Porous Chitin Microbeads for More Sustainable Cosmetics - ACS

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Porous Chitin Microbeads for More Sustainable Cosmetics Catherine A. King, Julia L. Shamshina, Oleksandra Zavgorodnya, Tatum Cutfield, Leah Elizabeth Block, and Robin D. Rogers ACS Sustainable Chem. Eng., Just Accepted Manuscript • DOI: 10.1021/ acssuschemeng.7b03053 • Publication Date (Web): 30 Oct 2017 Downloaded from http://pubs.acs.org on November 1, 2017

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ACS Sustainable Chemistry & Engineering

Porous Chitin Microbeads for More Sustainable Cosmetics*

Catherine A. King,1 Julia L. Shamshina,2 Oleksandra Zavgorodnya,3 Tatum Cutfield,1 Leah E. Block,3 and Robin D. Rogers1,3,4,*

1

Department of Chemistry, McGill University, 801 Sherbrooke Street West, Montreal, QC, H3A 0B8, Canada

2

Mari Signum, Ltd., 3205 Tower Oaks Boulevard, Rockville, MD 20852 USA

3

Department of Chemistry, The University of Alabama, Tuscaloosa, AL 35487, USA

4

525 Solutions, Inc., 720 2nd Street, Tuscaloosa, AL 35401, USA

KEYWORDS: Microspheres, porous beads, chitin, ionic liquids, cosmetic microbeads *Corresponding author: [email protected]

Abstract: The microbead form is a material architecture promising for use in biomedical and cosmetic applications; however, the use of petroleum-based microbeads (i.e., plastics) has raised significant environmental concerns in recent years. Microbeads prepared from renewable polymers could represent a sustainable alternative to these synthetic microbeads. This work explores the use of chitin in preparing biodegradable, biocompatible microbeads of low toxicity. Chitin microbeads were synthesized using the ionic liquid (IL) 1-ethyl-3-methylimidazolium acetate ([C2mim][OAc]); the IL was used to both extract chitin directly from waste shrimp shell

*

In honor of the groundbreaking work done by Prof. István T. Horváth in Green Chemistry and Sustainability

1 ACS Paragon Plus Environment


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

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and to prepare the porous microbeads by coagulation in polypropylene glycol (PPG). The effects of biopolymer source and bead preparation parameters on the formation of beads were investigated, as well as the effects of the drying conditions on the dry bead structure. It was found that IL-extracted chitin could be used to prepare beads of homogenous size distribution (with 60% of beads 125-250 µm) and shape, while commercially available practical grade chitin could not, suggesting that high molecular weight chitin is required for bead material formation. Supercritical CO2 drying and lyophilization of the wet beads led to dry chitin beads with an opaque appearance, porous interiors, and uniform shape. Loading and release studies of representative active compounds (indigo dye and sodium salicylate) into the chitin beads indicated that the dry beads could be easily loaded from an aqueous solution of active and could release 90% of the active compound within 7 h in DI water at room temperature.

INTRODUCTION Microbeads are spherical solid particles with diameters from 5 µm to 1 mm, which are generally produced from polyethylene (PE), polypropylene (PP), polyethylene terephthalate (PET), polymethyl methacrylate (PMMA), or nylon plastics.1 The use of microbeads is becoming more popular due to improvements in microsphere quality and functionality, and accordingly, they have increasingly been used in food science2 and separations,3 and as exfoliants in cosmetics and personal care products.3 Additionally, they are gaining popularity in biomedical applications, including uses in medical diagnostics as injectable biomaterials, as reagents in diagnostic devices, and as drug delivery vehicles.4 In biotechnological applications, microspheres are less prone to aggregation, providing significant advantages over non-spherical particles which tend to aggregate, complicating injection and delivery to the targeted sites. In


