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Use of microcapsules as controlled release devices for coatings
Use of microcapsules as controlled release devices for coatings Markus Andersson Trojer, Lars Nordstierna, Jonatan Bergek, Hans Blanck, Krister Holmberg, Magnus Nyd´en PII: DOI: Reference:
S0001-8686(14)00198-5 doi: 10.1016/j.cis.2014.06.003 CIS 1450
To appear in:
Advances in Colloid and Interface Science
Received date: Revised date: Accepted date:
2 February 2014 6 June 2014 6 June 2014
Please cite this article as: Trojer Markus Andersson, Nordstierna Lars, Bergek Jonatan, Blanck Hans, Holmberg Krister, Nyd´en Magnus, Use of microcapsules as controlled release devices for coatings, Advances in Colloid and Interface Science (2014), doi: 10.1016/j.cis.2014.06.003
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ACCEPTED MANUSCRIPT Use of microcapsules as controlled release devices for coatings Markus Andersson Trojer,*a,b Lars Nordstierna,a Jonatan Bergek,a Hans Blanck, c Krister Holmberga and
Department of Chemical and Biological Engineering, Applied Surface Chemistry, Chalmers University of
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Magnus Nydéna,d
Technology, Göteborg, Sweden.
Department of Colloid Chemistry, Max Planck Institute of Colloids and Interfaces, Potsdam, Germany. E-mail:
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[email protected]
Department of Biological and Environmental Sciences, University of Gothenburg, Göteborg, Sweden
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Ian Wark Research Institute, University of South Australia, Mawson Lakes Campus, Adelaide, South Australia
5095, Australia *E-mail: [email protected]
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Telephone: +49 331 567 9212
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Fax: +49 331 567 9202
Abstract
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Biofouling of surfaces is a considerable problem in many industrial sectors and for the public community in general. The problem is usually approached by the use of functional coatings and most such antifouling coatings
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rely on the effect of biocides. However, a substantial drawback is the poor control over the release of the biocide as well as its degradation in the paint. Encapsulation of the biocides in microcapsules is a promising approach that may overcome some of the problems associated with the more traditional ways of incorporating the antifouling agent into the formulation. In this review, we summarize more than a decade of microcapsule
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research from our lab as well as from other groups working on this topic. Focus will be on two coacervationbased encapsulation techniques; the internal phase separation method and the double emulsion method, which together enable the encapsulation of a broad spectrum of biocides with different physicochemical properties. The release of the biocide from core-shell particles and from encapsulated biocides in coatings is treated in detail. The release behaviour is interpreted in terms of the physicochemical properties of the core-shell particle and the coating matrix. In addition, special attention is given to the experimental release methodology and the implementation of proper diffusion models to describe the release. At the end of the review examples of antifouling properties of some coatings against common biofoulers are presented.
Keywords Antifouling, Marine paint, Exterior wall paint, Sustained release, Dyes, Diffusion
Contents of paper
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ACCEPTED MANUSCRIPT Abstract ................................................................................................................................................................... 1 Keywords ................................................................................................................................................................ 1 Contents of paper .................................................................................................................................................... 1 Introduction ..................................................................................................................................................... 3
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Formulation ..................................................................................................................................................... 6
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Microencapsulation ............................................................................................................................. 10
2.1.1
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2.1
Internal phase separation ................................................................................................................. 13
The spreading conditions .......................................................................................................................... 13 Predicting morphology using the van Oss formalism ............................................................................... 14 Double emulsion ............................................................................................................................. 16
2.1.3
Choice of dispersant ........................................................................................................................ 17
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2.1.2
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Internal phase separation ........................................................................................................................... 18 Double emulsion ....................................................................................................................................... 18 Post-encapsulation modifications .................................................................................................... 19
2.1.5
Solvent choice and environmentally friendly approaches ............................................................... 19
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2.2
Microcapsules in paints ....................................................................................................................... 20
3.1
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Controlled release ......................................................................................................................................... 23 Release from core-shell particles ......................................................................................................... 26
3.1.1
Release methodology and diffusion models .................................................................................... 26
Diffusion models ....................................................................................................................................... 29 3.1.2
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2.1.4
Physicochemical properties of core-shell particles ......................................................................... 31
The core..................................................................................................................................................... 31 The shell .................................................................................................................................................... 33 The outer dispersant layer and surface modifications ............................................................................... 36 3.2
Release from coatings .......................................................................................................................... 38
3.2.1
Methodology and diffusion models ................................................................................................. 38
Diffusion models ....................................................................................................................................... 40 3.2.2
Solvent-borne vs. water-borne paints .............................................................................................. 41
3.2.3
Drying time and macroscopic porosity............................................................................................ 43
3.3 3.3.1
Controlled release and antifouling ....................................................................................................... 44 Use of non-hazardous and efficient biocides ................................................................................... 44
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ACCEPTED MANUSCRIPT Optimized biocide cocktails ...................................................................................................................... 45 3.3.2 4
Use of rate-determining release systems ......................................................................................... 46
Conclusions ................................................................................................................................................... 48
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Acknowledgements ............................................................................................................................................... 49
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References ............................................................................................................................................................. 49
Introduction
Coatings , such as paints, varnishes, and lacquers , are used for a large variety of functions and can be divided
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into decorative, protective and functional coatings[1-3]. Functional and smart coatings; including self-healing, self-cleaning, antimicrobial/antifouling[3, 4], lubricating, self-polishing (SPC paints)[5], flame-retardant and piezo-electric coatings, are becoming increasingly important in a variety of industrial sectors[1]. Antifouling and
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antimicrobial coatings are probably the most widespread examples of functional or smart coatings. Fouling of organisms on surfaces, sometimes called biofouling, is a huge problem in many industrial applications such as wastewater plants, implants and wound care products. The perhaps most prominent examples are biofouling on
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ship hulls[6] and exterior walls (see Fig. 1)[7-9] and this review will focus on these applications. Regarding house façades, the biofouling (algae, moulds, lichens, mosses, etc.) results in discolouration and potential
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damage of the substrate[9-13]. The marine biofouling on ship hulls (microalgae, seaweeds, mussels, barnacles, etc.)[14, 15] will result in an increased friction drag leading to increased fuel consumption and therefore
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pollution of the environment[4, 6, 15-18]. As an example, a ship without an antifouling coating may increase its
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fuel consumption by 40 %[5, 17, 19].
