Biomineralization of Calcium Phosphate and Calcium Carbonate

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Interface-Rich Materials and Assemblies

Biomineralization of Calcium Phosphate and Calcium Carbonate within Iridescent Chitosan/Iota-carrageenan Multilayered Films Rayomand Shahlori, Daniel Rhys McDougall, Geoffrey I.N. Waterhouse, Feihong Yao, Jitendra P Mata, Andrew Nelson, and Duncan J. McGillivray Langmuir, Just Accepted Manuscript • DOI: 10.1021/acs.langmuir.8b00434 • Publication Date (Web): 01 Jul 2018 Downloaded from http://pubs.acs.org on July 6, 2018

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Biomineralization of Calcium Phosphate and Calcium Carbonate within Iridescent Chitosan/Iota-carrageenan

Multilayered

Films Rayomand Shahlori*a,b, Daniel R. McDougalla, Geoffrey I. N. Waterhousea,b, Feihong Yaoa, Jitendra P. Matac, Andrew R. J. Nelsonc, Duncan J. McGillivraya,b a

School of Chemical Sciences, University of Auckland, Private Bag 90219, Auckland,

New Zealand b c

The MacDiarmid Institute for Advanced Materials and Nanotechnology

Australian Nuclear Science and Technology Organization, Locked Bag 2001,

Kirrawee DC, NSW 2232, Australia

Corresponding author: Email: [email protected] Telephone number: +64 21 1403638

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ABSTRACT This work systematically explores the biomineralization of calcium phosphate (CaP) and carbonate (CaCO ) within chitosan/iota-carrageenan multilayer films. Multilayer 3

films of chitosan and iota-carrageenan (up to 128 coupled-layers) were prepared on glass substrates by a layer-by-layer dip coating technique. Cryo-SEM revealed dense interfaces between the chitosan and iota-carrageenan layers with thicknesses in the range 250 and 350 nm in the hydrated state, accounting for the iridescent nature of multilayer films when wet. Immersion of the multilayered films in simulated body fluid (SBF) or simulated seawater (SSW) at 25 °C resulted in the mineralization of CaP and CaCO , respectively, at the interfaces between the biopolymer layers and 3

modified the iridescence of the films. Lamellar scattering features in small-angle neutron scattering (SANS) measurements of the mineralized films provided evidence of the localized mineralization. Further evidence of this was found through the lack of change in the dynamic and static correlation lengths of the polymer networks within the bulk phase of the chitosan and iota-carrageenan layers. CaP mineralization occurred to a greater extent than CaCO mineralization within the films, evidenced by 3

the higher lamellar density and greater rigidity of the CaP mineralized films. Results provide valuable new insights into CaP and CaCO biomineralization in biopolymer 3

networks. KEYWORDS

Biomineralization,

chitosan,

iota-carrageenan,

iridescence,

biopolymer, small-angle neutron scattering, SAXS, SANS, calcium phosphate, calcium carbonate.


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INTRODUCTION Nature produces many types of biomineral structures that stimulate interdisciplinary research1-5 into the understanding and mimicking of biomineralization processes.6-10 For example, the remarkable mechanical and functional properties of bone and seashells are the result of the hierarchical structures composed of organic and inorganic components.11-14 Nacre consists of mutilayers of hexagonal calcium carbonate tiles of 300-500 nm thickness cemented together by proteins, with the submicron periodicity of the nacreous structure causing striking iridescence through selective diffraction of visible light. Researchers have attempted to reproduce nacre with varying degrees of success.6, 15-18 Li et al.15 induced slow crystallization of calcium carbonate within a suspension of carboxyl graphene. Scanning electron microscopy (SEM) revealed the formation of a multilayered carboxyl graphene/calcite structure with layer spacing similar to nacre (300-400 nm). However, the fabrication method used by Li et al. does not permit growth of continuous films on planar substrates (e.g. tens of square centimetre surface coverages). Finnemore et al.17 obtained multilayer films by layerby-layer deposition of the poly-electrolytes poly(acrylic acid) (PAA) and poly(4-vinyl pyridine) (PVP). Nano-pores were introduced into the films by immersion in a pH 10 solution. Calcium carbonate formation within the poly-electrolyte film was induced by the ammonium carbonate diffusion technique, resulting in a high-quality nacre mimic. Scanning electron microscopy analyses revealed strong similarity with nacre. However, the use of synthetic-polymer based templates meant that the nacre fabrication process reported was not a truly biomimetic process. Ideally, nacre and nacre-like materials could be fabricated using natural biodegradable and biocompatible template materials, such as biopolymer films, which could then open up a wide range of applications in drug-delivery and prosthetic development.

19-24


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Two examples of naturally-sourced biopolymers are chitosan and carrageenans. Chitosan is a positively charged linear polysaccharide composed of randomly distributed β-(1,4) linked D-glucosamine and N-acetyl-D-glucosamine units. Chitosan is readily obtained from shrimp chitin or crustacean chitin by alkali treatment, and is widely used in food, cosmetics, biomedical and pharmaceutical applications due to its biocompatibility and processability.25, 26 It is especially amenable to making coatings. Chitosan’s properties and functionality are highly dependent on the degree of acetylation

and

molecular

weight.

Carrageenans

are

negatively

charged

polysaccharides and typically are of high-molecular weight. This polysaccharide is made up of repeating galactose and 3, 6 anhydrogalactose (3, 6-AG) units (both sulfated and non-sulfated) connected by alternating α-1,3 and β-1,4 glycosidic linkages. Carrageenans are sourced from red seaweeds and are classified according to the number and position of the ester sulfate groups on the repeating galactose units which influence their gelation properties. Carrageenans are widely used as thickeners, to stabilize or gelate solutions in the food, cosmetic and pharmaceutical industries.8, 27, 28

On account of their opposing charges and high solubility in water, chitosan and carrageenans represent an ideal pair of starting materials for the fabrication of multilayer biopolymer films on planar substrates. Multilayered chitosan/iotacarrageenan films represent an ideal platform for studying biomineralization processes due to the diverse array of adsorption sites for anions and cations available on the chitosan and iota-carrageenan chains, respectively. In addition, the swelling and gelling behavior of the biopolymers, especially the carrageenans, in simulated body fluid (SBF) or seawater (SSW) is of fundamental interest to explore in tandem with the biomineralization process. The gelling and mineralization in the multilayer 4 ACS Paragon Plus Environment

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films in SBF or SSW could potentially give the multilayer films sub-micron scale periodicity and iridescence similar to nacre, though little work has been reported in this area to date, motivating a detailed investigation. This work aimed to systematically explore the biomineralization of CaP or CaCO in 3

multilayer films constructed from chitosan and iota-carrageenan prepared by dipcoating on glass. This method allows the biopolymer to form a nanostructured template rather than the typical gel templates in the literature.