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addition, the microsphere surface can be functionalized if necessary to induce a desired response within the surrounding tissue.5,6 Personal care companies (including Unilever, Target, Johnson & Johnson, Procter & Gamble, and L’Oréal) were some of the largest producers of microsphere containing products.7 In 2014, the global market for microspheres had attained $2.3 billion and was expected to reach $3.5 billion by 2020, registering a compound annual growth rate (CAGR) of 7.8% from 2015 to 2020. The medical technology market segment for microspheres alone was expected to grow from $504 million in 2015 to $810 million in 2020, at a CAGR of 10.0% from 2015-2020.7 However, as synthetic polymers have found widespread application in the global market, they have become more ubiquitous in the environment, leading to increasing concern about their environmental impacts.8 Although quantitative studies on routes of entry of microbeads into the environment are limited, it has been suggested that the effluent of waste water treatment plants acts as one such route.. One study, for example, demonstrated that though treatment plants retain a majority of the microplastic fragments, there was still substantial leakage of microplastics from the effluent water.9 The Australian Environment Protection Authority (EPA Australia) recently reported a UK case study that anticipated that between 4,594 and 94,500 microbeads can leak to the environment per a single use of a microbeads-containing face scrub.10,11 Another study suggested the average abundance of microplastic beads in some locations to be 43,000 microplastic particles/km2.12 Although their fate once released is not fully understood, it is known that microplastics can leach toxic additives, adsorb pollutants, enter the marine food chain, and contaminate large bodies of water, from lakes to oceans.13-15 All of this has raised public support for the banning of microbeads, and has prompted action from non-governmental organizations, multinational corporations, and policy-makers.16 Thus, in



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2015, the United States enacted federal legislation to ban microbeads in rinse-off cosmetics,17 causing many big players in the cosmetic industry to begin eliminating microbeads from their production lines. Companies such as L’Oréal, Johnson and Johnson, and Crest have all begun phasing our microbeads in their personal care products. In fact, both Johnson & Johnson and Crest have plans in place for a complete phase out of microbeads in their products globally by the end of 2017.18 Biopolymers have recently seen increased interest as a promising alternative to petroleumbased polymers,19 and could be applied here to the preparation of microbeads. Chitin, the second most abundant biopolymer, has many inherent properties such as natural biodegradability, biocompatibility, non-toxicity, and occurs in abundance in waste sources (such as crustacean shells),20 which can microbially degrade. Because of all of this, it is of particular interest for use in many of the areas in which microbeads are used, especially in cosmetics, personal care,21 pharmaceutical, and medical applications (e.g., drug delivery22 and enzyme immobilization23). In recent years, our group has reported extraction and regeneration of high molecular weight chitin directly from shrimp shell waste using the ionic liquid (IL) 1-ethyl-3-methylimidazolium acetate ([C2mim][OAc]). Using solution processing, we have prepared various material architectures such as fibers,24-26 films,27 and nanomats.28-30 We hypothesized, based on our work with these architectures and work we have done in the preparation of beads from IL/cellulose solutions,31 that chitin could be used for the preparation of microbeads which would be useful in specific applications such as the delivery of active ingredients within cosmetic applications. Not to mention, the use of such a biodegradable biopolymer in place of synthetic ones would avoid many of the environmental problems associated with synthetic microplastic beads.


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General preparation methods for synthetic microspheres include including spray-drying,32 emulsification,33 and ionic gelation.34,35 However, as chitin dissolution is difficult in most solvent systems, only spray-drying and emulsion methods have been used to make chitin beads, with very few studies being conducted with a commercial product in mind. Additionally, these methods require harsh solvents and lead to poor control of the sphere size and shape.36 The preparation of neat chitin beads has been done via toxic or corrosive solvents such as dimethylacetamide/lithium chloride (DMAc/LiCl),37 where chitin microspheres were obtained for enzyme immobilization. Methods using aqueous sodium hydroxide-urea (NaOH/urea) eutectic have been used as well, in one case for the direct preparation of beads. However, it should be noted that the chitin was pretreated with NaOH, which could lead to deacetylation and the formation of chitosan.38 This solvent system was also used for the preparation of chitin microspheres via the formation and processing of nanofibers (though again, the chitin was pretreated with NaOH).3940 Aside from neat chitin microspheres, composites of chitin with additives, and chitosan (the deacetylated form of chitin) based microspheres have been prepared. Composite microspheres of chitin and silica have been prepared from the IL 1-butyl-3-methylimidazolium acetate ([C4mim][OAc]), however, these were pressed into a mold, and never studied as individual beads.41 Composite microspheres from chitin and poly(D,L-lactide-co-glycolide) (PLGA) have also been prepared.42 Cross-linked chitosan beads have been reported as well, using DMAc/LiCl and aqueous acetic acid (CH3COOH), respectively.43 Here, we set out to demonstrate the possibility of obtaining pure chitin microbeads, uniform in size, using a nontoxic solvent and coagulation medium. We aimed to develop an approach which would allow control of microsphere size and shape and eliminate process waste generated