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Fig. 1. Examples of potential fouling organisms on house façades and aquatic surfaces. The A. niger image is reprinted from references[20, 21]. The exterior wall image is reprinted from[9]. The biofouling on exterior walls is not only an esthetic issue. Mold growth on houses results in a phenomenon called sick building syndrome. The release of mycotoxins and allergenic bioaerosols (e.g. spore fragments) is connected to the development of respiratory diseases and is consequently a public health problem[9, 13]. Regarding marine biofouling, a layer of slime will increase the fuel consumption by 1-2 %. Soft foulers such as seaweeds, hydroids or bryozans result in a further increase of the fuel consumption to 10%. Shell-forming organisms such as mussels and barnacles, called hard foulers, have such a large effect on the skin friction drag that the fuel consumption increases with 40 %[19].
The most common way to prevent fouling is to incorporate antifouling biocides (antifoulants) in the paint. Regarding marine paints, these biocides are usually termed booster biocides[4, 5, 15] indicating that they improve the performance of the usually copper-based paint. A major problem with the antifouling paints that are on the market today, in addition to the use of poisonous and potentially bioaccumulating biocides, is the poor control of the release of the antifoulants. Many of these coatings suffer from premature leakage or degradation of the biocides. Too fast release of the biocide will result in a loss of the coating’s antifouling function before the end of its intended life time. The desired lifetimes of façade coatings and of marine paints are at least 10 and 2 years, respectively, and achieving release profiles that cover such long times is a very demanding task[2, 22]. One approach to reach the required life times is to use high amounts of biocides in the coating but this may not be economically, environmentally, practically, or legally acceptable. Practically, the use of high concentrations of biocides in the formulation may result in macroscopic phase separation in the film.
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ACCEPTED MANUSCRIPT A very promising solution to many of the biocide-related problems mentioned above, which are discussed more in detail in Section 2, is microencapsulation of the biocides[22, 23]. Microencapsulation involves encapsulation of the biocide in miniature reservoirs called microcapsules or core-shell particles. It is worth mentioning that microcapsules have also found use in other types of functional coatings, including self-healing paints[1, 24].
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Figure 2 shows electron microscopy images and bright field microscopy photographies of various microcapsules.
Fig. 2. Different types of microcapsules based on poly(methyl methacrylate) (PMMA) and polystyrene (PS). a) SEM image of regular microcapsules of core-shell type based on PMMA with a dodecane core[25]. The core and the shell have been coloured in order to better display the internal morphology. b) SEM image of a PMMA-based microcapsule cut with a focused ion beam in order to display the multicore morphology[26]. The core and the shell have been coloured in order to better display the internal morphology c) Bright field microscopy image of a microsphere suspension dyed with Disperse Red 13[27]. d) Bright field microscopy image of microcapsules based on PS with an aqueous gelatine core which has been dyed with the natural dye betanin[28]. e) Bright field microscopy image of microcapsules based on a PMMA shell and a PS core[29]. The PS core has been dyed with β-carotene and the PMMA shell with Methylene blue.
In our group, which has been a part of the interdisciplinary research programme “Marine Paint”[17, 30] and the innovation company “Capeco” [10, 31], we have investigated antifouling and antimicrobial coatings[10, 11, 30, 32-39] for a decade and implemented microcapsule technology[10, 11, 22, 23, 25, 27, 31, 38-41]. It should be noted that a lot of antifouling research is focused on non-biocidal approaches[2, 18, 24, 42-59]. Some recent innovations regarding such biocide-free antifouling approaches comprises the use of anaerobic biofilms[54] and enzymatic peroxide formation[45, 48, 55-58]. However, approaches based on the release of biocides are still the most common and it is actually also a common strategy to combat fouling in biological systems[4, 60, 61]. In
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ACCEPTED MANUSCRIPT many natural systems the release of a bioactive substance is combined with a physicomechanical antifouling action[15, 16, 59]. In this review, we present our research during the past decade concerning encapsulation of biocides for use in antifouling coatings. The choice of encapsulation methods and of microcapsule material in relation to the
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physicochemical properties of the antifouling biocide is discussed. Furthermore, ways to control the release of
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the biocides by manipulating the physicochemical properties of the microcapsules are investigated and the resulting antifouling properties of the coatings are presented. A special focus will be given to the experimental
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methodology for controlled release measurements and to the subsequent implementation of diffusion models to describe the release. This is important because there are many examples of questionable setups for release experiments and of very pragmatic empirical release models that may not give reliable results that can be treated
Formulation
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in a scientific way.
In this section, the background to the problems associated with common antifouling formulations will be presented. These problems will be described in detail and the benefits of microencapsulation as a solution to the problems will be discussed. The following subsections will be devoted to the details of the encapsulation
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strategies implemented by us and others for use in antifouling coatings. The controlled release and the
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antifouling assessment of the release systems will be treated in Section 3. The typical practice when it comes to formulating biocides in coatings is to simply disperse them molecularly in the wet paint (see Scheme 1a)[4, 15, 16]. As mentioned in the Introduction, this approach is often not
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satisfactory because the antifouling function usually becomes quite short-lived. The biocidal action is over long before the desired life time of the coating, due to premature release or degradation of the active substance[16, 17]. Most biocides are small molecules and hence subject to fast self-diffusion in the coating matrix[11, 22, 36].
