Development of a drug delivery platform using multifunctional polymeric scaffold for scar therapy Vipul Agarwal, MApplSc This thesis is presented for the degree of Doctor of Philosophy at The University of Western Australia School of Chemistry and Biochemistry 2015
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Development of a drug delivery platform using
multifunctional polymeric scaffold for scar
therapy
Vipul Agarwal, MApplSc
This thesis is presented for the degree of Doctor of Philosophy at The University of
Western Australia
School of Chemistry and Biochemistry
2015
i
Contents Abbreviations .............................................................................................................. iv
Abstract ...................................................................................................................... vii
Acknowledgement ...................................................................................................... ix
Statement of candidate contribution ........................................................................... xi
1 Introduction and literature review ........................................................................ 1
Vipul Agarwal,a Dominic Ho,a Diwei Ho,a Yuriy Galabura,b Faizah M.D. Yasin,a Peijun Gong,c Weike Ye,d Ruhani Singh,a Alaa Munshi,a Martin Saunders,e Robert C. Woodward,f Timothy St. Pierre,f Fiona M. Wood,g Mark Fear,g Dirk Lorenser,c David D. Sampson,c,e Bogdan Zdyrkob, Nicole M. Smith,a Igor Luzinovb,*, and K. Swaminathan Iyera,*
Synthetic fractal materials have been regarded as a new class of hybrid materials with many potential applications. However, the lack of an efficient, reactive large-area fractal substrate has been one of the major limitations in the development of these materials as advanced functional platforms. Herein, we demonstrate the utility of electrospun polyglycidyl methacrylate (PGMA) fractal-like films as a highly versatile platform for the development of functional nanostructured fractal-like materials anchored to a surface. The utility of this platform as a reactive substrate is demonstrated by grafting poly (N-isopropyl acrylamide) to incorporate stimuli-responsive properties. Additionally, we demonstrate that functional fractal-like nanocomposites can be fabricated using this platform with properties for sensing, fluorescence imaging and magneto-responsiveness.
The development of nanostructured polymeric matrices to obtain organic-inorganic nanocomposites has been actively researched to produce hybrid materials for applications in electronics, optics, medical devices, sensors and catalysis.1-4 Of the various techniques developed to produce large area nanoscale polymeric matrices, one of the most researched, cost effective and facile method is electrospinning. It has been adapted to cover a wide range of polymers and optimized to regulate fiber diameter, alignment and shape.5-7 There have been numerous reports using this technique to develop matrices with enhanced mechanical strength,8 matrices with selective filtration/permeability,9 fire retarding material, optoelectronic devices10 and substrates for catalysis.11, 12 One of the key steps involved in the development of organic-inorganic nanocomposites is grafting to achieve excellent integration by minimizing interfacial tension of the nanoparticles in the
organic nanofiber matrix. Additionally, the ability to modify the surface of the nanofiber to alter the adhesion, lubrication, wettability and biocompatibility is pivotal in its customisation for end-use applications. The achievement of a certain degree of grafting universality requires the establishment of a controlled method of introducing the desired functional groups on a substrate.5 Currently, this is achieved by physisorption.13,
14 In contrast, chemisorption, which is difficult to achieve on a polymeric nanofibers, would result in permanent irreversible surface modification. Polymers containing epoxy groups are examples of functional polymers that are able to react with a wide range of substrates through ‘‘grafting to’’ interactions mediated by the epoxy groups.15, 16 The versatile chemistry of epoxy groups renders a polymer that is exceptionally suitable as a universal electrospun nanofiber matrix to provide reactive groups for further grafting reactions. To this end, poly(glycidyl methacrylate) (PGMA), which contains an epoxy group in every repeating unit, has been used extensively as a macromolecular anchoring layer for grafting of polymers to the surfaces.17-20 Upon electrospinning, epoxy groups in the polymer will undergo self-crosslinking upon heating, providing mechanical integrity to the matrix.21 Approximately 40% of the epoxy groups are still available for surface modification following a 12 hour treatment at 120°C.
The main advantage of using PGMA as a matrix for electrospinning, as opposed to modifying the surface using monolayers, is the high mobility of the epoxy groups located in the “loops” and “tails” of the polymer. The mobility of the free groups results in the formation of a highly effective interpenetrating anchoring zone.22 In this article, we report that PGMA can be directly electrospun (ES-PGMA) to form large area nanofibers. We demonstrate that this polymer nanofiber matrix can be used as an effective platform to graft polymers to impart switchability, and can be used to produce
nanocomposites with upconverting properties, with hydrogen sensing capability or with magneto-responsive properties.
The ES-PGMA nanofibers (see Supporting Information for method of synthesis) were uniform over a large area and had an average diameter of 0.69 ± 0.04 µm (average ± standard error mean) (Fig. 1A). The average thickness of the 1 x 1 cm2 ES-PGMA generated was 127 ± 3 µm in 7 hours (Fig. 1B). In order to test the efficacy of the ES-PGMA nanofiber matrix as an anchoring platform, carboxylic acid-terminated poly (N-isopropyl acrylamide) (PNIPAM-COOH) was end grafted to the nanofibers via a ring opening reaction with the epoxy groups to yield ES-PGMA-g-PNIPAM-COOH.23 PNIPAM is a thermo-responsive polymer, which has been utilized in various
forms, such as thermo-responsive hydrogels, particles, brushes, spheres and micelles.24-27 Importantly, PNIPAM exhibits a
temperature-sensitive phase transition in water at what is known as a lower critical solution temperature (LCST), 32 °C.28
The transition is due to the coil-to-globule transition at the critical temperature resulting in switching from hydrophilic to hydrophobic behavior.29 At temperatures below the LCST, PNIPAM chains arrange into an expanded and hydrated conformation. Conversely, at temperatures above the LCST, PNIPAM chains collapse and arrange into a compact, dehydrated conformation.29 This thermo-responsive behavior is retained post-grafting and post-end group functionalization. The ES-PGMA-g-PNIPAM-COOH nanofibers demonstrated thermo-responsive behavior as monitored using contact angle measurements. The contact angle changed from 60 ± 2° at 70°C (Fig. 1C) to 15 ± 2° at room temperature (Fig. 1D). The contact angle of the unmodified ES-PGMA remained unchanged at 100 ± 2° both at room temperature and at 70°C. The ability of the ES-PGMA nanofiber matrix to produce nanocomposites was further evaluated using three distinct classes of nanoparticles: upconverting fluorescent particles of NaGdF4:Yb, Er (UCNP), palladium (Pd) and magnetite (Fe3O4). The nanoparticles synthesized (see Supporting Information for methods) had a narrow size distribution of 7.4 ± 1.4 nm (average ± standard error mean) for UCNP, 19.3 ± 0.2 nm for Pd and 6.7 ± 1.4 nm for Fe3O4 respectively (Fig. 2A, C and E). One of the major hurdles in developing functional materials by electrospinning nanocomposites is the lack of control in attaining a homogeneous distribution of nanoparticles throughout the polymer matrix. In the present case, electrospinning PGMA with the aforementioned nanoparticles resulted in relatively uniform distributions of the nanoparticles throughout the fiber matrix (Fig. 2B, D and F) which was observed through various images obtained at similar fields of view. It has been reported that variations in solution properties such as surface tension and solution conductivity in the presence of nanoparticles result in changes in the nanofiber diameter.30-32 In the present case, the electrospun fiber diameter increased in the presence of nanoparticles to 2.56 ± 0.16 µm (average ± standard error mean) for UCNP, 1.75 ± 0.07 µm for Pd and 4.37 ± 0.44 µm for Fe3O4 (Insets in Fig. 2G, H and I, respectively). Small fibers were chosen for TEM analysis because of the contrast problems related to thicker samples. The ability of the nanocomposites to be used as functional materials was evaluated by testing the upconverting properties, hydrogen sensing properties and magnetic properties of the, UCNP/ES-PGMA, Pd/ES-PGMA and Fe3O4/ES-PGMA fibers, respectively.
In the case of UCNP/ES-PGMA fibers, the ability to convert near-infrared excitation into visible emission was evaluated (see supporting information for methods). UCNP have been successfully used as ultrasensitive magnetic/upconversion fluorescent dual-modal molecular probes for MRI and upconversion fluorescence imaging.33-35 In the present case the UCNP/ES-PGMA fibers demonstrated excellent upconversion properties upon 974 nm laser excitation (Fig. 3A). The three major emissions were located at 521, 541, and 655 nm. Green emission from 500- 600 nm was attributed to 2H11/2 → 4I15/2 and 4S3/2 → 4I15/2 transitions, respectively, and the red emission from 635-670 nm was attributed to the 4F9/2 → 4I15/2 transition (Fig. 3A). The green to red (G/R) ratio for the fibers was 1.35:1. The UCNP/ES-PGMA composite fibers retained the upconversion signal levels to the pure NaGdF4:Yb,Er nanoparticle samples (G/R ratio = 1.37:1).
Fig. 1: (A) SEM secondary electron image of the electrospun PGMA (ES-PGMA) fibers, (B) cross-sectional image of ES-PGMA, (C) Water contact angle θ = 60° at 70°C, (D) Water contact angle θ = 15° at room temperature respectively measured on ES-PGMA -g-PNIPAM-COOH.
The Pd/ES-PGMA nanofibers were evaluated for sensing hydrogen (see Supporting Information for methods). Palladium has emerged as an important candidate for hydrogen gas sensing because of its ability to absorb high quantities of hydrogen and its highly selective response.36 Sensing herein is based on the well-established principle that palladium spontaneously absorbs H2 gas as atomic hydrogen which diffuses into the lattice to form palladium hydride, PdHx, resulting in an to phase transition and a corresponding change in the lattice spacing.37 The change in phase and lattice spacing leads to a measurable resistance change of the palladium material. However, either replacing precious noble metals with cheaper materials or alternatively development of methods that result in the reduction of material used by several orders of magnitude, especially in applications that require large amounts of material, would be beneficial. Currently, hydrogen sensing platforms are based on all-palladium constructs or hybrids with high Pd loading to stimulate an effective sensing response. Herein, using the electrospun polymer/nanoparticle nanocomposite material we demonstrate a
response is obtainable for as low as 0.6 ng of Pd dispersed across a 650 m x 900 m area over interdigitated electrodes (IDE). The ability of Pd/ES-PGMA nanofibers to sense different hydrogen concentrations (between 1 and 10% in N2 as a carrier gas) was tested38 (Fig. 3B). An increase in resistance with hydrogen gas-flow and a return to the original state in the absence of a hydrogen gas flow was observed for hydrogen concentrations (1-10%) with a response time 90 of ~14 seconds (Fig. 3B).
Finally, the magnetization properties of the Fe3O4/ES-PGMA fibers were measured by SQUID magnetometry
Fig. 2: TEM images of the (A) Upconverting nanoparticles (UCNP), (B) UCNP/ES-PGMA composite fibers, (C) Pd nanoparticles, (D) Pd/ES-PGMA composite fibers, (E) magnetite (Fe3O4) nanoparticles, (F) Fe3O4/ES-PGMA composite fibers. X-ray microanalysis spectrum obtained on: (G) UCNP/ES-PGMA composite fibers showing the presence of Gd and Yb among other elements (Inset: SEM micrograph of UCNP/ES-PGMA composite fibers): (H) Pd/ES-PGMA composite fibers showing the presence of Pd (Inset: SEM micrograph of Pd/ES-PGMA composite fibers) and (I) Fe3O4/ES-PGMA composite fibers (Inset SEM micrographs of Fe3O4/ES-PGMA composite fibers). Scale bars for images A, C and E 10 nm, for images B, D and F 1 µm and for inset images G, H and I 20 µm
Fig. 3: (A) Upconversion fluorescence spectrum of both UCNP’s (blue) and UCNP/ES-PGMA fiber composite (black) showing three main emissions green at 521 and 541 nm and red between 635 and 670 nm upon 974 nm laser excitation, (B) Current response of the Pd/ES-PGMA matrix sensor to 1-10% hydrogen gas, with alternating 4 min hydrogen and 20 min. nitrogen exposure, (C) Zero-field cooled (orange) and field cooled (blue) curves for Fe3O4/ES-PGMA composite (Inset: hysteresis loop at 5 K (pink) and 300 K (green) for Fe3O4/ES-PGMA composite) as measured by SQUID magnetometry.
(Superconducting Quantum Interference Device) (see Supporting Information). The Fe3O4/ES-PGMA fibers are superparamagnetic at room temperature with the zero field cooled/field cooled curves showing a maximum blocking temperature of 30 K (where the two curves merge) and the absence of hysteresis at 300K (Fig. 3C). The mass specific saturation magnetization, Ms of the fibers was 4.0 emu g−1. Particle loading was estimated to be ~7% by weight as determined from the Ms values of the Fe3O4 nanoparticles and Fe3O4/ES-PGMA fibers.
Conclusions In summary, we have developed a robust polymeric platform for the large scale production of electrospun nanofibers based on poly(glycidyl methacrylate) (PGMA). We have demonstrated that the epoxy groups of the polymeric matrix can be effectively used as a grafting platform for surface modifications and the polymer serves as an excellent platform to fabricate functional nanocomposites. We believe our findings presented herein will aid in the design of novel electrospun materials with tailorable surfaces for applications as scaffolds in regenerative medicine, optoelectronics, magnetic filtration and catalysis. Notes and references a School of Chemistry and Biochemistry, The University of Western Australia, WA 6009, Australia b Department of Materials Science and Engineering, Clemson University, Clemson, SC 29634, USA c School of Electrical, Electronic and Computer Engineering, The University of Western Australia, WA 6009 Australia d School of Chemistry and Chemical Engineering, Nanjing University, China e Centre for Microscopy, Characterisation and Analysis, The University of Western Australia, WA 6009 Australia f School of Physics, The University of Western Australia, WA 6009 Australia g Burn Injury Research Unit, School of Surgery, The University of Western Australia, WA 6009, Australia E-mail: [email protected], [email protected] The authors would like to acknowledge the Australian Microscopy & Microanalysis Research Facility at the Centre for Microscopy, Characterization & Analysis, The University of Western Australia, funded by the University, State and Commonwealth Governments. Authors would also like to thank Dr C.W.Evans and Dr Peter R. T. Munro for assistance with valuable experimental and result discussions. Peijun Gong is supported by The University of Western Australia and the China Scholarship Council. † Footnotes should appear here. These might include comments relevant to but not central to the matter under discussion, limited experimental and spectral data, and crystallographic data. Electronic Supplementary Information (ESI) available: [details of materials and methods]. See DOI: 10.1039/c000000x/ 1. S. Komarneni, J. Mater. Chem., 1992, 2, 1219-1230. 2. D. R. Paul and L. M. Robeson, Polymer, 2008, 49, 3187-3204.
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Intracellular Drug Delivery
DOI: 10.1002/anie.201((will be filled in by the editorial staff))
Enhancing the Efficacy of Cation-Independent Mannose 6-Phosphate Receptor Inhibitors by Intracellular Delivery** Vipul Agarwal, Priyanka Toshniwal,# Natalie E Smith,# Nicole M Smith, Binbin Li, Tristan D Clemons, Lindsay T Byrne, Foteini Hassiotou, Fiona M Wood, Mark Fear, Ben Corry* and K Swaminathan Iyer*
Abstract: Extracellular targeting of the cation-independent mannose 6-phosphate/insulin-like growth factor II (M6P/IGFII) receptors has been an attractive approach for the development of antifibrotic drugs. Several M6P analogues have been developed to regulate the activity of transforming growth factor-β1 (TGFβ1) by inhibiting its conversion from the latent to the active form. Herein, we adopt a combinatorial approach using an in vitro wound healing model and molecular dynamic simulations, to reveal that the efficacy of M6P/IGFII inhibitors can be significantly enhanced by adopting an intracellular approach. We demonstate this using systematic analysis of a bioisosteric M6P analogue, a lipophilic prodrug which upon cellular internalization undergoes ester hydrolysis to yield an
active M6P analogue to effectively downregulate collagen 1 expression in primary human dermal skin fibroblasts. In mammalian cells, the cation-independent mannose 6-phosphate/insulin-like growth factor II (M6P/IGFII) and cation-dependent mannose 6-phosphate (CD-MPR) receptors, have been identified as pivotal targets that modulate cellular response because of their role in protein trafficking. Both these receptors are functionally complimentary and can partially compensate for the absence of the other.[1] These sorting receptors play an important role of transporting M6P-bearing glycoproteins from the trans-Golgi network (TGN) to lysosomes mediated through their M6P binding sites.[2] Both receptors transport important enzymes to the intracellular acidic pre-lysosomal compartments where low pH leads to the release of the enzymes from the complex. The receptor then gets recycled into the Golgi apparatus.[3] However, only the M6P/IGFII receptor is anchored to the cell surface membrane and has been implicated in the internalization of M6P bearing compounds.[4] Importantly, it modulates the activity of a variety of extracellular M6P bearing glycoproteins including latent transforming growth factor-β (LTGFβ) precursor, urokinase-type plasminogen activator receptor, glycoprotein D of the herpes virus, granzyme B an essential factor for T cell-mediated apoptosis and proliferin.[4] This has resulted in an enormous interest in the design of M6P bearing compounds that target the M6P/IGFII receptor as it offers an efficient means for internalization of high specificity therapeutics.[5] This approach has been used to deliver therapeutic compounds in enzyme replacement therapies in lysosomal diseases like Fabry disease, aid wound healing, as a treatment for breast cancer, and to combat viral infections.[4] However, the approach suffers a major drawback as the phosphomonoester bond of M6P is prone to hydrolysis by various phosphatase enzymes.[6] This dramatically reduces its binding efficiency to the receptor thereby compromising its potency. This problem has been circumvented by
Figure 1: a) Chemical structure of mannose-6-phosphate (M6P and the two analogues, b) Schematic representation of the cLogP of three compounds
[] V. Agarwal, P. Toshniwal, Dr. N.M. Smith, Dr. T.D. Clemons, Dr. F. Hassiotou, Prof. Dr. K. S. Iyer School of Chemistry and Biochemistry, The University of Western Australia, Crawley, Western Australia, Australia E-mail: [email protected]
B. Li State Key Laboratory of Advanced Technology for Materials
Synthesis and Processing, Wuhan University of Technology, Wuhan, PR China
Dr. L. T. Byrne Centre for Microscopy, Characterization & Analysis, The University of Western Australia, Australia
Dr. N. E. Smith, Dr. B. Corry Research School of Biology, Australian National University, Canberra, Australian Capital Territory, Australia
Burn Injury Research Unit, School of Surgery, The University of Western Australia, Crawley, Western Australia, Australia
[] The authors would like to thank the Australian Research Council for funding the work, Pharmaxis Pvt Ltd for kindly providing analogues 1 and 2. The authors would also like to acknowledge the Australian Microscopy & Microanalysis Research Facility at the Centre for Microscopy, Characterization & Analysis, The University of Western Australia, funded by the University, State and Commonwealth Governments. Authors would like to thank Dr Bernard Callus, Dr Cameron Evans and Dr Megan Finch for their assistance with experimental analysis.
[#] These authors contributed equally
Supporting information for this article is available on the WWW under http://dx.doi.org/10.1002/anie.201xxxxxx.
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the design of several isosteric M6P analogues with phosphonate, carboxylate or malonate groups, which have higher affinity to the receptor and a stronger stability in human serum than M6P.[7] This approach is successful in overcoming the issues with hydrolysis of the phosphomonoester bond, yet falls short as these analogues can only target the receptors present on the cell surface. In the steady state, ~90 % of the M6P/IGFII receptors are localized in the transmembrane compartments while the remainder stays on the cell surface.[8] The receptor has a relatively long half-life (t1/2 ~ 20 hours) and recycles between the trans-Golgi network, endosomes and the plasma membrane.[9] In this communication, we report a novel approach to improve ligand-receptor protein interaction in cells whilst overcoming stability issues associated with M6P. We demonstrate this by exploring a prodrug (analogue 2) that undergoes intracellular chemical modification by esterases to yield an active M6P analogue (analogue 1) (Figure 1a),[10] resulting in a sustained and focused therapeutic strategy in an in vitro model of wound healing.
Table 1. Ligand-Receptor Protein interaction energies obtained for M6P and each of the two analogues in domain 3 and domain 5 as determined from 100 ns of molecular dynamics simulation. Two ligands were placed into the dimer binding pocket, because the receptor is secreted as a dimer.