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during current production methods in the design of more environmentally-friendly processes, in order to develop a scalable process for microbeads suitable for commercial use. Scalable continuous processes have been developed for the production of cellulose beads,44 but as of yet there has been no scalable, reliable method reported for the production of neat chitin microspheres. Here, the IL [C2mim][OAc] was used for the extraction of high molecular weight chitin from shrimp shells, and also for the solution processing of the biopolymer into the microsphere architecture. We also studied the effect of the source of chitin (chitin extracted from biomass with [C2mim][OAc] and commercially available chitin) on bead synthesis using IL processing. Batch processing parameters such as the chitin solution properties, coagulation phase, and formation parameters were optimized for the preparation of uniform chitin beads. Loading and release of the chitin beads with representative active ingredients were tested with indigo dye and sodium salicylate.

RESULTS AND DISCUSSION Bead Preparation The preparation of neat chitin beads was attempted with both IL-extracted shrimp shell chitin and PG chitin from IL solution. The formation of homogenous, viable beads, as well as the size and shape of the beads formed is dependent on the chitin-IL solution properties, properties of the coagulation phase, and the process parameters. Our previous publications have demonstrated that the ability to form a material depends on solution properties of the chitin-IL solution, such as the chitin source used, viscosity of the chitin/IL solution, and temperature of the chitin/IL


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solution.27,30 Other literature has suggested that the process parameters (such as rate of addition of the polymeric solution to the coagulation phase) are crucial for the size control of beads.45,46 The method used here for bead preparation was based on that of cellulose in IL.31 As our previous studies with cellulose had suggested that the viscosity of the biopolymer/IL solution has a direct effect on the ability to form the bead architecture,31 the first focus in the attempt to prepare neat chitin beads was on the properties of the biopolymer/IL solution. Both the formation and the size and shape of the beads is strongly dependent on chain entanglement of the polymer, the polymer concentration, and the polymer shear viscosity.47-49Thus, the loading of chitin into IL was based on the viscosity matching that of microcrystalline cellulose (MCC) in 1-butyl-3methylimidazolium chloride ([C4mim][Cl]) solutions which were successful in bead formation.31 In the cellulose studies it was found that the ideal viscosity for bead formation was around 180 cP at 100 oC, which resulted from a 5.5 wt% solution of MCC in IL. Therefore, solutions of both PG chitin and IL-extracted chitin ranging from 1 through 3.5 wt% were prepared, and the viscosity measured to obtain chitin solutions with a viscosity around 180 cP at 100 oC. Viscosity measurements of chitin solutions revealed that a loading of PG chitin and IL-extracted chitin in IL of 1.65 wt% and 3 wt%, respectively, in IL yielded this desired viscosity (see ESI for complete experimental details and Figure S2). With the loading of the solutions selected, the formation of beads from the two different chitin sources was studied, using the 1.65 wt% PG chitin solution and the 3 wt% IL-extracted chitin solution. Bead formation was attempted using a liquid-liquid biphasic system of [C2mim][OAc] IL and PPG. The chitin was initially dissolved in IL, then the biopolymeric solution was dispensed as droplets into the PPG using a syringe pump. The liquids were mixed using an overhead stirrer as the IL phase was agitated in the hydrophobic PPG phase under a



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constant stirring rate. All initial preparation parameters, (i.e., flow rate, PPG molecular weight (MW), temperature(s) of the solutions) were first based on those used in reference [31], and were then optimized individually by holding all other parameters constant. The first processing parameters studied were those relating to the PPG coagulation bath (i.e., molecular weight of the PPG, temperature of the bath, and stirring method and speed) for the formation of uniform beads of desirable size and shape. PPG 425 was suitable for bead formation with cellulose in reference [31], however, here we assessed a range of MWs from 425-2000. The optimal PPG MW was found to be 2000 (with a viscosity of 22 cP at 100 oC), as its viscosity and shear rate led to the formation of relatively uniform microspheres. PPG baths with MW below 2000 yielded only large clusters of IL/chitin/PPG, but not microspheres. The next parameter of the PPG bath to be studied was the temperature of coagulation for bead formation. It was experimentally determined that when the temperature of the PPG bath started at 100 °C and slowly dropped to 55 °C over a period of 30 min, beads of uniform size and shape were formed. When the PPG bath fell below 50 °C, a collapse of beads was observed. When the initial bath temperature rose above 100 oC, the bead size decreased to

Porous Chitin Microbeads for More Sustainable Cosmetics - ACS

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