29-33

Of particular interest

was the extent of CaP and CaCO mineralization when the multilayered chitosan/iota3

carrageenan films were immersed in SBF or SSW. A wide range of techniques including, Cryo-SEM, TEM, small-angle neutron scattering (SANS) and small-angle X-ray scattering (SAXS) were applied to document and corroborate the changes within the internal structure of the multilayer film following mineralization. The main objective of the study was better understanding of the CaP and CaCO mineralization 3

processes within complex biopolymer networks. EXPERIMENTAL SECTION Materials Chitosan (CAS 9012-76-4) and iota-carrageenan (CAS 9062-07-1) were obtained from Sigma Aldrich and used without further purification. Aqueous chitosan and iotacarrageenan solutions were prepared at concentrations of 1 mg mL-1 and 5 mg mL-1. To achieve complete dissolution of chitosan, acetic acid was added (to a final concentration of 2 vol.%). Complete dissolution of the iota-carrageenan powder required heating of the solution to 65 °C for 30 min. The iota-carrageenan solution was stored in an oven at 42 °C until use.


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Multilayer chitosan/iota-carrageenan films were prepared by dip-coating glass microscope slides sequentially into the chitosan and iota-carrageenan solutions at 20°C. The glass microscope slides were pre-treated in piranha solution, following the method described by Canning et al.34 The immersion time for each dip in chitosan solution or iota-carrageenan solution was approximately 20 s, after which the slides were withdrawn and excess solution removed by gentle dabbing with paper towel. A total of 16 coupled-layers were coated onto each slide before allowing the films to dry in air for 4 h. Following drying, another 16 coupled-layers were added by dip coating, followed by a 4 h drying step. The intermediate drying step was necessary to the prevent films bulging excessively and to achieve better film uniformity. This same strategy can be implemented to produce multilayered films comprising hundreds of coupled-layers. Calcium phosphate mineralization was conducted using a simulated body fluid (SBF) of composition as reported by Tas et al35 (Table 1). To induce uniform calcium phosphate mineralization, the multilayer film coated slides were placed vertically in H2O for 10 min to hydrate the films, after which they were transferred into a 2.5 mM CaCl2 solution (with the same Ca2+ concentration as SBF) for another 10 min. At this point, the slides were then immersed in the SBF solution to initiate calcium phosphate mineralization at 22°C. A similar method was used for the calcium carbonate mineralization studies, except that the SBF was replaced by simulated seawater (SSW) made accordingly to the methods described by Kester et al.36 The ionic composition and concentration of the SSW used is shown in Table 2. In order for CaCO3 mineralization to occur on a timescale similar to SBF mineralization, the SSW was diluted 16-fold with 0.2 mM Na2HCO3. After mineralizing in either SBF or SSW the films were placed in a beaker


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of type 1 water for 2 h in order to wash away loosely bound mineral and prevent salt precipitation on the film. The films were dried in a fumehood overnight. The films were swollen in either SBF or SSW (without Ca2+ to prevent further mineralization) for 30 min for a measurement that required a swollen film.

Film Characterization Scanning Electron Microscopy (SEM) SEM images were taken using a FEI Quanta 200 F ESEM operating at a voltage of 10 kV or a Philips XL-30S Field Emission Gun scanning electron microscope (FEGSEM) with Cryo-SEM attachment operated at a voltage of 5 kV in high vacuum. Prior to analysis, samples were mounted on black carbon adhesive tape and Pt sputter coated for 60 s using a Quorum Q150RS to reduce specimen charging during analysis. The biopolymer films used for SEM comprised 32 coupled-layers of chitosan and iota-carrageenan and were mineralized in SBF or SSW for time periods of 5 min, 30 min, 3 h and 18 h. The films were then cut into small sections for the SEM analysis. The biopolymer films used for Cryo-SEM comprised 60 coupledlayers of chitosan and iota-carrageenan films. A thicker film was required to improve sample preparation for cryo-SEM. A slush of solid and liquid nitrogen was used to rapidly cool the swollen biopolymer film. The sample was then placed into the sample chamber of the SEM with the temperature maintained below -140 oC. A scalpel that was built into the sample chamber was used to cut a cross-section of the frozen swollen film. Before imaging the sample was sputtered with Pt for 1 min within the sample chamber.


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The amount of coupled-layers (32 and 60) was lower than those used for small angle neutron scattering experiment as at higher layers the films began to peel from the glass substrate under the vacuum system.

Quartz-Crystal Microbalance with Dissipation (QCM-D) Quartz-crystal microbalance measurements were taken using a Q-sense E4 instrument (Q-sense, Gothenburg, Sweden) that allowed the flow of solution through four samples cells mediated by a peristaltic pump (Ismatec SA, Glattbrug, Switzerland). The sensor crystals used were coated with SiO2 (QSX-303) from Q-sense. QSoft software was used to operate the instrument and record the data. Data analysis was conducted using QTools software, the collected data was fitted using an inertial loading model (Sauerbrey equation).

Small-Angle X-ray Scattering (SAXS) Small-angle X-ray scattering measurements were conducted using a Bruker SAXS instrument, employing a rotating anode Cu Kα source (1.541Å). Scattering was obtained from a q-range of 0.01 – 0.4 Å-2. Biopolymer films containing 120 coupledlayers were used to obtain adequate scattering statistics during data collection. A 10 ´ 5 mm section of each biopolymer film was cut and placed into a sample holder. The X-ray measurements were done in transmission (without a quartz capillary) with the beam positioned perpendicular to the biopolymer film.