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Therefore, this simple approach requires the use of very high concentrations of the biocide in order to maintain the antifouling function for a long time (see Section 3 for a theoretical argumentation)[7]. The use of excessive amounts of biocide may pollute the environment, and it may also lead to technical problems such as macroscopic phase separation in the paint film if the biocide is incompatible with the binder-solvent system used[11]. This is valid for most biocides except for biocides which are present in the coating as pigments (inorganic crystals) such as copper (CuO or Cu2O). Pigment based biocides are released very slowly; however, the release profile is significantly influenced by the paint system. Yet, this is outside the scope of this review and the interested reader is referred to reviews which are solely focused on antifouling paints[15, 16].
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Scheme 1. An antifouling biocide can be introduced into a paint formulation by molecular dispersion (left) which is the most common approach, by immobilization to a large component in the formulation such as the binder (SPC paints) or a pigment (middle), or by encapsulation (right).
The issue of incompatibility can also prevent the use of different types of biocides in a paint formulation. Most fouling problems are multifaceted and involve a spectrum of different fouling organisms (see Fig. 1) with
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varying sensitivity to biocides[17, 62]. The use of very potent biocides with a broad efficacy towards fouling organisms will also affect non-target organisms[4, 16, 63, 64]. In the past, organometallic compounds were commonly used as antifoulants, with tributyl tin oxide, TBTO, being particularly popular. These biocides were
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very efficient in preventing fouling but, as it turned out, also very harmful to the local ecosystems. Such antifouling agents were banned by the governmental authorities starting in the late 1980’s and the beginning of
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the 2000’s and were phased out from the market during a relatively long transition period. As a response to the new regulations, considerable research efforts were made to implement and evaluate alternative antifoulants with
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better environmental profile. As a summary of the efforts made during this time period one may conclude that it seems very unlikely to find one single biocide that can replace TBTO and similar efficient compounds and at the same time be sufficiently benign to the environment. Instead, there is now a general consensus that an efficient and environmentally friendly antifouling coating requires a cocktail of complementary biocides with different
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efficacy profiles and with a rapid and efficient deactivation in the environment [17, 65, 66]. The biocides that are relevant for this review are presented in Chart 1 and are categorized according to their physicochemical properties. However, it is also common to categorize biocides according to their biocidal mechanism, efficacy profile, target organisms, etc. Triclosan and triclocarban, are antimicrobial compounds used as preservatives in personal care products. Also silver has antimicrobial effects and is presently used to make antimicrobial surfaces. Bronopol is used as a desinfectant in hospitals, but also in industry to avoid bacterial growth, e.g. in paper mills and cooling towers. Clotrimazole[67] is used to treat fungal infections in man and animals, while propiconazole is mainly used in agriculture for plant protection. IPBC is a fungicide used to protect painted surfaces from growth of fungi. The metals copper, zinc and silver, and the metalloid arsenic have broad-spectrum effects on micro-organisms, plants and animals, and have been used for wood preservation. Copper and zinc are also used as antifouling agents with effects on a variety of organisms. The hydrophobic isothiazolinone DCOIT is an antifouling compound effective against bacteria and algae and some animals. Irgarol and medetomidine are antifouling compounds with a narrow efficacy profile, targeting algal photosynthesis and the settling of barnacle larvae, respectively. DINCH is a new plasticizer often used together with antimicrobial agents.
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Chart 1. Biocides and dyes discussed in this review. The logarithm of the octanol-water partition coefficient (logKow values) and the water solubility are given for each substance, as obtained from Chemical databases[68-70].
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ACCEPTED MANUSCRIPT The problem of premature release has been approached by immobilization of the biocide on a more or less immobile object such as binders and pigments[30]. In more advanced paint systems, such as self-polishing marine coatings (SPC-paints)[4, 5], the biocide is attached to the binder by a covalent linkage (see Scheme 1) and the release rate is controlled by the rate of hydrolysis of the linkage. The release kinetics for such systems is
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of zeroth order or pseudo-zeroth order since the concentration of unhydrolyzed ester bonds in the paint and of
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water in the sea is constant. The resulting constant release rate is optimal with respect to environmental and economical aspects as discussed more in detail in Section 3. Note that the self-polishing hydrolysis itself is a
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mechanical antifouling mechanism. This strategy is, due to practical reasons (synthesis procedure, compatibility and solubility issues), only feasible for one or a few biocides per paint system and therefore only applicable to very potent and hence potentially environmentally hazardous biocides. The most well-known example is the selfpolishing coating with tributyl tin covalently attached, a SPC-TBT antifouling paint. This paint system, usually
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with added copper pigment, was very efficient and dominated the market before the ban as discussed above[66, 71-73]. To meet the new environmental regulations, a number of tin-free SPC paints have been developed and the organotin group is in these paints replaced by for example copper, silicon or zinc compounds [4, 5].
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Although these tin-free side groups possess some biocidal properties, the antifouling function is far from sufficiently efficient and booster biocides are therefore required. Note that these booster biocides are molecularly dispersed in the paint and hence subjected to fast release of non-zeroth order kinetics as mentioned earlier in this
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section and in the introduction. Moreover, the selection of suitable biocides is, at least in Europe, restricted due to the Biocidal Products Regulations which are controlled by the European Chemical Agency. Most used
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antifouling biocides are therefore conventional approved agrochemicals and efforts to introduce new antifouling 3.3.1).
