these substances to the surface of cellulose-based membranes was achieved by several mechanisms including covalent attachment, biochemical affinity or physical adsorption.3134,36-40

The method that potentially offers the most stable level of protein attachment to the

cellulose surface is covalent attachment. This method, however, is not straightforward and can be particularly tedious.38-39 Also, it usually involves the use of toxic chemical substances,38 which represent a problem from an environmental point of view. An alternative strategy that we recently proposed is to produce cellulose-based membranes with proteins covalently attached to their surface is to use cellulose fibrils that naturally possess covalently attached proteins to their surface.41 This has been achieved by controlled chemical isolation of cellulose fibrils from a tunicate (Pyura chilensis) since this cellulose source is naturally associated to relatively high amounts of proteins.41-45 We referred this strategy to as top-down approach.41 In the present work, protein functionalized cellulose fibrils were used to produce flat and dense 2D cellulose-based membranes that possess proteins covalently attached to their surface. Bleached cellulose membranes were also produced and used as reference material. As a result, the purpose of the present investigation is to study the effect of denatured residual proteins on the biocompatibility of cellulose-based membranes, which could be potentially used for high value biomedical application. The presence of denatured proteins at the surface of these cellulose-based membranes could potentially offer bioactivity and additional chemical functionality, which could serve for further chemical surface modification. The cellulose fibrils used to fabricate these membranes are derived from a side stream of local sea food industry.



MATERIALS AND METHODS Materials and reagents. All chemicals, including potassium hydroxide, sodium hypochlorite and acetic acid, were of analytical grade and purchased either from Merck (Germany) or Sigma Aldrich (USA). Paper filters discs having a grammage of 84 g/m2 and retention of 12-15 μm were purchased from Munktell-Ahlstrom (Sweden) and used for filtration and for the fabrication of the cellulose-based membranes. Pyura chilensis tunicates were purchased from a local fish market (Terminal Pesquero Metropolitano de Santiago de Chile).

Chemical isolation of protein functionalized cellulose fibrils. Cellulose fibrils were extracted using our previously reported method.41 Briefly, tunics obtained from Pyura Chilensis tunicates were washed three times using aqueous KOH, at a concentration of 5 % w/v for 24 h. The tunics were subsequently dried using a freeze dryer (Christ Alpha 1-2 LDplus, Germany) and cryogenically grounded using liquid nitrogen and a knife mill (Retsch Grindomix GM 2000, Germany). At the end of the grinding process, the tunic powder was obtained, which was sieved and a powder with a particle size < 208 μm was obtained. The amino acid profile of the tunic powder was determined by reverse-phase high performance liquid chromatography as reported before in the literature.46 The corresponding results are reported in Table S1. The tunic powder was first submitted to one day of alkaline hydrolysis. The resulting cellulose-based materials is referred to as H1. In a separate experiment, the tunic powder was


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submitted to a total of three days of alkaline hydrolyses (H3). All hydrolyses were performed using a 5 wt.% KOH aqueous solution at 85 °C for 24 h using a reflux column. H1 and H3 cellulose-based materials were subsequently washed with acetone, distilled water and finally freeze dried for 48 h (Christ Alpha 1-2 LDplus, Germany). A fraction of the H3 material was then submitted to a final bleaching step. This was performed for 24 h at a temperature of 80 °C using a 100 mL solution containing sodium hypochlorite (>4% chlorine, ~670 µL) and acetic acid (~330 µL). The bleached cellulose material, named B, was then washed using acetone and distilled water and finally freeze dried for 48 h. The H1, H3 and B cellulose materials were stored in hermetic plastic bags at ambient temperature in dry and dark condition until further use.

Fabrication of cellulose-based membranes. ~1.5 g of H1, H3 or B, respectively, were added individually into 150 mL glass beakers, containing 100 mL of distilled water and allowed to soak for 24 h. The mixture was then processed using a kitchen blender (Thomas Premium TH-850D, Germany) and submitted to the following processing conditions. The obtained suspension was processed using a high shear homogenizer (T18 Digital Ultraturrax IKA, Germany) for 5 min at 10 000 rpm. Three separate suspensions containing ~1.5 wt.% of H1, H3 and B, respectively, were obtained. To fabricate H1, H3 and B cellulose-based membranes, first the ~1 wt.% suspensions were diluted to 0.5 wt.% by adding distilled water. 40 mL of the suspension was first vacuum filtered for ~30 min using a Büchner funnel having a disc diameter of 45 mm connected to a vacuum pump (Rocker 400, Taiwan). A wet filter cake was obtained at the end of the vacuum filtration step, which was subsequently inserted in between a series of layers; first in between two very fine mesh metallic grids, then two filter papers and finally two metallic plates. The



sandwich was pressed at a pressure of 12 MPa and then dried under load at 85 °C for 30 min using a heat press (HP3802, Xinhong, China) to remove residual water. At the end of the process, flat and dense cellulose-based membranes were obtained (H1, H3 and B) having thicknesses within the range of 90-110 μm and a diameter of ~54 mm. The B membrane was used as reference material in all subsequent characterizations.

Characterization of cellulose-based membranes. Molecular structure analysis by ATR-FTIR spectroscopy The molecular composition at the surface of H1, H3 and B cellulose-based membranes was investigated by attenuated total reflectance (ATR)-Fourier-transform infrared spectroscopy (FTIR). The spectra were obtained using a Thermo Scientific FTIR spectrometer (Nicolet iS10, USA) and the spectra were processed using software OMNIC version 9.8.286. The spectra were acquired using a resolution of 4 cm-1 and 32 scans in the wavenumber range of 400–4000 cm-1. Spectra were baseline corrected and normalized. The surface of each sample was analyzed in duplicate by obtaining spectra from two different locations.

Molecular structure analysis by Raman spectroscopy Raman spectroscopy was used to study the molecular composition at the surface of H1, H3 and B cellulose membranes. Spectra were recorded from 500 to 1600 cm-1 with a spectral resolution of 1 cm-1 using a Witec Raman spectrometer (Alpha 300, Ulm, Germany) equipped with a green laser having a wavelength of 532 nm. Each spectrum was obtained using 10 accumulations and an exposure time of 30 s. A grating of 1800 g/mm was used to improve


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signal to noise ratio. Three spectra were acquired at different surface location for each sample, which were found to be equal for H1, H3 and B cellulose-based membranes.

Surface protein quantification by X-ray photoelectron spectroscopy X-ray photoelectron spectroscopy (XPS) was used to probe the surface elemental composition of H1, H3 and B cellulose-based membranes. More particularly, the amount of nitrogen atoms available at the surface was quantified to derive the amount of protein available at the surface of the membranes. Initially, full range low-resolution XPS spectra were acquired using a hemispheric analyzer (Physical Electronics 1257 System, USA). The system was equipped with a twin Al K anode, emitting an X-ray Al Kα radiation (1486.6 eV) having a constant power of 400 W. An emission angle of 90° was used to reach electrons that belong to deeper energy levels (down to ~4-5 nm below the sample’s surface). Experiments were performed under ultra-high vacuum, at a pressure of ~10-6 Pa and spectra were acquired within the binding energy range of 1200 – 0 eV using steps of 44.75 eV and 50 scans. Highresolution XPS spectra were obtained using a spectral resolution of 0.2 eV/step. Spectra for nitrogen were obtained within the range of 408 – 395 eV, using up to 30 scans. Prior to experiments, the adventitious C1s (C-C) carbon signal was used for calibrating the bonding energy. Chemical element quantification was determined by fitting all spectra using a Gaussian function. The crude protein content (P) present at the surface of H1, H3 and B cellulose-based membranes was then derived from the concentration values of nitrogen, which was calculated using the equation