Domain 3 Ligand-Receptor Protein interaction Energy (kcal/mol)
M6P Analogue 1 Analogue 2
Ligand 1 -368.4 -309.3 -81.6
Ligand 2 -347.4 -304.3 -79.6
Domain 5 Ligand-Receptor Protein interaction Energy (kcal/mol)
M6P Analogue 1 Analogue 2
Ligand 1 -128.2 -44.1 -43.6
Ligand 2 -118.3 -74.0 -47.8
The design of phosphonate analogue 1 is based on established principles of bioisosteric M6P analogues by replacing the P-O bond at C6 by a methylene bridge. Moreover, the replacement of the hydroxyl group at the anomeric position by an aromatic subtituent slightly improves recognition by the M6P/IGFII receptor.[11] This could be due to the hydrophobic interactions between the aromatic moiety of analogue 1 and the binding pocket of the M6P/IGFII receptor. Previous studies have demonstrated that neutral ester prodrugs are relatively benign towards enzymatic degradation, thereby altering their apparent elimination and half-life.[12] Hence analogue 2 was designed by masking analogue 1 via esterification of the phosphate group to yield a non-charged bis(pivaloyloxymethyl) (POM) derivative. Importantly, derivatization of phosphates decreases the polarity of the parent drug thereby promoting its cellular internalization and altering the elimination/distribution mechanism.[12-13] Notably, the clogP (calculated-logP evaluated using Chem Draw) for M6P, analogue 1 and 2 are -3.28, 0.10 and 3.29 respectively (Figure 1b). LogP is an estimate of a compound's overall lipophilicity, a value that influences its physiological properties such as solubility, permeability through biological membranes, hepatic clearance, and non-specific toxicity.[14] Polar compounds with low logP have very low cellular permeability due to their low affinity for the lipid bilayers. Alternatively, lipophilic compounds with high logP have high affinity for the phospholipid phase facilitating their internalization and prohibiting their escape into the aqueous basolateral side.[14] Herein the lipophilic prodrug,
analogue 2, will have improved cellular internalization compared to its charged parent analogue, 1. Once internalized the bis(pivaloyloxymethyl) linkers of analogue 2 will be gradually prone to ester hydrolysis by microsomal esterases present within the intracellular compartments,[15] resulting in the conversion to the charged parent analogue, analogue 1. Analogue 1 on the contrary would only target extracellular M6P/IGFII receptors, when administered directly, due to its low cellular permeability deemed to its low logP value. The extracellular region of the M6P/IGFII receptor is comprised of 15 repetitive domains and contains three distinct M6P binding sites located in domains 3, 5, and 9, with only domain 5 exhibiting preference for phosphodiesters.[16] In order to assess our strategy to use the intracellular conversion of the produg analogue 2 to a high receptor binding phosphonate analogue 1, it is pivotal to examine the ligand-receptor interactions to validate the hypothesis that analogue 2 will have minimal interaction with the extracellular receptors. In the current study, we used six independent molecular dynamics simulations to study the ligand-receptor protein interactions of M6P, analogues 1 and 2 with domains 3 and 5 of the extracellular M6P/IGFII receptor (see Supporting Information for experimental details, section S8.1). Domain structures were adopted from previously reported studies and two ligands were placed into the dimer binding pocket, because the receptor is secreted as a dimer.[17] Analogue 1 showed similar ligand-receptor protein interaction energies to M6P in domain 3 (Table 1). Importantly, the m-xylene ring of analogue 1 was positioned in the middle of the binding pocket further stabilizing the binding of this compound in comparison to M6P (see Supporting Information, Figures S1 and S2). This is in accordance with the previous studies of other phosphonate analogues of M6P, which are reported to display higher affinity and stronger stability in human serum than M6P.[6, 7c] The domain 5 binding pocket is larger than in domain 3, hence all the
Figure 2: Cell viability assays showing percentage of live cells in the culture post incubation with M6P, analogue 1 and 2. First and second column in each condition is representing 24 h and 72 h respectively. Data presented as average ± SEM (n=4). Significance was set at * p < 0.05 using bonferroni post-hoc test in one way ANOVA analysis.
Figure 3: Cell body area showing change in cell area post TGFβ1 stimulation and subsequent analogues treatment. Cell area was measured from the fluorescent images of live cells taken for viability assay (cells from minimum 40 images per group were measured). Significant increase in cell body area was observed for cells treated with TGFβ1 (2 ng/mL), however no such increase was observed in cells treated with analogues +/- TGFβ1 (2 ng/mL). Data presented as average ± SEM (n > 40). Significance was set at * p < 0.05 using bonferroni test in one way ANOVA.
3
compounds displayed weaker interactions with the receptor and occupied more diverse positions in domain 5 due to the increased space (see Supporting Information, Figure S2). Furthermore, in the case of analogue 1 in domain 5, the simulations suggested that one of the two analogue 1 ligands (ligand 1) bound to the protein dimer has weaker interactions with the protein as it primarily interacts with the second molecule of analogue 1 (ligand 2). Overall, the simulations suggested that analogue 1 has high affinity towards domain 3 similar to M6P whilst the prodrug 2 has weak interactions with both domains of the receptor (Table 1 and see Supporting Information, Figure S3). The molecular dynamics simulations further validated our aforementioned hypothesis that the prodrug will be internalized with minimal extracellular receptor-ligand interactions. We next validated our hypothesis in a well-established in vitro model for wound healing using primary human dermal skin fibroblasts (HDF). In mammals, wound healing is not a regenerative process that restores normal tissue architecture, but a reparative process that results in scar formation.[18] This process occurs in all tissues of the body in response to physical, chemical and biological stressors. Scar tissue is functionally and aesthetically inferior to normal tissue. It is a result of the excessive production of extracellular matrix (ECM) that occurs after injury.[19] One of the most important proteins influencing the ECM architecture during wound healing is collagen I. Collagen I is synthesized predominantly by fibroblasts and its synthesis is largely regulated by cytokine transforming growth factor β1 (TGFβ1).[20] TGFβ1 is secreted in an inactive form (LTGFβ1), requiring enzymatic conversion to active TGFβ1 to effect a change in cell function. One of the methods of TGFβ1 activation involves binding of M6P residues within the N-linked oligosaccharides on latent TGFβ1 to the M6P/IGFII receptor.[21] Since the M6P binding sites are involved in various steps of TGFβ1 activation and inactivation, it is believed that small molecule inhibitors that block the binding of M6P residues could present an opportunity to block the activity of TGFβ thereby reducing overproduction of an important profibrotic extracellular matrix protein collagen I. Cytotoxicity and cell viability of the analogues were initially assessed using MTS and live/dead assays (see experimental details in Supporting Information, sections S2.1, S3.1 and S4.1). Previous studies characterizing M6P binding affinity towards the M6P/IGFII receptor reported significant binding affinity at a concentration of 10 µM.[7b, 22] This concentration was therefore selected for our in vitro studies. All compounds showed no effect on cell viability and proliferation both in the presence and absence of TGFβ1 (Figure 2 and see Supporting Information, Figure S4 respectively). Exposure to TGFβ1 in the absence of analogues 1 and 2 resulted in a reduction in HDF proliferation (see Supporting Information, Figure S4). This growth suppressive response has been previously reported in many cell types.[23] The observed change in cell proliferation upon exposure to TGFβ1 influenced HDF cell morphology (and see Supporting Information, Figure S5b). Fibroblasts alter their morphology from dendritic to stellate upon exposure to various external cues caused by changes in actin polarisation and focal adhesion.[24] TGFβ1 has been shown to alter the morphology of many cell types including fibroblasts, potentially by inducing polymerisation of the actin cytoskeleton from globular to filamentous.[24a] Different factors such as cell motility and mechanical strain have also been reported to cause this alteration.[25] In the present case, we observed a reversal of HDF cell morphology back to initial cell morphology without TGFβ1 stimulation when treated with the analogues 1 and 2 (Figure 3 for quantification of cell body area and see Supporting Information, Figure S5c-e for
images). We next assessed if the observed change in morphology is correlated to collagen I gene expression using qRT-PCR, and if changes in collagen I gene expression could be altered by inhibition of TGFβ1 activity by targeting the M6P/IGFII receptor in the presence of the analogues (refer to Supporting Information for method, section S5.1). Indeed as previously reported, exposure of HDF to TGFβ1 (2 ng/mL) resulted in a significant increase in collagen I mRNA expression at 48 hours post-stimulation.[20, 26] It is noteworthy that although collagen I gene expression was upregulated throughout the study period (72 hours), the optimal response was observed after 48 hours exposure to TGFβ1 (see Supporting Information, Figure S6). Therefore, the efficacy of the aformentioned compounds was assessed in the presence of TGFβ1 at 48 hours. TGFβ1 induced collagen I mRNA expression was downregulated significantly (p < 0.05) with the addition of prodrug analogue 2 (10 µM) with levels returning to that of normal untreated cells (Figure 4a). Downregulation was also observed for M6P however, the change did not reach stastistical significance (Figure 4a). This suggests that the variable responses that have been reported in the use of hydrolytically unstable M6P may be due to its realtive instability and that the development of stable analogues may resolve this issue. Importantly, in the present case we observed no significant change in collagen I gene expression in HDF cells treated with analogue 1 (Figure 4a). This was expected given the low cellular permeability which is believed to affect the ligand-receptor protein interactions in cells. Next, we investigated if the observed change at transcription level would have a corresponding influence on protein translation. Changes in collagen I protein expression were quantified using immunoblotting (refer to Supporting Information for method, section S6.1). All protein expression studies were carried out at 72 h post-stimulation. Significant upregulation in collagen I protein expression was observed post TGFβ1 stimulation (Figure 4b; column 2; p < 0.05) which is consistent with previous reports.[27] Analogue 2 (10 µM) was observed to reduce TGFβ1
Figure 4: Change in Collagen 1 a) mRNA levels and b) protein levels post TGFβ1 stimulation in the presence and absennce of M6P, analogues 1 and 2 compared to untreated (negative) control. Collagen I protein expression was normalised against β-actin levels. Data are presented as average ± SEM (n = 3). Significance was set at * p < 0.05 using bonferroni post-hoc test in one way ANOVA analysis
4
mediated upregulation of collagen I protein to non-stimulated levels (Figure 4b; column 5; p < 0.05). No significant changes were observed in the case of M6P or analogue 1. This further confirms that analogue 2 is a potent repressor of TGFβ1 induced collagen I synthesis and thus can ameliorate the profibrotic effects of TGFβ1 in human skin dermal fibroblasts. In summary, we have developed a novel approach using an intracellular prodrug of M6P, analogue 2, to target M6P receptors. This approach overcomes the physiological problems associated with the hydrolysis of M6P whilst successfully targeting the receptors using an intracellular coversion of the analogue. We believe that this approach of intracellular drug coversion for receptor targeting will have far reaching implications in the design of highly potent drug candidates for enzyme replacement therapies of lysosomal storage diseases, to aid wound healing and in cancer therapy.
Received: ((will be filled in by the editorial staff)) Published online on ((will be filled in by the editorial staff))
Keywords: drug development • intracellular drug delivery • Mannose-6-Phosphate analogue
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Entry for the Table of Contents (Please choose one layout) Layout 2:
Intracellular Drug Delivery
Vipul Agarwal, Priyanka Toshniwal,# Natalie E Smith,# Nicole M Smith, Binbin Li, Tristan D Clemons, Lindsay T Byrne, Foteini Hassiotou, Fiona M Wood, Mark Fear, Ben Corry* and K Swaminathan Iyer* __________ Page – Page
Enhancing the Efficacy of Cation-Independent Mannose 6-Phosphate Receptor Inhibitors by Intracellular Delivery Intracellular delivery of M6P/IGFII receptor inhibitors exhibits better efficacy than
extracellular inhibitors to regulate TGFβ1 mediated upregulation of profibrotic marker, collagen I.
Inhibiting the activation of transforming growth factor-β using a polymeric nanofiber scaffold
Vipul Agarwal,a Fiona M. Wood,b,c Mark Fearb and K. Swaminathan Iyer*a
Electrospun poly(glycidyl-methacrylate) (PGMA) nanofibers were fabricated in the presence of a hydrophobic analogue of mannose-6-phosphate, (PXS64). The nanofibers were tested for biocompatibility as a tissue engineering scaffold and for their efficacy to inhibit transforming growth factor β1 (TGFβ1) activation in human dermal skin fibroblasts.
The principle of tissue engineering involves the development of advanced functional biomaterials that incorporate biochemical cues to aid in the regeneration of tissues to restore and maintain normal organ function.1 This approach has resulted in the fabrication of several advanced biomaterial platforms that have been used to repair damaged or diseased tissues and to create therapeutic approaches for entire tissue replacement.2-4 While tissue regeneration has successfully been demonstrated in the presence of biocompatible scaffolds, scarring remains one of the unresolved issues. In mammals postnatal wound healing results in scar formation, characterised by excessive collagen deposition and dysfunctional extracellular matrix formation. This is also a hallmark of fibrotic disease which occurs in many tissues. Central to wound repair is transforming growth factor β1 (TGFβ1), a cytokine secreted by several different cell types involved in wound healing.5 In the case of skin, scarring during the wound healing process is a result of TGFβ1 mediated imbalance in the fibroblast activity resulting in an architecturally disorganised extracellular matrix.6 One of the most important proteins influencing the ECM architecture during wound healing is collagen I. Collagen I is synthesized predominantly by fibroblasts and its synthesis is largely regulated by TGFβ1 signalling.7 Following wound healing in skin, a scar is not only aesthetically and psychologically detrimental but can also cause functional disability and pain.8 Importantly, inhibition of TGFβ1 activity has been documented to reduce scar formation9 whereas subcutaneous delivery of exogenous TGFβ1 in newborn mice was shown to promote fibrosis and angiogenesis at the site of injection.10 During wound healing TGFβ1 activation from its latent state involves binding of the mannose 6-phosphate (M6P) residues within
the N-linked oligosaccharides on the latent TGFβ to the M6P/IGFII receptor.11 Since the M6P binding sites are involved in various steps of the TGFβ1 activation and inactivation route, it is believed that the presence of small molecule analogues of M6P that competitively bind to the receptor could present an opportunity to block the activation of TGFβ1 thereby reducing overproduction of extracellular matrix protein collagen and potentially reducing scarring.12 The major drawback with using M6P as a drug in the reduction of scarring is that the phosphomonoester bond of M6P is prone to hydrolysis by various phosphatase enzymes.13 Isosteric M6P analogues with phosphonate, carboxylate or malonate groups have been shown to circumvent the aforementioned issues and have been reported to have greater stability in human serum than M6P.14-17 The analogue PXS64 reported in the present study is a lipophilic bioisosteric phosphonate analogue developed by [(bis(pivaloyloxymethyl)) (POM)] ester derivatization of M6P. Importantly, the high lipophilicity of PXS64 limits its solubility in aqueous solutions thereby reducing its bioavailability.18 In this communication we report that PXS64 can be incorporated in an electrospun poly(glycidyl methacrylate) (PGMA) nanofibrous scaffold. Furthermore, we demonstrate the utility of this scaffold as a biomaterial platform for wound healing using human dermal skin fibroblasts (HDF). We demonstrate its effectiveness as a drug delivery platform to mitigate TGFβ1 mediated upregulation of collagen I. Electrospinning is a widely used technique to fabricate large area nanofibrous scaffolds19 mimicking the architecture of extracellular matrix (ECM).20 They have been used as tissue engineering scaffolds to promote cell growth and migration and to achieve controlled delivery of drugs and growth factors.21 They have been widely used in skin, bone, cartilage, vascular and neural tissue engineering.22 For example, Yang et al. employed emulsion electrospinning to fabricate ultrafine core sheath poly(ethylene glycol)-poly(D,L-lactide) fibers loaded with basic fibroblast growth factor (bFGF) and demonstrated its gradual release over 4 weeks.23 In vitro studies on mouse embryonic fibroblasts showed enhanced cell adhesion, proliferation and secretion of ECM when cultured on the nanofibrous scaffold.24
In the in vivo studies on dorsal wound model in diabetic rats, bFGF/ poly(ethylene glycol)-poly(D,L-lactide) mats showed elevated healing with complete re-epithelialization and regeneration of skin appendages such as hair and sebum, while bFGF promoted collagen deposition and its remodelling similar to normal architecture.23 In the case of electrospun scaffolds the ability to modify the surface properties of the nanofibers such as adhesion, wettability and biocompatibility is pivotal in its integration as a tissue engineering platform. Currently one of the biggest challenges using this technique to produce scaffolds is that the polymers used thus far to develop nanofibers need specific approaches for surface modification that are complex and laborious.25 The achievement of a certain degree of grafting universality requires the establishment of a controlled method of introducing the desired functional groups. Currently, this is achieved by physisorption.26 In contrast, chemisorption which is difficult to achieve on polymeric nanofibers would result in covalent attachment. Polymers containing epoxy groups are examples of functional polymers that are able to react with a wide range of biologically relevant molecules through
‘‘grafting to’’ interactions mediated by the epoxy groups.27, 28 To this end, poly(glycidyl methacrylate) (PGMA) used in the current study, has an epoxy group in every repeating unit and has been used extensively as a macromolecular anchoring layer.29, 30 In the present study, PGMA was electrospun in the presence of PXS64 and in the absence of PXS64 as a control (see supporting information for experimental conditions). PXS64 loading in the electrospun PGMA fibers was measured using high pressure liquid chromatography (HPLC) to be ~76 % w/w. Electrospun fibrous scaffolds were characterized using scanning electron microscopy (SEM). Fiber diameter was measured from the SEM images to be 0.69 ± 0.31 µm (average ± standard deviation) for ES-PGMA and 2.24 ± 1.06 µm in the case of ES-PGMA + PXS64 (Figure 1). Biocompatibility of the scaffold was investigated using the colorimetric MTS assay and live/dead cell viability assay (see Supporting Information for method) and were imaged using fluorescence and scanning electron microscopy.31 Human dermal fibroblasts (HDF) cells were cultured on the scaffolds and incubated
Figure 1: SEM secondary electron image of electrospun fibers of a) PGMA, b) PGMA + PXS64, C) showing the chemical structure of PXS64. Scale bars: a) and b) 10 µm.
Figure 2: a) Cell viability assay showing percentage of live cells in culture post incubation on ES-PGMA and ES-PGMA + PXS64, in the presence and absence of TGFβ1 (2 ng/mL), b) and c) showing HDF cell morphology on ES-PGMA + PXS64 + TGFβ1 (2 ng/mL) using fluorescent microscopy and scanning electron microscopy respectively. Red arrows highlighting the cell attachment and adhere points on the fibers. Scale bars: b) and c) 2 µm.
Figure 3: Collagen I gene expression analysis showing the percentage change in gene expression as compared to non-treated negative control (column 1). HDF cells incubated on both scaffolds and plastic tissue culture plate both in presence and absence of TGFβ1 and release media. Data presented as average ± SEM (n = 3). Significance was set at * p < 0.05 using bonferroni post-hoc test in one way ANOVA analysis.
both with and without TGFβ1 over a period of 72 hours. No significant changes were observed on cell proliferation (see Supporting Information; Figure S1) or cell viability (Figure 2a) on both scaffolds with or without TGFβ1. To study the cell-matrix interactions, scaffolds were incubated with HDF cells and imaged using fluorescence and electron microscopy (see Supporting Information for method). HDF cells were observed to adopt their characteristic spindle shape morphology on both ES-PGMA and ES-PGMA + PXS64 scaffolds in the presence and absence of TGFβ1 (Figure 2b, c and see Supporting Information; Figure S2 and S3). Finally, the ability of ES-PGMA + PXS64 scaffold in regulating the over expression of collagen I gene in HDF cells post TGFβ1 stimulation was evaluated using RT-qPCR (see Supporting Information). HDF cells cultured on the biocompatible ES-PGMA control scaffold showed significant increase in collagen I gene expression in the presence of TGFβ1 (Figure 3). Cells incubated on ES-PGMA + PXS64 scaffold under similar conditions showed reduced expression of collagen I gene compared to ES-PGMA scaffold, which reached significance in the presence of supplemented release media. The use of DMSO in release media was adopted from previously reported methodology32 to accelerate the release of hydrophobic drugs in vitro and was shown to have no effect on collagen expression alone (Figure 3). Here we developed a PGMA scaffold and encapsulated an anti-scarring drug, PXS64. The biocompatibility of the scaffold was demonstrated in human dermal skin fibroblasts. Finally, the efficacy of scaffold and drug in mitigating the increased expression of collagen I in response to TGFβ1 stimulation was also demonstrated. We believe that this potential proof of principle approach can easily be adapted in the design of scaffolds for tissue regeneration in the presence of other anti-fibrotic agents. The authors would like to thank the Australian Research Council for funding the work. The authors would also like to acknowledge the Australian Microscopy & Microanalysis Research Facility at the Centre for Microscopy, Characterization & Analysis, The University of Western Australia, funded by the University, State and Commonwealth Governments. The authors would like to thank Dr Foteini Hassiotou to kindly provide the RT-qPCR instrument and Pharmaxis Ltd, Sydney for providing PXS64. MF is supported by Chevron Australia. This work was partly funded by the Fiona Wood Foundation. Notes and references a School of Chemistry and Biochemistry, The University of Western Australia, Crawley, Western Australia, Australia. E-mail: [email protected] b Burn Injury Research Unit, School of Surgery, The University of Western Australia, Crawley, Western Australia, Australia. c Burns Service Western Australia, Department of Health, Perth, Western Australia, Australia † Footnotes should appear here. These might include comments relevant to but not central to the matter under discussion, limited experimental and spectral data, and crystallographic data. Electronic Supplementary Information (ESI) available: [details of any supplementary information available should be included here]. See DOI: 10.1039/c000000x/ 1. T. Dvir, B. P. Timko, D. S. Kohane and R. Langer, Nat.