Small-Angle Neutron Scattering (SANS) Small-angle neutron scattering measurements were conducted using the 40 m QUOKKA37 SANS instrument at the OPAL reactor at ANSTO, Australia. Sections of


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biopolymer film were taken from the same microscope slide as the sample taken for the SAXS measurements. Each biopolymer film was carefully lifted from the glass slide and cut into circular sections of 12 mm diameter. These sections were inserted between two quartz slides and placed into a demountable cell for the SANS measurements. Prior to the measurements, the films were swollen for 1 h using 600 µL of D2O SBF or SSW buffer without Ca2+. The demountable cells were positioned such that the neutron beam is aligned perpendicular to the biopolymer films. The SANS patterns were recorded with neutron wavelengths of 5.0 and 8.1 Å. Sample-to-detector distances of 1.3 and 12 m were used for the 5.0 Å wavelength and 20 m (with the lens optics guide37) for the 8.1 Å wavelength. SANS data for the empty cell, and D2O buffer samples were measured as backgrounds. Data reduction and analysis was performed using NCNR SANS macros modified for the QUOKKA instrument38. Momentum transfer (q) was obtained after calculating the scattering angle (θ) from the detector profile data (Equation 1), using the the neutron wavelength (l). 𝑞 =

4πsinθ (1) 𝜆

The data from the samples were transformed to an absolute scale by normalizing the scattered intensity to the incident beam flux. Finally, the data were radially averaged to produce I(q) versus q plots. In all cases, the instrument and buffer backgrounds were subtracted from the sample scattering. Physical parameters were obtained from the scattering data of the multilayer films by utilizing three models to account for the scattering features at different q ranges: Power law, lamellar model, and Gaussian-Lorentz gel model. The Power law38 model (Equation 2) was used to fit low-q scattering; the slope, m, of I(q) vs q (log-log plot) provides information on the overall shape of the scattering object. 9 ACS Paragon Plus Environment

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𝐼(𝑞 ) = 𝑠𝑐𝑎𝑙𝑒 ∗ |𝑞|67 + 𝑏𝑎𝑐𝑘𝑔𝑟𝑜𝑢𝑛𝑑 (2) The lamellar model39 was used to fit the mid-q region. This model was selected due to the multilayered nature of the biopolymer films and accounts for the correlation peak observed in the mid to high q-range. Physical parameters of the model include lamellar thickness and scattering length density (SLD) (Equation 3 & 4) 𝐼 (𝑞 ) = 2𝜋

𝑃 (𝑞 ) =

𝑃(𝑞) (3) 𝛿𝑞 E

2∆𝑝E (1 − cos(𝑞𝛿 )) (4) 𝑞E

where: ∆𝑝 is the neutron scattering length density contrast between the D2O solvent and the lamellae within the biopolymer film and 𝛿 is the bilayer thickness of the lamellae within the film. The Gaussian Lorentz Gel model40 was used to describe scattering in the high-q region. This model was selected to account for the polymer network within the gel component of the film. Physical parameters obtained from this model are the static and dynamic correlation lengths, as Ξ and 𝛇 , respectively. These parameters correspond to the fixed,(Ξ),and fluctuating, (𝛇 ), crosslinks between polymer chains within a physical gel system. 𝐼 (𝑞 ) = 𝐼O (0)𝑒 (

6QR 𝚵𝟐 ) 𝟐

+

𝐼U (0) (5) 1 + 𝑞 E 𝛇E

Ig and IL are non-physical scale factors used in calculations to fit the model with experimental data. These models were combined and fitted against the the SANS data using SasView41.


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X-ray Diffraction (XRD) XRD measurements were conducted using a PANalytical Empyrean instrument. Cu Kα X-rays were generated using a rotating Cu anode that was operated at a voltage of 45 kV and a current of 40 mA. Diffracted X-rays were detected using a 1-D PIXCEL detector, and processed using the X’Pert Data Collector software. Diffraction patterns were collected for the unmineralized, SBF and SSW mineralized films, after washing in type 1 water for 2 h and drying in a fumehood.

Fourier transform infrared spectroscopy (FTIR) Fourier transform infrared spectroscopy measurements were conducted using a PerkinElmer UATR Two instrument. All the dry films were carefully transferred from the glass slide onto the diamond crystal of the ATR instrument. Spectra were collected from 500 – 4000 cm-1 with 4 cm-1 resolution.

RESULTS The chitosan/iota-carrageenan films prepared by dip coating displayed iridescence when wet, as shown in Figure 1. This iridescent colour results from dense interface regions formed between the chitosan and iota-carrageenan layers in the films, as discussed below (cryo-electron microscopy, Figure 4). The multilayer films uniformly covered approximately 12 cm2 on each side of the glass microscope slide. The unmineralized 120 coupled-layer film was golden and transitioned to an orange colour after 5 min of mineralization in SBF. Conversely, the unmineralized 120 coupledlayer film had a purple colouration after immersion in SSW for 5 min. After 30 min immersion in SBF or SSW, the multilayer films appeared blue. Mineralization for 3 h caused the intensity of this blue iridescence to fade, with only a very faint blue colour 11 ACS Paragon Plus Environment

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observed after 18 h of mineralization. No iridescence was observed for the unmineralized or the mineralized films in the dry state, indicated that hydration and a swollen film state was necessary to achieve the sub-micron periodicity needed to realize iridescence at visible wavelengths.

Quartz Crystal Microbalance with Dissipation (QCM-D) QCM-D measurements were used to follow the stepwise deposition of chitosan and iota-carrageenan layers on QCM-D chips. The biopolymer solutions (both 0.1 mg mL1

) were introduced sequentially into the QCM cells. The concentration of both

biopolymer solutions was low to prevent the co-acervation of chitosan and iotacarrageenan upon solvent exchange, which can cause blockages within the tubes of the QCM cells. Figure 2A shows synchronous decreases in the frequency and increases in dissipation as each biopolymer solution was successively introduced, indicating multilayer film formation. In the experiment, the formation of three coupled-layers was followed. The data allows estimates to be made of the adsorbed mass and kinematic viscosity of the adsorbed multilayer films. Photos of the QCM-D chips before and after biopolymer adsorption (Figure 2B) verified successful film formation. The data in Figure 2C revealed that the first chitosan layer had an adsorbed amount of approximately 3.3 μg cm-2, which was followed by an iota-carrageenan layer with an adsorption of 2.6 μg cm . This first coupled-layer thus had a large area -2

density of 5.9 μg cm , whilst the area density of two subsequent coupled-layers were -2

much smaller, 1.2 and 1.8 μg cm , respectively. The kinematic viscosity also changed -2

dramatically after the first layer was introduced. A steady 0.89 mm2 s-1 was measured for H2O, which increased in to 1.7 mm2 s-1 after passing the chitosan solution for the first time. Additional adsorbed layers caused further increases in the kinematic 12 ACS Paragon Plus Environment

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viscosity, with an approximate increase of 0.15 mm2 s-1 in viscosity observed for each subsequent biopolymer layer added.