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(1)

𝑃 (%) = 𝑁(%) × 𝐾

where N is the concentration of nitrogen in percentage (%) and K corresponds to a conversion factor that relates nitrogen content to protein content. For protein of animal origin, a conversation factor of 6.25 is usually used47 and has been reported before to quantify the amount of protein present in the tunic of various tunicate specie.42

Crystalline structure analysis of cellulose-based membranes by powder X-ray diffraction The crystalline structure of H1, H3 and B cellulose-based membranes was investigated by powder X-ray diffraction (XRD). The samples were analyzed using a D8 Advance powder X-ray diffractometer (Bruker, UK), equipped with a copper X-ray source producing a CuKα radiation having a wavelength of 0.154 nm operated at 40 kV and 30 mA. The patterns were obtained within the diffraction angle range of 5º to 40º at an angle step of 0.02º per 3 s and using a rotational speed of 60 rpm, which were baseline corrected using the software Origin 8 SR0 V8.0724 (BT24, USA). The samples were analyzed in duplicate and typical XRD patterns are reported. The crystallinity index (χc) for H1, H3 and B cellulose-based membranes was estimated from these patterns by using an Integration Method48 and the equation

𝐴𝐶

(2)

𝜒𝑐 = 𝐴𝐶 + 𝐴𝐴 × 100


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where Ac and Aa correspond to the area under the X-ray diffraction pattern generated from the crystalline and amorphous regions respectively. The crystallite size for H1, H3 and B cellulose-based membranes was estimated using Scherrer’s equation49

𝐾𝜆

(3)

𝐿002 = 𝛽002cos 𝜃

where L002 is the crystallite size of the 002 reflection, K = 0.91 and β002 is the full width at half maximum of the 002 reflection.

Surface morphology of cellulose-based membranes by scanning electron microscopy The surface of H1, H3 and B cellulose-based membranes was observed using a field emission high-resolution scanning electron microscope (INSPECT-F50, FEI, Holland). Images were obtained at an acceleration voltage of 10 kV, spot size of 3 and a working distance of 6 mm. Prior to imaging, the samples were fixed onto metal stubs using double sided adhesive carbon tabs and subsequently gold-coated using a sputtering coater (Cressington 208HR, UK). The thickness of the gold coating was 5 nm. The width size distributions of the fibrils constituting the H1, H3 and B cellulose-based membranes were obtained using the Image J software version 1.45 (National Institutes of Health, USA). At least 150 measurements were performed on each SEM images having a magnification of 40 000 to obtain the respective width size distributions. The data are presented as frequency (%) as a function of the fibril width (nm).



Tensile mechanical properties and thermal stability of cellulose-based membranes The tensile mechanical properties of H1, H3 and B membranes were determined using a mechanical tensile tester (WDW-S5, SX, China). Membranes were cut into strips having width and length of 1 and 30 mm, respectively. Prior to measurements, the strips were secured onto window cards by fixing using two-part epoxy glue. A force range of 0-50 N was selected for the load cell. The gauge length and the displacement speed of the mobile clamp were set to 20 mm and 50 mm min-1, respectively. Experiments were performed at a temperature of ~22 °C. A total of 6 samples were tested for H1, H3 and B cellulose-based membranes. Average values are reported along with their associated standard deviations, used as error values. The thermal stability as well as the relative evaluation (or semi-quantification) of the amount of residual sand grains in the membranes H1, H3 and B cellulose-based membranes was determined using a thermogravimetric analyzer TGA Q50 (TA Instruments, USA). Samples having with an initial mass of ~8 mg were submitted to a temperature ramp of 5 °C min-1 from 30 up to 600 °C under a nitrogen atmosphere at a flow rate of 40 mL min-1. The onset and peak degradation temperatures were determined from the first derivative of the weight loss as a function of temperature using the software Universal Analysis 2000 (TA Instruments, USA). The measurements were performed in triplicate and average values are reported along with their associated standard deviations, used as error values.

Surface charge assessment of cellulose-based membranes by ζ-potential The surface charges present at the surface of H1, H3 and B cellulose-based membranes were determined by performing streaming ζ-potential measurements as a function of pH. A Surpass electrokinetic analyzer from Anton Paar (Graz, Austria) fitted with an adjustable gap


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cell (gap width 100 μm) was used to perform the measurements. The electrolyte solution (1 mM KCl) was pumped through the cell while steadily increasing the pressure up to 300 mbar. The pH was set by adding controlled amounts of KOH and HCl solutions (0.05 mol L-1) into the electrolyte solution. The calculation of the ζ-potential was performed with the streaming current. The measurements were performed in triplicate and average values are reported along with their associated standard deviations, used as error values.

Myoblasts proliferation and morphology at the surface of cellulose-based membranes The myoblast cell line C2C12 was used as model of mesenchymal cells. The cell line was purchased from the European Collection of Cell Cultures (ECACC) and supplied by SigmaAldrich (USA). The cells were cultured using standard conditions for cell culture (37 °C and 5% CO2 in a humidified atmosphere) in a CO2 incubator (Heracell Vios 160i, Thermo Scientific, Germany). Cells were seeded onto the films at 1x104 cells/cm2. The medium used to culture the cells was DMEM high glucose (Gibco, USA), supplemented with 10% fetal bovine serum (Biologicals Industries, Israel), L-glutamine (2 mM), and antibiotics (100 U/mL of penicillin and 100 μg/mL of streptomycin). Cell morphology, adhesion and growth were determined such as described before in the literature.50 The degree of cell adhesion and growth was quantified by estimating the viable biomass at different times by using the commercial colorimetric assay WST-1 (Roche, Germany). Cell adhesion was assessed after 4 h of incubation and compared against control (cells adhered to commercial plastic for cell culture). For cell growth, the membranes were sampled at 24, 48, and 72 h; then the data were fitted using the classic exponential model to obtain the doubling time. Experiments were performed in triplicate and average values are reported along with their associated standard deviations used as error values. Cell morphology was observed after



48 h of their seeding onto the surface of H1, H3 and B cellulose-based membranes using standard fluorescence microscopy techniques. Glass coverslip for cell culture was used as surface control. Rhodamine-phalloidin (Thermo Fisher Scientific, USA) was used to stain polymerized actin and Hoechst 33342 (Thermo Fisher Scientific, USA) for nuclear staining. The cells were observed using an inverted fluorescence microscope (Nikon, Eclipse TS2FL, Japan). RESULTS AND DISCUSSION Molecular structure characterization of cellulose-based membranes. Figure 1a shows typical FTIR spectra obtained from the surface of H1, H3 and B cellulosebased membranes within the wavenumber range of 450-4000 cm-1. In all spectra, six typical bands that correspond to the vibrational motions of chemical moieties belonging to the molecular structure of cellulose can be observed. These bands are located at wavenumber positions of 1033, 1055, 1110, 1160, 2907 and 3333 cm-1 and have been reported to occur due to the vibrational motions of COC, COC, COH, CO, CH and OH moieties5152,

respectively and as reported in Table S2. Interestingly, the relative intensity of the bands

located at wavenumber positions of 1055, 1110 and 1160 cm-1 increases upon using cellulose fibrils submitted to one (H1), three (H3) alkaline treatments and finally a bleaching (B) treatment in order to fabricate these cellulose-based membranes. This is because the cellulose fibrils used to make these membranes is getting purer upon applying more chemical treatments as reported before in our previous work.41 For H1, H3 and B cellulose-based membranes, a low intensity band that do not belong to cellulose can be seen at a wavenumber position of ~1651 cm-1. This band is due to the vibrational motions of CO and NH moieties (amide I vibrational mode)42,53 that corresponds to the presence of proteins at the surface of