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2. Agarwal, V., Toshniwal, P., Smith N. E., Smith, N. M., Li, B., Clemons, T. D.,
Byrne, L. T., Hassiotou, F., Wood, F. M., Fear, M., Corry, B., Iyer, K.S., Intracellular
Enhancing the Efficacy of Cation-Independent Mannose 6-Phosphate Receptor Inhibitors
by Intracellular Delivery, Angewandte Chemie International Edition (submitted)
3. Agarwal, V., Wood, F. M., Fear, M. and Iyer, K. S., Inhibiting the activation of
transforming growth factor-β using a polymeric nanofiber scaffold, Nanoscale (Submitted)
80
Supporting Information
A Functional Reactive Polymer Nanofiber Matrix
Vipul Agarwal,a Dominic Ho,a Diwei Ho,a Yuriy Galabura,b Faizah M.D. Yasin,a Peijun Gong,c Weike Ye,d Ruhani Singh,a Alaa Munshi,a Martin Saunders,e Robert C. Woodward,f Timothy St. Pierre,f Fiona M. Wood,g Mark Fear,g Dirk Lorenser,c David D. Sampson,c, e Bogdan Zdyrkob, Nicole M. Smith,a Igor Luzinovb, *, and K. Swaminathan Iyera,*
Materials
Polyglycidyl methacrylate (PGMA) with Mn = 220515 and Mw = 433730 was synthesized
by radical polymerisation as reported previously,1 Carboxy terminated N-isopropyl
The magnetic properties of the dried magnetite/ES-PGMA composite (88.2 mg) were
measured using a SQUID magnetometer (Quantum Design 7 Tesla MPMS) operating
between 5K and 300K. Magnetite nanoparticles (22.5 mg) were lyophilised prior to their
measurement and their saturation magnetisation was recorded at 70 kOe at 5K. The zero
field cooled and field cooled measurements were conducted in a field of 0.1 kOe.
Contact Angle:
Static contact angles of MilliQ water on the surface of the electrospun polymer matrix
were measured using a home-built goniometer with Rame- Hart scope attachment.7 The
polymer was electrospun on the 12 mm glass coverslips (Cat. # G401-12, ProSciTech)
both with and without nanoparticles (refer PNIPAM attachment on ES-PGMA). 5 µL of
water was pipetted onto the membrane and images were taken after the drop edges came to
rest (~2 min) using a Canon EOS450D and the angle was measured using the calibrated
85
Rame-Hart scope. Measurements were carried out at room temperature and again at 40 °C.
Images were processed using ImageJ software. Measurements were done in duplicates and
reported as average ± standard deviation.
NIR Room Temperature Emission Spectroscopy (λexc
= 974 nm):
Upconversion spectra measurements were carried out as previously described.8 Briefly,
upconverting particles were suspended in chloroform and were air dried on glass slides.
The upconversion spectra were obtained with an optical set up incorporating a 980 nm
laser diode. The peak wavelength of the laser diode is 974.5 nm. The optical excitation
intensity for obtaining the spectrum shown in Fig. 3A was 7665 W/mm2
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Am. Chem. Soc., 2003, 126, 273-279.
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87
Enhancing the Efficacy of Cation-Independent Mannose 6-Phosphate Receptor Inhibitors by Intracellular Delivery** Vipul Agarwal, Priyanka Toshniwal,# Natalie E Smith,# Nicole M Smith, Binbin Li, Tristan D Clemons, Lindsay T Byrne, Foteini Hassiotou, Fiona M Wood, Mark Fear, Ben Corry* and K Swaminathan Iyer*
Supporting Information
S1.1 Cell culture
Human primary dermal fibroblast cell cultures from normal skin were cultured in
Dulbecco’s Modified Eagle’s Medium (DMEM/F12 - GlutaMAX; Invitrogen Gibco)
supplemented with 10 % fetal bovine serum (FBS; Invitrogen Gibco) and 1%
penicillin/streptomycin (Invitrogen Gibco). Analogues 1 and 2 were provided by
Pharmaxis Ltd after stringent QC analysis. M6P and analogue 1 were dissolved in
PBS filter sterilized, aliquoted and stored at -20 °C. Each aliquot was used for
maximum 2 freeze thaw cycles. Analogue 2 was dissolved in DMSO and treated the
similar way as other analogues. The cells were incubated at 37 °C in a humidified
atmosphere of 5 % CO2. All experiments were carried out with cells between
passages 3-6.
S2.1 Cell Viability
Cell viability was determined using a LIVE/DEAD viability/cytotoxicity Kit
(Invitrogen, UK) which measures the membrane integrity of cells, as per
manufacturer’s protocol. In brief, 20000 cells were seeded in each well in a 24 well
plate and incubated with analogues at 10 µM concentration in cell culture media
(DMEM F-12 containing 10% FBS and 1% Penicillin/ Streptomycin) for 30 min
(media was added in case of controls). Following which human recombinant TGFβ1
88
(2 ng/mL) (Cat.# 14-8348-62, eBioscience) was added to the wells (fresh media was
added in controls) and plates were incubated for 24 h and 72 h in the humidified
incubator at 37 °C with 5 % CO2. At the stipulated time (24 h and 72 h), cells were
washed with PBS (3 times) and then stained with calcein (100 µL, 1 µM)/ ethidium
bromide (100 µL, 2 µM) in PBS and incubated in the humidified incubator at 37 °C
and 5 % CO2 for 30 min. Images were captured using an Olympus IX71 inverted
microscope with a 20 x objective with fixed exposure time. Both live and dead cells
were counted using Image J software (NIH) with cell counter plug in. Data presented
as mean ± standard error mean (n = 4).
S3.1 Cell Body Area
Cell size was measured using Image J software (NIH).[1] Minimum of 40 cells were
randomly selected from the fluorescence images and their area was measured.
Values reported as mean ± standard error mean.
S4.1 Cell Proliferation
Cell proliferation was measured using the MTS assay (Cell Titer 96 ®Aqueous,
Promega, Madison, USA) as per the manufacturer’s protocol. Briefly, 1500 cells
were seeded in each well of a 96 well plate and treated with analogues at 10 µM
concentration in cell culture media (DMEM F-12 containing 10 % FBS and 1 %
Penicillin/ Streptomycin) for 30 min (media was added in case of controls) prior to
the addition of human recombinant TGFβ1 (2 ng/mL) to each well (fresh media was
added in controls) and the plates (individual for each time point) were incubated for
72 h in the humidified incubator at 37 °C with 5 % CO2. MTS solution (40 µL) was
added in each well the next day, and was considered as 0 h time point, and incubated
for 3 h in the humidified incubator at 37 °C with 5 % CO2. Following which 80 µL
89
from each well was transferred into a new 96 well plates and read under a plate
reader at 490 nm excitation wavelength. Same protocol was followed at every time
point for next 72 h. Data presented as mean ± standard error mean (n = 5).
S5.1 Gene Expression
Gene expression was measured using real time quantitative polymerase chain
reaction. Cells (50000) were seeded in 24 well plates and incubated for 24 h in the
humidified incubator at 37 °C and 5% CO2. Next day culture media (10 % FBS) was
replaced with starve media (0.1 % FBS) and incubated for further 24 h in the
incubator to bring all the cells under same physiological cycle. Next day, cells were
treated with required concentration of M6P analogue 30 min prior to TGFβ1 (2
ng/mL in starve media) stimulation and further incubated for 48 h in the humidified
incubator at 37 °C and 5% CO2. mRNA was extracted using RNeasy mini kit®
according to manufacturers’ protocol (Qiagen GmbH). For reverse transcription 1.5
µg of total mRNA was converted to cDNA using Superscript VILO (Cat.# 11754,
Applied Biosystems) according to manufacturers’ protocol. 150 ng of cDNA was
analysed by ABI 7500 fast analysis real-time PCR system using TaqMan® master
mix and col1a1 probes (Hs01076777_m1, Life Technologies). GAPDH was used as
a reference gene (Cat.# 4326317E, Life Technologies). Analysis was carried out
using the instrument software. Data presented as mean ± standard error mean (n =
3).
S6.1 Protein Expression
90
Protein expression was measured using western blotting. Cells (1 x 105) were seeded
in each well of a 6 well plate and incubated for 24 h in the humidified incubator at
37 °C and 5% CO2. Next day culture media (10 % FBS) was replaced with starve
media (0.1 % FBS) and incubated for further 24 h in the incubator to bring all the
cells under same physiological cycle. Next day, cells were treated with required
concentration of M6P analogue 30 min prior to TGFβ1 (2 ng/mL in starve media)
stimulation and further incubated for 72 h in the humidified incubator at 37 °C and
5% CO2. Whole cell lysates were prepared from treated cells. 35 µg of protein was
denatured and subjected to SDS-PAGE, and transferred to nitrocellulose membrane
(Cat.# 10600007, Amersham, General Healthcare Lifesciences) by standard transfer.
Post transfer, membrane was blocked with 5% skim milk/0.1% TBST for 30 min at
room temperature, then incubated overnight with rabbit anti-human collagen I
antibody (1:2000 in 5% skim milk/0.1% TBST, Cat. # NB600-408, Novus) at 4 °C.
Membranes were washed with 0.1% TBST and incubated with peroxide conjugated
mouse anti-rabbit (1:5000 in 5% skim milk/0.1% TBST, Cat.# NA934VS, GE
Healthcare Lifesciences) for 1 h in 5% skim milk/0.1% TBST at room temperature.
Immunoreactivity was detected using the chemiluminescent HRP substrate (Cat.#
WBKLS0100, Millipore IMMobilon) and the signal was captured with the
Chemidoc (BioRad, Model #731BR02144) and analysed using ImageJ software
(NIH). To confirm equality of protein loading, all membranes were stripped and
reanalysed for β-actin expression using 1° antibody (1:50000 in 5% skim milk/0.1%
TBST, Cat.# A1978, Sigma) and 2° (1:5000 in 5% skim milk/0.1% TBST, Cat.#
NA9310V, GE Healthcare Lifesciences). Data presented as mean ± standard error
mean (n = 3).
91
S7.1 Statistics
The results for cell viability, cell body area, cell proliferation, gene and protein
expression are expressed as mean ± standard error mean (SEM) and analysed for
analysis of variance (ANOVA). Significance was evaluated using Bonferroni and
Turkey’s post-hoc analysis and set at 95% confidence (p < 0.05).
S8.1 Simulation Systems:
Experimental Methods:
6 simulation systems of the M6P/IGFII receptor were investigated; these included
the domain 3 and domain 5 dimers with a ligand in each binding site (2 binding sites
per dimer). These ligands were M6P, analogues 1 and 2. The coordinates for the
domain 3 dimer with M6P bound (pdb accession code 1SYO)[2] and the domain 5
monomer (pdb accession code 2KVB)[3] were obtained from the protein database. In
order to obtain the dimer for domain 5 and position M6P in the binding pocket, the
domain 5 beta sheet regions were aligned with the corresponding beta sheet regions
of domain 3. Analogue 1 and 2 were positioned in each binding pocket by aligning
the mannose ring and phosphate group to the M6P coordinates obtained for domain
3.[2] Each system was then solvated in a TIP3P water box of dimensions 65 x 60 x
114 Å and ionised with 150 mM KCl. All simulations were run with periodic
boundary conditions, constant temperature (310 K) maintained using Langevin
dynamics and constant pressure (1 atm) maintained with a Langevin piston, and the
particle mesh Ewald method was used to compute full system electrostatics.[4] The
CHARMM 36 force field was used for protein, water and M6P parameters.[5] The
ion parameters were obtained from Joung and Cheatham.[6] Missing parameters for
analogues 1 and 2 were obtained using ab-initio techniques [7] with the program
92
Gaussian 09.[8] All molecular dynamics simulations were run with the program
NAMD[9] using rigid bonds to hydrogen and 2 fs time steps. Molecular graphics
were generated using VMD.[10]
Water and ions were energy minimized for 5000 steps and equilibrated for 20 ps
with the protein and substrate held fixed. A harmonic restraint with a force constant
of 20 kcal/mol was then applied to the backbone atoms of the protein and on each
ligand and the system was minimized for a further 10,000 steps prior to 500 ps of
equilibration. This step was repeated with gradual reductions in the force constant
with values of 10, 5, 2.5 and 1 kcal/mol. Finally, to replicate the influence of the
surrounding protein domains on the individual domain being simulated, a harmonic
restraint with a force constant of 0.1 kcal/mol was placed on all of the protein Cα
atoms which were located more than 10 Å from the binding pocket in order to
ensure no loss of secondary structure throughout the simulation. The system was
then minimized for a further 10,000 steps prior to 10 ns of equilibration.
Subsequently 100 ns of equilibrium simulation were obtained for each of the 6
systems.
Data Analysis:
Cluster analysis was performed on the final 100 ns of equilibrium simulation for
each ligand in order to determine the most occupied binding positions. Each ligand
was clustered according to the RMSD of its coordinates with a cut-off of 3 Å.
Subsequently the NAMDEnergy plugin of VMD was utilized to determine the
interaction energy of each ligand with the protein for the entire time and for the most
populated clusters.
93
Figure S1: Most populated binding positions (clusters) of each drug in the domain 3 and domain 5 binding domains as determined from 100 ns of molecular dynamics simulation. In each case the protein is shown in grey, and the most to the least populated clusters are for ligand 1 (initially bound to dimer 1): cyan, pink, mauve, white and green and for ligand 2 (initially bound to dimer 2): dark blue, red, orange, yellow and green. a) M6P in domain 3: Ligands 1 and 2 both remain in the vicinity of the binding site determined in the crystal structure. However as there are multiple hydrogen bond acceptors and donors, both ligands sample multiple positions as the mannose ring hydroxyl groups form hydrogen bonds with various residues and these change over the 100 ns simulation. b) Analogue 1 in domain 3: Ligand 1 has only one binding position (cyan) and ligand 2 has only two clusters indicating that in both cases it binds stably to the protein and remains in the binding pocket throughout the simulation. This binding appears to be stabilized by the interaction between the m-xylene rings of the 2 ligands. c) Analogue 2 Domain 3: Both ligand 1 and 2 are oriented such that the benzyl groups associate in the center of the dimerization domain, hence while the ligands do have more than one binding position they remain in the vicinity of the predicted binding site. d) Domain 5 M6P: In this case the ligands occupy positions which are not asymmetrical, while ligand 1 favors a horizontal orientation with one major binding position (cyan) ligand 2 favors a vertical orientation where it has more flexibility to sample new positions. e) Analogue 1 in domain 5: Ligand 1 moves away from its initial position in the binding site and occupies a position where it is in close contact with ligand 2. f) Analogue 2 in domain 5: In this case both ligands remain in the vicinity of the original binding position. However, while ligand 1 has one major cluster implying its motion is restricted, ligand 2 has more flexibility to move occupying multiple clusters.
94
Figure S2: Snapshots from 100 ns molecular dynamics simulations representative of the most heavily occupied binding positions for each ligand in the domain 3 and domain 5 dimers. The two protein subunits are shown in pink and grey respectively and residues which form common hydrogen bonds are labelled. a) M6P and b) Analogue 1 in domain 3: Ligand 1 and 2 form the strongest energetic interactions with Lys350, Lys358, Ser386 and Arg391. c) Analogue 2 in domain 3: Ligand 1 and 2 form the strongest interactions with Gln348, Arg391 and Glu416. d) M6P in domain 5: Ligand 1 and 2 are oriented differently to each other in the binding pocket. M6P forms its strongest interactions with Arg687. e) Analogue 1 in domain 5: interacts most favorably with Asn680 andArg687. f) Analogue 2 in domain 5: Ligand 1 and 2 form the strongest interactions with Gln644, Trp653, Arg687 and Tyr714.
Figure S3: Average interaction energies obtained for the most occupied positions of each ligand in domains 3 and 5. R391 in domain 3 is equivalent to R687 in domain 5. M6P (a) and analogue 1 (b) in domain 3 both show similar interactions for each ligand and the protein subunit it is most closely associated with (for example Protein 1 with Ligand 1 as compared to Ligand 2 with Protein 2). This is because the binding mode is similar for each ligand in its respective binding site (Figure S2). Similarly, in each case the ligands also interact closely with Lys350 which is actually located on the alternate protein subunit (for example Ligand 1 to Protein 2). When compared to (a) and (b) analogue 2 (c) in domain 3 shows significant decreases in the interaction energies which is reflected by its lower overall interaction energy. M6P (d) and analogue 1 (e) in domain 5 have two major interactions the first one to Arg687 and the second to Glu709. Overall the interaction energies for domain 5 are much lower than those observed for domain 3. Analogue 2 (f) in domain 5 shows a similar pattern with a further reduction in the observed interaction energies.
95
Figure S4: Cell Proliferation assay showing cell growth over the period of 72 h post incubation with analogues both in the presence and absence of TGFβ1. First and second column in each condition is representing 24 h and 72 h respectively. Significant proliferation was observed in all groups despite the reduction in proliferation in TGFβ1 treated groups. Data presented as Mean ± SEM. Significance was set at * p < 0.05 using one way ANOVA with Bonferroni post-hoc analysis.
Figure S5: HDF cell morphology post calcein AM staining imaged using fluorescent microscopy. Cells were treated for 72h, stained and imaged: a) untreated (control), b) TGFβ1 (2 ng/mL) treatment, c) M6P (10 µM) + TGFβ1 (2 ng/mL), d) Analogue 1 (10 µM) + TGFβ1 (2 ng/mL) and e) Analogue 2 (10 µM) + TGFβ1 (2 ng/mL). Scale bar 2 µm.
Figure S6: Collagen I gene time response curve post TGFβ1 (2 ng/mL) stimulation. Collagen I gene expression was significantly upregulated at 24 and 48 h post TGFβ1 stimulation compared to non-treated control. The 48 h stimulation was selected for all further experiments as it yielded significantly higher response. Data presented as average ± SEM (n = 3). Significance was set at * p < 0.05 using bonferroni post-hoc test in one way ANOVA analysis.
96
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W. E. Reiher, B. Roux, M. Schlenkrich, J. C. Smith, R. Stote, J. Straub, M.
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[6] I. S. Joung, T. E. Cheatham, 3rd, J. Phys. Chem. B 2008, 112, 9020-9041.
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Cheeseman, G. Scalmani, V. Barone, B. Mennucci, G. A. Petersson, H. Nakatsuji,
M. Caricato, X. Li, H. P. Hratchian, A. F. Izmaylov, J. Bloino, G. Zheng, J. L.
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Tomasi, M. Cossi, N. Rega, J. M. Millam, M. Klene, J. E. Knox, J. B. Cross, V.
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98
Inhibiting the activation of transforming growth factor-β using a polymeric nanofiber scaffold Vipul Agarwal,a Fiona M. Wood,b,c Mark Fearb and K. Swaminathan Iyer*a
Supporting Information
S1. Materials
Polyglycidyl methacrylate (PGMA) with Mn = 220515 and Mw = 433730 was synthesized by radical
polymerisation as reported previously.1 MEK was purchased from Merck. PXS64 was kindly provided by
Pharmaxis Ltd after intensive QC testing.
S2. Electrospinning Procedure:
PGMA (15 wt%) was dissolved in MEK overnight at room temperature with constant stirring. In case of PGMA
+ PXS64, PXS64 (2.95 mg) was mixed with PGMA and dissolved in MEK overnight at room temperature.
PGMA polymer and PGMA + PXS64 fibrous scaffold were obtained via electrospinning (Nanofiber
Electrospinning Unit, Cat. # NEU-010, Kato Tech, Japan) onto 12 mm and 6 mm glass coverslips (Cat. # G401-
12, and G401-06 respectively, ProSciTech). The electrospinning parameters after optimisation were a voltage of
9.1 kV, working distance of 9 cm, syringe pump speed of 0.04 mm/min (1 mL/h), and the traverse and collection
drum speeds were set at 0 cm/min and 0 m/min respectively. Fibers were dried and crosslinked by annealing at
80 °C for 5 h and stored at room temperature for maximum of a month. Scaffolds were UV sterilised in the tissue
culture hood for 15 min and further washed with 2 x PBS prior to cell experiments.