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compounds are rare and limited to extraordinarily promising new biocides such as medetomidine (see Section In addition to covalent linkages, biocides may be immobilized by physical interactions[35-37, 74]. We have investigated immobilization; using coordination chemistry, of biocides containing a suitable ligand, with a special focus on the biocide medetomidine (see Chart 1), on common pigments (usually metal oxides
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nanoparticles) used in paints [30, 36]. However, this approach is also limited to a restricted number of biocides, which must contain a suitable ligand[30] and is outside the scope of this review. Microencapsulation can solve both the problem of premature release and the issue of compatibility when using several biocides in the formulation[17]. The release rate of the actives from the coating is determined by the slow release from the microcapsules to the surrounding coating matrix, and the encapsulated biocide is protected from degradation[11, 38, 39] (see Scheme 1 and Section 3.2). Moreover, compatibility is ensured and macroscopic phase separation prevented by the use of microcapsule shell materials that are compatible with the binder used in the formulation. Note that the only interface of importance between the paint and the formulated biocide is that between the microcapsule surface and the surrounding paint matrix. This allows for the use of carefully selected cocktails of biocides[65], containing biocides with high specificity[75-77] towards target organisms and low toxicity towards non-target organisms[75, 78-81], such as medetomidine (see Chart 1 and Section 3.3.2),
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Microencapsulation
The term “microcapsules” covers a broad range of colloidal particles of different sizes, morphologies and chemical compositions (see Scheme 2). Sub-micron particles are typically referred to as nanoparticles. Likewise, the encapsulation is typically termed nanoencapsulation for nanocapsules whereas microencapsulation usually
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regardless of the size and morphology of the corresponding colloidal particle.
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refers to microcapsules. However, from here on, we will use the terms microcapsules and microencapsulation
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Scheme 2. Microcapsule morphologies; a) core-shell, b) microsphere (monolithic/matrix), c) multicore-shell and d) core-multishell
There are numerous methods available to encapsulate biocides, many of which have recently been reviewed by
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us[23]. The choice of encapsulation method depends on the intended release profile and on the physicochemical properties of the biocide (polarity, size, charge, surface activity etc.). This interrelationship is theoretically
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discussed in detail in Section 3 and experimental examples are provided in Section 3.1.2. As illustrated in Chart 1, antimicrobial actives cover a very broad spectrum of chemicals with different physicochemical properties. In
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addition to biocides, we have also investigated the encapsulation of a variety of dyes (see Chart 1). The use of dyes with different physicochemical properties is an effective way to study the controlled release with UVvis[27] or fluorescence spectroscopy. In addition, the distribution and the partition of molecules with given physicochemical properties, into the different compartments of the microcapsule, can be investigated using
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bright field or fluorescence microscopy techniques (see Fig. 2c-e and Chart 1)[28, 29]. We have mainly investigated coacervation-based methods including the internal phase separation which is suitable for encapsulation of hydrophobic actives, and multiple emulsion routes which are suitable for encapsulation of hydrophilic actives. For both methods the coacervation is induced by solvent evaporation from an emulsion (see Scheme 3), and the barrier material is typically polymeric.
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Scheme 3. Encapsulation process in an aqueous continuous medium via the internal phase separation route and the double emulsion route. Reprinted from[22] .
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These methods are popular for a number of reasons. The multitude of polymers available with different physicochemical properties allows for the tailoring of the release according to need, and any core material can in principle be encapsulated[22]. Various types of core and shell materials with different physicochemical properties investigated by us are presented in Chart 2.The experimental procedure is straight-forward and
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feasible for industrial scale up[22, 23] which is discussed in more detail in Section 2.2. The resulting microcapsules contain no by-products such as monomers or initiators which can be present in microcapsules prepared using polymerisation techniques[23]. Even though the two coacervation processes are methodologically similar (see Scheme 3), the interfacial processes involved in the encapsulation are phenomenologically very different (see Sections 2.1.1 and 2.1.2). The theoretical details of these coacervation methods have recently been reviewed by us[22] and we will in this contribution focus on the results obtained in our group including some very recent innovations.
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Chart 2. Polymers used for shells, oils/polymers used for cores and polymers and surfactants used as dispersants. Here, the polyesters (PCL, PLLA and PLGA) and the cellulosics (CA and CTA) belong to the class biopolymers. PS and PMMA are considered commodity plastics whereas PU is an engineering/high performance plastic. The advantage (+) and the drawbacks (-), unrelated to the physicochemical properties, regarding the choice of a specific polymer class is also provided. Solubility parameters (δ [MPa1/2) are given for the shell polymers[82] and the logarithm of the octanol-water partition coefficient (logKow values) and the water solubility are given for each substance, as obtained from Chemical databases[68-70].
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ACCEPTED MANUSCRIPT 2.1.1 Internal phase separation The internal phase separation method was developed by the Vincent group in Bristol in the late 1990’s [83] and has since then been implemented and expanded by a number of research groups[12, 25, 26, 40, 84-87]. Both the
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core and the shell material are dissolved in a water insoluble and volatile solvent or solvent mixture (see Scheme
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3). As the solvent evaporates, the core and the shell materials phase separate inside the emulsion droplet (see Scheme 3). Since we are considering encapsulation in aqueous dispersion, which is the most appropriate system
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for most applications, the core material including the biocide needs to be more hydrophobic than the shell material. However, this simple requirement is not sufficient for ensuring proper microencapsulation as discussed in detail in the following paragraphs. It should be noted that this method has some advantages over the double emulsion route. The shell of the resulting microcapsule is homogenous[83, 88] with an easily controllable shell
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thickness. Moreover, the particles are relatively monodisperse[89] with almost 100% encapsulated active substance[88, 90]. Even though the discussion so far concerns microcapsules in aqueous dispersion, it should be noted that it is possible to produce microcapsules with aqueous cores using the internal phase separation method.
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Atkin and Vincent developed a method based on W/O emulsion systems, or rather acetone/O emulsion systems, for the fabrication of polymeric core-shell particles with aqueous cores[91]. However, as mentioned above, most applications require the microcapsules to be dispersed in aqueous medium. Yet, this can be accomplished by
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extracting the microcapsules from the oil phase to an aqueous phase using a proper choice of surfactants[91]. The choice of core oil can sometimes be restricted. Some oils that are good solvents for the biocide will also dissolve the shell polymer and subsequently hinder the shell solidification. Examples of such solvents are the
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aromatic hydrocarbons mesitylene and toluene, which cannot be used in combination with hydrophobic and semihydrophobic shell polymers (see Chart 2). A way to circumvent this is to use polymers of similar chemical nature as core material. The resulting microcapsule is consequently consisting of both a polymeric core and a polymeric shell. The practically non-existing entropy of mixing two polymers ensures segregation of the
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polymeric core and the polymeric shell and therefore microcapsule formation. As an example, we have investigated the use of polystyrene cores as replacement for aromatic oils (see Fig. 2e[29]. It is however important to realize that the use of a solid instead of a liquid core will have a profound effect on the diffusivity of the biocide as discussed in Section 3.1.2.