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the cellulose-based membranes. This is indicative of the presence of amine and carboxylic acid moieties present at the surface of the cellulose-based membranes. This result agrees with the amino acid profile of the tunic powder reported in Table S1. Due to the low intensity of that band it was, however, not possible to obtain semi-quantitative information about the residual protein content of each membrane. FTIR spectroscopy results suggest that H1, H3 and B membranes mainly contain cellulose and residual amounts of proteins, which is likely to decrease upon applying successive alkaline and bleaching chemical treatments. Figure 1b reports Raman spectra obtained from the surface of H1, H3 and B cellulosebased membranes within the Raman shift range of 850-1550 cm-1. The four main Raman bands located at Raman shift positions of ~1097, 1122, 1152 and 1509 cm-1 were related to the vibrational motions of chemical moieties that belong to the molecular structure of cellulose.52,54 The detailed chemical assignment of these bands is reported in Table S1. The relative intensity of the bands located at ~1097 and 1122 cm-1 is found to increase upon using cellulose fibrils submitted to one (H1) or three (H3) alkaline treatments and finally a bleaching (B) treatment. These bands are due to the vibrational motions of CO moieties that belong to the molecular structure of cellulose.54 This suggests that the increasing order of cellulose purity of these membranes is H1 < H3 < B. The intensity of the bands located at ~1152 and 1509 cm-1 is, however, found to decrease. These bands have been reported to occur due to the vibrational motions of heavy atoms that belong to the molecular structure of carotenoid pigments55-57, that may be responsible for the orangish color of the tunic of Pyura chilensis. In our previous study, the relative intensity decrease of these two bands has been associated to the loss of organic pigments due the successive alkaline and bleaching chemical treatments applied to Pyura chilensis tunics.41 These pigments have been reported to be



present in other tunicates and to be a type of carotenoid.58 For the membrane constituted of bleached cellulose fibrils, the band located at a Raman shift of ~1509 cm-1 completely vanished, which suggests that this membrane is pigment free as confirmed by its white color. As previously suggested by FTIR spectroscopy, Raman spectroscopy results confirm that H1, H3 and B membranes mainly contain cellulose. In addition to residual proteins (as identified by FTIR), however, H1 and H3 cellulose-based membranes possessed pigments, a type of carotenoid, at their surface. Both techniques, however, did not provide quantitative information on the amount of residual protein present at the surface of the cellulose-based membranes.

Figure 1. Typical (a) Fourier-transform infrared and (b) Raman spectra obtained for H1, H3 and B cellulose-based membranes.

Atomic surface composition of cellulose-based membranes. X-ray photoelectron spectroscopy (XPS) was used to determine the atomic composition at the surface of H1, H3 and B cellulose-based membranes. Figure S1 reports low resolution XPS spectra for H1, H3 and B cellulose-based membranes. From these spectra, one can


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observe two main peaks located at binding energy positions of ~534 and 287 eV that correspond to the binding energy of the O1s and C1s energy states, respectively. This observation agrees with the fact that H1, H3 and B membranes are essentially constituted of cellulose that primarily possesses oxygen and carbon atoms in its molecular structure. For H1 and H3 membranes, two low intensity peaks located at binding energy positions of ~401 and 100 eV are also present in those spectra. These binding energies correspond to the N1s and Si2p energy states, respectively. The signal for nitrogen atoms originates from the existence of proteins (constituted of the amino acids reported in Table S1) that naturally possess nitrogen atoms in their atomic composition. On the other hand, the presence of Si may originate from the presence of residual sand grains. The presence of C, O, N and Si present at the surface of H1, H3 and B cellulose-based membranes was then quantified by recording high-resolution XPS spectra. The surface elemental composition in C, O, N and Si is reported in Table 1. As expected, the amount of C present at the surface of the membranes was found to increase whereas the concentration of N, O and Si (associated to the gradual removal of sand grains) was found to decrease upon using cellulose fibrils submitted to one (H1) or three (H3) alkaline treatments, respectively, and finally a bleaching (B) treatment to fabricate these cellulose-based membranes. For nitrogen atoms, it was actually not possible to quantify their amount at the surface of H3 and B cellulose-based membranes due to the weak signal. As a result, it was not possible to quantify the amount of protein present at the surface of H3 and B membranes. This was, however, calculated for H1 membranes using Equation 3 and the amount of residual proteins was found to be ~3.1%. This value is close but lower than our previous value of ~3.8% reported before for H1 cellulose fibrils.41 The presence of a relatively high amount of N atoms at the surface of H1 cellulose-based membranes (0.5%) suggests the presence of amine moieties -NH2. The higher amount of O



quantified for H1 cellulose-based membranes (47.4%) compared to H3 (39.4%) and B (40.9%) cellulose-based membranes may also indicate the presence of carboxylic moieties. The increase in O content at the surface of B membranes compared to H3 membranes may originate from the bleaching treatment, which is known to introduce carboxylic acid moieties at the surface of cellulose by oxidation.59 Due to the gradual removal of amino acids, it is likely that the number of carboxylic moieties is lower for H3 cellulose-based membranes and even lower for B cellulose-based membranes.

Table 1. Surface elemental composition for carbon (C), oxygen (O), nitrogen (N) and silicon (Si) obtained from high resolution X-ray photoelectron spectra for H1, H3 and B cellulosebased membranes. The calculated protein content is also reported.

Surface elemental composition (%) Sample H1 H3 B

C 50.5 59.8 58.6

N 0.5 -

O 47.4 39.4 40.9

Si 1.7 0.5 0.3

Protein content (%) 3.1 -

Crystalline structure analysis. Figure 2 reports typical powder X-ray diffraction patterns for H1, H3 and B cellulose-based membranes. Four diffraction peaks located at diffraction angle 2θ positions of ~14.7°, 16.6°, 22.5° and 34.5° corresponding to the diffraction planes (110), (110), (200) and (040), respectively, are present. These diffraction planes are typical for cellulose I and have been reported before for cellulose obtained from tunicates.42 One can observe that the intensity of these reflections gradually increases upon using a purer source of cellulose to fabricate these


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membranes, following the increasing order of cellulose purity of these membranes is H1 < H2 < B, which support FTIR and Raman spectra previously reported in the present study. From Figure 2, one can also observe three reflections in the diffraction angle 2θ range of 25-30°. These possibly correspond to the presence of residual sand grains that are fully or partly crystalline. The intensity of these reflections was found to decrease, also following the order B > H3 > H1, which suggests that sand grains were gradually eliminated when more alkaline and bleaching chemical treatments were applied to the tunics. It is important to note that no diffraction peak was observed in the diffraction angle range of 8-9. This suggests that the residual protein available at the surface of the membranes (especially H1) do not possess a collagen-like or renatured protein structure. This might suggest that the proteins that are available at the surface of H1 and H3 cellulose-based membranes present a denatured or unfolded structure. This agrees with the alkaline extraction conditions (5% KOH, 85 C for 24 h) that were used to gradually remove proteins. The use of a basic pH is known to destabilize the folded structure of proteins by deprotonation of the acidic groups, including carboxylic acids moieties (COOH to COO-), that belong to the molecular structure of amino acid residues responsible for the stabilization of the folded structure of proteins by hydrogen bonding. In addition, in the present study, no special conditions were used to try to renature or fold back the structure of the residual proteins. As a result, it is very likely that the residual proteins that are present at the surface of H1 and H3 membranes are denatured and possess an unfolded structure.