S3. Scanning Electron Microscopy:
Dried electrospun PGMA fibres both with and without PXS64 were coated with 4 nm of platinum and imaged
using scanning electron microscope (Zeiss 1555, VP-FESEM) at an accelerating voltage of 4-5 kV. The fiber
diameter was calculated using the image analysis software ImageJ (NIH). A minimum of 50 random fibers were
measured. The data is reported as an average ± standard deviation. In the cell experiments, cells were incubated
as per cell viability protocol. At the stipulated time point, culture media was removed and coverslips loaded with
scaffold and cells were washed with 2 x PBS and fixed for 10 min using glutaraldehyde (2.5 % in PBS),
99
followed by 2 x PBS washes. Next serial dehydration steps were performed with increasing concentration of
ethanol to replace all the water in the samples with dry ethanol before critical point drying step. Post critical
point drying samples were coated with 4 nm of platinum and imaged using aforementioned parameters.
S4. High Pressure Liquid Chromatography
Drug loading analysis was carried out using HPLC (solvent A: water with 0.1 % TFA, solvent B: acetonitrile
with 0.1 % TFA, Phenomenex-C18 (2) 100A column (4.6 × 150 mm, 5 microns) at room temperature, 1 mL/ min,
λ = 280 nm, gradient: 100 % solvent B in 17 min) with Water 2695 pumping system and 2489 UV/Vis detector.
Drug loading was calculated by dissolving 35 mg of fibers (PGMA + PXS64) in acetonitrile (0.5 mL) and
calculated from the standard curve developed by dissolving free PXS64 in acetonitrile.
S5. Cell culture
Human primary dermal fibroblast cell cultures from normal skin were cultured in Dulbecco’s Modified Eagle’s
Medium (DMEM/F12 - GlutaMAX; Invitrogen Gibco) supplemented with 10% fetal bovine serum (FBS;
Invitrogen Gibco) and 1 % penicillin/streptomycin (Invitrogen Gibco). The cells were incubated at 37 °C in a
humidified atmosphere with 5 % CO2. All experiments were carried out with cells between passages 3-6.
S6. Cell Viability
Cell viability was determined using a LIVE/DEAD Viability/Cytotoxicity Kit (Invitrogen, UK) which measures
the membrane integrity of cells, as per manufacturer’s protocol. In brief, 20000 cells were seeded on the
sterilised electrospun scaffold on glass coverslips in a 24 well plate and supplemented with cell culture media
(DMEM F-12 containing 10 % FBS and 1 % Penicillin/ Streptomycin) for 30 min. Next, human recombinant
TGFβ1 (2 ng/mL, Cat. # 14-8348-62, eBioscience) was added to the wells (fresh media was added in controls)
and plates were incubated for 24 h and 72 h in the humidified incubator at 37 °C with 5 % CO2. At the stipulated
time (24 h and 72 h), cells were washed with PBS (3 times) and then stained with calcein AM (100 µL, 1
µM)/ethidium bromide I (100 µL, 2 µM) in PBS and further incubated for 30 min in the incubator. Images were
captured using an Olympus IX71 inverted microscope with a 20 x objective with fixed exposure time. Both live
and dead cells were counted using Image J software (NIH) with cell counter plug in. Data presented as mean ±
standard error mean (n = 4).
100
S7. Cell Proliferation
Cell proliferation was measured using the MTS assay (Cell Titer 96 ®Aqueous, Promega, Madison, USA) as per
the manufacturer’s protocol. Briefly, 1500 cells were seeded on the sterilised electrospun scaffold on 6 mm glass
coverslips in each well of a 96 well plate and supplemented with cell culture media (DMEM F-12 containing 10
% FBS and 1 % Penicillin/ Streptomycin) for 30 min followed by the addition of human recombinant TGFβ1 (2
ng/mL) to the wells (fresh media was added in controls) and the plates (individual for each time point) were
incubated for 72 h in the humidified incubator at 37 °C with 5 % CO2. MTS solution (40 µL) was added in each
well the next day, and was considered as 0 h time point, and incubated for 3 h in the humidified incubator at 37
°C with 5 % CO2. Following which 80 µL from each well was transferred into a new 96 well plates and read
under a plate reader at 490 nm excitation wavelength. Same protocol was followed at every time point for next
72 h. Data presented as mean ± standard error mean (n = 5).
S8. Gene Expression
Gene expression was measured using real time quantitative polymerase chain reaction. Cells (50000, in
DMEM/F12 media containing 10% FBS) were seeded on sterilised scaffolds (electrospun on coverslips) in 24
well plates and incubated for 24 h in the humidified incubator at 37 °C and 5% CO2. Next day culture media
(containing 10 % FBS) was replaced with starve media (containing 0.1 % FBS) and incubated for further 24 h in
the incubator to bring all the cells under same physiological cycle. Next day, TGFβ1 (2 ng/mL in starve media)
was added in the culture and further incubated for 48 h in the incubator. mRNA was extracted using RNeasy
mini kit® according to manufacturers’ protocol (Qiagen GmbH). For reverse transcription 1.5 µg of total mRNA
was converted to cDNA using Superscript VILO (Cat. # 11754, Applied Biosystems) according to
manufacturers’ protocol. 150 ng of cDNA was analysed by ABI 7500 fast analysis real-time PCR system using
TaqMan® master mix and col1a1 probes (Hs01076777_m1, Life Technologies). GAPDH was used as a reference
gene (Cat. # 4326317E, Life Technologies). Analysis was carried out using the instrument software. Release
media was prepared using DMEM/F-12 media (containing 10 % FBS and 1 % Penicillin/ Streptomycin)
supplemented with 0.2 % DMSO. Data presented as mean ± standard error mean (n = 3).
S9. Statistics
101
The results for cell viability, cell proliferation and gene expression are expressed as mean ± standard error mean
(SEM) and analysed for analysis of variance (ANOVA). Significance was evaluated using Bonferroni and
Turkey’s post-hoc analysis and set at 95% confidence (p < 0.05).
Figure S1: Cell proliferation assay showing cell growth over the period of 72 h post incubation with the two scaffolds both with and without TGFβ1 (2 ng/mL). Data presented as Mean ± SEM. Significance was set at * p < 0.05 using one way ANOVA and Bonferroni post-hoc analysis.
Figure S2: A representative florescent images showing cell morphology and the number of live and dead cells in culture. HDF cells were incubated on a) ES-PGMA scaffold, b) ES-PGMA + TGFβ1 (2 ng/mL) and c) ES-PGMA + PXS64 scaffold. Scale bar 2 µm. Cells were stained using Calcein AM/Ethidium bromide I staining where live cells fluoresce green while dead cells fluoresce red.
102
Figure S3: A representative image showing cell adherence on the two scaffolds. HDF cells were incubated a) ES-PGMA scaffold, b) ES-PGMA + TGFβ1 (2 ng/mL) and c) ES-PGMA + PXS64 scaffold. Scale bar: a) 2 µm, b) and c) 1 µm. Red arrows highlighting the cells.
References
1. K. S. Iyer, B. Zdyrko, H. Malz, J. Pionteck and I. Luzinov, Macromolecules, 2003, 36, 6519-6526.
103
Appendix B
Published work not directly included in the thesis
Peer reviewed publications contributed by the candidate are listed below. However, only
published articles are attached in the thesis.
1. Agarwal, V., Tjandra, E.S., Iyer, K. S., Humfrey, B., Fear, M., Wood, F. W.,
Dunlop, S. and Raston, C. L., Evaluating the effects of nacre on human skin and scar cells
in culture, Toxicology Research, 3, 223-227 (2014)
2. Eroglu, E., Chen, X., Bradshaw, M., Agarwal, V. , Zou, J., Stewart, S.G., Duan, X.,
Lamb, R.N., Smith, S.M., Raston, C. and Iyer, K. S., Biogenic production of palladium
nanocrystals using microalgae and their immobilization on chitosan nanofibers for
COMMUNICATION Colin L. Raston et al.Evaluating the eff ects of nacre on human skin and scar cells in culture
Volume 3 Number 4 July 2014 Pages 217–292
Toxicology Research
COMMUNICATION
Cite this: Toxicol. Res., 2014, 3, 223
Received 6th January 2014,Accepted 2nd May 2014
DOI: 10.1039/c4tx00004h
www.rsc.org/toxicology
Evaluating the effects of nacre on human skin andscar cells in culture†
Vipul Agarwal,a Edwin S. Tjandra,a K. Swaminathan Iyer,a Barry Humfrey,b Mark Fear,c
Fiona M. Wood,c Sarah Dunlopd and Colin L. Raston*e
Pearl nacre, a biomineralisation product of molluscs, has growing
applications in cosmetics, as well as dental and bone restoration,
yet a systematic evaluation of its biosafety is lacking. Here, we
assessed the biocompatibility of nacre with two human primary
dermal fibroblast cell cultures and an immortalised epidermal cell
line and found no adverse effects.
There are three main types of pearl oysters of the genusPinctada: the “Akoya” pearl oyster called Pinctada fucata, the“Golden lipped” oyster Pinctada maxima and the “Blacklipped” oyster named Pinctada margaritifera. Mollusc shells aremainly made up of two layers of calcium carbonate, compris-ing an outer layer of calcite and an inner layer of aragonite.Nacre (mother of pearl) in all oyster shells is a calcified struc-ture that forms the lustrous inner layer. It is mainly composedof aragonite (∼95–97%) tablets oriented in multiple layers,each surrounded by organic matrix.1,2 This organic matrixmakes up ∼5% of the nacre composition and is mainly com-prised of polysaccharides and proteins.3 According to a Euro-pean Commission report published in 2007 the cosmetic andtoiletries industry in the EU, Japan, China and the US had atotal market value of €136.2 billion.4 The cosmetics industrymaintains its edge by constantly developing novel topical skintreatments. A popular example is the use of all-natural ororganic ingredients, such as fruit and plant extracts to offerwrinkle relief that mimics the painful and potentially danger-ous side effects associated with invasive chemical remedies.5
Clinically, topical treatments containing, for example, aloevera, vitamin C, corticosteroids and tacrolimus are used with
the aim of minimizing scarring.6 Recently, there has beeninterest in the cosmetics industry in the use of nacre as a keyingredient.7 Most of the formulations are reported to eitheruse powdered pearl shell or powdered nacreous layer shell.Powdered shell and powdered nacre comprises of both organicand inorganic components. It is reported that nacre stores inits mineral-based organic structure a variety of bioactive mole-cules. Efficacy of this water soluble matrix (WSM) has beentested in a porcine burn injury model.8 WSM was obtained bysuspending powdered nacre in ultra-pure water and collectingthe supernatant via precipitation of insoluble components bycentrifugation. It was concluded that the active mineral basedorganic component has beneficial effects on the skin withenhanced wound healing.8,9
Nacre has also attracted attention for its potential in sup-porting bone grafting and bone regeneration. In culture underphysiological conditions, nacre can transform to hydroxy-apatite, the phosphorous rich main constituent of the mamma-lian bone framework.10,11 Nacre and its WSM can also aid inosteogenic regeneration.9,12–17 High phosphorous rich domainshave been described at the interface between bone andimplants made from Margaritifera shells which are biocompa-tible, biodegradable and osteoconductive and thus are thoughtto promote bone formation.18 Furthermore, nacre powder hasbeen used as an implantable material for reconstruction andregeneration of maxillary alveolar ridge bone in humans.19 Inthis example, the implanted nacre dissolves gradually and iseventually replaced by the mature lamellar bone suggestingthat the nacre acts as a biocompatible substrate for bone repla-cement.19 The water soluble components of the crushed nacrehave also been investigated for their potential in bone regener-ation in a similar vein.20,21 Lee et al., demonstrated the woundhealing potential of WSM component in a deep burn porcineskin model and showed enhanced collagen secretion anddeposition at the injury site resulting in enhanced healing.8 Inanother in vivo study using a rat skin incisional injury model,powdered nacre was implanted between the epidermis anddermis at the incisional site, with an aim of studying the effectof nacre on the synthesis of certain constituents of the dermal
†Electronic supplementary information (ESI) available. See DOI:10.1039/c4tx00004h
aSchool of Chemistry and Biochemistry, The University of Western Australia, Crawley,
AustraliabPearl Technology Pty Ltd, Geraldton, AustraliacBurn Injury Research Unit, School of Surgery, The University of Western Australia,
Crawley, AustraliadSchool of Animal Biology, The University of Western Australia, Crawley, AustraliaeSchool of Chemical and Physical Sciences, Flinders University, Bedford Park,
extracellular matrix. It was concluded that implanted nacreincreased collagen synthesis by dermal fibroblasts.22 Whileextensive investigations have been carried out in bone, theevaluation of the biocompatibility of nacre with human skincells is lacking. Thus the growing number of cosmetic formu-lations in the market with nacre as a key ingredient7,23,24
clearly warrants a thorough assessment with human skin cells.Since scars are also common, and contain cells with a pheno-type distinct from normal skin,25 it is also important to testpotential cosmetic ingredients with both cell types.
In the present study, we use nacre from the inner calcifiedlayer of the shell of Pinctada margaritifera and report thein vitro toxicity assessment of the material on three cell typesrepresenting both epidermal and dermal layers of humanskin. These were HaCaT cells, a human derived immortalisedkeratinocytes cell line, primary human dermal skin fibroblasts(HDF) and primary human scar fibroblasts (HSF).
Nacre used in the study was gently scraped26,27 from theinner layer of the shell to avoid the post processing required inthe case of powdered shell. SEM images (Fig. 1) confirmedthat the nacre was composed of pseudo-hexagonal shaped ara-gonite tablets which have basal plane dimensions of 2–6 µm,and a thickness of 300–400 nm.28 This structure is character-istic of previously reported nacre, which is a compositematerial consisting of alternating layers of mineral tablets sepa-rated by thin layers of biomacromolecular “glue”.29,30
To test the cytotoxicity of nacre, a live/dead assay wascarried out (see ESI S1.4†). Cells were incubated with nacre inculture media at physiological conditions for 24 h to 72 h andwere then stained for viability using calcein AM/ethidiumbromide I solutions. Viable cells fluoresce green through thereaction of calcein AM with intracellular esterase, whereasnon-viable cells fluoresce red due to the diffusion of ethidiumhomodimer across damaged cell membranes and binding withnucleic acids.
Fig. 2 shows live cells as the percentage of the total cells inhuman primary dermal skin fibroblast (HDF), human primaryscar fibroblast (HSF) and human derived immortalised HaCaTcell cultures when exposed to various concentrations of nacrefor 24 h and 72 h. Cytotoxicity of nacre was not observed forany of the concentrations examined in HDF cells (Fig. 2a).However, interestingly at a concentration of 2.5 mg ml−1 ofnacre (highest concentration tested) there was a significantreduction in viability at both 24 and 72 hours in the HSF cells(Fig. 2b). This underlines the importance of testing both scarand normal skin cell types for cosmetic application. Toxicitywas also observed at a concentration of 0.5 mg ml−1 in HaCaTcells (Fig. 2c), although this was only observed at 24 hours and
Fig. 1 Top view of the scrapped nacre, imaged using scanning electronmicroscopy (SEM). Scale bar (a) 1 µM and (b) 2 µM respectively. Samplewas coated with 4 nm platinum prior to imaging.
Fig. 2 Cell viability assays showing percentage of live cells in theculture post incubation with nacre. (a) Human dermal skin fibroblastscells, (b) human scar fibroblasts cells and (c) human derived immorta-lised HaCaT cells were incubated with various concentrations ofscrapped nacre and treated with calcein AM/ethidium bromide I to stainfor live and dead cells. Both live and dead cells were counted using fluo-rescence microscopy. ‘None’ is the untreated control. Data presented asaverage ± SEM (n = 4). Significance was set at *p < 0.05 using bonferronitest in one way ANNOVA.
was no longer present at the 72 hour time point. These resultsare in line with the results obtained previously where constitu-ents of nacre were shown to promote wound healing in a ratmodel22 and deep burn porcine skin.8 In both these studies,nacre has been shown to promote the recruitment of fibro-blasts for restoration and coverage of the injury site whileshowing no apparent signs of cytotoxicity. It has also beenshown to promote bone formation when implanted in thefemur of sheep with midshaft hemidiaphysis resection of theirfemur in vivo reiterating the non-cytotoxic advantage ofnacre.31
Fibroblasts have been reported to undergo morphologicalchanges from dendritic to stellate shapes upon exposure toexternal cues caused by changes in actin polarisation andadhesion.32,33 Cell morphology in fibroblasts is known to beinfluenced by cytokines such as transforming growth factor β
which can potentially induce polymerisation of globular tofilamentous actin.34 Fibroblast morphology can also be modu-lated by extracellular matrix architecture during woundhealing via cell–matrix interaction.32 Such morphologicalchange has been observed in cells undergoing oxidativestress.35,36 In our study, we found similar changes in fibroblastmorphology for both HDF and HSF cells at the highest concen-tration of nacre of 2.5 mg mL−1 (Fig. 3). Similar altered mor-
phology was also observed for HaCaT cells (see ESI Fig. S1†). Itcould be postulated that the high concentration of nacreinduces cellular stress, resulting in changes in the actin cyto-skeleton and a more stellate morphology (Fig. 3i, iii, ii and iv).Cell area was calculated from the fluorescent images shown inFig. 3 using Image J software.37 It was found that both HDFand HSF had significantly larger cell areas (p < 0.05) whentreated with 2.5 mg mL−1 of nacre (HDF: 2.79 ± 0.13 µm andHSF: 3.0 ± 0.19 µm respectively) as compared to the non-treated controls (HDF: 1.56 ± 0.08 µm and HSF: 1.54 ± 0.10 µmrespectively) (see ESI Fig. S2†).
Altered fibroblast morphology has been thought to occur inresponse to various factors including aging,38 strength of theextracellular matrix39 or other etiologies that induce mechan-ical stress on the cell. Changes in morphology also commonlyindicate oxidative as well as mechanical stress.39 Therefore, weexplored whether the morphological changes and increase in
Fig. 3 Cell morphology post calcein AM staining and imaged usingfluorescent microscopy. Cells were treated with various concentrationsof nacre for 24 h, stained and imaged. (i) untreated (control) primaryhuman dermal skin fibroblasts (HDF), (ii) HDF treated with 0.05 mg mL−1
nacre, (iii) HDF treated with 2.5 mg mL−1 nacre, and (iv), untreated(control) primary human dermal scar fibroblasts (HSF), (v) HSF treatedwith 0.05 mg mL−1 nacre and (vi) HSF treated with 2.5 mg mL−1 nacre.Scale bar 1 µm.
Fig. 4 Reactive oxygen species (ROS) assay showing ROS levels in cellsstressed with various concentrations of nacre for 24 h. No significantstress was observed as a result of calcium (from nacre) induced oxidativestress at the concentrations studied. (a) Human dermal skin fibroblastscells, (b) human scar fibroblasts cells and (c) human derived immorta-lised HaCaT cells were incubated with various concentrations ofscrapped nacre for the specified period of time. Cells were then incu-bated with 2’,7’-dichlorodihydrofluorescein diacetate (DCFH-DA) solu-tion which fluoresce in the presence of reactive oxygen species. ‘None’is the untreated control. Data presented as average ± SEM (n = 3). Sig-nificance was set at *p < 0.05 using bonferroni test in one way ANNOVA.
cell area at the highest concentration of nacre in culture was aresult of, or induced oxidative stress in, the cells. Oxidativestress was tested using the cell permeable fluorogenic probe2′,7′-dichlorodihydrofluorescein diacetate (DCFH-DA).DCFH-DA is taken up by cells and is deacetylated by cellularesterases to non-fluorescent 2′,7′-dichlorodihydrofluorescein(DCFH) which is rapidly oxidised by reactive oxygen species(ROS) to highly fluorescent 2′,7′-dichlorodihydrofluorescein(DCF). The fluorescent intensity is proportional to the ROSlevels within the cytosol (see ESI S1.5†). Cell responsiveness tothe assay was carried out by stressing the cells with the H2O2
solution provided in the kit which was also used to generatethe calibration curve (see ESI Fig. S3†). No changes in levels ofreactive oxygen species were observed in any cell type at anyconcentration of nacre (Fig. 4). This is important as oxidativestress is known to be a significant contributor to skin damageand excessive scarring in previous studies.40 It has beenknown that cells alter their morphology depending on theirenvironment.41,42 It is therefore hypothesized that the alteredfibroblast morphology in the present case is mainly due to theregulation of cell motility through geometrical constraint inthe presence of nacre. Indeed, it has been previously reportedthat when cells probe their physical surroundings, they acquiremechanical information or signals that help to determine thedirection of migration, with a consequential change in cellmorphology.43
Conclusions
We have established the biocompatibility of nacre using threehuman cell types representing the two primary layers ofhuman skin, using immortalised keratinocytes from the epi-dermal layer and two primary human dermal cell cultures. Thenacre used in the present study showed limited cytotoxicity athigh concentrations in scar derived cells, with the morphologyof the cells significantly changed by exposure at such concen-trations of nacre. No apparent oxidative stress was evident inany of the cell types. Overall, the data support the use of lowconcentrations of nacre in aesthetic formulations, with thepotential for high concentrations to cause changes in skinand/or scar cells which may have impact on efficacy.