The spreading conditions So far, only the most fundamental aspects of the internal phase separation have been considered. A key element with respect to the final particle morphology is the interfacial tensions between the core, the shell and the continuous medium and in particular their spreading coefficients (see Equation 1). We will in the following discussion, for convenience, consider an aqueous dispersion of core-shell particles. This theory, based on spreading coefficients, was developed by Torza and Mazon[92] and later expanded by Loxley and Vincent[83] to predict the morphology of the core-shell particles obtained.
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ACCEPTED MANUSCRIPT Core-shell formation requires that the shell-forming polymer (index p) wets the core material (index o) in the aqueous phase (index w), i.e. spreads between the core and the water (see Equation 1). The spreading coefficient[93] Sp is defined by the interfacial tensions γij between phases i and j as;
Sp Gpc Gpa ow pw op
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ΔGcp and ΔGap are the work of cohesion and adhesion, respectively. If Sp>0, the polymer will spread between the
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water and the oil. The spreading coefficients for the oil and the water phases may be derived in a similar manner. It may be assumed that γow>γop which leads to three possible sets of spreading conditions[83, 92];
So<0;
Sw<0;
So<0;
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Sp>0,
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Sw<0;
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So<0;
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The core-shell morphology can only be obtained if the conditions in Equation 2 are satisfied. Equation 3 will generate so called acorn particles [83] and Equation 4 will result in separate oil and polymer droplets. The formation of acorn particles is not necessarily an unsuccessful encapsulation. It is also a popular route to produce so called Janus particles[94]. However, this is outside the scope of this review and will not be further treated. If
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no core material, often termed non-volatile-non-solvent, is added during the encapsulation, a homogenous
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polymer particle or microsphere[90, 95] will form and the spreading conditions have no meaning. The different types of morphologies and outcomes from the encapsulation process are depicted in Scheme 4. Since the morphology is highly dependent on the interfacial tensions between the three phases, it is clear that the choice of
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dispersant will be of paramount importance for the end result. This is treated in Section 2.1.3.
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Scheme 4. Morphology of a two-phase droplet system as a function of spreading coefficients; a) engulfed core (core-shell particle), b) partially engulfed core (acorn particle) and c) droplet separation.
Predicting morphology using the van Oss formalism The correct way to predict the microcapsule morphology is to experimentally measure all interfacial tensions between the core phase, the shell polymer and the aqueous dispersant solution using optical[25, 26] or force tensiometry[25, 26, 83]. However, this experimental procedure is rather time-consuming and arduous if the intention is merely to find a suitable core shell pair for encapsulation. The interfacial tension between the solid polymer and the liquid phases is determined by measuring the contact angle in air or in a liquid medium and
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ACCEPTED MANUSCRIPT subsequently applying Young’s equation. This requires that the surface free energy of the polymer is known. If not, the surface free energy of the polymer has to be determined by measuring the contact angle of a set of wellcharacterized liquids on the polymer surface and subsequently applying theoretical approaches, such as the Fowkes[96] or the van Oss-Chaudhury-Good formalism (see below)[97, 98]. For microcapsules that consist of a
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polymeric core as well as a polymeric shell, measuring the interfacial tension between two polymers is difficult
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and is usually performed on polymer melts.
A more straight-forward route to predict the microcapsule morphology is to theoretically calculate the interfacial
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tensions using theoretical approaches such as those mentioned earlier. The interfacial energy is a result of intermolecular forces and can therefore be separated into additive contributions (ion-ion, ion-dipole, Keesom, Debey, London dispersion)[97] in a similar manner as for the bulk[99] (see Section 2.1.5). The most simple representation is the separation of the interfacial energy into a dispersive part γd and a polar part γp as proposed
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by Fowkes[96]. However, it has been demonstrated that the Debey and Keesom contributions to the polar term γp are very small[97, 98]. Moreover, the polar contribution is usually overestimated and neglects the more complicated but important contribution from H-bonding[97, 98]. One popular approach is the van Oss-
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Chaudhury-Good formalism (Equations 5-8), which takes asymmetrical interactions; such as hydrogen-bonding, into account [26, 97, 98]. The apolar γLW contribution is expressed as the total Lifshitz or van der Waals force and the polar term γAB accounts only for Lewis acid-base interaction. The polar term is expressed by an acid
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component γ+ and a base component γ- (see Equation 6)[97].
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i iLW iAB
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iAB 2 i i
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The interfacial tension components can now be expressed by the geometric mean of the usually known surface tension components (Equations 7 and 8) and combined according to Equation 5 to give the total interfacial
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tension.
ijLW
LW i
LW j
ijAB 2 i j
i
2
j
7 8
Clearly, γijLW will always be positive. However, γijAB may attain negative values if i is a strong Lewis base and j is a strong Lewis acid or vice versa[26, 97]. The surface energy components; γLW, γ+ and γ- , can for most common polymers and solvents be found in the literature and the interfacial tensions are conveniently calculated from Equations 7 and 8. It is important to note that this simplistic approach does not take biocide or dispersant effects into account, which can be profound as discussed in Section 2.1.3. The calculated spreading coefficients based on the van Oss approach for the system water-PMMA-alkane (dodecane or hexadecane) are very similar to the experimentally measured counterparts which corroborates the validity of the approach (see Table 1). The more simple Fowkes approach does also predict the correct morphology; however, the values of the coefficients deviate significantly. Moreover, the effect of the dispersant can be significant and for very surface active ones, the deviation from the experimentally or modelled values is large. The acorn morphology is predicted for surfactant-stabilized systems (such as the SDS-based system in Table 1). However, the PVA-stabilized system displays core-shell morphology although acorn morphology is predicted. It should be noted that PVA-stabilized PMMA-based microcapsules occasionally 15
ACCEPTED MANUSCRIPT exhibit an uneven surface with protrusion and indentations which might be an indication of the partially fulfilled wetting conditions. Moreover, certain conditions may result in multi- rather than single core-shell morphology. Regarding the TC4-PDADMAC system, the multiple core droplets are stabilized by the surfactant TC4. The sorbitan monooleate system is a W/O emulsion and the fast evaporation of the solvent acetone freezes the initial
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multicore morphology (see Scheme 3) in the solidified PMMA matrix before the coalescence of the water
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droplets is completed. Note that the prediction does not distinguish between single core-shell and multicore-shell morphology, although attempts have been made to correlate this with the magnitude of Sp[100]. Overall, the
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predicted morphologies agree well with the observed ones in Table 1. Moreover, the theoretical van Oss approach will provide an insight into whether core-shell formation is at all conceivable for a given core-shell pair and has been used by us to predict the morphology of microcapsules with polystyrene cores[29] (see Fig. 2e).