Figure 2. Typical powder X-ray diffraction patterns obtained for H1, H3 and B cellulosebased membranes.


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Table 2 reports values of crystallinity index and crystallite size for H1, H3 and B cellulosebased membranes. The crystallinity index of these membranes was found to increase (from 70 to 87%), following the order H1 < H3 < B. These values are, however, higher than values reported in our previous work41, which may originate from the fact that a different batch of tunic was used. This confirms that the cellulose fibrils present in B membranes are purer that the cellulose fibrils constituting H3 and H1 cellulose-based membranes. This result agrees with FTIR and Raman spectroscopy data discussed earlier. As reported in Table 2, crystallite size values obtained for H1, H3 and B cellulose-based membranes were found to increase (from 81.7 to 92.5 Å) when using purer cellulose fibrils. This suggests that alkaline and bleaching chemical treatment remove non-cellulosic components, gradually revealing the crystalline fraction of H1, H3 and B cellulose fibrils utilized to fabricate these membranes.

Table 2. Crystallinity index and crystallite size obtained for H1, H3 and B cellulose-based membranes.

Cellulose-based membrane Crystallinity index (%) Crystallite size (Å) H1 70 81.7 H3 74 83.9 B 87 92.5

Surface morphology observation and width size distribution. Figure 3 shows scanning electron microscopy images obtained from the surface of H1, H3 and B cellulose-based membranes. All membrane surfaces are constituted of randomly oriented cellulose fibrils having inhomogeneous width size. The width size distribution for the cellulose fibrils that formed H1, H3 and B cellulose-based membranes are reported in



Figure S2. H1, H3 and B membranes are mostly constituted of cellulose fibrils having a width size in the range of 20-50 nm, with some cellulose fibrils having width sizes ranging from 50 to 300 nm. These images depict the presence of pores, irregular in size and shape and formed between cellulose fibrils during the membrane fabrication.

Figure 3. Scanning electron microscopy images obtained from the surface of (a) H1, (b) H3 and (c) B cellulose-based membranes (magnification ×10 000). The white arrows indicate sand grains.


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Residual sand grains, as shown by the white arrows in Figure 3a, are also present on the surface of the H1 cellulose membrane. On the surface of H3 (Figure 3b), there are also a few sand grains but less compared to the surface of H1. Opposed to that, on the surface of B (Figure 3c), no sand grains were found. These microscopic observations are in agreement with the results obtained by powder X-ray diffraction, where the intensity of the diffractions peaks corresponding to sand grains decreased where purer cellulose fibrils are used to fabricate cellulose-based membranes (from H1 to B).

Tensile mechanical properties and thermal stability. Figure S3 reports typical tensile stress-strain curves for H1, H3 and B cellulose-based membranes. The detailed tensile mechanical properties data, including Young’s modulus, stress and strain at failure are reported in Table S3. From these data, one can observe that H1, H3 and B cellulose-based membranes possess similar mechanical properties, with no significant difference between them with respect to Young’s modulus, stress and strain at failure. Figures 4a and 4b report TGA thermograms and DTGA curves for H1, H3 and B cellulosebased membranes in the temperature range of 30-600 °C, respectively. TGA analysis was performed to evaluate the thermal stability of the H1, H3 and B cellulose-based membranes but also to semi-quantify the amount of residual sand grains that were previously identified by SEM imaging. It was found that TGA curves shifted towards higher temperatures (from H1 to B cellulose-based membranes). This is also supported by onset and peak degradation temperature values reported in Table 3, suggesting that the thermal stability of these cellulose-based membranes significantly improves upon gradual removal of both proteins and pigments by applying successive alkaline and chemical bleaching treatments. Table 3



also reports the residual weight value for H1, H3 and B cellulose-based membranes. For H1, H3 and B cellulose-based membranes, weight residue values of 19  3, 13  5 and 4  4 % were obtained, respectively. This suggests that the sand grain content of the cellulose-based membranes (from H1 to B) was significantly lower when longer and more intense chemical treatment was performed. Figure 4b reports typical DTGA curves for H1, H3 and B cellulose-based membranes. For H1 and H3 membranes the presence of a shoulder at a temperature of ~250 °C was observed (see black arrow). This might be explained by the thermal degradation of non-cellulosic substances such as proteins and lipids. Similar shoulders have been reported for cellulose obtained from macroalgae.60 From 250 to 380 °C, the main peak that corresponds to the thermal degradation of cellulose occurred. The maximum of this peak shifts significantly towards higher temperature upon using cellulose fibrils submitted to longer and more intense chemical treatments.

Figure 4. Typical (a) thermogravimetric analysis (TGA) and (b) DTGA traces obtained for H1, H3, and B cellulose-based membranes.


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Table 3. Detailed thermal stability data obtained for H1, H3 and B cellulose-based membranes. Tonset and Tpeak, correspond to the onset degradation temperature and to the peak degradation temperature, respectively.

Cellulose-based membrane Tonset (°C) Tpeak (°C) Weight residue at 600 °C (%) H1 282 ± 1 318 ± 3 19 ± 3 H3 297 ± 1 336 ± 1 13 ± 5 B 313 ± 3 348 ± 10 4±4

Surface charge assessment. Figure 5 reports streaming ζ-potential measurements as a function of pH for H1, H3 and B cellulose-based membranes. The trend of the streaming ζ-potential behavior of B, which is almost pure cellulose is similar to the behavior of bacterial cellulose TEMPO-modified CNF and phosphorylated nanocellulose membranes with a plateau at high pH and the isoelectric point (iep) at pH 3.16,61-62 Above pH 3, the surface charge was negative, reaching a maximum value of -45 mV at pH = 10. This negatively charged surface was the result of the full deprotonation of OH and also possibly due to the presence of acidic groups (carboxylic acids) present at the surface of B that established during the bleaching treatment. Upon



decrease of the pH, protonation of the acidic groups may occur and as a result, the surface charges became significantly less negative. For example, at physiological pH of 7.4, the ζpotential was -39 mV. This increase in surface charge occurred until reaching pH 3, where charge neutrality was reached (isoelectric point) as shown in Figure 6. Below pH 3, the surface charge at the membrane surface became positive, reaching a maximum ζ-potential of 7 mV due to the full protonation of OH and residual acidic groups including amine and carboxylic acid moieties. For the H1 and H3 membranes, at pH 10, the ζ-potential values were negative, with a plateau value of about -30 mV. This ζ-potential value is significantly less negative than for the B membrane (-45 mV). This may be due to the higher number of carboxylic groups present at the surface of B membrane as consequence of the bleaching treatment59 as evidenced from surface elemental analysis by XPS in the present study and in our previous report.41 Another contribution might originate from the presence of pigments present at the surface of H1 and H3 membranes as evidenced by Raman spectroscopy. The exact mechanism is, however, unclear. Upon decrease of the pH, the ζ-potential became less negative due to protonation of the acidic groups (-NH2 to NH3+ and COO- to COOH) present at the membrane surface. At physiological pH, the surface charge of both membranes reached a value of -30 mV, which is 9 mV lower than for the B cellulose-based membranes. Upon further pH decrease, the surface charge still became less negative until reaching a value of 12 mV at pH 4. However, below pH 4, the ζ-potential became more negative again, until reaching a value of almost -30 mV at pH 2. This behavior observed for both H1 and H3 membranes differs completely from B membranes. This may probably not originate from the cellulose itself and neither from the presence of residual proteins. As measured by XPS, H1 possessed much higher protein content than H3 (which was not quantifiable) but both H1 and


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H3 membranes possessed similar ζ-potential values below pH 4. If there was an impact of the proteins, a significantly different ζ-potential would have to be found for H1 and H3. Since the ζ-potential was equal within the error of the measurement, no significant impact of the proteins onto this behavior can be confirmed. The results of Raman spectroscopy spectra, however, suggested the presence of a pigment (carotenoid), present at the surface of H1 and H3 cellulose-based membranes. In a previous work, it has been reported that carotenoids possess negative ζ-potential values over the whole pH range between 2 and 10, thus also at acidic pH.63 This suggests that the pigments present at the surface of H1 and H3 cellulosebased membranes might be responsible for the negative surface charge measured at pH < 4, although no direct evidence for this can be reported in the present work.