The authors would like to thank the Australian ResearchCouncil and Pearl Technologies for funding the work underthe grant number LP100100812. The authors would also liketo acknowledge the Australian Microscopy & MicroanalysisResearch Facility at the Centre for Microscopy, Characteri-zation & Analysis, The University of Western Australia, fundedby the University, State and Commonwealth Governments.
Notes and references
1 J. H. E. Cartwright and A. G. Checa, J. R. Soc. Interface,2007, 4, 491–504.
2 F. Nudelman, E. Shimoni, E. Klein, M. Rousseau,X. Bourrat, E. Lopez, L. Addadi and S. Weiner, J. Struct.Biol., 2008, 162, 290–300.
3 Y. Oaki and H. Imai, Angew. Chem., Int. Ed., 2005, 44,6571–6575.
4 D. G. f. E. a. I. European Commission, A Study of the Euro-pean Cosmetics Industry, Report, European Commission,2007.
5 P. M. Reddy, M. Gobinath, K. M. Rao, P. Venugopalaiah andN. Reena, Int. J. Adv. Pharm. Nanotechnol., 2011, 1, 121–139.
6 J. M. Zurada, D. Kriegel and I. C. Davis, J. Am. Acad. Derma-tol., 2006, 55, 1024–1031.
7 Pearlcium, Genesis Framework – WordPress, vol. 2013, p. US.8 K. Lee, H. Kim, J. Kim, Y. Chung, T. Lee, H.-S. Lim,
J.-H. Lim, T. Kim, J. Bae, C.-H. Woo, K.-J. Kim andD. Jeong, Mol. Biol. Rep., 2012, 39, 3211–3218.
9 M. J. Almeida, C. Milet, J. Peduzzi, L. Pereira, J. Haigle,M. Barthélemy and E. Lopez, J. Exp. Zool., 2000, 288,327–334.
10 Y. Guo and Y. Zhou, J. Biomed. Mater. Res., Part A, 2008, 86,510–521.
11 P. Westbroek and F. Marin, Nature, 1998, 392, 861–862.12 M. Lamghari, M. J. Almeida, S. Berland, H. Huet,
A. Laurent, C. Milet and E. Lopez, Bone, 1999, 25, 91S–94S.13 M. Lamghari, H. Huet, A. Laurent, S. Berland and E. Lopez,
Biomaterials, 1999, 20, 2107–2114.14 M. Lamghari, S. Berland, A. Laurent, H. Huet and E. Lopez,
Biomaterials, 2001, 22, 555–562.15 F. Moutahir-belqasmi, N. Balmain, M. Lieberrher,
S. Borzeix, S. Berland, M. Barthelemy, J. Peduzzi, C. Miletand E. Lopez, J. Mater. Sci.: Mater. Med., 2001, 12, 1–6.
16 D. Duplat, A. Chabadel, M. Gallet, S. Berland, L. Bédouet,M. Rousseau, S. Kamel, C. Milet, P. Jurdic, M. Brazier andE. Lopez, Biomaterials, 2007, 28, 2155–2162.
17 C. Silve, E. Lopez, B. Vidal, D. Smith, S. Camprasse,G. Camprasse and G. Couly, Calcif. Tissue Int., 1992, 51,363–369.
18 H. Liao, H. Mutvei, M. Sjöström, L. Hammarström andJ. Li, Biomaterials, 2000, 21, 457–468.
19 G. Atlan, N. Balmain, S. Berland, B. Vidal and É. Lopez,C. R. l’Academie. Sci., Ser. III, 1997, 320, 253–258.
20 L. Pereira Mouriès, M.-J. Almeida, C. Milet, S. Berland andE. Lopez, Comp. Biochem. Physiol., Part B: Biochem. Mol.Biol., 2002, 132, 217–229.
21 M. Rousseau, H. Boulzaguet, J. Biagianti, D. Duplat,C. Milet, E. Lopez and L. Bédouet, J. Biomed. Mater. Res.,Part A, 2008, 85, 487–497.
22 E. Lopez, A. L. Faou, S. Borzeix and S. Berland, Tissue Cell,2000, 32, 95–101.
23 E. Lopez and A. E. Chemouni, US Pat, US10/089,982, 2005.24 D. M. DeLaRosa, US Pat, US2008/0199533 A1, 2008.25 C. Chipev and M. Simon, BMC Dermatol., 2002, 2, 13.26 M. Norizuki and T. Samata, Mar. Biotechnol., 2008, 10,
234–241.27 C. Ma, C. Zhang, Y. Nie, L. Xie and R. Zhang, Tsinghua Sci.
Evaluating the effects of nacre on human skin and scar cells in culture
Vipul Agarwal,a Edwin S. Tjandra,a K. Swaminathan Iyer,a Barry Humfrey,b Mark Fear,c Fiona M. Wood,c Sarah Dunlopd and Colin L. Rastone, *
Supporting Information
S1. Experimental
S1.1 Materials
Pinctada margaritifera shells were provided by Pearl Technologies Pty Ltd, which were grown in the waters of the Abrolhos Islands, Western Australia. The decalcified organic conchiolin layer was removed by wet sand blasting of the shell followed by gentle brushing to remove any dust particles that might otherwise contaminate the samples. Inner nacreous layer was then scraped using a surgical scalpel and stored at room temperature for a maximum of 2 weeks.
S1.2 Scanning Electron Microscopy
Scraped nacre above from the inner layer of the shell was mounted on SEM stubs (ProSciTech, Cat.# G040). Samples were coated with 4 nm of platinum. Images were taken using scanning electron microscope (Zeiss 1555, VP-FESEM) at 4-5 kV at 30µm aperture. Images were analyzed with the image analysis software ImageJ (NIH).1
S1.3 Cell culture
A human derived immortalized keratinocyte cell line, HaCaT 2 and two human primary dermal (fibroblast) cell cultures from normal skin and normal scar were used. All three cell types were cultured in Dulbecco’s Modified Eagle’s Medium (DMEM/F12 - GlutaMAX; Invitrogen Gibco) supplemented with 10% fetal bovine serum (FBS; Invitrogen Gibco) and 1% penicillin/streptomycin (Invitrogen Gibco). The cells were incubated at 37°C in an atmosphere of 5% CO2. Primary cells used were between the passages 7-10. Nacre was sterilized using UV sterilization technique in the tissue culture hood for 15 min prior its dissolution in media. Fresh nacre solution was prepared before every experiment.
S1.4 Cell Viability
Cell viability was determined using a LIVE/ DEAD Viability/Cytotoxicity Kit (Invitrogen, UK) which measures the membrane integrity of cells,3, 4 as per manufacturer’s protocol. In brief, 20000 cells were seeded in each well in a 24 well plate and treated with scraped nacre at various concentrations in cell culture media (DMEM F-12 containing 10% FBS and 1% Penicillin/ Streptomycin) and incubated for 24 h or 72 h in the humidified incubator at 37°C with 5% CO2. At the stipulated time (24h and 72h), cells were washed with PBS (3 times) and then stained with calcein (100 µL, 1µM)/ ethidium bromide (100 µL, 2 µM) in PBS and incubated in the humidified incubator at 37°C and 5% CO2 for 30 min. Images were captured using an Olympus IX71 inverted microscope with a 20 x objective with fixed exposure time. Both live and dead cells were counted using Image J with cell counter plug in. Experiments were performed in triplicate. Minimum of fifty images were captured per condition.
S1.5 Reactive Oxygen Species (ROS)
ROS was measured using the ROS assay kit (Oxiselect ROS assay kit, Cat.# STA 342, Cell Biolabs) following manufacturer’s protocol. In brief, 6000 cells were seeded in a 96 well plate and incubated in the humidified incubator at 37°C with 5% CO2 for 24h. Next day, wells were washed with PBS (3 times) and incubated with 2’, 7’- dichlorodihydrofluorescin diacetate (DCFH-DA) solution (0.1 x, 100 µL/ per well) for 1 h in the humidified incubator at 37°C with 5% CO2. DCFH-DA solution was removed and the wells washed with 3 x PBS. Cells were then treated with scraped nacre solution in culture media at a specified concentration for 24 h, wells were washed with 3 x PBS and cells were lysed using the lysis buffer provided (1 x, 100µL/ per well, incubated for 20 min. at room temperature) before reading the plate at 480 nm excitation/ 530 nm emissions using the plate reader. Experiments were performed in triplicate.
Cell size was measured using Image J software (NIH).1 A minimum of 25 cells were randomly selected from the fluorescence images and their area was measured. Values reported as mean ± standard error mean.
S1.7 Statistics
The results for cell viability, ROS experiments and cell area are expressed as mean ± standard error mean (SEM) and analysed by analysis of variance (ANOVA). Significance was evaluated using Bonferroni and Turkey’s post-hoc analysis and set at 95% confidence (p < 0.05).
Supporting Figures
Figure S1: Cell morphology post calcein AM/ ethidium bromide staining and imaged using fluorescent microscopy. HaCaT cells were treated with various concentrations of nacre for 24h, stained and imaged. a) Untreated (control), b) HaCaT cells treated with 2.5 mg/mL nacre. Scale bar 1µm.
Figure S2: Cell area size showing increase in the cell area post incubation with nacre. Cell area was measured from the fluorescent images of live cells taken for viability assay. ‘None’ is the untreated control. Data presented as average ± SEM (n>25). Significance was set at * p < 0.05 using bonferroni post hoc test in one way ANNOVA
Figure S3: Reactive oxygen species (ROS) assay showing increase in ROS levels in cells stressed with various concentrations of H2O2 in dose dependent manner. Human dermal skin fibroblasts cells were incubated with various concentrations of H2O2 for the specified period of time to generate the standard curve. Cells were then incubated with 2’, 7’- dichlorodihydrofluorescin diacetate (DCFH-DA) solution which fluoresce in the presence of reactive oxygen species. ‘None’ is the untreated control. Data presented as average ± SEM (n=3).
References
1. T. J. Collins, BioTechniques, 2007, 43, S25-S30.2. P. Boukamp, R. T. Petrussevska, D. Breitkreutz, J. Hornung, A. Markham and N. E. Fusenig, J. Cell Biol., 1988, 106, 761-771.3. L. W. Zhang and N. A. Monteiro-Riviere, Toxicol. Sci., 2009, 110, 138-155.4. I. Gotman and S. Fuchs, in Active Implants and Scaffolds for Tissue Regeneration, ed. M. Zilberman, Springer Berlin Heidelberg,
2011, vol. 8, pp. 225-258.
Cite this: RSC Advances, 2013, 3, 1009
Received 4th October 2012,Accepted 12th November 2012
Biogenic production of palladium nanocrystals usingmicroalgae and their immobilization on chitosannanofibers for catalytic applications3
DOI: 10.1039/c2ra22402j
www.rsc.org/advances
Ela Eroglu,ab Xianjue Chen,a Michael Bradshaw,a Vipul Agarwal,a Jianli Zou,a ScottG. Stewart,c Xiaofei Duan,d Robert N. Lamb,d Steven M. Smith,*b Colin L. Raston*a
and K. Swaminathan Iyera
Spherical palladium nanocrystals were generated from aqueous
Na2[PdCl4] via photosynthetic reactions within green microalgae
(Chlorella vulgaris). Electrospun chitosan mats were effective for
immobilizing these biogenic nanocrystals, as a material for
recycling as a catalyst for the Mizoroki–Heck cross-coupling
reaction. This photosynthetically-driven metal transformation
system can serve as a good candidate for an environmentally-
friendly method for the synthesis of metal nanocatalysts.
Biogenic systems can control the phase, structure, and topographyof inorganic nanocrystals with a level of precision similar tosynthetic approaches.1 Various organisms such as bacteria,2
yeast,3 fungi4 and algae5 are capable of processing a wide rangeof metals. Such bioprocessing has been effectively used in thereduction of environmental pollution and also for the recovery ofmetals from waste.6 The formation of metal nanoparticles in thepresence of microorganisms primarily involves the reduction ofmetal ions in solution by enzymes generated by microbial cellactivities, which can be intracellular or extracellular. This dependson the nanoparticle crystallization site, as being either inside thecell or on the cell surface.7,8 When considering palladium, thebiosynthesis of nanoparticles of the metal have been reported inbacteria (Desulfovibrio desulfuricans, Shewanella oneidensis, Bacillussphaericus),9 cyanobacteria (Plectonema boryanum UTEX 485),10,11
plants,12–14 and viruses (e.g., tobacco mosaic virus).15,16 Herein wereport a biogenic synthesis of palladium nanocrystals in thepresence of Chlorella vulgaris with the photoautotrophic micro-
algal metabolism most likely providing the necessary reducingagents. Studies on the biogenic reduction of palladium by variousorganisms are usually limited to the generation of Pd(0). We haveused a non-toxic and environmentally-available microorganism toachieve this, and then collected the particles from the liquidculture via immobilizing on electrospun chitosan nanofibers. Inaddition, we have established the utility of this composite materialas a recyclable catalyst which is functional even at very low loadingrates. The overall integrated approach is without precedent, as arethe individual components.
Algae are a large group of photosynthetic organisms that areubiquitous in various aquatic habitats including sea, freshwater,wastewater and also in moist solids.17 Microalgal remediation hasbeen reported for several metal ions,18,19 and involves bothintracellular (polyphosphates) and extracellular (polysaccharideson algal cell wall) metal binding groups.20–22 Based on theseobservations, we initially evaluated the ability of a common greenmicroalga, Chlorella vulgaris, to reduce palladium(II) to Pd(0)starting with a solution of Na2[PdCl4]. Various concentrations ofPd salt were investigated (100, 50, 25, 12.5, 0 mg L21) in order todetect their effects on cell viability. The cells were grown insolutions of Na2[PdCl4] added to algal freshwater media (MLAmedia)23 at 25 uC and a rotation speed of 120 rpm, under diurnalillumination of 16 h light/8 h dark cycles, with the total chlorophyllcontent used to validate the viability of the cell cultures.24
Accumulation of total chlorophyll pigment (Chl a + Chl b) as afunction of growth time was monitored (Fig. 1a), and thethreshold concentration for toxicity on the cells establishedamongst 25–50 mg L21. Within the first five days, the cultureflasks containing less than this concentration clearly showed anincrease in cellular growth with solution retaining the green colorwhich is associated with the chlorophyll content of the cells. Incontrast, higher concentrations of Na2[PdCl4] resulted in the colorfading and loss of cell viability (Fig. 1b). The solution containing25 mg L21 of Na2[PdCl4] had the highest amount of palladiumprecursor while showing little reduction in growth rates comparedto the control. Thus, it was chosen as the prototype for furtherexperiments.
aCentre for Strategic Nano-Fabrication, School of Chemistry and Biochemistry, The
University of Western Australia, M313, 35 Stirling Highway, Crawley, WA 6009,
Tel: +61 8 6488 3045bARC Centre of Excellence in Plant Energy Biology, The University of Western
Australia, M313, 35 Stirling Highway, Crawley, WA 6009, Australia.
E-mail: [email protected]; Fax: +61 8 6488 4401; Tel: +61 8 6488 4403cSchool of Chemistry and Biochemistry, The University of Western Australia, 35
Stirling Highway, Crawley, WA 6009, AustraliadSurface Science & Technology Group, School of Chemistry, The University of
Melbourne, VIC 3010, Australia
3 Electronic supplementary information (ESI) available: Experimental details,characterization, and an additional TEM image. See DOI: 10.1039/c2ra22402j
RSC Advances
COMMUNICATION
This journal is � The Royal Society of Chemistry 2013 RSC Adv., 2013, 3, 1009–1012 | 1009
Transmission electron microscopy (TEM) of the liquid culturemedia revealed the presence of crystalline spherical palladiumnanoparticles with an average particle size of around 7 nm, with arange of 2 to 15 nm in diameter (Fig. 2a). The correspondingelectron diffraction pattern further confirmed the presence ofelemental Pd nanocrystals with a characteristic face-centered cubic(fcc) structure and average d-spacing values of 0.22, 0.19, 0.14, and0.12 nm for the (111), (200), (220) and (311) planes, respectively(Fig. 2b). High resolution TEM analysis further confirmed thepresence of single palladium nanocrystals (Fig. 2c and d).
Reducing agents produced by photosynthesis are the keycomponents for the reduction of Pd(II) into Pd(0) nanoparticles(Scheme 1). Photosynthetic processes in green microalgae can takeplace either under oxygenic or anoxic environments.17,25 Greenalgae have chlorophyll-a and chlorophyll-b pigments, and can
accomplish oxygenic (oxygen-evolving) photoautotrophic reactionswhile using H2O as an electron donor.17 Oxygenic photoauto-trophic processes include two sets of photochemical reactions: (i)light reactions conserving chemical energy in the form ofadenosine triphosphate (ATP), and the reduced form of nicotina-mide adenine dinucleotide phosphate (NADPH), and (ii) ‘dark’reactions in which CO2 is reduced to organic compounds usingATP and NADPH. Light reactions have two separate sets ofphotosystems (PS), namely PS I and PS II, in which the electrontransfer follows a Z-scheme between these two photosystems. PS IIis mainly responsible for the splitting of water (H2O A KO2 + 2e2
+ 2H+) as a first stage of cyclic electron flow. While electron flowfollows the Z-scheme between PS II and PS I, a proton motive forceis created and used for the generation of ATP. CO2 is fixed by theenzyme ribulose bisphosphate carboxylase (RuBisCO) and reducedin the Calvin cycle using NADPH.17 Reducing equivalents can beexported from the chloroplast in the form of carbon metabolites,particularly triose phosphates (triose-P).26 Their oxidation bydehydrogenases in the cytosol can also generate NADH andNADPH for other reduction reactions. We postulate that suchreducing agents promote the reduction of Pd(II) into crystallinePd(0) nanoparticles, which is further partially oxidized due to theaerobic culture conditions. An hypothesis for the reduction ofpalladium(II) by photoheterotrophic bacteria (Rhodobacter sphaer-oides) cultures involves the reduced electron carriers (such asferredoxin, NADH, and FADH) as the electron donors.27
Consistent with this hypothesis, we found that addition ofNADPH to the MLA growth medium23 resulted in reduction ofNa2[PdCl4] to produce Pd nanocrystals in solution which wereeffectively recovered on exposure to cross-linked electrospunchitosan nanofiber mats (see ESI3, Fig. S1).
Fig. 2 (a) TEM image of palladium nanoparticles precipitated in the four week oldmicroalgae solution (scale bar: 20 nm). (b) Selective area electron diffraction patternof the palladium nanoparticles in 2(a), giving d-spacings of 0.22, 0.19, 0.14, and 0.12nm which correspond to 111, 200, 220, and 311 reflections, respectively, for Pd(0).(c) High resolution TEM image of palladium nanoparticles. (d) Pd(0) nanocrystalswith (111) lattice spacings of around 0.22 nm. Inset shows the Fast FourierTransform (FFT) pattern corresponding to the area shown within the yellowrectangle in 2(c), indicating the crystal structure of palladium nanoparticles.
Scheme 1 Palladium nanoparticle synthesis by photosynthetic green microalgae,and their uptake on an electrospun chitosan mat for use as a catalyst in Mizoroki–Heck reactions. The left stage shows a combination of mechanisms taking placewithin the photosynthetic organisms, resulting in the production of reducingagents.17,25,26 (ADP: adenosine diphosphate, ATP: adenosine triphosphate, Fd:ferredoxin, NADP+: oxidized form of nicotinamide adenine dinucleotide phosphate,NADPH: reduced form of nicotinamide adenine dinucleotide phosphate, PGA:phosphoglycolic acid, RuBisCO: rubilose biphosphate carboxylase). NADPH is likelyone of the main reducing agents for the reduction of Na2[PdCl4], which is partiallyoxidised as a result of aerobic culture conditions.