Scanning Electron Microscopy (SEM) & Cryo-SEM SEM analyses revealed the multilayered nature of the chitosan/iota-carrageenan films in the dry state. SEM images of films containing 32 coupled-layers are shown in Figure 3. Panels A and B show the unmineralized film, which contained repeating coupled-layers of chitosan and carrageenan. The iota-carrageenan (bright) layers are thicker than the chitosan (dark) layers (72 ± 3 nm and 31 ± 2 nm, respectively). The individual layers appear to be uniform and smooth. The film is continuous over large areas (tens of micrometers) and is approximately 3.3 µm in thickness. Figure 3C and 3D show the multilayer film after 30 mins of mineralization in SBF, whereas panels E and F show the multilayer film after 30 mins of mineralization in SSW. The individual biopolymer layers have additional texture following mineralization, with the minerlization being uniform and the films maintaining their multilayered structure. The latter explains why the films retained their iridescence upon wetting. Mineralization appeared to occur on both the chitosan or iota-carrageenan layers. There does not appear to be any distinction between mineralization on the chitosan or iota-carrageenan layers. The SBF and SSW mineralized film thicknesses after 30 min exposure were determined to be ~4.6 μm and ~4.2 μm in the dry state, respectively. This represented an increase of 1.3 µm (39%) and 0.9 μm (27%), respectively, compared to the unmineralized multilayer films. SEM images for the 32 coupledlayer film after 3 h of mineralization in SBF (Figure 2G and 2H) or SSW (Figure 2I and 2J) revealed significant mineralization within the film, though, the multilayered nature of the film is less obvious. The films obtained after 18 h SBF or SSW 13 ACS Paragon Plus Environment

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treatment contained even more mineral, which further diminished the uniformity of the multilayered structure, resulting in the loss of iridescence that was characteristic of the unmineralized film (Figure 1.). Cryo-SEM images for a 60 coupled-layer film of chitosan and iota carrageenan swollen by immersion in H2O are shown in Figure 4. On hydration, the 60 coupledlayer film mass increases more than a hundred-fold due to absorbed H2O adsorption (Supporting Figures S1, S2 & S3). Figure 4 shows the vast volume expansion that occurs upon swelling. Panels A, B and C also show the multilayered nature of these films in the swollen state, with an average coupled-layer thickness, in the wet state, of approximately 4.3 μm. This is thicker than the entire 32 coupled-layers film in its dry state (3.3 μm). The coupled-layers are generally considered to exist as horizontal stacked sheets within the film to maximise charge neutralization. However, the individual sheets appear to be vertical to the film in the cryo-SEM images, which reveal compressed regions of biopolymer resulting from ice-crystal formation during sample preparation. These artefacts remain after the sublimation of the ice. Figure 4D, 4E and 4F show the 60 coupled-layer film after 180 min of exposure to SSW. This swollen film is also significantly increased in thickness compared to the dry state film. The film is less-ordered compared to the unmineralized film, with the film folding on itself and not showing distinct layers. Figure 4G, 4H and 4I show the film after exposure to SBF for 180 min (supplementary figure S5 shows the calcium phosphate particles formed within these layers). The layers appear more textured when compared to the unmineralized films and films mineralized in SSW. Cryo-TEM images of the unmineralized film (Supporting Figure S4) also show the multilayered nature of the film upon swelling. The black particles are ice crystal related artefacts.


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Small-Angle Neutron and X-ray Scattering The SANS profiles for the 120 coupled-layer chitosan and iota-carrageenan films are displayed in Figure 5. These films were exposed to SBF or SSW for 30 min and 180 min to assess the internal structure upon mineralization. The SANS profiles show distinct structural features, which can be attributed to both the gel phase and the mineral component in the chitosan/iota-carrageenan multilayer films. Upon swelling the unmineralized multilayer film shows a small correlation peak at q = 0.022 Å-1, which is characteristic of polymer crosslinks found within the gel structure of biopolymers42, 43, this feature is modelled by the Guassian-Lorentz gel component of the model and was present in all measurements. An increase in low q scattering was observed with increased mineralization. The film exposed to SBF for 180 min shows another correlation peak at q = 0.003 Å-1. We attribute this peak to the regular lamellar structure of the coupled-layered system. Films exposed to SBF for 30 min or SSW for 30 and 180 min all show a similar, though less pronouced feature. We attribute the intensity of this particular feature to the neutron scattering length density (nSLD) contrast between the interface layers and the swollen bulk biopolymer. A combined model using the power law, lamellar, and Gaussian Lorentz gel models, was used to model the SANS data and infer overall film rigidity, lamellar thickness and nSLD, along with static and dynamic correlation lengths within the multilayer film. SAXS measurements were also taken on the same sample films used for SANS, though by SAXS the films could only be measured in the dry-state. The SAXS data only required the power law in order to account for the scattering present. This provided a dry state comparison with the slope obtained using the SANS power law for the swollen sample.


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SANS and SAXS data for the unmineralized and mineralized 120-coupled-layer films and the fitted parameters for each type of treated film is summarized in Table 3.

X-ray Diffraction (XRD) and Fourier Transform Infrared Spectroscopy (FTIR) XRD patterns and FTIR spectra for the unmineralized and SBF or SSW treated films are shown in Figure 6. The XRD patterns do not show any distinct reflections from calcium phosphate or calcium carbonate within the mineralized films, suggesting any mineral components were amorphous. The unmineralized film shows a broad amorphous feature from 20° to 35° in 2θ along with two sharp reflections occurring at 2θ values of 8.2° and 16.9°, which correspond to d-spacings of 10.8 Å and 5.2 Å, respectively in the multilayered dry film. The feature at approximately 8.2° was also present in XRD patterns of the mineralized films; with a shift towards lower 2θ angles for the SSW exposed films. This reflection was observed at a 2θ of 7.9° (d spacing of 11.2 Å) for the 3 h SSW exposed film. The sharp feature observed at 16.9° for the unmineralized film was strongly attenuated by exposure to SBF for 30 min and 3 h. This feature was not observed at all in the XRD patterns of SSW exposed films, suggesting that mineralization in the multilayer film may have occurred preferentially on sites giving rise to the 16.9o feature, resulting in a disruption of the crystalline order at such sites (hence the attenuation or disappearance of the peak). The FTIR spectra for the unmineralized and

SBF

and

SSW exposed

chitosan/iotacarrageenen films are shown in Figure 6. The spectrum of the unmineralized film was characteristic for the two component biopolymers and dominated by bands in the 900-1100 cm-1 region associated with the saccharide linkages in the biopolymers. Adsorbed water gave rise to band at 3300 and 1640 cm-1.