Figure 5. Streaming ζ-potential as a function of pH as measured by streaming potential measurements for H1, H3 and B cellulose-based membranes.



Myoblasts proliferation evaluation and morphology observation The biocompatibility of H1, H3 and B cellulose-based membranes was evaluated by seeding myoblasts (muscle cells) onto their surface. First, cell proliferation onto the surface of the membranes was evaluated by following their adhesion, doubling time and growth. Table 4 reports values of adhesion and doubling time of myoblasts for H1, H3 and B cellulose-based membranes and for a control commercial plastic. With respect to adhesion, all the membranes were found to show good levels of adhesion, despite their negative charged surface at pH 7.4 as measured by streaming ζ-potential. The B cellulose membranes were found to possess the highest degree of adhesion with a value of 78.1 ± 2.4 % (although lower than the control experiment), which is significantly higher than for the H1 and H3 membranes, with adhesion values of 63.0 ± 2.1 % and 66.7 ± 7.2 %, respectively. This suggests that the B cellulose membranes may better promote adhesion of myoblasts onto their surface due to its higher degree of purity when compared to H1 and H3 cellulose-based membranes, as suggested by FTIR, Raman and powder X-ray diffraction. Figure 6a reports the cell growth kinetics of the myoblasts seeded on the surface of H1, H3 and B membranes. The data corresponding to cell growth on the control substrate are reported in Table S4. The trend of cell growth kinetics curves is similar for all membranes with a slight decrease due to non-adherent cells, followed by an increase in cell growth. After 24 h of cell culture, one can observe that the B cellulose membrane induce a significant increase in cell growth when compared to H1 and H3. No significant difference in cell growth was, however, observed after 48 and 72 h of cell culture when comparing H1, H3 and B cellulosebased membranes. H3 membranes had the shortest doubling time with a value of 18.6 ± 4.2 h, followed by the H1 membranes with a value of 22.0 ± 2.9 h and the B membranes with a


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value of 26.3 ± 3.5 h (Table 4). All these values are at large within the range of 12-24 h, as commonly reported for muscle cells.64 The shorter doubling time of both H1 and H3 membranes might be related to their less negative surface charge compared to B membranes, as measured by streaming ζ-potential at pH 7.4. Figure 6b reports fluorescence images of the morphology of myoblasts after being maintained for 48 h in the culture medium. One can observe that the cells seeded onto H1 membrane spreaded out onto the surface and possess elongated or oriented morphology. Cells present at the surface of H3 membranes appeared agglomerated (no spreading) and did not possess an elongated or oriented morphology, similarly to the control. As for the B membrane, cells were found to possess an elongated or oriented morphology possibly due to fibril morphology, but cell density was much lower than for the H1 and H3 membranes as well as the control. This suggests that the presence of a higher relative amount of residual proteins present at the surface of the membranes (3.1 % for H1), even if denatured, may promote higher muscle cell density, cell spreading as well as an orientated shape cell morphology due to fibrils’ morphology. This might be favorable for the differentiation of proliferated myoblasts into fiber muscle. In a previous study, amino acid modified cellulose surfaces were found to modify the morphology of fibroblast cells depending on the amino acid type that was grafted on the cellulose surface.37 Amino acid type was also found to affect the spreading of cells at the surface of cellulose modified by amino acids.37 Aromatic amino acids were found to favor cell spreading whereas aliphatic amino acids had the opposite effect (rounded shape).37 In the present study, a wide range of amino acid types is present at the surface of the cellulose fibrils as reported in Table S1. In another study, it has been reported that the morphology of C2C12 myoblasts cells depends on the surface type onto where cells



are being cultured. Myoblasts having star-like and fibroblastic morphology have been observed on plastic surfaces65 as well as rounded shapes on Matrigel® matrix.66

Table 4. Cell adhesion and doubling time of myoblasts seeded onto the surface of H1, H3 and B cellulose-based membranes. Commercial plastic for cell culture was used as control.

Cellulose-based membrane H1 H3 B Control

Cell adhesion (%) 63.0 ± 2.1 66.7 ± 7.2 78.1 ± 2.4 100.0 ± 4.6

Doubling time (h) 22.0 ± 2.9 18.6 ± 4.2 26.3 ± 3.5 17.5 ± 3.1

Figure 6. (a) Cell growth kinetics of myoblasts seeded onto the surface of H1, H3 and B cellulose-based membranes. Percentage of growth is expressed relative to the initial adhered cells. (b) Fluorescence microscopy images showing the differentiation of myoblasts that


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adhered and grew at the surface of H1, H3 and B cellulose-based membranes after being maintained during 48 h in the culture medium (glass coverslip for cell culture was used as control).

CONCLUSIONS In this study, protein functionalized cellulose-based membranes were successfully fabricated using protein functionalized cellulose fibrils obtained by a top-down approach. HPLC was utilized to determine the amino acid composition of the tunic powder. FTIR and Raman spectroscopy confirmed that the membranes mainly contained cellulose along with residual proteins and a form of carotenoid pigment. XPS allowed quantifying the amount of protein present at the surface of mildly treated H1 cellulose-based membranes, whereas for membranes treated for prolonged times (H3) and being bleached (B) the protein content could not be accurately quantified. Powder XRD confirmed the H1, H3 and B cellulose-based membranes are constituted of highly crystalline cellulose fibrils. Also, the presence of residual sand grains was identified. This was also supported by scanning electron microscopy, where the presence of sand grains was observed for H1 and H3 membranes. The membranes were constituted of randomly oriented cellulose fibrils that possess inhomogeneous

size

width

and

possess

similar

tensile

mechanical

properties.

Thermogravimetric analysis also confirmed the presence of residual sand grains present within the structure of H1 and H3 cellulose-based membranes. The data suggested that H1 membranes possess the highest relative sand grain content followed by H3 and B membranes. The surface charges of the membranes were measured by streaming ζ-potential. The B membrane followed the typical trend reported before for nanocellulose membranes. H1 and H3 membranes, however, were found to possess highly negative surface charges at pH < 4.



Also, at pH 7.4 (physiological pH), H1 and H3 membranes were found to possess less negative surface charges when compared to the B membranes, possibly due to the presence of a kind of carotenoid pigments present at their surface. Myoblasts were found to adhere and proliferate onto the surface of all cellulose-based membranes. No significant different was reported with respect to cell growth. The surface of H1 membrane, owing to the presence of higher amount of denatured residual protein was, however, found to promote higher cell density and orientated shape cell morphology when compared to H3 and B membranes.