Fig. 1 (a) Total amount of accumulated chlorophyll (Chl a + Chl b) measured as afunction of growth time for Chlorella vulgaris cultures in the presence of variousconcentrations of Na2[PdCl4] (100, 50, 25, 12.5 mg L21), and control cultures in theabsence of Na2[PdCl4] (0 mg L21, hollow circles), with the total volume of solution inall flasks at 40 mL during the growth experiments. (b) Chlorella vulgaris culture flaskscontaining various concentrations of Na2[PdCl4].
1010 | RSC Adv., 2013, 3, 1009–1012 This journal is � The Royal Society of Chemistry 2013
Chitosan is derived from chitin present in the exoskeletonsof crustaceans,28 and is an attractive, renewable feedstockwhich is effective in binding a number of metal ions,29 andeffective as a support for Pd, Ni, Cu, Cr and Zn based catalysts.30
Cross-linked chitosan mats (3 6 2 cm rectangular shape)generated using electrospinning31,32 were placed into a fourweek old microalgal culture containing 25 mg L21 Na2[PdCl4]. Acontrol experiment involved exposing chitosan mats to 25 mgL21 Na2[PdCl4] solution in the absence of microalgae. Chitosanmats were kept in the solution for about two weeks as to achievesufficient particle adsorption from the liquid media. Scanningelectron microscopy (SEM) revealed the nanofiber structure ofthe chitosan mats with an average diameter of 100 nm, whichwas maintained after exposure to the growth media (Fig. 3a). Itis noteworthy that there was a detectable difference in color ofthe recovered chitosan mats from the microalgae and thecontrol solutions. The initial light yellow chitosan mat turneddark brown after being incubated with algae (Fig. 3b), whereasno color change was observed for the control experimentwithout algae (Fig. 3c). The darker color is considered to beconsistent with the immobilization of the reduced Pd nano-crystals from the growth media, and corresponds to approxi-mately 1.03 wt% Pd loading on each rectangular piece ofelectrospun chitosan mat (3 6 2 cm), which was establishedusing inductively coupled plasma-optical emission spectro-scopy (ICP-OES). X-ray photoelectron spectrometric (XPS)analysis of the Pd loaded chitosan mats showed a dominatingpeak in the PdO 3d5/2 region (binding energy of 337.9 eV) andalso the presence of the Pd(0) peak in the Pd 3d5/2 region
(binding energy of 336.0 eV) (Fig. 3d). Analysis of the samplefollowing argon ion etching primarily showed a dominant Pd(0)peak in the Pd 3d5/2 region (binding energy of 335.9 eV) with anadditional lower peak of PdO in the 3d5/2 region (binding energyof 337.7 eV) (Fig. 3e). Thus the XPS data establish that thesurface of the palladium nanoparticles is partially oxidizedunder the aerobic culture conditions. Slight oxidation was alsoreported when palladium containing samples were exposed toan oxygen containing atmosphere before XPS analysis.33
Chlorine was not detected during these XPS analyses, indicatingthe absence of palladium(II) chloride species. The bindingenergies of both Pd 3d5/2 and PdO 3d5/2 were found to be slightlyhigher than the values for pure Pd metal,34 and PdO samples.35
This slight shift is reported to be common when the Pd/PdOnanoparticles are embedded in an insulating substrate,36 whichis the chitosan mat in the present case. In their study,Schildenberger and his colleagues also reported similar peaksfor Pd 3d5/2 (336.0 eV) and PdO 3d5/2 (337.9 eV) due to theisolated arrangement of metal nanoparticles on the layers ofoxidized silicon wafers.36
The utility of these palladium loaded chitosan mats were testedas catalyst supports for the standard Mizoroki–Heck reaction.37,38
Six dried mats (a total of 0.23% mol Pd per mol of iodobenzene)were introduced into a solution containing iodobenzene, butylacrylate, triethylamine and dimethylformamide (DMF). Thereaction temperature was kept constant at 80 uC for 16 h, andafter each reaction, the supported catalyst was recovered forrecycling studies after washing with DMF under nitrogen gas toavoid any oxygen induced regrowth of Pd(0) nanoparticles.39 Thecatalyst can be recycled at least four times with quantitativeconversion for each cycle, with the conversion yields of: 68% (1stcycle), 62% (2nd cycle), 45% (3rd cycle) and 36% (4th cycle) byweight. For the control comparison, a mat exposed to a Na2[PdCl4]solution without algae resulted in a conversion yield of only 5%.We have previously reported that a chitosan mat containingpalladium nanoparticles generated by reduction of pre-absorbedNa2[PdCl4] is also an effective catalytic support with no appreciableleaching of the metal.31 Conventional palladium catalysts usuallyrequire a 1–5 mol% loading rate for effective Mizoroki–Heck cross-coupling reactions.40 Our biogenic palladium nanocatalysts havehigh catalytic activity (68%) with respect to commercial material(5%), even at low palladium loadings (0.23 mol%), which issignificant for applications in the fine chemical industries.
In conclusion we have established a biogenic synthesis ofpalladium nanocrystals in the presence of Chlorella vulgaris andtheir subsequent immobilization on an electrospun chitosan matas a novel support for application in catalysis. In addition, we havedemonstrated that NADPH involved in the photoautotrophicmicroalgal metabolism is likely to play a role in the biogenicsynthesis. Utilization of easy-to-grow, nontoxic and environmen-tally-available microalgae for the synthesis of palladium haspotential for the development of green chemistry processes forother metals.
Fig. 3 (a) SEM image of the as-prepared electrospun chitosan nanofibers beforebeing exposed to palladium solution (scale bar: 1 mm). Mats of this material placedinto 4 week old 25 mg L21 Na2[PdCl4] solutions with and without microalgae whichwere collected after two weeks are shown in (b) and (c) respectively. Curve-fitted Pd3d XPS spectra for (d) as collected and (e) etched internal surface of a chitosan matunder argon ions for 60 s, both showing Pd(0) (blue curve) and PdO (red curve).
This journal is � The Royal Society of Chemistry 2013 RSC Adv., 2013, 3, 1009–1012 | 1011
We kindly acknowledge support of this work by the AustralianResearch Council (ARC) and internal grants of The Universityof Western Australia. The microscopy analysis was carried outusing the facilities in the Centre for Microscopy,Characterization and Analysis (The University of WesternAustralia). The authors also would like to thank Dr L. Byrne forhis kind help during the NMR analysis.
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7 K. Simkiss and K. Wilbur, Biomineralization. Cell Biology andMineral Deposition, Academic Press, Inc., San Diego, 1989.
8 S. Mann, Biomimetic Materials Chemistry, VCH, Weinheim, NewYork, 1996.
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10 J. R. Lloyd, P. Yong and L. E. Macaskie, Appl. Environ. Microbiol.,1998, 64, 4607–4609.
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1577–1581.23 R. A. Andersen, Algal Culturing Techniques, Elsevier Academic
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25 E. Eroglu and A. Melis, Bioresour. Technol., 2011, 102,8403–8413.
26 M. Taniguchi and H. Miyake, Curr. Opin. Plant Biol., 2012, 15,252–260.
27 M. D. Redwood, Bio-hydrogen and biomass supported palladiumcatalyst for energy production and waste minimisation, PhDThesis, The University of Birmingham, 2007.
28 M. Rinaudo, Prog. Polym. Sci., 2006, 31, 603–632.29 E. Guibal, Sep. Purif. Technol., 2004, 38, 43–74.30 A. B. Sorokin, F. Quignard, R. Valentin and S. Mangematin,
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1012 | RSC Adv., 2013, 3, 1009–1012 This journal is � The Royal Society of Chemistry 2013
Biogenic production of palladium nanocrystals using microalgae and
their immobilization on chitosan nanofibers for catalytic applications
Ela Eroglua,b, Xianjue Chena, Michael Bradshawa, Vipul Agarwala , Jianli Zoua, Scott G. Stewartc,
Xiaofei Duand, Robert N. Lambd, Steven M. Smithb*, Colin L. Rastona*, and K. Swaminathan Iyera
a Centre for Strategic Nano-Fabrication, School of Chemistry and Biochemistry, The University of Western
Australia, Crawley, WA 6009, Australia, E-mail: [email protected] b ARC Centre of Excellence in Plant Energy Biology, The University of Western Australia M313, 35 Stirling
Highway, Crawley, WA 6009, Australia, E-mail: [email protected] c School of Chemistry and Biochemistry, The University of Western Australia, 35 Stirling Highway, Crawley,
WA 6009, Australia d Surface Science & Technology Group, School of Chemistry, The University of Melbourne, VIC 3010, Australia
S1. Biosynthesis of Palladium Nanoparticles
Wild type Chlorella vulgaris cultures (from the Australian National Algae Culture Collection
at CSIRO, Tasmania) were used as the green microalgae source for the biosynthesis of
palladium. Sterile algal freshwater media (MLA media)[1] containing standard micronutrients,
nitrate, phosphate, carbonate buffer and vitamins was used as the substrate source. The
microalgae cultures were mixed with Na2[PdCl4] solution at various concentrations (100, 50,
25, 12.5, 0 mg/L), towards reaching an initial total-chlorophyll content of around 1.8 mg/L
(Figure 1a).
Na2[PdCl4] solution was prepared by dissolving Na2[PdCl4] powder (Sigma-Aldrich) in
distilled and sterilized water overnight, and subsequently filtered through a sterile Pall®
Acrodisc® 32 mm syringe filter (0.2 µm membrane) for further sterilization. Concentrations
of Na2[PdCl4] in microalgae solutions are reported here as the initial concentrations
measured by the gravimetric analysis before the filtration process. The experiments were
conducted under batch conditions and cyclic diurnal conditions (16 h light/8 h dark) at a
constant temperature (25 °C). Algae cultures (total liquid volume of 40 mL) were grown in
250 mL Erlenmeyer flasks, under continuous cool-white fluorescent illumination at an
incident intensity of around 200 µmol photons m-2s-1(PAR) upon orbital shaking (Thermoline
Nitrate removal from liquid effluents using microalgae immobilized onchitosan nanofiber mats
Ela Eroglu,a,b Vipul Agarwal,a Michael Bradshaw,a Xianjue Chen,a Steven M. Smith,*b Colin L. Raston*a
and K. Swaminathan Iyera
Received 25th June 2012, Accepted 8th August 2012DOI: 10.1039/c2gc35970g
Mats of electrospun chitosan nanofibers were found effectivein immobilizing microalgal cells. These immobilized micro-algal cells were also used as a durable model system forwastewater treatment which has been demonstrated by theremoval of around 87% nitrate from liquid effluents ([NO3
Wastewater treatment focuses on eliminating unwanted chemi-cals and/or biological impurities from contaminated water. Ingeneral the treatment methods are mainly based on the separationof pollutants from the wastewater with a requirement for afurther processing stage to eliminate these pollutants.1 Integratedwastewater treatment processes are important in eliminatingundesired species, ideally converting them into valuable pro-ducts. As relatively recent bioprocesses, algal cultivation inwastewaters has a combination of several advantages such aswastewater treatment and simultaneous algal biomass pro-duction, which can be further exploited for biofuel production,food additives, fertilizers, cosmetics, pharmaceuticals, and othervaluable chemicals.2 However, there are inherent difficultiesassociated with algae-based bioprocessing in the harvesting,dewatering and processing of the algal biomass.
Immobilization of the cells on solid surfaces confer advan-tages over free cells in suspension, namely the immobilized cel-lular matter occupy less space, require smaller volume of growthmedium, are easier to handle, and can be used repeatedly forproduct generation.3,4 Moreover, immobilization can alsoincrease the resistance of cell cultures to harsh environmentalconditions such as salinity, metal toxicity and variations inpH.3,4 Entrapment is one of the most common immobilizationmethods which involves capturing the cells in a three dimen-sional gel matrix, made of polymeric materials or inorganicspheres.4 Both synthetic polymers (e.g., acrylamide,
polyurethane, polyvinyl) and natural polymers (e.g., collagen,agar, cellulose, alginate, carrageenan) have been used for thispurpose.5 Several studies have been reported on wastewater treat-ment involving the entrapment of microalgae cultures insidealginate beads, porous glass, and several synthetic polymers.6–8
However, most attempts to immobilize viable algae cells insidesuch insoluble materials have limitations, with the encapsulatingmaterials having volume/surface ratios usually orders of magni-tude larger than thin films. As a consequence, algal viability ismostly reported to decrease which relates to the need for thenutrients or reactants to diffuse far into the material to reach thealgal cells.5 We have developed a new technique to overcomethese problems using electrospun nanofiber mats as the matrixfor immobilizing the algal cells, with an overall strategy tocombine wastewater treatment processing with algal harvestingin a single process.
Electrospun nanofibers of chitosan were employed as apolymer/matrix support for green microalgae in the currentstudy. Chitosan is composed of D-glucosamine and N-acetyl-D-glucosamine, and is formed by the deacetylation of chitin (β-N-acetyl-D-glucosamine polymer).9 Chitin is usually extracted fromthe exoskeletons of crustaceans (e.g., crab, lobster and shrimp)and even from the cell walls of fungi.9,10 Chitosan is non-toxicand biodegradable, and can be used as an animal feed.9 Further-more, it can be used as a coagulant for wastewater treatment andfor the recovery of waste sludge.11
The removal of nitrate ions is regulated by law which relatesto its hazardous effects on human health and the environment.Several methods have been reported for the removal of nitratefrom water bodies, including biological denitrification,12 chemi-cal reduction,13 electrodialysis,14 and a combined bioelectro-chemical/adsorption process.15 In this study, we aimed tosuperimpose the treatment efficiencies of microalgae in reducingnitrate and electrostatic binding of the nitrate ion by chitosan, i.e.combining biological and chemical processing.
Chlorella vulgaris cultures were used as the green microalgaewhich were originally obtained from the Australian NationalAlgae Culture Collection at CSIRO, Tasmania. Cell growth wascarried out under artificial diurnal-illumination (16 h light/8 hdark cycle) at around 22 °C. Electrospun chitosan mats werefabricated following the procedure optimized in previousstudies.16,17 This involved dissolving chitosan powder (6 wt%)in a mixture of trifluoroacetic acid (TFA) and dichloromethane(DCM) (70 : 30 v/v), with 5.4% (v/v) addition of glutaraldehyde
aCentre for Strategic Nano-Fabrication, School of Chemistry andBiochemistry, The University of Western Australia, M313, 35 StirlingHighway, Crawley, WA 6009, Australia. E-mail: [email protected];Tel: +61 8 6488 3045; Fax: +61 8 6488 8683bARC Centre of Excellence in Plant Energy Biology, The University ofWestern Australia M313, 35 Stirling Highway, Crawley, WA 6009,Australia. E-mail: [email protected]; Tel: +61 8 6488 4403;Fax: +61 8 6488 4401
solution (25% in H2O) immediately prior to electrospinning.17
This was deemed necessary to cross-link the chitosan to avoidpolymer breakup and dissolution. The experimental settings ofelectrospinning processes were as follows: (i) syringe pumpspeed: 0.1 mm min−1, (ii) voltage: 18 kV, (iii) distance betweenthe target and the tip of the syringe: 11 cm, (iv) target speed:1 m min−1, (v) traverse speed: 0.5 cm min−1. Fiber mats wereannealed overnight to remove any remaining solvent and storeduntil required. After electrospinning, 2 mL of algae solution inits exponential phase of growth with a total chlorophyll content(Chl a and b) of ∼2 mg L−1, was placed onto a cut out rectangu-lar chitosan-mat (3 × 2 cm) and left at room temperature forapproximately 48 hours to allow sufficient attachment of algaecells to the surface of the mat. Microalgae cell walls containvarious polysaccharides, which are compatible with the surfaceof the chitosan nanofibers.18,19 The presence of negative surfacecharge on the surface of Chlorella cells, arising from dissociationof uronic acid groups, and/or the presence of sulfate groups forexample,18 provide electrostatic attraction to the positivelycharged primary amine groups of chitosan not involved in theabove cross linking. Moreover, the negatively charged surface ofthe microalgae can also result in binding metal ions, thereby pro-viding an opportunity for biosorption applications, along withthe removal of nitrate ions as functional algal cells.18–20
This bionano-composite material was then placed into nitratecontaining artificial growth medium which contained mainlyphosphates, nitrates, carbonate buffer, micronutrients and vita-mins,21 with an initial nitrate-nitrogen concentration of around30 mg L−1. Nitrate-nitrogen (NO3
−-N) term refers to the amountof nitrogen (N) in liquid solutions coming from nitrate ions(NO3
−). This nitrate-nitrogen concentration is within the rangeof other algal nitrate removal studies.2,22 Furthermore, it simu-lates the range of nitrate content present in ground-waters(∼0.1–50 mg L−1 NO3
−-N)23 and sewage treatment planteffluents. The regulatory limit for the maximum contaminantlevels of [NO3
−-N] in public drinking water is 10 mg L−1, asestablished by the United States Environmental ProtectionAgency (EPA).24,25
During the algal growth, the pH of the medium was keptaround 6.5–7.0 by the addition of dilute hydrochloric acid (HCl)when necessary. The colorimetric “cadmium reduction method”was employed for the nitrate-nitrogen analysis, using chemical-kits in the form of powder pillows (HACH®, NitraVer NitrateReagent) and a colorimeter (HACH® DR/870).26 For compara-tive purposes, a control experiment with a chitosan nanofibermat devoid of algae culture was also treated under the same con-ditions. Scanning electron microscopy (SEM) analyses wereacquired using a Zeiss 1555 VP-FESEM, while the acceleratingvoltage was changed between 3 to 5 kV. The air-dried sampleswere coated with approximately 3 nm layer of platinum beforeimaging. A NanoMan AFM system (Veeco Instruments Inc.)was used for the atomic force microscopy (AFM) analysis, oper-ating under the tapping mode. Chlorophyll content of the cellswas analyzed using spectrophotometric measurements of metha-nol extracts obtained from the algal culture pellets.27
A challenge in the present work was to fabricate an insoluble,fibrous structure with sufficient porosity, which can facilitate thediffusion of nutrients and cellular products between the environ-ment and the algae. Fig. 1a and b show scanning electron
microscopic (SEM) images of the nanofiber structure of thechitosan mats after the electrospinning process. The diameter ofthe fibers was between 50 to 180 nm, with an average diameterof around 91 nm. After placing the nanofiber mat into aqueoussolution, the fibers gradually swelled with a significant increasein porosity of the material, Fig. 2a, and became an effectivesupport matrix for the C. vulgaris cells, which have the expecteddiameter from SEM images around 3–4 μm, Fig. 2b and c. Thisporous structure has an advantage for facilitating the diffusion ofmaterials such as nutrients and waste products between theenvironment and the algae, while the replication of algal cells isaccomplished on the surface of the nanofiber mat, Fig. 2c. Theheight profile measurements obtained by AFM analyses esta-blished the thickness of the chitosan mat at close to 400 nm,Fig. 3a, with the overall thickness for the algae attached chitosanmat at 4.3 μm, Fig. 3b. The difference in height is consistentwith the attachment of a single layer of individual C. vulgariscells on the nanofiber mats.
Fig. 4 shows the optical images of algae cells attached to thesurface of chitosan mats (a) initially, (b) after 3 days, and (c)after 10 days from the start of the growth experiments in 40 mLliquid media. Note the increasing green color on the surface isdue to the increased concentration of algal cells on the chitosanmat with respect to time. Detailed imaging is given as SEMimages in Fig. 5. The amount of algae cells on a chitosan matwith same dimensions yielded around 4 cells per 100 μm2 for a3 day old sample, Fig. 5c, whereas this increased to around
Fig. 2 (a) SEM images of porous and swollen chitosan nanofiber mats,after exposure to nitrate containing media for two days, (b) SEM imagesof chitosan nanofibers surrounding individual, and (c) multiple algaecells. Scale bars are given as 1 μm.
Fig. 1 SEM images of as prepared electrospun chitosan nanofibers atlow and high magnifications with scale bars of (a) 1 μm, and (b) 10 μm,respectively.
20 cells per 100 μm2 by the 10th day of the treatment process,Fig. 5d.
Fig. 6 shows the nitrate-nitrogen concentration versus time inthe absence of algae (triangles) or algae attached (rectangles)nanofiber mats. After the insertion of this bionano-compositeinto the liquid media (Vtotal: 40 mL), around 30% of the initialnitrate value was decreased within the first 2 days. This reductionin nitrate is mainly caused through the uptake by the chitosannanofibers rather than the algal cultures, as a physicochemicaladsorption process. A similar pattern was also observed for themats devoid of algal cells where after the second day there wasno further nitrate removal. In contrast the algae containing matscontinued their nitrate uptake, being used in their cellular meta-bolism for replication, building more biomass and energyproducts.