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Mineralization using both SBF and SSW introduced a number of new features in the FTIR spectra, which could readily be assigned to phosphate and carbonate ions, respectively. For example, SBF exposed films showed the characteristic v3 (950-1150 cm-1) and v4 (450-650 cm-1) bands of phosphate. However, for both SBF and SSW exposed films, it is not possible to unambiguously identify the particular phase of mineral present due to the amorphous nature of the mineral deposited. In order to confirm mineral formation within these films, the SBF and SSW treated chitosan/iota-carrageenan films, as well as the unmineralized films, were heated to 600 °C in N2 for 2 h (Supplementary Information Figure S7). The unmineralized film after heating gave an XRD pattern containing many reflections, suggesting some form of crystalline carbonaceous product. The heat-treated SBF exposed films showed fewer reflections but distinct features at 2θ values of 23° and 34°, which were more prominent in the 24 h mineralized film. The 2θ values suggest the formation of Ca3(PO4)2. The heat-treated SSW exposed films showed distinct features at 2θ values of 29°, 39°, 43°, 47° and 49°, which can be attributed to the [104], [113], [202], [018] and [116] planes microcrystalline particles of calcite44 polymorph of CaCO3, respectively. These features are also of greater intensity after 24 h SSW exposure, suggesting CaCO3 formation. The FTIR spectra of heated treated SBF and SSW treated films confirmed the formation of Ca3(PO4)2 and CaCO3, respectively. It is likely that the as-prepared mineralized films also contained similar mineralized species, albeit in amorphous form.


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DISCUSSION Biopolymer film formation The multilayer biopolymer films were prepared by dip-coated on glass microscope slides. The piranha treated slides exposed a hydrophilic negatively charged SiO2 surface (SiO2 has an isoelectric point of 2, which is well below the pH of the chitosan and iota-carrageenan solutions, pH 3.9 and 4.5, respectively). On immersion in the chitosan solution, the positively charged NH3+ groups on the poly-saccharide chains will be electrostatically attracted to the slide surface. QCM-D (Figure 2) showed that chitosan adsorption occurred within 20 seconds of the chitosan solution being introduced to the cell. The chitosan adsorption causes a drop in frequency (adsorbed mass) and increase in dissipation (layer is viscoelastic). The data shows that approximately 20 sec is sufficient for complete adsorption as the frequency and dissipation do not change with longer exposure. Similarly, the time required for iotacarrageenan adsorption on the chitosan layer was also ~20 sec, with no further accumulation on the chitosan surface occuring after this time. The rate of adsorption of subsequent chitosan and iota-carrageenan layers was similar (~20 s to completion), thus allowing 16 coupled-layers to be deposited in approximately 20 min. Similar dipcoating procedures have exposed slides to each polymer solution for up to 15 minutes, individually, in order to achieve uniform coating.45 Our work reveals that for the chitosan/iota-carrageenan system, such lengthy immersions of the inidvidual biopolymer solutions are not necessary. SEM and TEM images confirmed the formation of uniform multilayer structures composed of alternating chitosan and iotacarrageenan layers. The uniformity of the multilayer coatings was especially evident when the biopolymer films are swollen with H2O (absorbing ~150 times the dry


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weight), whereupon iridescence could be observed due to the periodic structures in the films.

Biopolymer film mineralization Mineralization of amorphous calcium phosphate and amorphous carbonate was induced by exposing the films to SBF and SSW, respectively (Figure 6). Summarized SANS and SAXS data for the mineralized films is provided in Table 3, and plotted in Figures 6, 7 and 8. The slopes obtained from the power law fits for both the SAXS and SANS data are shown in Figure 7. Plotting these parameters shows groupings of SBF (red) and SSW (blue), with the SBF samples tending to have larger a SANS power law gradient (m) values and smaller SAXS gradient (m), revealing the striking difference in overall structure upon swelling. We would expect the SBF films to contain more mineral than the SSW exposed films, since the Ksp of calcium phosphate46 relative to amount of Ca2+ and PO43- in SBF is lower than the Ksp of calcium carbonate relative to the Ca2+ and CO32- in SSW. The unmineralized film containing organic material had the lowest SANS m value and a relatively high SAXS m value, as a result of only containing organic material (no mineral). This indicates that the unmineralized film shows scattering characteristic of mass fractals (SANS m = 2.26) in the swollen state. Mineralization causes the film to transition towards a smooth interface, with the most mineralized system approaching scattering characteristic of a smooth film (SBF 30 min, 180 min SANS m = 3.70 & 3.83). The dashed line through the plot shows where the data would be if the SAXS and SANS provided the same slopes. The parameters obtained from the lamellar model fitting for the films that showed a significant scattering feature at 0.003 Å-2 are shown in Figure 8. This plot shows the 19 ACS Paragon Plus Environment

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lamellar thickness and neutron scattering length density (nSLD) that arise from the interface layers seen by Cryo-SEM (Figure 4). Only the heavily mineralized samples exhibit this feature due to a threshold of mineralization required to create an interface layer or lamellar with enough nSLD contrast in order to scatter. This is supported by the SANS scattering profiles, wherein a lamellar scattering feature at 0.003 Å-2 emerges abrutly without a gradual rise from high q, which would indicate the growth of a particle. The same lamellar features were present in the unmineralized films, as seen in cryo-SEM, but do not give rise to the lamellar feature in the SANS due to the low contrast between the multilayers. The dashed vertical line on Figure 8 is placed on the nSLD of D2O (6.36´10-6 Å-2), which was the value used for the solvent in the modelling. Upon mineralization the nSLD of the lamellar/interface layers start to decrease due to a decrease in moisture and increase in mineral content. Figure 8 shows that the SBF exposed films, which are more heavily mineralized than the SSW exposed films, display the lowest lamellar nSLD (or highest nSLD contrast against D2O). The most heavily mineralized films (i.e. 30 and 180 min SBF films) show the highest contrast against D2O with lamellar nSLDs of 2.7´10-6 and 2.0´10-6 Å-2, respectively. The polydispersity of the film thickness obtained from the SANS fitting ranged from 14 to 20 %, with the average lamellar thickness for the films between 1500 and 2500 Å (150-250 nm), which is in excellent accordance with the Cryo-SEM and TEM results. A plot of the lamellar nSLD against the SANS power law slope in (Figure 9) shows an inverse correlation between the two variables. Again the vertical dashed line represents the nSLD of D2O (6.36´10-6 Å-2), with the horizontal line showing the SANS power law slope for the unmineralized 120 coupled-layer film (m = 2.26). The point where these lines converge can be viewed as approximately where the 20 ACS Paragon Plus Environment