So
Sw
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Table 1. Calculated spreading coefficients for the system water-PMMA-alkane (dodecane or hexadecane) containing (or not containing) different types of dispersants. The pure systems (not containing dispersants) in the table include experimentally measured values as well as theoretically predicted ones according to Equations 5-8. All dispersant-based systems are prepared from O/W emulsions where the dispersant is dissolved in the aqueous phase. The exceptions are TC4-PDADMAC and sorbitan monooleate. Regarding TC4-PDADMAC, the dispersant is residing in the PMMA phase (TC4) and is dissolved in water (PDADMAC) respectively. The sorbitan monooleate system is prepared from a W/O emulsion where the dispersant is dissolved in the alkane. For this system, the spreading coefficients have been estimated from interfacial tensions obtained from the literature.
Sp
Morphology
Predicted
-43.5
Van Oss[29]
-34.1
Fowkes[29]
-23.8
-60.5
26.5
Core-shell
-68.2
31.0
Core-shell
-78.8
16.6
Core-shell
Dispersants SDS[83]
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Experimental[25, 26]
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Pure system
Observed
-9.80
-3.60
-19.4
Acorn
Acorn
-17.4
-25.8
-11.8
Acorn
Core-shell
-31.1
-39.7
1.90
Core-shell
Core-shell
600-4600[25]
-45.4
-29.3
28.4
Core-shell
Core-shell
TC4-PDADMAC[26]
-59.7
-43.7
50.8
Core-shell
Multicore-shell
Sorbitan monooleate[91]
-6.96
-37.8
3.84
Core-shell
Multicore-shell
PVA[83]
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PMAA[25, 83]
2.1.2 Double emulsion The double emulsion route may, in contrast to the internal phase separation route, produce particles with aqueous cores directly in an aqueous suspension (see Scheme 3). The most common type of double emulsion for encapsulation applications is W1/O/W2[22, 101-104], a system that has been extensively reviewed by Garti and coworkers[102, 104-108]. The most explored procedure to produce double emulsions is a two-step homogenization method[109] which shares many similarities with the internal phase separation method (see Scheme 3)[28, 102]. However, the corresponding pre-emulsion is in this case a W/O emulsion formed under high shear[28, 101, 102]. The aqueous phase contains the active substance and the oil phase contains the shell forming polymer and a volatile solvent. The prepared W1/O pre-emulsion is subsequently dispersed in the 16
ACCEPTED MANUSCRIPT continuous W2 phase under moderate shearing and slow evaporation of the solvent in order to solidify the oil phase that forms the shell. This procedure usually results in quite large, polydisperse and multicore-shell particles (see Scheme 2c and Fig. 2d)[28, 101]. However if multicore-shell particles suffice, it is preferable to attain as small inner droplets as possible in order to slow down the coalescence kinetics[101]. The water droplets
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of microemulsions are therefore suitable as inner phases. The microcapsule-forming dispersed phase may also
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consist of a dispersed mesophase, such as cubosomes (dispersed cubic liquid crystals), swelled liposomes (dispersed lamellar phases), etc.[101, 102]. Another way to reduce the kinetics of the segregation of the inner
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droplets is to add thickeners in order to increase the viscosity of the inner phase (see Chart 2 and Fig. 2d)[28]. Modern techniques such as membrane and in particular microfluidic devices[110] enable the formation of monodispersed droplets with controllable inner morphology[111]. Yet, the particles are still relatively large and the shell is occasionally inhomogeneous. For a more detailed investigation of the properties, technical aspects
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and recent developments of microcapsules prepared using microfluidics, the interested reader is referred to a recent review by Datta [112]. However, in contrast to the two-step homogenization method, the microfluidics
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method is as of yet unsuitable for mass production of microcapsules.
2.1.3 Choice of dispersant
The main function of the dispersant is to facilitate the emulsification and to provide colloidal stability of the
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core-shell particle suspension. Polymeric stabilizers are the primary choice for core-shell particle production
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using the internal phase separation or the double emulsion route as will be explained below. The polymer dispersant can provide colloidal stability through steric, electrostatic or depletion stabilization[113]. In many
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cases (PVA or PMAA stabilization), the latter mechanism is predominant and a large excess of dispersants is required to prevent irreversible coalescence[40, 114]. The adsorbed dispersant layer is usually very thin in comparison with the core-shell particle (see Fig. 3) but may nonetheless be viewed as an additional outer shell. This implies that the choice of dispersant type should have a
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large impact on the release rate which is indeed the case[113] as discussed in Section 3.1.2.
Fig. 3. Adsorption isotherms given by the thickness of the adsorbed layer δ as a function of the concentration c of the dispersants PVA (▲), PMAA (▼) and 600-4600 (■) on PMMA surfaces as measured using QCM-D[27].