Amino groups in chitosan are protonated at acidic to neutralpH conditions,28 which enhance the adhesive properties of chito-san by increasing its tendency to attach negatively charged enti-ties which in this case are algal cell walls and nitrates. Chlorellavulgaris cell walls are known to be highly negatively chargedwith a zeta potential of around −30 mV in neutral water.29 Onthe other hand, the zeta potential of the positively charged chito-san nanofibers is +20 mV at neutral pH. Matsumoto et al.30
reported the zeta potential values of chitosan nanofibers to behighly dependent on the pH of the media, with it increasingto +30 mV in pH around 5–6, whereas it drops to zero for pHvalues above 8.30 Algal growth tends to alkalify its medium, asthe cellular uptake of anions (such as nitrates, phosphates, car-bonates, etc.) is stabilized with equivalent amounts of hydroxyl(OH−) anion efflux.31 For this reason, we maintained the pH ofthe culture around 6.5–7 by the regular addition of dilute HClduring the current study. Due to the nature of HCl, several otheracidifying agents (such as CO2) can be considered for any futuredevelopments and advanced scale-up processes for municipaland/or industrial wastewater samples.
Clearly the presence of the nanofiber mat in the liquidenvironment is responsible for the initial removal of nitrate whilethe continued growth of algae subsequently consumes theremaining nitrate in further stages with a slower rate. Overallnitrate removal rates were calculated as 32 ± 3%, and 87 ± 4%,for the “microalgae-absent” and “microalgae-attached” chitosanmats, respectively. Several studies have already been reported onwastewater treatment with immobilized microorganisms. Fierroet al.22 investigated the effect of nitrate removal by Scenedesmusspp. cyanobacterial cells immobilized within spherical chitosanbeads. They achieved 70% nitrate removal for the immobilizedcultures, while 20% of the initial nitrate content was removed bythe blank chitosan beads. In another study, Mallick and Rai32
also achieved relatively higher nitrate removal rates (73%) byAnabaena doliolum and Chlorella vulgaris cells immobilized inchitosan beads compared to the cells immobilized in other typesof gels made of alginate, carrageenan or agar. De-Bashan et al.33
achieved only 15% nitrate removal for co-immobilized
Fig. 3 AFM topographic mapping of chitosan nanofiber mats(4 × 4 μm) without, (a) and with, (b) algal cells.
Fig. 4 Progress of the algal growth on the surface of chitosan mats: (a)initially, (b) after 3 days, (c) after 10 days of the growth experiment.
Fig. 5 SEM images of immobilized C. vulgaris cells on the surface ofchitosan nanofiber mats after different time intervals. (a and c) are formats after 3 days, and (b and d) are for mats 10 days old.
Fig. 6 Nitrate-nitrogen (NO3−-N) concentration (mg L−1) of algal
medium versus time. Chitosan nanofibers devoid of algal cells are rep-resented with triangles, whereas those with immobilized algal cells areshown as rectangles.
microorganisms (Chlorella vulgaris with a growth-promotingbacterium Azospirillum brasilense) within alginate beads. At theother end of the scale, Tam and Yong34 reported complete nitrateremoval using immobilized C. vulgaris cells within calcium algi-nate beads. The treatment efficiency of our current method iscomparable with that of these aforementioned methods, althoughlarge variations among the experimental parameters; includingwastewater composition, microbial species, duration of theprocess, type of bioreactor, chemical composition and shape ofthe immobilization matrix, make direct comparison difficult.
In summary, we have established the use of cross-linked chito-san nanofiber mat as a water-insoluble and non-toxic support foralgal growth and nitrate removal from waters. Algal growth on asupport material can lead to combine algal harvesting, dewater-ing, and processing steps in a single stage. This bionano-compo-site material is potentially an attractive, simple and highlydurable polymer, with the mats still retaining their integrity aftersix months in contact with an aqueous solution, and has promisefor industrial and/or municipal wastewater treatment processes.
Acknowledgements
This work has been supported by The University of WesternAustralia and the Australian Research Council. We would liketo acknowledge the facilities of the Australian Microscopy &Microanalysis Research Facility at the Centre for Microscopy,Characterization & Analysis, The University of Western Austra-lia, which was funded by the University, State and Common-wealth Governments. AFM images were performed at CurtinUniversity.
Notes and references
1 Metcalf and Eddy, Inc, Wastewater Engineering: Treatment and Reuse,McGraw-Hill, New York, 4th edn, 2003.
2 N. Mallick, BioMetals, 2002, 15, 377.3 L. Hall-Stoodley, J. W. Costerton and P. Stoodley, Nat. Rev. Microbiol.,2004, 2(2), 95.
4 Y. Liu, M. H. Rafailovich, R. Malal, D. Cohn and D. Chidambaram,Proc. Natl. Acad. Sci. U. S. A., 2009, 106(34), 14201.
5 M. S. A. Hameed and O. H. Ebrahim, Int. J. Agri. Biol., 2007, 9, 183.
6 K. Abe, E. Takahashi and M. Hirano, J. Appl. Phycol., 2008, 20, 283.7 G. Schumacher and I. Sekoulov, Water Sci. Technol., 2002, 46, 83.8 E. Delahaye, R. Boussahel, T. Petitgand, J. P. Duguet and A. Montiel,Desalination, 2005, 177, 273.
9 A. Lavoie and J. de La Noue, J. World Maricul. Soc., 1983, 14, 685.10 A. Zamani, L. Edebo, B. Sjöström and M. J. Taherzadeh, Biomacromole-
cules, 2007, 8, 3786.11 W. A. Bough, Process Biochem., 1976, 11, 13.12 E. Wasik, J. Bohdziewcz and M. Blaszczyk, Process Biochem., 2001, 37,
57.13 H.-Y. Hu, N. Goto and K. Fujie, Water Res., 2001, 35, 2789.14 A. Elmidaoui, F. Elhannouni, S. M. A. Menkouchi Sahli, L. Chay,
E. Elabbassi, M. Hafsi and D. Largeteau, Desalination, 2001, 136, 325.15 Z. Feleke and Y. Sakakibara, Water Sci. Technol., 2001, 43, 25.16 K. Ohkawa, D. I. Cha, H. Kim, A. Nishida and H. Yamamoto, Macromol.
Rapid Commun., 2004, 25, 1600.17 M. Bradshaw, J. Zou, L. Byrne, K. S. Iyer, S. G. Stewart and
C. L. Raston, Chem. Commun., 2011, 47, 12292.18 D. Kaplan, D. Christiaen and S. M. Arad, Appl. Environ. Microbiol.,
1987, 53, 2953.19 R. H. Crist, K. Oberholser, N. Shank and M. Nguyen, Environ. Sci.
Technol., 1981, 15, 1212.20 B. Volesky and Z. R. Holan, Biotechnol. Prog., 1995, 11, 235.21 C. J. S. Bolch and S. I. Blackburn, J. Appl. Phycol., 1996, 8, 5.22 S. Fierro, M. del P. Sanchez-Saavedra and C. Copalcua, Bioresour.
Technol., 2008, 99, 1274.23 U. N. Dwivedi, S. Mishra, P. Singh and R. D. Tripathi, in Environmental
Bioremediation Technologies, ed. S. N. Singh and R. D. Tripathi,Springer, New York, 2007, ch. 16, pp. 353–389.
24 EPA, National Pesticide Survey: Project Summary, U.S. EnvironmentalProtection Agency, Washington DC, 1990.
25 S. Ghafari, M. Hasan and M. K. Aroua, Bioresour. Technol., 2008, 99,3965.
26 APHA, Standard Methods for the Examination of Water andWastewater, American Public Health Association, Washington, DC,18th edn, 1992.
27 H. K. Lichtenthaler and C. Buschmann, in Current Protocols in FoodAnalytical Chemistry, ed. R. E. Wrolstad, John Wiley & Sons Inc.,New York, 2001, pp. F4.3.1–F4.3.8.
28 L.-Q. Wu, P. Gadre Anand, H. Yi, M. J. Kastantin, W. Rubloff Gary,E. Bentley William, F. Gregory Payne and R. Ghodssi, Langmuir, 2002,18, 8620.
29 B.-M. Hsu, Parasitol. Res., 2006, 99, 357.30 H. Matsumoto, H. Yako, M. Minagawa and A. Tanioka, J. Colloid Inter-
face Sci., 2007, 310, 678.31 J. Naus and A. Melis, Plant Cell Physiol., 1991, 32, 569.32 N. Mallick and L. C. Rai, World J. Microbiol. Biotechnol., 1994, 10,
439.33 L. E. de-Bashan, Y. Bashan, M. Moreno, V. K. Lebsky and J. J. Bustillos,
Can. J. Microbiol., 2002, 48, 514.34 N. F. Y. Tam and Y. S. Wong, Environ. Pollut., 2000, 107, 145.
Hierarchical Patterning ofMultifunctional Conducting PolymerNanoparticles as a Bionic Platform forTopographic Contact GuidanceDominic Ho,†,‡ Jianli Zou,§ Xianjue Chen,†,0 Alaa Munshi,† Nicole M. Smith,†, ) Vipul Agarwal,†
Stuart I.Hodgetts,‡GilesW.Plant,^Anthony J.Bakker,‡AlanR.Harvey,‡ Igor Luzinov,# andK.Swaminathan Iyer*,†
†School of Chemistry and Biochemistry, The University of Western Australia, Crawley, Western Australia 6009, Australia, ‡School of Anatomy, Physiology and HumanBiology, The University of Western Australia, Crawley, Western Australia 6009, Australia, §Institute for Integrated Cell-Material Sciences (iCeMS), iCeMS Complex 2,Kyoto University, Yoshida-Honmachi, Sakyo-ku, Kyoto, 606-8501, Japan, )Experimental and Regenerative Neurosciences, School of Animal Biology, The University ofWestern Australia, Crawley, Western Australia 6009, Australia, ^Stanford Partnership for Spinal Cord Injury and Repair, Department of Neurosurgery,Stanford University School of Medicine, Stanford, California 94305, United States, and #School of Materials Science and Engineering, Clemson University, Clemson,South Carolina 29634, United States. 0Present address: Centre for NanoScale Science and Technology, School of Chemical and Physical Sciences, Flinders University,Bedford Park, Adelaide, SA 5042, Australia.
Exogenous electrical stimulation hasbeen effectively used both in clinicalpractice and in laboratory research
to regulate cell-type-dependent adhesion,differentiation, and growth.1 This phenom-enon of introducing programmed electricalsignals locally to influence biological eventshas resulted in the pursuit of sophisticatedmedical bionic devices.2 An important prop-erty that dictates the performance of mostbionic electrodes is the electrode/cellularinterface and its ability to transmit chargeacross the biointerface.3 Traditionally metal-lic electrodesmadeofplatinum, gold, iridiumoxide, tungsten, their alloys, and morerecently carbon fibers have been effectively
employed in bionic devices.4 They havebeen employed for deep brain stimulation,as cochlear implants, for vagus nerve stimu-lation to treat epilepsy, and for stimulatingregeneration in the central nervous sys-tem.2 However, stiff metal electrodes sufferamajor drawback of eliciting tissue damageover long-term implantation.2 Importantly,it is now recognized that nanoscale patternsprovide topographic guidance cues forcells. This has been widely exploited toengineer sophisticated regenerative plat-forms for nerves, muscles, skin, and bones.5
The need to incorporate large-area nano-scale patterns for bionic applications coupledwith the demand toward miniaturization of
Received for review November 20, 2014and accepted January 26, 2015.
Published online10.1021/nn506607x
ABSTRACT The use of programmed electrical signals to influence
biological events has been a widely accepted clinical methodology for
neurostimulation. An optimal biocompatible platform for neural
activation efficiently transfers electrical signals across the electrode�cell interface and also incorporates large-area neural guidance
conduits. Inherently conducting polymers (ICPs) have emerged as
frontrunners as soft biocompatible alternatives to traditionally used
metal electrodes, which are highly invasive and elicit tissue damage
over long-term implantation. However, fabrication techniques for the
ICPs suffer a major bottleneck, which limits their usability and medical translation. Herein, we report that these limitations can be overcome using colloidal
chemistry to fabricate multimodal conducting polymer nanoparticles. Furthermore, we demonstrate that these polymer nanoparticles can be precisely
assembled into large-area linear conduits using surface chemistry. Finally, we validate that this platform can act as guidance conduits for neurostimulation,
whereby the presence of electrical current induces remarkable dendritic axonal sprouting of cells.
biocompatible implantable devices has resulted inthe emergence of inherently conducting polymers asfrontrunners for fabricating flexible organic electrodematerials. However, advances in the applicability ofpatterned surfaces of inherently conducting polymersin bionic devices have been limited due to the diffi-culties of transferring printing techniques and theirintegration under physiological conditions. In thisarticle, we report a transferable method to fabri-cate multifunctional poly(3,4-ethylenedioxythiophene)-poly(styrenesulfonate) (PEDOT:PSS) nanoparticles anddirect their self-assembly by electrostatic interactionsinto large-area patterns. Using the rat pheochromocy-toma cell line (PC12), we demonstrate the suitabilityof the assembly as a bionic platform for exogenouselectrical stimulation.The three primary classes of conducting polymers
that have been studied are polyanilines, polypyrroles,and polythiophenes.6 The ease of functionalization ofpolythiophenes and maintenance of conductivity un-der physiological conditions has made them primarycandidates for multifunctional organic bionic devices.7
Themost widely explored processes for the fabricationof organic conducting polymer patterns are electro-polymerization, extrusion printing, inkjet printing,microcontact printing, electrospinning, and morerecently high-precision Dip Pen Nanolithography(DPN).4,6 Electropolymerization has been widely usedfor coating metal/carbon substrates, following whichpatterning is achieved by top-down lithography onpolymer thin films covering larger area electrodes. Thistechnique results in controlled, high-resolution nano-scale patterns but is limited by the ability to regulatepolymerization of monomers on nanoscale implanta-ble electrodes.8 Similarly, printing techniques haveachieved significant advances in recent years, reachinghigh-throughput patterns, but are limited in resolutionby the liquid dispensing techniques, which operatewithin the limit of tens of micrometers.9 Electrospin-ning techniques have offered simple processable solu-tions to generate 3D scaffolds at resolutionsmimickingthe extracellular matrix architecture but are limitedby the inability to generate patterned conductingconduits for the development of bionic guidancechannels.4 The aforementioned shortfalls have beenrecently overcome by the advances of DPN, whichenables precise deposition, patterning down to nano-scale resolution, and most importantly applicabilityover a wide range of substrates.10 However, advancesare limited by their cost, need for specialized equip-ment, and low throughput. In the present paper weadopt a bottom-up self-assembly process to preciselypattern conducting polymer nanoparticles into pat-terns as conduits for guidance. The approach is easilyadoptable over multiple substrates, needs no specia-lized equipment, and affords large-area patterns. Impor-tantly, this approach enables drug encapsulation and
sustained release from the nanoparticles once pat-terned and multimodal imaging of the nanoparticleconstructs once implanted.
RESULTS AND DISCUSSION
Patterned Multifunctional PEDOT:PSS Nanoparticle Arrays.In this study poly(glycidal methacrylate) (PGMA) isused as a reactive macromolecular anchoring platformboth on the substrate as a nanoscale layer and as acolloidal nanoparticle to enable multilayer assembly(Figure 1). A polymer with epoxy functionality waschosen, since the reactions of epoxy groups are uni-versal and easily transferable to various substrates,affording ease of attachment of functional molecules.Furthermore, the epoxy groups of the polymer cancross-link to provide structural integrity to the patternand nanoparticle constructs.11 The mobility of thereactive loops of PGMA ensures greater access toanchoring, resulting in a 2�3-fold greater graftingdensity when compared to a monolayer of epoxygroups on a nanoparticle surface of similar dimension,enabling high loading using a layer by layer approachthat is adopted in the current study.11 Polymer nano-spheres were initially prepared using an oil in wateremulsion methodology from PGMA modified with arhodamine-B (RhB) dye, encapsulated with magnetite(Fe3O4) nanoparticles to form the core platform(Figure 1a,b). Not only does the incorporation ofmagnetite and RhB render these constructs multi-modal for both MRI and fluorescence imaging, butimportantly in the present case magnetite provides ameans to separate, wash, and purify the nanoparticlesusing a magnetic fractionation column during eachstep of layered assembly. Polyethylenimine (PEI) wasthen covalently bound to the RhB-PGMA core to facil-itate a cationic layer for electrostatic conjugation of ananionic conducting polymer, PEDOT:PSS (Figure 1c,d).Capillary force lithography (CFL) was then usedto generate large-area nanoscale conduits in whichPEDOT:PSS nanoparticles are electrostatically directedto self-assemble as linear channels from solution(Figure e,f). Capillarity allows the polymer melt to fillup the void space between the polymer and theapplied mold when the temperature is above theglass-transition temperature (Tg), thereby generatinga large-area pattern that depends on the size of stamp.Importantly, the technique needs no specializedinstrumentation for generation of large-area patterns.Patterns can easily be generated using polydimethyl-siloxane (PDMS) stamps, which in turn can be fabri-cated using the ubiquitous optical storage discs as amaster. An optical data storage disc is typicallymade ofa polymer (polycarbonate) disc, onwhich a single spiraltrack is drilled. The typical width and depth of each linein the spiral track are 800 and 130 nm, respectively, andthe periodicity of the track is∼1.5 μm (Figure S1). In thepresent study, an indium tin oxide (ITO) substrate was
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modified first by spin coating a thin film of PGMAfollowed by a second spin-coated layer of polystyrene(PS) using previously reported conditions.12 The PSlayer acts as a chemical resist to selectively react theepoxy groups of PGMA following patterning. ThePS/PGMA bilayer was annealed with the PDMS maskat 130 �C (T> Tg of PS) to induce patterning via capillaryflow. The reusable PDMS stamp was peeled off fol-lowing heat treatment to obtain a patterned sur-face resulting in alternating PGMA and PS stripes.
Ethylenediamine (EA) was then grafted to PGMA toresult in cationic linear patterns. PEDOT:PSS nano-particles were then electrostatically assembled ontothe patterned surface, followed by washing steps toremove PS to obtain linear arrays of assembled PEDOT:PSS nanoparticles. A detailed schematic of the fabrica-tion process is shown in Figure S2. The nanoparticleand the patterns were characterized at each step of theassembly (Figure 2). The PEDOT:PSS nanoparticleswere an average size of 200 nm (Z-average) with a
Figure 1. Schematic illustration of the fabrication protocol to pattern multifunctional PEDOT:PSS nanoparticle arrays forexogenous electrical stimulation. (a�d) Multilayer assembly of conducting PEDOT:PSS nanoparticle fabrication via non-spontaneous emulsification. (a) An organic phase is initially formed by dissolving RhB-modified PGMA (yellow) and Fe3O4
(purple) in a 1:3mixture of CHCl3 andMEK. (b) Colloidal fluorescent PGMA-Fe3O4 nanoparticles are fabricated upon dropwiseaddition of the organic phase to an aqueous solution of Pluronic F-108. (c) Cationic second layer via covalent attachment ofPEI (green) to the PGMA-Fe3O4 core. (d) Anionic conducting polymer layer via electrostatic attachment of PEDOT:PSS (blue).(e�g) Patterning of the multilayered PEDOT:PSS nanoparticles for exogenous electrical stimulation of PC12 cells. (e) Linearnanoparticle conduits patternedon a substrate via capillary force lithography (CFL) using charge complementarity. A detailedschematic of the CFL procedure can be found in Figure S2. (f) PC12 cells (green) were cultured onto the biocompatibleplatform, followed by (g) exogenous electrical modulation.
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Figure 2. Characterization of the multilayered PEDOT:PSS conducting nanoparticles and their assembly as linear conduits.(a) TEMmicrograph of the multilayered PEDOT:PSS nanoparticles. Scale bar = 200 nm. (Inset: high-magnification TEM imageof PEDOT:PSS-coated nanoparticles showing encapsulated Fe3O4 nanoparticles. Scale bar = 10 nm.) (b) DLS particle sizedistributions of the PEDOT:PSS nanoparticles in solution. (c) Zeta potential distributions of the nanoparticles: PGMA-Fe3O4
core (black) with an average zeta potential of 3.9 ( 1.3 mV, cationic PEI-coated (red) with an average zeta potential of 37 (1.2 mV, and anionic PEDOT:PSS-coated (blue) with an average zeta potential of �29 ( 6.15 mV. (d) Current vs voltageresponse for the nonconducting PEI-coated nanoparticles (red) and conducting PEDOT:PSS-coated (black) nanoparticles.(e�g) Tappingmode AFM topography images of the nanoparticle patterns at each stage of fabrication: PGMA and PS stripes(e), EA-modified PGMA and PS stripes (f), PEDOT:PSS nanoparticle patterns (g). (h�j) Corresponding height profiles of thenanoparticle patterns at each stage of fabrication.: PGMA and PS stripes (h), EA-grafted PGMA and PS stripes (i), PEDOT:PSSnanoparticles patterns (j). The AFM line scans corresponding to the height profiles are indicated on the topography images in(e)�(g). (k, l) SEM micrographs of the nanoparticle patterns at a magnification of 25k� (k) and 11k� (l) indicating theformation of tightly packed and highly ordered nanoparticle arrays. (m) Confocal fluorescence image of the RhB-functionalized PEDOT:PSS nanoparticle arrays at 20� magnification.