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unmineralized film would be positioned if its lamellar nSLD was low enough to produce enough contrast for lamellar scattering. This point and the region around it is empty for this reason, however a trend from this region towards the high SANS power law slope and low nSLD is observed. This supports our assertion that the scattering arises from lamellae in the form of interfaces within the coupled-layers of the biopolymer film. It also shows that with increasing lamellar density, as a result of mineralization, the overall internal structure of the film tends to a smoother flat surface. The dynamic and static correlation lengths (Figure S7) for each of the treated 120coupled-layer films were not grouped with respect to SBF or SSW, additionally these correlation lengths do not seem to relate to extent of mineralization. The values for the dynamic and static correlation lengths range from 35 to 60 Å and 10 to 40 Å, respectively. The obtained range of polymer-polymer spacing within the swollen film correlates well with the dry-state spacing found by XRD reflections (Figure S7) corresponding to 11 and 5 Å. It is expected that this spacing would increase upon interaction with D2O through swelling, however, there is no effect of mineralization on the spacing between the polymer network within the film. With this in mind, it is possible that the 5 Å spacing found in the XRD pattern of the unmineralized film arises from the periodic ordering of the biopolymer within the interfaces of chitosan and iota-carrageenan layers and the 11 Å spacing from layers between the interfaces. This would explain the difference in length scales of the two spacings, since the interface contains oppositely charged biopolymers, leading to stronger interactions and shorter distances between molecules. Additionally, the disappearance of this reflection upon mineralization is explained by the exclusive mineralization at these interfaces, causing a disruption of the periodic ordering of the biopolymers in this 21 ACS Paragon Plus Environment

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region when unmineralized. This finding provides additional indication that mineralization is occuring exclusively at the interfaces between the individual chitosan and iota-carrageenan layers. Therefore, the polymer-polymer spacing within the film remains unaltered by mineralization within the interface layers.

CONCLUSION We have successfully demonstrated that multilayered mineral films can be fabricated by immersing chitosan/iota-carrageenan multilayered films in SBF or SSW. The unmineralized and mineralized chitosan/iota-carrageenan films that were produced show striking iridescence exclusively when swollen with H2O. Calcium phosphate and carbonate mineralization through exposure to SBF and SSW altered the density and changed the colour of the multilayer films Extensive mineralization resulted in a loss of the iridescence. SANS profiles from a 120 coupled-layer film showed scattering features from the biopolymer gel phase, the mineralized lamellae and the overall film structure. We observed a tendency for these films to show scattering characteristic of smooth sheets and increasingly dense lamellae upon exposure to SBF and SSW. The rapid and simple dip coating method described here to fabricate the chitosan/iota-carrageenan multilayers is expected to be applicable to other charged biopolymer systems, allowing the fabrication of various biopolymer lamellae systems for drug delivery and biomineralization studies.


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ASSOCIATED CONTENT Supporting Information Experimental methods and 7 figures AUTHOR INFORMATION Corresponding Author: E-mail: [email protected] Phone: +64 21 1403638. Notes: The authors declare no competing financial interest. ACKNOWLEDGEMENTS The authors acknowledge funding to support the experiments and travel provided by the MacDiarmid Institute for Advanced Materials and Nanotechnology and the Australian Institute for Nuclear Science and Engineering (AINSE). The authors would also like to thank the Australian Nuclear Science and Technology Organization (ANSTO) for the awards of beamtime (experiment numbers P4932 & P5426). RS was supported by a PhD scholarship from the MacDiarmid Institute and a Postgraduate Research award from AINSE.


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REFERENCES 1. Bergmann, C.; Lindner, M.; Zhang, W.; Koczur, K.; Kirsten, A.; Telle, R.; Fischer, H., 3D printing of bone substitute implants using calcium phosphate and bioactive glasses. Journal of the European Ceramic Society 2010, 30 (12), 25632567. 2. Nico, A. D.; Sommerdijk, A. J. M., Advanced Transmission Electron Microscopy to Explore Early Stages of Bio(mimetic) mineralization. In Materials Design Inspired by Nature: Function through Inner Architecture, Fratzl, P.; Dunlop, J. W. C.; Weinkamer, R., Eds. Royal Soc Chemistry: Cambridge, 2013; pp 74-106. 3. Uysal, A.; Stripe, B.; Lin, B.; Meron, M.; Dutta, P., Assembly of Amorphous Clusters under Floating Monolayers: A Comparison of in Situ and ex Situ Techniques. Langmuir 2013, 29 (47), 14361-14368. 4. Zollfrank, C., Bioinspired material surfaces - Science or engineering? Scripta Materialia 2014, 74, 3-8. 5. Halthur, T. J.; Claesson, P. M.; Elofsson, U. M., Immobilization of enamel matrix derivate protein onto polypeptide multilayers. Comparative in situ measurements using ellipsometry, quartz crystal microbalance with dissipation, and dual-polarization interferometry. Langmuir 2006, 22 (26), 11065-11071. 6. Qin, Z.; Ren, X.; Shan, L.; Guo, H.; Geng, C.; Zhang, G.; Ji, S.; Liang, Y., Nacre like-structured multilayered polyelectrolyte/calcium carbonate nanocomposite membrane via Ca-incorporated layer-by-layer-assembly and CO2-induced biomineralization. Journal of Membrane Science 2016, 498, 180-191. 7. Chen, Y. J.; Xiao, J. W.; Wang, Z. N.; Yang, S. H., Observation of an Amorphous Calcium Carbonate Precursor on a Stearic Acid Monolayer Formed during the Biomimetic Mineralization of CaCO3. Langmuir 2009, 25 (2), 1054-1059. 8. Diaz-Dosque, M.; Aranda, P.; Darder, M.; Retuert, J.; Yazdani-Pedram, M.; Arias, J. L.; Ruiz-Hitzky, E., Use of biopolymers as oriented supports for the stabilization of different polymorphs of biomineralized calcium carbonate with complex shape. Journal of Crystal Growth 2008, 310 (24), 5331-5340. 9. Abdelkebir, K.; Morin-Grognet, S.; Gaudiere, F.; Coquerel, G.; Labat, B.; Atmani, H.; Ladam, G., Biomimetic layer-by-layer templates for calcium phosphate biomineralization. Acta Biomaterialia 2012, 8 (9), 3419-3428. 10. Shahlori, R.; Waterhouse, G. I. N.; Nelson, A. R. J.; McGillivray, D. J., Morphological, chemical and kinetic characterisation of zein protein-induced biomimetic calcium phosphate films. Journal of Materials Chemistry B 2015. 11. Maestro, A.; Kotsmar, C.; Javadi, A.; Miller, R.; Ortega, F.; Rubio, R. G., Adsorption of beta-Casein-Surfactant Mixed Layers at the Air-Water Interface Evaluated by Interfacial Rheology. Journal of Physical Chemistry B 2012, 116 (16), 4898-4907. 12. Rey, C.; Combes, C.; Drouet, C.; Glimcher, M. J., Bone mineral: update on chemical composition and structure. Osteoporosis International 2009, 20 (6), 10131021. 13. Fratzl, P.; Weinkamer, R., Nature's hierarchical materials. Progress in Materials Science 2007, 52 (8), 1263-1334. 14. Snow, M. R.; Pring, A.; Self, P.; Losic, D.; Shapter, J., The origin of the color of pearls in iridescence from nano-composite structures of the nacre. American Mineralogist 2004, 89 (10), 1353-1358.