SUPPORTING INFORMATION Detailed information on HPLC, FTIR and Raman band assignment, XPS spectra, size width distribution of cellulose fibrils, control experiments of cell growth kinetics of myoblasts and tensile mechanical properties can be found in the supporting information.

AUTHOR INFORMATION (*)Corresponding

Authors. F. Quero, Email: [email protected].

Notes. The authors declare no competing financial interest. Author Contributions. The manuscript was written through contributions of all authors. All authors have given approval to the final version of the manuscript.

ACKNOWLEDGMENT


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The authors gratefully acknowledge Francisco Melo and Victor Carrasco for providing access to their Raman spectroscopy facility. F.Q. acknowledges financial support from CONICYT/FONDECYT (No. 11160139), from the Program UINICIA VID 2016, GRANT UI-14/2016, and CONICYT/FONDEQUIP, (No. EQM130149). Acknowledgments are also extended to the “Nuclei for Smart Soft Mechanical Metamaterials” grant funded by the Millennium Science Initiative of the Ministry of Economy, Development and Tourism. Funding from University of Vienna for A.M. is also acknowledged.

REFERENCES (1)

Mano, J. F.; Silva, G. A.; Azevedo, H. S.; Malafaya, P. B.; Sousa, R. A.; Silva, S. S.;

Boesel, L. F.; Oliveira, J. M.; Santos, T. C.; Marques, A. P.; Neves, N. M.; Reis, R. L., Natural Origin Biodegradable Systems in Tissue Engineering and Regenerative Medicine: Present Status and Some Moving Trends. Journal of The Royal Society Interface 2007, 4, (17), 999-1030. DOI: 10.1098/rsif.2007.0220. (2)

Malafaya, P. B.; Silva, G. A.; Reis, R. L., Natural–Origin Polymers as Carriers and

Scaffolds for Biomolecules and Cell Delivery in Tissue Engineering Applications. Advanced Drug Delivery Reviews 2007, 59, (4), 207-233. DOI: 10.1016/j.addr.2007.03.012. (3) Lin, N.; Dufresne, A., Nanocellulose in Biomedicine: Current Status and Future Prospect.

European

Polymer

Journal

2014,

59,

302-325.

DOI:

10.1016/j.eurpolymj.2014.07.025. (4) Eichhorn, S. J.; Gandini, A., Materials from Renewable Resources. MRS Bulletin 2010, 35, (03), 187-193. DOI: 10.1557/mrs2010.650.



(5)

Nechyporchuk, O.; Belgacem, M. N.; Bras, J., Production of Cellulose Nanofibrils: A

Review of Recent Advances. Industrial Crops and Products 2016, 93, 2-25. DOI: 10.1016/j.indcrop.2016.02.016. (6)

Kargarzadeh, H.; Ioelovich, M.; Ahmad, I.; Thomas, S.; Dufresne, A., Methods for

Extraction of Nanocellulose from Various Sources. Handbook of Nanocellulose and Cellulose Nanocomposites 2017. DOI: 10.1002/9783527689972. (7)

Mondal, S., Preparation, Properties and Applications of Nanocellulosic Materials.

Carbohydr. Polym. 2017, 163, 301-316. DOI: 10.1016/j.carbpol.2016.12.050. (8)

Dufresne, A., Nanocellulose: A New Ageless Bionanomaterial. Materials Today 2013,

16, (6), 220-227. DOI: 10.1016/j.mattod.2013.06.004. (9)

Jorfi, M.; Foster, E. J., Recent Advances in Nanocellulose for Biomedical Applications.

Journal of Applied Polymer Science 2015, 132, (14), 41719. DOI: 10.1002/app.41719. (10) Brown, A. J., On An Acetic Ferment Which Forms Cellulose. Journal of the Chemical Society 1886, 49, 432-439. DOI: 10.1039/CT8864900432. (11) Gama, M.; Gatenholm, P.; Klemm, D., Bacterial NanoCellulose: A Sophisticated Multifunctional Material. CRC Press: 2016. DOI: 10.1201/b12936. (12) Rånby, B. G., Physico-Chemical Investigations on Animal Cellulose (Tunicin). Arkiv Kemi 1952, 4, 241-248. DOI: 20000736493. (13) Dugan, J. M.; Collins, R. F.; Gough, J. E.; Eichhorn, S. J., Oriented Surfaces of Adsorbed Cellulose Nanowhiskers Promote Skeletal Muscle Myogenesis. Acta Biomaterialia 2013, 9, (1), 4707-4715. DOI: 10.1016/j.actbio.2012.08.050. (14) Dugan, J. M.; Gough, J. E.; Eichhorn, S. J., Directing the Morphology and Differentiation of Skeletal Muscle Cells Using Oriented Cellulose Nanowhiskers. Biomacromolecules 2010, 11, (9), 2498-2504. DOI: 10.1021/bm100684k.


Page 34 of 43

Page 35 of 43 1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60


(15) De France, K. J.; Hoare, T.; Cranston, E. D., Review of Hydrogels and Aerogels Containing Nanocellulose. Chemistry of Materials 2017, 29, (11), 4609-4631. DOI: 10.1021/acs.chemmater.7b00531. (16) Mautner, A.; Maples, H. A.; Kobkeatthawin, T.; Kokol, V.; Karim, Z.; Li, K.; Bismarck, A., Phosphorylated Nanocellulose Papers for Copper Adsorption from Aqueous Solutions. Int. J. Environ. Sci. and Technol. 2016, 13, (8), 1861-1872. DOI: 10.1007/s13762016-1026-z. (17) Mautner, A.; Kobkeatthawin, T.; Bismarck, A., Efficient Continuous Removal of Nitrates from Water with Cationic Cellulose Nanopaper Membranes. Resource-Efficient Technologies 2017, 3, (1), 22-28. DOI: 10.1016/j.reffit.2017.01.005. (18) Metreveli, G.; Wågberg, L.; Emmoth, E.; Belák, S.; Strømme, M.; Mihranyan, A., A Size-Exclusion Nanocellulose Filter Paper for Virus Removal. Adv. Healthcare Mater. 2014, 3, (10), 1546-1550. DOI: 10.1002/adhm.201300641. (19) Khan, S.; Ul-Islam, M.; Ullah, M. W.; Israr, M.; Jang, J. H.; Park, J. K., Nano-Gold Assisted Highly Conducting and Biocompatible Bacterial Cellulose-PEDOT:PSS Films for Biology-Device Interface Applications. International Journal of Biological Macromolecules 2018, 107, 865-873. DOI: 10.1016/j.ijbiomac.2017.09.064. (20) Suárez, I.; Hassanabadi, E.; Maulu, A.; Carlino, N.; Maestri, C. A.; Latifi, M.; Bettotti, P.; Mora-Seró, I.; Martínez-Pastor, J. P., Integrated Optical Amplifier–Photodetector on a Wearable Nanocellulose Substrate. Advanced Optical Materials 2018, 6, (12), 1800201. DOI: 10.1002/adom.201800201. (21) Golmohammadi, H.; Morales-Narváez, E.; Naghdi, T.; Merkoçi, A., Nanocellulose in Sensing

and

Biosensing.

Chem.

Mater.

2017,

29,

10.1021/acs.chemmater.7b01170.


(13),

5426-5446.