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polydispersity index (PDI) of 0.07, a zeta potential of�29( 6.15mV, and a conductivity of 2.5� 10�12 S/cm.Themeasured conductivity is in accordance with othervalues reported in the literature for polymer blends.13
Importantly, this low conductivity is important underphysiological conditions to induce local cellular stimu-lation and avoid tissue damage due to toxic over-stimulation.14 The final self-assembled linear arraysof PEDOT:PSS nanoparticles were of large-area high-density packing, as confirmed at various length scalesusing AFM, SEM, and fluorescence imaging.
Biocompatibility Assessment of the PEDOT:PSS NanoparticleArrays. Topographic modulation of tissue response isone of the most important considerations in develop-ing bionic implants. Topographic contact guidanceusing micropatterns has been widely exploited toinfluence cell migration, adhesion, and prolif-eration.15,16 One of the pivotal first steps in the pres-ent study was to establish biocompatibility of thepatterned structures. PC12 cells were chosen in thepresent case, as they have been demonstrated to show
enhanced neurite outgrowth and spreading uponexogenous stimulation on a conducting polymersubstrate.17 MTS (3-(4,5-dimethylthiazol-2-yl)-5-(3-carboxymethoxyphenyl)-2-(4-sulfophenyl)-2H-tetrazo-lium, inner salt) assays, cell viability assays, and SEMimaging were performed after exogenous electricalstimulation and in the absence of electrical stimulationto determine effects on cell viability and cell adhesion(Figure 3a,b and Figure S3). The stimulation conditionsused in the present study involved a monophasicpulsed current at a frequency of 250 Hz with a 2 mspulse width and an amplitude of 1mA for 2 h, similar toprotocols previously reported for similar cell lines.18,19
Importantly, we observed no changes in cell viabilityupon exogenous stimulation and observed preferen-tial adhesion of the PC12 cells to the patterned sur-face over a nonpatterned surface in both cases(( stimulation). High-magnification SEM imaging (nostimulation) further revealed preferential interaction ofthe PC12 cells to the PEDOT:PSS nanoparticle arrays,confirming not only biocompatibility with the large
Figure 3. Biocompatibility of the PEDOT:PSS nanoparticle arrays with PC12 cells. (a) Cell viability determined using MTScalorimetric assay obtained at 72 h after an initial exogenous electrical stimulation for 2 h and in the absence of stimulationshowing no significant changes. (b) SEM micrograph demonstrating preferential cell adhesion to the pattern area (yellowbox). Image acquired at 323�magnification 72 h after the addition of NGFwithout exogenous electrical stimulation. (c) High-magnification (12k� magnification) SEM images demonstrating specific and preferential interactions of neurites (whitearrows) with the PEDOT:PSS linear conduits (red arrows).
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area of the pattern but also potential applicabilityof the nanoscale linear arrays as guidance conduits(Figure 3c).
Exogenous Electrical Stimulation Induced Dendritic Sproutingof the PC12 Cells. Electrical stimulation has been effec-tively used to modulate growth and differentiation of
anchorage-dependent cells such as neurons, fibro-blasts, and epithelium cells.17,20,21 In the central ner-vous system, brief stimulation to the proximal end oftransected peripheral nerves has been shown to aug-ment preferential motor reinnervation,22 improve thespecificity of sensory reinnervation,23 and accelerate
Figure 4. Exogenous electrical stimulation induced dendritic sprouting of the PC12 cells guided by the PEDOT:PSS linearconduits. (a) Significant increase in the average cell area is observed 72 h after exogenous electrical stimulation on thenanoparticle platform in comparison to unstimulated and nonpatterned controls. (b) Corresponding decrease in PC12 cellproliferation observed 72 h after exogenous electrical stimulation on the nanoparticle platform in comparison tounstimulated and nonpatterned controls. (c�f) Representative confocal images (40� magnification) of β-III tubulinimmunohistochemically stained cells 72 h after the following treatments: {(þ) pattern, (�) stimulation} (c); {(þ) pattern,(þ) stimulation} (d); {(�) pattern, (�) stimulation} (e); {(�) pattern, (þ) stimulation} (f), demonstrating modulation of cellmorphology. (g) High-magnification SEM image (magnification 8k�) indicating the formationof extensive dendritic networks(white arrows) guided by the PEDOT:PSS arrays. Inset: The corresponding low-magnification image of the area (yellow box)analyzed (magnification 3k�).
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the reinnervation of distal target tissues.24 These havebeen reported to depend on depolarization of theneuronal soma and its axon, involvement of axonguidance factors such as polysilylated neural celladhesion molecule,25 the L2/HNK-1 carbohydrate,26
and brain-derived neurotrophic factor.27 Finally elec-trical stimulation induced neurite outgrowth wasrecently reported to be dependent on calcium influxthrough L- and N-type voltage-dependent calciumchannels and calcium mobilization from IP3R andRYR-sensitive calcium stores.28 In the present case,we analyzed the morphological modulation of PC12cells following electrical stimulation having deter-mined no change in cell viability using the MTS assay.Nerve growth factor (NGF) induces PC12 cells tochange their phenotype and acquire a number ofproperties that are similar to sympathetic neurons.Importantly, although they can acquire propertiessimilar to sympathetic neurons upon NGF treatment,they do not develop definitive dendritic axons or formtrue synapses with each other in the absence ofexogenous stimulation.29 This change in phenotypeupon NGF treatment is associated with a retardation inproliferation, the extension of neurites making themelectrically excitable. Monitoring the cell numbers andcell area can assess this change from the proliferationstate to a differentiation state. Furthermore, micro-tubule levels correlate precisely with the neurite
extension during NGF-induced PC12 cell differ-entiation.29,30 Using immunohistochemical stainingfor β-III tubulin it was determined that stimulation onthe patterned surface resulted in a significant increasein the cell area and lower number of cells per unit area,indicating exogenous electrical stimulation induceddifferentiation of PC12 cells (Figure 4a�f, Figure S4).High-magnification SEM (Figure 4g) also revealedthat stimulation resulted in an extensive dendriticnetwork guided by the linear conduits of PEDOT:PSSnanoparticles.
CONCLUSION
In summary, we have demonstrated a practical andtransferable protocol to fabricate self-assembled large-area patterns of conducting polymers from solution.This overcomes some of the shortfalls in the currentfabrication techniques in developing patterned organicbionic devices. The patterns generated have demon-strated excellent biocompatibility. At the same time,they have been shown to induce exogenous electricalstimulation under physiological conditions to elicit ameasurable and consistent cellular response. Impor-tantly the methodology permits the design of bionicdevices capable of inducing local electrical stimulationfor in vivo applications while integrating multimodalimaging and simultaneous drug delivery capabilities ofnanoparticles.
METHODS SUMMARY
Nanoparticle Synthesis. The conducting nanoparticles wereprepared via a nonspontaneous emulsification route. Briefly,rhodamine Bwas attached to PGMA inMEK at 80 �Cunder N2 for5 h. The modified PGMA was then precipitated in diethyl etherand dried under N2. This was dispersed in a 1:3 mixture of CHCl3and MEK along with 25 mg of Fe3O4 to form the organic phase.This organic phase was added dropwise into a rapidly stirringaqueous solution of Pluronic F-108. The emulsion was homo-genized with a probe-type ultrasonic wand for 1 min. Theorganic solvents were then evaporated off under N2. Largeaggregates of Fe3O4 and excess polymer were separated viacentrifugation. The nanoparticles in the supernatant were thenmixed with PEI and heated to 80 �C for 16 h to facilitateattachment. The PEI-coated nanoparticles were isolated andwashed on a magnetic separation column. Next, a dilutedsolution of PEDOT:PSS was added dropwise under rapid stirringto nanoparticles at a concentration of 0.5 mg/mL to facilitateelectrostatic attachment. This was followed by sonication for10min and stirring for 18 h. The nanoparticles were thenwashedmultiple times inwater before being stored at 4 �C for further use.
Platform Fabrication. To direct the self-assembly of the nano-particles, a template was fabricated by CFL. A 0.2%w/v PGMA inCHCl3 solution was spin coated on ITO coverslips and annealedat 120 �C for 20 min. Next, 1.3% w/v PS in toluene was spincoated onto the PGMA surface. A PDMS stamp was then placedonto the PS layer, followed by heat treatment in an oven at130 �C for 1 h. Once cooled, the stamp was peeled off. This wasfollowed by exposure to EA at room temperature for 5 h.The pattern was next washed multiple times with water toremove unreacted EA. A 50 μL amount of 4mg/mL nanoparticlesuspension was drop casted onto the patterned area of thecoverslip. The setup was then placed in a sealed vial, facilitating
controlled evaporation, which allowed for electrostatic nano-particle attachment onto the EA surface. The PS mask was thenremoved by washing with toluene. The resulting patternedPEDOT:PSS nanoparticle array was then used for furtherexperimentation.
Electrical Stimulation Protocol. For electrical stimulation experi-ments, two silver epoxy electrodes were painted onto the endsof the patterned nanoparticle arrays. Prior to cell culture, thewhole platform was UV and ethanol sterilized. Wells werecoated with poly(L-lysine) and 15 μg/mL of laminin followedby cell seeding at a density of 50 000 cells/well. Cells were left toadhere for 18 h. Immediately prior to stimulation, the prolifera-tion media was replaced with low-serum nerve growth factorcontaining differentiation media. For stimulation, the cells weresubjected to a monophasic pulsed current at a frequency of250 Hzwith a 2ms pulsewidth and an amplitude of 1mA for 2 h,after which they were left for an additional 72 h before analysis.
Cell Viability Assessment. Cell viability was measured using theMTS assay as per the manufacturer protocols (Invitrogen, UK).For measurements, 80 μL from each well was transferred into anew 96-well plate and read under a plate reader at 490 nmexcitation wavelength. To analyze cell morphology, cells wereimmunohistochemically stained for β-III tubulin.
Material Characterization. AFMwas performed on a Dimension3100 AFM systemwith a Nanoscope IV controller used to obtainthe AFM images in tapping mode, using Pt/Ir-coated contactmode probes with a spring constant of 0.2 N/m (type SCM-PIC,Bruker). TEM was performed on a JEOL 2100 transmissionelectron microscope at an accelerating voltage of 80 kV. SEMwas performed on a Zeiss 1555 VP-FESEM, and all samples werecoated with 5 nm of Pt. Biological samples were initially fixed in2.5% glutaraldehyde and dehydrated in increasing concen-trations of ethanol followed by critical point drying prior toPt coating. Immunohistochemically stained samples were
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analyzed using a Leica TCS SP2 AOBS multiphoton confocalmicroscope.
Conflict of Interest: The authors declare no competingfinancial interest.
Supporting Information Available: Detailed materials andmethods: synthesis, characterization (TEM, SEM, AFM), cellculture, and electrical stimulation experiments. This material isavailable free of charge via the Internet at http://pubs.acs.org.
Acknowledgment. D.H., I.L., and K.S.I. designed the experi-ments, developed the concept, and analyzed the data. D.H., J.Z.,and N.M.S. optimized the capillary force lithography experi-ments. D.H., X.C., V.A., and A.M. performed image acquisitionusing confocal microscopy, transmission electron microscopy,scanning electron microscopy, and atomic force microscopy.D.H., A.R.H., G.W.P., S.I.H., and A.B. optimized and designed theelectrical stimulation experiments. This work was funded bythe Australian Research Council (ARC), the National Health &Medical Research Council (NHMRC) of Australia, and the NationalScience Foundation (CBET-0756457). The authors acknowledgetheAustralianMicroscopy&Microanalysis Research Facility at theCentre for Microscopy, Characterization & Analysis, and TheUniversity of Western Australia, funded by the University, Stateand Commonwealth Governments. The authors also wish tothank Margaret Pollett and Chrisna LeVaillant for their invaluablecontribution in assisting with the PC12 cell cultures and immu-nohistochemistry, and Ella Marushchenko (www.scientific-illustrations.com) for assistance with Figure 1.
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3. Wallace, G. G.; Moulton, S. E.; Clark, G. M. Electrode-CellularInterface. Science 2009, 324, 185–186.
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ARTIC
LE
S1
Hierarchical Patterning of Multifunctional Conducting Polymer Nanoparticles as a Bionic
v/v), L-glutamine (1 % v/v) and nonessential amino acids (1 % v/v). For PC12 differentiation,
cells were cultured in differentiation media consisting of DMEM, L-glutamine (1 % v/v), horse
serum (1 % v/v) and nerve growth factor (50 ng/mL).
Electrical Stimulation Experiments. Prior to stimulation experiments, two silver epoxy
electrodes were painted onto the ends of the prepared NP array and platinum wires attached to
allow for connections with the stimulator. Next, a cell culture well was created by first cutting a
1.5 mL microcentrifuge tube in half and then sealing the capped end with silicon vacuum grease.
This was stuck onto the ITO glass with the patterned arrays in the centre of the well. This was
done to ensure that the electrodes did not come into direct contact with the cell culture media.
The array was then placed in a Petri dish to maintain sterility throughout the course of the
experiment (Fig S5a). Prior to culturing cells on the arrays, the coverslips were UV sterilised (20
mins) and then washed with 70 % ethanol three times. Wells were then coated with poly-(L-
lysine) and 15 µg/mL of laminin. Cells were then seeded at a density of 50 000 cells/well and left
to adhere for 18 h. Prior to stimulation, the proliferation media was replaced with differentiation
media. The cells were then stimulated according to protocols as listed below. Following
stimulation, the cells were left for a further 72 h with fresh differentiating media added every 48
S6
h. Photographs of the electrical stimulation setup are described in Figure S5.
Stimulation Protocol. The electrical signals were supplied by Grass S44 Stimulator (Quincy,
Massachusetts, USA). The stimulation regime is similar to that used by Wallace et al.3-5 Briefly,
the cells were subjected to a monophasic pulsed current at a frequency of 250 Hz with a 2 ms
pulse width and an amplitude of 1 mA for 2 h.
Cell Viability Assays. Cell viability was measured using the (3-(4,5-dimethylthiazol-2-yl)-5-(3-
carboxymethoxyphenyl)-2-(4-sulfophenyl)-2H-tetrazolium, inner salt) (MTS) assay as per the
manufacturer protocols (Invitrogen, UK). Cells were plated as per “electrical stimulation
protocol” stated above. Viability was to be determined at 3 time points: (i) 0 h (immediately
prior to electrical stimulation), (ii) 72 h after the addition of differentiation media and (iii) 72 h
after the addition of differentiation media and electrical stimulation. For measurements, 80 µL
from each well was transferred into a new 96 well plate and read under a plate reader at 490 nm
excitation wavelength. The same protocol was followed for every sample and each measurement
was carried out in triplicate.
Immunohistochemical Staining. The PC12 cells were immunohistochemically stained for ß-III
tubulin. The cells were fixed in 4 % paraformaldehyde for 10 mins. Cells were first incubated
with a primary antibody solution containing PBS, 10 % Normal Goat Serum, 0.1 % Triton X-100
and the anti-β-III tubulin antibody (1:1000, anti-rabbit, Covance) at room temperature for 30
mins. After 3 PBS washes, the antibody binding was visualised with anti-rabbit FITC (1:100,
Sigma) following incubation for 30 mins at room temperature. Coverslips were mounted on glass
slides covered with Dako Fluorescent Mounting Medium (Dako, USA). All experiments were
S7
performed in triplicate.
Confocal and Fluorescence Microscopy Analysis. Immunohistochemically stained samples
were analysed using confocal and fluorescence microscopy. Confocal microscopy was carried
out using a Leica TCS SP2 AOBS Multiphoton Confocal microscope and fluorescence
microscopy with a Diaplan fluorescence microscope.
Image and Statistical Analysis. To determine the effects both stimulation and the NP arrays had
on the PC12 cells, the average area of each cell was determined. 3 randomly selected areas on
each sample was visualised at 40 x magnification. The average area covered by each cell was
assessed using Image J analysis software (version 1.48a, NIH). All immunohistochemical
analyses were conducted by a single investigator, ensuring constant selection criteria, and results
expressed as means ± SD. Data were analysed using Origin data management software to
conduct ANOVA on groups of data. Statistically significant differences between each treatment
were determined using Bonferroni/Dunn post hoc tests (p≤0.05).
Scanning Electron Microscopy (SEM). Prior to SEM imaging, samples without cells were
coated with 5 nm of Pt. Samples with cells were fixed in 2.5 % glutaraldehyde for 2 h at 4 oC and
dehydrated. Samples were washed with deionized water and dehydrated in a microwave in serial
concentrations of ethanol (50 %, 70 % and 90 % once then 3x in absolute ethanol), before critical
point drying with carbon dioxide for 1h and then coating with 5 nm of Pt. Samples were imaged
using a Zeiss 1555 VP-FESEM.
Transmission Electron Microscopy (TEM). Synthesized polymer nanoparticles were drop-
casted on carbon coated TEM grids and imaged with an accelerating voltage of 100 kV on a
S8
JEOL 2100 transmission electron microscope.
Atomic Force Microscopy. A Dimension 3100 AFM system (Bruker) with a Nanoscope IV
controller (Bruker) was used to obtain the AFM images in Contact Mode, using Pt/Ir coated
contact mode probes with a spring constant of 0.2 N/m (type SCM-PIC, Bruker). The scan
parameters were adjusted to ensure reliable imaging with the smallest possible contact force
setpoint. Data analysis was performed using the SPM analysis freeware Gwyddion
(http://gwyddion.net).
Dynamic light scattering (DLS) and zeta potential measurements. DLS experiments were
performed using a Malvern Zetasizer Nano series. For measuring the size distribution, 5
measurements were taken and in each measurement there were 10 data acquisitions. Zeta
potential (ζ) measurements were performed using the same instrument. Measurements for each
sample were recorded in triplicate and 100 data acquisitions were recorded in each measurement.
All measurements were recorded at 25 oC in Malvern disposable clear Folded Capillary Cells.
S9
Supplementary Figures
Figure S1. (a) The PDMS stamp used in the study. SEM micrograph of the grooved structure of
the PDMS. Image taken at 6k x magnification; (b) Photograph of the polycarbonate disc peeled
from a CD. PDMS was cast on the grooved surface and stamps of the desired size were cut out.
(a) (b)
S10
Figure S2. Schematic of CFL procedure. Briefly, ITO substrate was modified with a thin layer
of PGMA followed by second layer of PS; a PDMS stamp was placed over the PS film and heat
treated at 130 oC; PDMS stamp was peeled off after cooling; EA was selective reacted to the
exposed PGMA stripes to produce cationic stripes to enable charge complementarity to assemble
the anionic PEDOT:PSS nanoparticles. The PS mask was removed by washing with toluene, to
obtain linear PEDOT:PSS conduits.
T > Tg (PS) (130 oC)
EA grafting onto PGMA Conducting NPs
PS removal
Peel off PDMS Stamp
Electrostatic attachment
S11
Figure S3. SEM micrograph of PC12 cells 18 h after plating and immediately prior to electrical
stimulation. PC12 cells on the patterned surface (yellow box) were evenly spread out, in
comparison to the rounded cells on non-patterned areas of the substrate demonstrating
preferential adhesion. Image taken at 1000 x magnification.
S12
Figure S4. Representative low magnification (magnification 400 x) SEM micrographs of PC12
cells 72 h after the following treatments: (a) (+) pattern, (-) stimulation and (b) (+) pattern, (+)
stimulation demonstrating lower coverage due to reduction in proliferation upon stimulation.
.
(b) (a)
S13
Figure S5. Photographs of the electrical stimulation setup: (a) A sterile Petri dish containing the
modified cell culture well (red arrow) and the Platinum wires which allow for connections to the
stimulator (green arrows), (b) The stimulator (yellow arrow) was placed next to an incubator and
the wires from the machine leading into the stimulator (blue arrows), (c) The wires from the
stimulator were connected to the platinum wires via alligator clips.
(a)
(c)
(b)
S14
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