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15. Li, J.; Liu, D.; Li, B.; Wang, J.; Han, S.; Liu, L.; Wei, H., A bio-inspired nacre-like layered hybrid structure of calcium carbonate under the control of carboxyl graphene. Crystengcomm 2015, 17 (3), 520-525. 16. Voet, V. S. D.; Kumar, K.; ten Brinke, G.; Loos, K., Bioinspired Synthesis of Well-Ordered Layered Organic-Inorganic Nanohybrids: Mimicking the Natural Processing of Nacre by Mineralization of Block Copolymer Templates. Macromolecular Rapid Communications 2015, 36 (19), 1756-1760. 17. Finnemore, A.; Cunha, P.; Shean, T.; Vignolini, S.; Guldin, S.; Oyen, M.; Steiner, U., Biomimetic layer-by-layer assembly of artificial nacre. Nature Communications 2012, 3. 18. Li, X. Q.; Zeng, H. C., Calcium Carbonate Nanotablets: Bridging Artificial to Natural Nacre. Advanced Materials 2012, 24 (47), 6277-6282. 19. Zhang, C. Y.; Zhang, W.; Mao, L. B.; Zhao, Y.; Yu, S. H., Biomimetic mineralization of zein/calcium phosphate nanocomposite nanofibrous mats for bone tissue scaffolds. Crystengcomm 2014, 16 (40), 9513-9519. 20. Biswas, A.; Nagaraja, A. T.; McShane, M. J., Fabrication of Nanocapsule Carriers from Multilayer-Coated Vaterite Calcium Carbonate Nanoparticles. Acs Applied Materials & Interfaces 2014, 6 (23), 21193-21201. 21. Elyada, A.; Garti, N.; Furedi-Milhofer, H., Polyelectrolyte Multilayer-Calcium Phosphate Composite Coatings for Metal Implants. Biomacromolecules 2014, 15 (10), 3511-3521. 22. Hsu, B. B.; Hagerman, S. R.; Hammond, P. T., Rapid and efficient sprayed multilayer films for controlled drug delivery. Journal of Applied Polymer Science 2016, 133 (25). 23. Tas, A. C.; Bhaduri, S. B., Rapid coating of Ti6A14V at room temperature with a calcium phosphate solution similar to 10x simulated body fluid. Journal of Materials Research 2004, 19 (9), 2742-2749. 24. Popescu, L. M.; Piticescu, R. M.; Antonelli, A.; Rusti, C. F.; Carboni, E.; Sfara, C.; Magnani, M.; Badilita, V.; Vasile, E.; Trusca, R.; Buruiana, T., Recent advances in synthesis, characterization of hydroxyapatite/polyurethane composites and study of their biocompatible properties. Journal of Materials Science-Materials in Medicine 2013, 24 (11), 2491-2503. 25. Kato, T., Polymer/calcium carbonate layered thin-film composites. Advanced Materials 2000, 12 (20), 1543-1546. 26. Leite, A. J.; Sher, P.; Mano, J. F., Chitosan/chondroitin sulfate multilayers as supports for calcium phosphate biomineralization. Materials Letters 2014, 121, 62-65. 27. Kosanovic, C.; Falini, G.; Kralj, D., Mineralization of Calcium Carbonates in Gelling Media. Crystal Growth & Design 2011, 11 (1), 269-277. 28. Paula, G. A.; Benevides, N. M. B.; Cunha, A. P.; de Oliveira, A. V.; Pinto, A. M. B.; Morais, J. P. S.; Azeredo, H. M. C., Development and characterization of edible films from mixtures of kappa-carrageenan, i-carrageenan, and alginate. Food Hydrocolloids 2015, 47, 140-145. 29. Gashti, M. P.; Helali, M.; Karimi, S., Biomineralization-Inspired Green Synthesis of Zinc Phosphate-Based Nanosheets in Gelatin Hydrogel. International Journal of Applied Ceramic Technology 2016, 13 (6), 1069-1073. 30. Gashti, M. P.; Stir, M.; Hulliger, J., Growth of strontium hydrogen phosphate/gelatin composites: a biomimetic approach. New Journal of Chemistry 2016, 40 (6), 5495-5500.


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31. Chauhan, C. K.; Joshi, M. J., In vitro crystallization, characterization and growth-inhibition study of urinary type struvite crystals. Journal of Crystal Growth 2013, 362, 330-337. 32. Gashti, M. P.; Stir, M.; Bourquin, M.; Hulliger, J., Mineralization of Calcium Phosphate Crystals in Starch Template Inducing a Brushite Kidney Stone Biomimetic Composite. Crystal Growth & Design 2013, 13 (5), 2166-2173. 33. Gashti, M. P.; Stir, M.; Hulliger, J., Synthesis of bone-like micro-porous calcium phosphate/iota-carrageenan composites by gel diffusion. Colloids and Surfaces B-Biointerfaces 2013, 110, 426-433. 34. Canning, J.; Petermann, I.; Cook, K., Surface treatment of silicate based glass: base Piranha treatment versus 193nm laser processing. In Third Asia Pacific Optical Sensors Conference, Canning, J.; Peng, G. D., Eds. Spie-Int Soc Optical Engineering: Bellingham, 2012; Vol. 8351. 35. Tas, A. C., The use of physiological solutions or media in calcium phosphate synthesis and processing. Acta Biomaterialia 2014, 10 (5), 1771-1792. 36. Kester, D. R.; Duedall, I. W.; Connors, D. N.; Pytkowicz, R. M., Preparation of Artificial Seawater. Limnology and Oceanography 1967, 12 (1), 176-+. 37. Gilbert, E. P.; Schulz, J. C.; Noakes, T. J., 'Quokka' - the small-angle neutron scattering instrument at OPAL. Physica B-Condensed Matter 2006, 385-86, 11801182. 38. Kline, S. R., Reduction and analysis of SANS and USANS data using IGOR Pro. Journal of Applied Crystallography 2006, 39, 895-900. 39. Berghausen, J.; Zipfel, J.; Lindner, P.; Richtering, W., Influence of watersoluble polymers on the shear-induced structure formation in lyotropic lamellar phases. Journal of Physical Chemistry B 2001, 105 (45), 11081-11088. 40. G. Evmenenko, E. T., K. Mortensen, H. Reynaers., SANS study of surfactant ordering in κ-carrageenan/cetylpyridinium chloride complexes. Polymer 2001, 42, 2907-2913. 41. http://www.sasview.org/about.html. 42. da Silva, M. A.; Bode, F.; Grillo, I.; Dreiss, C. A., Exploring the Kinetics of Gelation and Final Architecture of Enzymatically Cross-Linked Chitosan/Gelatin Gels. Biomacromolecules 2015, 16 (4), 1401-1409. 43. Berts, I.; Gerelli, Y.; Hilborn, J.; Rennie, A. R., Structure of polymer and particle aggregates in hydrogel composites. Journal of Polymer Science Part BPolymer Physics 2013, 51 (6), 421-429. 44. Munro, N. H.; Green, D. W.; Dangerfield, A.; McGrath, K. M., Biomimetic mineralisation of polymeric scaffolds using a combined soaking and Kitano approach. Dalton Trans. 2011, 40 (36), 9259-9268. 45. Patel, I. F.; Kiryukhin, M. V.; Yakovlev, N. L.; Gupta, H. S.; Sukhorukov, G. B., Naturally inspired polyelectrolyte multilayer composite films synthesised through layer-by-layer assembly and chemically infiltrated with CaCO3. Journal of Materials Chemistry B 2015, 3 (24), 4821-4830. 46. Ito, A.; Maekawa, K.; Tsutsumi, S.; Ikazaki, F.; Tateishi, T., Solubility product of OH-carbonated hydroxyapatite. Journal of Biomedical Materials Research 1997, 36 (4), 522-528.