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ACCEPTED MANUSCRIPT Internal phase separation Normally, the use of surfactants as dispersants during microencapsulation using the internal phase separation method prevents the formation of core-shell particles[22]. From Equations 1-4 it is clear that the interfacial tension γow must remain high for polymer spreading and subsequent core-shell particle formation to occur. Small
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surfactants will significantly reduce γow and this is the reason why the use of surfactants as dispersants prevents
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core-shell particle formation. As an example, Loxley and Vincent studied the effect of using the water-soluble surfactants cetyltrimethylammonium bromide (CTAB) and sodium dodecyl sulfate (SDS) as dispersants which
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resulted in partially engulfed oil cores termed acorn particles[83]. However, contrary to the current knowledge, we have observed that very hydrophobic surfactants can be used for the purpose.
The oil-soluble surfactant TC4
(see Chart 2) enabled core-shell formation when used in combination with the polycation rather than single core-shell (see Fig. 2b)[25, 26].
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poly(diallyldimethylammonium chloride) (PDADMAC)[26]. Yet, the observed morphology was multicore-shell As previously mentioned, the appropriate choice of dispersants are polymeric stabilizers[40, 83]. The ideal surface activity of the polymeric dispersant is a compromise between low surface pressure and stabilization of
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the colloidal surface[40]. We have investigated novel types of ionic dispersant which provided PMMA-based core-shell particles with strong electrostatic stabilization, avoiding the use of excess dispersant in the bulk. These included poly(methacrylic acid) (PMAA) treated with base after adsorption[114], the hydrophobic anionic
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surfactant TC4 in combination with PDADMAC[25, 26] and charged amphiphilic block copolymers of PMMA-
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b-PMANa type (see Chart 2)[40]. In particular the latter dispersant type provided the core-shell particle suspensions with long-term stability by a combination of electrostatic and steric stabilization[25, 40]. This was achieved by using a block copolymer with a hydrophobic block of the same monomer composition as the shell-
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forming polymer[40]. We have shown that the hydrophobic block is dissolved in the polymer phase during the emulsification and becomes amalgamated with the shell as the volatile solvent evaporates[40].
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Double emulsion
In contrast to the internal phase separation route, the core-shell formation from double emulsions is not relying on the spreading of a polymer coacervate per se. This broadens the choice of combinations of shell forming polymers, dispersants and volatile solvents to be used. Nevertheless, double emulsions have a limited shelf life , even more so than regular emulsions[102]. Therefore, successful core-shell formation requires proper stabilization of the W1/O and O/W2 interfaces. Typically, two sets of dispersants are needed; an oil soluble one to stabilize the W1/O interface and a water soluble one to stabilize the O/W 2 interface. Once again, polymeric stabilizers are more suitable than small surfactants[28, 102] but for different reasons compared with the internal phase separation case. Small surfactants have a tendency to migrate from one interface to the other while amphiphilic polymers (graft or block copolymers) tend to adsorb more “irreversibly” at the interface. Therefore, ethylene oxide-propylene oxide block copolymers of Pluronic type are common choices. A very promising type of block copolymers for these purposes are diblock polymers with a large hydrophobic block of the same type as the shell forming polymer and a small hydrophilic polyethylene oxide block (see Chart 2). For example, the inner gelatinous water droplets of the PS based microcapsules in Fig. 2d are stabilised by the P(S-b-EO) blockcopolymer shown in Chart 2. The hydrophobic block will amalgamate with the shell during the evaporation stage
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ACCEPTED MANUSCRIPT as described above[40] and the poly(ethylene oxide) block allows for solvation in organic solvents in contrast to hydrophilic polyelectrolyte blocks[28]. In addition, the polymeric film at the interface prevents coalescence of droplets and migration of water from W 1 to W2.
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2.1.4 Post-encapsulation modifications
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The core-shell particles can, after the encapsulation process, be modified in order to tailor their release
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behaviour. There are many techniques available for post-encapsulation modifications, and in this section we will consider ways to affect the microporosity of shell. We will also discuss surface modifications that are able to introduce additional barriers (shells) on the microcapsule.
The microporosity (or free volume) of the shell can be reduced by annealing or cross-linking[22]. The Vincent
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group investigated ways to cross-link PS and poly(2-vinylpyridine) based microcapsules by simply mixing monomers and initiators in the oil phase[115]. The shell of the microcapsule was cross-linked after the encapsulation by exposing the particle suspension to UV-radiation or heat. The alternate method “annealing” is
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perhaps a more convenient route to reduce the microporosity. Annealing implies relaxing the polymer chains from a kinetically arrested state to a more thermodynamically stable state. It has been shown by the Vincent group, and our group [38, 88], that the encapsulation procedure can result in microcapsules with an expanded shell structure. However, this can be counteracted by heating the microcapsule suspension above the glass
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transition temperature of the shell-forming polymer followed by slow cooling.
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Regarding the surface assembly of additional barriers (shells) on the microcapsule surface, we have investigated the adsorption of polyelectrolyte multilayers (PEMs)[27, 39, 116], as well as of lipid bilayers[116]. The PEMs
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are assembled using the well-known Layer-by-Layer (LbL) technique that for colloidal systems, was developed by the Möhwald group in the late 1990’s[117-119]. The lipid bilayer is assembled by spreading charged liposomes on a microcapsule surface of opposite charge[120, 121]. Both surface modifications require a stable and highly charged surface. The surface charges can be introduced by using an intrinsically charged colloid[25,
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40, 114, 122] or by chemical means[122]. We have investigated the use of an amphiphilic ionic dispersant in order to introduce a high and stable surface charge density as described in Section 2.1.3. The PEM and the lipid bilayer represent two very different barriers in terms of physicochemical properties. The PEM is hydrophilic, highly charged and relatively swollen with solvent. On the other hand, the lipid bilayer is very hydrophobic and; for saturated symmetrical hydrocarbon chains longer than ca 14-16 carbons, also highly dense and crystalline[22, 23].