DOI:


(22) Morales-Narváez, E.; Golmohammadi, H.; Naghdi, T.; Yousefi, H.; Kostiv, U.; Horák, D.; Pourreza, N.; Merkoçi, A., Nanopaper as an Optical Sensing Platform. ACS Nano 2015, 9, (7), 7296-7305. DOI: 10.1021/acsnano.5b03097. (23) Yang, J.; Du, M.; Wang, L.; Li, S.; Wang, G.; Yang, X.; Zhang, L.; Fang, Y.; Zheng, W.; Yang, G.; Jiang, X., Bacterial Cellulose as a Supersoft Neural Interfacing Substrate. ACS Appl. Mater. & Interfaces 2018, 10, (39), 33049-33059. DOI: 10.1021/acsami.8b12083. (24) Credou, J.; Berthelot, T., Cellulose: From Biocompatible to Bioactive Material. Journal of Materials Chemistry B 2014, 2, (30), 4767-4788. DOI: 10.1039/C4TB00431K. (25) Guo, Y.; Fang, Z.; Du, M.; Yang, L.; Shao, L.; Zhang, X.; Li, L.; Shi, J.; Tao, J.; Wang, J.; Li, H.; Fang, Y., Flexible and Biocompatible Nanopaper-Based Electrode Arrays for Neural Activity Recording. Nano Research 2018, 11, (10), 5604-5614. DOI: 10.1007/s12274-018-2005-0. (26) Domingues, R. M. A.; Gomes, M. E.; Reis, R. L., The Potential of Cellulose Nanocrystals in Tissue Engineering Strategies. Biomacromolecules 2014, 15, (7), 2327-2346. DOI: 10.1021/bm500524s. (27) Shunqing, T.; Wei, Y.; Xuan, M., Agarose/Collagen Composite Scaffold as An AntiAdhesive Sheet. Biomed. Mat. 2007, 2, (3), 129-134. DOI: 10.1088/1748-6041/2/3/S09. (28) Steele, T. W. J.; Huang, C. L.; Nguyen, E.; Sarig, U.; Kumar, S.; Widjaja, E.; Loo, J. S. C.; Machluf, M.; Boey, F.; Vukadinovic, Z.; Hilfiker, A.; Venkatraman, S. S., Collagen– Cellulose Composite Thin Films That Mimic Soft-Tissue and Allow Stem-Cell Orientation. J. Mater. Sci.: Mater. Med. 2013, 24, (8), 2013-2027. DOI: 10.1007/s10856-013-4940-3. (29) Xiong, G.; Luo, H.; Zhang, C.; Zhu, Y.; Wan, Y., Enhanced Biological Behavior of Bacterial Cellulose Scaffold by Creation of Macropores and Surface Immobilization of Collagen. Macromol. Res. 2015, 23, (8), 734-740. DOI: 10.1007/s13233-015-3099-9.


Page 36 of 43

Page 37 of 43 1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60


(30) Rouabhia, M.; Asselin, J.; Tazi, N.; Messaddeq, Y.; Levinson, D.; Zhang, Z., Production of Biocompatible and Antimicrobial Bacterial Cellulose Polymers Functionalized by RGDC Grafting Groups and Gentamicin. ACS Appl. Mater. & Interfaces 2014, 6, (3), 1439-1446. DOI: 10.1021/am4027983. (31) Fritz, C.; Jeuck, B.; Salas, C.; Gonzalez, R.; Jameel, H.; Rojas, O. J., Nanocellulose and Proteins: Exploiting Their Interactions for Production, Immobilization, and Synthesis of Biocompatible Materials. In Cellulose Chemistry and Properties: Fibers, Nanocelluloses and Advanced Materials, Rojas, O. J., Ed. Springer International Publishing: Cham, 2016; pp 207-224. DOI: 10.1007/12_2015_322. (32) Andrade, F. K.; Costa, R.; Domingues, L.; Soares, R.; Gama, M., Improving Bacterial Cellulose for Blood Vessel Replacement: Functionalization With a Chimeric Protein Containing a Cellulose-Binding Module and An Adhesion Peptide. Acta Biomaterialia 2010, 6, (10), 4034-4041. DOI: 10.1016/j.actbio.2010.04.023. (33) Bodin, A.; Ahrenstedt, L.; Fink, H.; Brumer, H.; Risberg, B.; Gatenholm, P., Modification of Nanocellulose with a Xyloglucan–RGD Conjugate Enhances Adhesion and Proliferation of Endothelial Cells: Implications for Tissue Engineering. Biomacromolecules 2007, 8, (12), 3697-3704. DOI: 10.1021/bm070343q. (34) Pertile, R. A. N.; Andrade, F. K.; Alves, C.; Gama, M., Surface Modification of Bacterial Cellulose by Nitrogen-Containing Plasma for Improved Interaction With Cells. Carbohydrate Polymers 2010, 82, (3), 692-698. DOI: 10.1016/j.carbpol.2010.05.037. (35) Abitbol, T.; Rivkin, A.; Cao, Y.; Nevo, Y.; Abraham, E.; Ben-Shalom, T.; Lapidot, S.; Shoseyov, O., Nanocellulose, A Tiny Fiber With Huge Applications. Current Opinion in Biotechnology 2016, 39, 76-88. DOI: 10.1016/j.copbio.2016.01.002.



Page 38 of 43

(36) Zhang, Y.; Carbonell, R. G.; Rojas, O. J., Bioactive Cellulose Nanofibrils for Specific Human

IgG

Binding.

Biomacromolecules

2013,

14,

(12),

4161-4168.

DOI:

doi.org/10.1021/bm4007979. (37) Kalaskar, D. M.; Gough, J. E.; Ulijn, R. V.; Sampson, W. W.; Scurr, D. J.; Rutten, F. J.; Alexander, M. R.; Merry, C. L. R.; Eichhorn, S. J., Controlling Cell Morphology on Amino Acid-Modified Cellulose. Soft Matter 2008, 4, (5), 1059-1065. DOI: 10.1039/B719706N. (38) Arola, S.; Tammelin, T.; Setälä, H.; Tullila, A.; Linder, M. B., Immobilization– Stabilization of Proteins on Nanofibrillated Cellulose Derivatives and Their Bioactive Film Formation. Biomacromolecules 2012, 13, (3), 594-603. DOI: 10.1021/bm201676q. (39) Rusmini, F.; Zhong, Z.; Feijen, J., Protein Immobilization Strategies for Protein Biochips. Biomacromolecules 2007, 8, (6), 1775-1789. DOI: 10.1021/bm061197b. (40) Shoseyov, O.; Shani, Z.; Levy, I., Carbohydrate Binding Modules: Biochemical Properties and Novel Applications. Microbiology and Molecular Biology Reviews 2006, 70, (2), 283-295. DOI: 10.1128/MMBR.00028-05. (41) Quero, F.; Opazo, G.; Zhao, Y.; Feschotte-Parazon, A.; Fernandez, J.; Quintro, A.; Flores, M., Top-down Approach to Produce Protein Functionalized and Highly Thermally Stable

Cellulose

Fibrils.

Biomacromolecules

2018,

19,

(8),

3549-3559.