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Table 1. Concentration of ions in SBF solution, pH = 7.4, Ionic strength = 193 mmol L. -1

Film

SANS Power law/Q-m

SANS Lamellar thickness/Å

Lamellar neutron SLD/10-6Å-2

Dynamic correlation length/Å

Static correlation length/Å

SAXS Power law/Q-x

0 SSW 30 SSW 180 SBF 30 SBF 180

2.26 ± 0.04 2.91 ± 0.07 3.19 ± 0.03 3.70 ± 0.02 3.83 ± 0.02

1500 ± 100 1480 ± 40 1870 ± 50 2120 ± 30

5.2 ± 0.1 3.6 ± 0.1 2.7 ± 0.1 2.0 ± 0.1

41 ± 1 42 ± 2 48 ± 2 36 ± 1 40 ± 2

22 ± 1 19 ± 1 22 ± 1 17 ± 1 18 ± 1

2.60 ± 0.01 2.44 ± 0.01 2.56 ± 0.01 2.32 ± 0.01 2.21 ± 0.01

Table 2. Concentration of ions in SSW solution, pH = 6.8, Ionic strength = 577 mmol L. -1

Ion mmol L

-1

Na 160

K 5

+

Ca 3

+

Mg 1

2+

Cl 152

2+

HPO 1

-

CO 10

24

23

Table 3. Fitted parameters of the measured SANS data, a combined model using the power law, lamellar and Gaussian Lorentz gel model. The power law of the SAXS data is also included. Ion mmol L-1

Na 411 +

K 9

+

Ca 10

2+

Mg 53

2+

Cl 474 -

CO 2

23


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Table of Contents Graphic


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Figure Captions Figure 1. Digital photographs showing chitosan/iota-carrageenan multilayer films containing 120 coupled-layers. A, B show the unmineralized (UM) film in both the dry and swollen hydrated state, respectively. Subsequent rows show the multilayer film after exposure for 5 min, 30 min, 180 min and 18 h to SBF (dry, C, F, I, L), (swollen, D, G, J, M) and SSW (swollen, E, H, K, N). Figure 2. QCM-D data (A) showing changes in frequency and dissipation on successive introduction of chitosan and iota-carrageenan solutions. QCM-D chips before (top) and after (bottom) biopolymer film formation (B). Modeled thickness and kinematic viscosity with chitosan and iota-carrageenan addition (C). Figure 3. SEM images showing chitosan/iota-carrageenan multilayer films containing 32 coupled-layers in the dry state. Panel A shows a cross-sectional view of the unmineralized (UM) film, panel B is the same film at higher magnification. The subsequent rows show the biopolymer film after 30 min, 180 min and 18 h exposure to SBF (C, D, G, H, K, L) or SSW (E, F, I, J, M, N). Figure 4. Cryo-SEM images showing chitosan/iota-carrageenan multilayer films containing 60 coupled-layers in the swollen hydrated state. Panel A shows a crosssectional view of the unmineralized film, panels B and C show the same film at higher magnification. The subsequent rows are the biopolymer film after 180 min exposure to SSW (D, E, F) or SBF (G, H, I). Figure 5. Small angle neutron scattering (SANS, top) and small angle X-ray scattering (SAXS, bottom) profiles for a chitosan/iota-carrageenan multilayer film containing 120 coupled-layers. Data for the unmineralized film and film after


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exposure 30 min and 180 min exposure to SBF or SSW is shown. All data was collected with the films in the wet state. Figure 6. XRD patterns (top) and FTIR spectra (bottom) of a 128 coupled-layer film in the unmineralised state before and after exposing to SBF and SSW for 30 min and 3 h. Figure 7. Plot showing the SAXS power slope against the SANS power slope for 120-coupled-layer chitosan/iota-carrageenan films mineralized in SBF (red) or SSW (blue) for 30 and 180 min. Data for unmineralized film is labelled 0. The diagonal dashed line shows the 1:1 SAXS and SANS m values. Figure 8. Plot showing the lamellar nSLD against the lamellar coupled-layer thickness for the 120-coupled-layer chitosan/iota-carrageenan films mineralized in SBF (red) or SSW (blue) for 30 and 180 min. The unmineralized film did not show significant lamellar scattering and thus is not plotted. The vertical dashed line shows the nSLD of the bulk biopolymer layer. Figure 9. Plot showing the lamellar nSLD against the SANS power slope for the 120coupled-layer chitosan/iota-carrageenan films mineralized in SBF (red) and SSW (blue) for 30 and 180 min, the unmineralized film did not show significant lamellar scattering to be plotted. The horizontal dashed line shows the SANS power law value for the unmineralized multilayer film, whereas the vertical dashed line shows the nSLD of the bulk biopolymer layer.


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Figure 1


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Figure 2


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Figure 3


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Figure 4


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Langmuir

Figure 5


Langmuir 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

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Figure 6


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Langmuir

Figure 7


Langmuir 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

Page 38 of 39

Figure 8


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Langmuir

Figure 9.


Biomineralization of Calcium Phosphate and Calcium Carbonate

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