DOI:

10.1021/acs.biomac.8b00831. (42) Zhao, Y.; Li, J., Excellent Chemical and Material Cellulose From Tunicates: Diversity in Cellulose Production Yield and Chemical and Morphological Structures From Different Tunicate Species. Cellulose 2014, 21, (5), 3427-3441. DOI: 10.1007/s10570-014-0348-6. (43) Zhao, Y.; Li, J., Ascidian Bioresources: Common and Variant Chemical Compositions and Exploitation Strategy – Examples of Halocynthia roretzi, Styela plicata, Ascidia sp. and


Page 39 of 43 1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60


Ciona intestinalis. In Zeitschrift für Naturforschung C, 2016; Vol. 71, p 165. DOI: 10.1515/znc-2016-0012. (44) Smith, M. J.; Dehnel, P. A., The Composition of Tunic From Four Species of Ascidians. Comparative Biochemistry and Physiology Part B: Comparative Biochemistry 1971, 40, (3), 615-622. DOI: 10.1016/0305-0491(71)90136-2. (45) van Name, W. G., The Tunicata, with an Account of the British Species. N. J. Berrill. The Quarterly Review of Biology 1951, 26, (3), 299-300. DOI: 10.1086/398289. (46) Díaz-Calderón, P.; Flores, E.; González-Muñoz, A.; Pepczynska, M.; Quero, F.; Enrione, J., Influence of Extraction Variables on the Structure and Physical Properties of Salmon

Gelatin.

Food

Hydrocolloids

2017,

71,

118-128.

DOI:

10.1016/j.foodhyd.2017.05.004. (47) Merrill, A. L.; Watt, B. K., Energy Value of Foods - Basis and Derivation. USDA: 1955; pp 105. DOI: 19561401372. (48) Park, S.; Baker, J. O.; Himmel, M. E.; Parilla, P. A.; Johnson, D. K., Cellulose Crystallinity Index: Measurement Techniques and Their Impact on Interpreting Cellulase Performance. Biotechnology for Biofuels 2010, 3, (1), 10. DOI: 10.1186/1754-6834-3-10. (49) Scherrer, P., Bestimmung der inneren Struktur und der Größe von Kolloidteilchen mittels Röntgenstrahlen. In Kolloidchemie Ein Lehrbuch, Zsigmondy, R., Ed. Springer Berlin Heidelberg: Berlin, Heidelberg, 1912; pp 387-409. DOI: 10.1007/978-3-662-33915-2_7. (50) Acevedo, C. A.; Orellana, N.; Avarias, K.; Ortiz, R.; Benavente, D.; Prieto, P., Micropatterning Technology to Design an Edible Film for In Vitro Meat Production. Food Bioprocess Technol. 2018, 11, (7), 1267-1273. DOI: 10.1007/s11947-018-2095-4.



(51) Nakashima, K.; Sugiyama, J.; Satoh, N., A Spectroscopic Assessment of Cellulose and the Molecular Mechanisms of Cellulose Biosynthesis in the Ascidian Ciona intestinalis. Marine Genomics 2008, 1, (1), 9-14. DOI: 10.1016/j.margen.2008.01.001. (52) Blackwell, J., Infrared and Raman Spectroscopy of Cellulose. In Cellulose Chemistry and Technology, American Chemical Society: 1977; Vol. 48, pp 206-218. DOI: 10.1021/bk1977-0048.ch014. (53) Pancake, S. J.; Karnovsky, M. L., The Isolation and Characterization of a Mucopolysaccharide Secreted by the Snail, Otella lactea. Journal of Biological Chemistry 1971, 246, (1), 253-262. DOI: http://www.jbc.org/content/246/1/253.long. (54) Wiley, J. H.; Atalla, R. H., Band Assignments in the Raman Spectra of Celluloses. Carbohydrate Research 1987, 160, (Supplement C), 113-129. DOI: 10.1016/00086215(87)80306-3. (55) Butnariu, M., Methods of Analysis (Extraction, Separation, Identification and Quantification) of Carotenoids from Natural Products. Journal of Ecosystem & Ecography 2016, 6, (2). DOI: 10.4172/2157-7625.1000193. (56) Kirti, K.; Jyoti, S.; Brajesh Kumar, T.; Mukesh Kumar, A., Detection of Carotenoids in Psychrotrophic Bacteria by Spectroscopic Approach. Journal of BioScience and Biotechnology 2014, 3, (3), 253-260. (57) Withnall, R.; Chowdhry, B. Z.; Silver, J.; Edwards, H. G. M.; de Oliveira, L. F. C., Raman Spectra of Carotenoids in Natural Products. Spectrochimica Acta Part A: Molecular and Biomolecular Spectroscopy 2003, 59, (10), 2207-2212. DOI: 10.1016/S13861425(03)00064-7.


Page 40 of 43

Page 41 of 43 1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60


(58) Lyerla, T. A.; Lyerla, J. H.; Fisher, M., Pigmentation In The Orange Tunicate, Ecteinascidia Turbinata. The Biological Bulletin 1975, 149, (1), 178-185. DOI: 10.2307/1540488. (59) Lewin, M.; Epstein, J. A., Functional Groups and Degradation of Cotton Oxidized by Hypochlorite. Journal of Polymer Science 1962, 58, (166), 1023-1037. DOI: 10.1002/pol.1962.1205816663. (60) Guo, J.; Uddin, K. M. A.; Mihhels, K.; Fang, W.; Laaksonen, P.; Zhu, J. Y.; Rojas, O. J., Contribution of Residual Proteins to the Thermomechanical Performance of Cellulosic Nanofibrils Isolated from Green Macroalgae. ACS Sustainable Chemistry & Engineering 2017, 5, (8), 6978-6985. DOI: 10.1021/acssuschemeng.7b01169. (61) Mautner, A.; Lee, K.-Y.; Tammelin, T.; Mathew, A. P.; Nedoma, A. J.; Li, K.; Bismarck, A., Cellulose Nanopapers as Tight Aqueous Ultra-Filtration Membranes. React. Funct. Polym. 2015, 86, 209-214. DOI: 10.1016/j.reactfunctpolym.2014.09.014. (62) Mautner, A.; Mayer, F.; Hervy, M.; Lee, K.-Y.; Bismarck, A., Better Together: Synergy in Nanocellulose Blends. Philos. Trans. R. Soc., A 2018, 376, (2112), 20170043. DOI: 10.1098/rsta.2017.0043. (63) Zhang, Y.; He, S.; Ma, Y.; Xu, W.; Tang, H., Characterization and Bioaccessibility of β-Carotene in Re-Assembled Casein. RSC Advances 2015, 5, (95), 77595-77600. DOI: 10.1039/C5RA07736B. (64) Kurth, F.; Franco-Obregón, A.; Bärtschi, C. A.; Dittrich, P. S., An Adaptable Stage Perfusion Incubator for the Controlled Cultivation of C2C12 Myoblasts. Analyst 2015, 140, (1), 127-133. DOI: 10.1039/c4an01758g.



(65) Burattini, S.; Ferri, P.; Battistelli, M.; Curci, R.; Luchetti, F.; Falcieri, E., C2C12 Murine Myoblasts as a Model of Skeletal Muscle Development: Morpho-Functional Characterization. Europ. J. Histochem. 2004, 48, (3), 223-33. DOI: 10.4081/891. (66) Grabowska, I.; Szeliga, A.; Moraczewski, J.; Czaplicka, I.; Brzóska, E., Comparison of Satellite Cell-Derived Myoblasts and C2C12 Differentiation in Two- and ThreeDimensional Cultures: Changes in Adhesion Protein Expression. Cell Biol. Int. 2011, 35, (2), 125-133. DOI: 10.1042/CBI20090335.

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Production of Biocompatible Protein Functionalized Cellulose Membranes by a Topdown Approach Franck Quero, Abraham Quintro, Nicole Orellana, Genesis Opazo, Andreas Mautner, Alonso Jaque, Fabiola Valdebenito, Marcos Flores and Cristian Acevedo


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Production of Biocompatible Protein Functionalized Cellulose

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