2.1.5 Solvent choice and environmentally friendly approaches The properties of the volatile solvent used in solvent evaporation-based methods and in particular the internal phase separation method must fulfil certain requirements, which has been reviewed by us in detail elsewhere[12, 22]. Perhaps, the most obvious requirement is that the solvent must dissolve the shell forming polymer and the core. The prediction of solubility of a polymer in a solvent can be done using the Hildebrand’s solubility parameter concept. The Hildebrand solubility parameter δ is based on the cohesive energy density E/V which employs the semi-empirical theorem, like dissolves like (Equation 9-11)[123, 124]. The solubility parameter is
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ACCEPTED MANUSCRIPT related to the interaction parameter χ according to Equation 10 where V0 is the molar volume (assumed to be the same for both components).
EV
9
ij V0 RT ij
10
ijH i j
11
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2
Clearly, substances with similar cohesive energies, i.e. similar intermolecular interactions strengths, will be
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mutually soluble as χ→0. This is valid for non-polar systems but does not account for asymmetric interactions in strongly polar or hydrogen-bonding systems. Here, the extended Hildebrand approach may be used (Equation 12). The solubility parameter is separated into additive components of intermolecular forces (Equation 13) in a similar fashion as for the surface tension (see Section 2.1.1); e.g. apolar, Lewis acid and Lewis base components
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(with indexes “LW”, “+” and “-”)[125, 126]. Moreover, the extended Hildebrand approach allows for negative values of χ. Note the similarity of Equations 12 and 13 with the van Oss approach.
2 2
ijEH iLW jLW
LW 2 i
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i2
2
i
j
i
j
i i
12 13
The most common solvents used as volatile solvents are chlorinated compounds such as dichloromethane or
D
chloroform. These chlorinated compounds can dissolve most hydrophobic and semihydrophobic polymers and
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possess some Lewis acid properties[97]. For more hydrophilic polymers such as cellulose acetate (see Chart 2), acetone can be added to the oil phase to increase its polarity and add some additional Lewis base properties[28].
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Acetone is also the solvent of choice regarding encapsulation of aqueous cores using the internal phase separation method[22, 91]. However, for O/W emulsions it is important to note that acetone will be distributed between the water and the oil phase and therefore evaporate faster than the more hydrophobic solvent. This will affect the point of internal separation and shell solidification.
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For the implementation of solvent evaporation based-methods on an industrial scale the environmental and toxicity aspects of the solvent must also be taken into account. The use of chlorinated solvents in the industry is highly restricted today[12]. Replacement of the chlorinated solvents with more environmentally friendly ones is not easy since they, in addition to the above mentioned property requirements, must be cheap and/or recyclable for industrial large-scale production[127]. However, we have found that ethyl acetate fulfils these requirements[31]. We and others have subsequently used ethyl acetate to prepare microcapsules using the internal phase separation method[11, 12, 31], as well as the double emulsion route[128].
2.2
Microcapsules in paints
The incorporation of microcapsules into a paint system is simple and requires a minor workload and equipment. The microcapsules are formulated into wet paint as an additive, which mean that no specific preceding formulation of the paint is needed. Hence, the addition of microcapsules can be performed at any time prior to the application of the coating and, moreover, commercially available paints can be used. Our standard procedure of microcapsule addition involves two steps.
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ACCEPTED MANUSCRIPT First, water should be removed from the aqueous microcapsule suspension. Section 2.1 describes the formulation of microcapsules and the final suspension contains a microcapsule-to-water ratio of approximately 1:20. Before the addition of the microcapsules to the wet paint, the water content of the microcapsule suspension needs to be significantly decreased, using for instance centrifugation or ultrafiltration. A large amount of the continuous
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solvent phase may cause undesirable effects in the paint and must in some cases be exchanged with a solvent
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which is more compatible with the paint solvent. Completely dry microcapsules should be avoided as the microcapsule agglomeration in the dry state can be difficult to break by mere stirring of a highly viscous paint
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system. However, the agglomeration is strongly dependent on the type of dispersant used in the microcapsule suspension. The use of conventional dispersants such as PVA and PMAA results in irreversible aggregation if the excess dispersant in the water phase is removed. However, microcapsule suspensions prepared using the highly charged dispersants described in Section 2.1.3 are fully dispersible also from a completely dried state
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when all excess dispersant has been removed. For the microcapsule suspension stabilized by PVA or PMAA, a microcapsule-to-water ratio of approximately 1:1 is convenient since it produces a manageable fluid material, which will allow for completely homogenous dispersion of the microcapsules in the wet paint system. Water
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removal from the suspension is, from our experience, a critical stage in the scale-up for industrial production. The microcapsule-to-water ratio should, at industrial quantities in a relatively fast step, increase from approximately 1:20 to 1:1 but where the latter ratio is important not to stretch further to avoid agglomeration.
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Moreover, recycling of the removed aqueous containing dispersant is most probably preferred.
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In the second step, the concentrated microcapsules suspension is added to the wet paint. If a waterborne paint is used, the formulation only requires a simple, but thorough, mechanical stirring of the paint system after addition
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of the manageable fluid material. In the case of solvent-borne paint, the situation is different. The water content of the concentrated suspension might need to be exchanged to a solvent that is more compatible with the solvent of the paint. Moreover, attention should be made with regard to the paint solvent and its influence on the microcapsule material. The various microcapsules presented in this article are all suitable for at least water-borne
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paints and varnishes. Fig. 4 displays electron micrographs of the area inside dried paint matrices revealed by scratching the coating with a thin needle [11]. It is clear that the microcapsules are stable and properly dispersed in both solvent-borne (xylene-based paint in Fig. 4a and b) and latex paint systems (Fig. 4c and d). Further information regarding the application of coatings for laboratory studies is presented in Section 3.2.1.
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Fig. 4. SEM images of solvent-borne (a and b) and water-borne (c and d) paint matrixes containing well-dispersed microcapsules (c and d)[11]. The microcapsules are homogeneously embedded in the coating and their geometry is unaffected by the mechanical stirring during the addition of manageable fluid material of microcapsules to the paint, the drying of the applied paint, and by the presence of the surrounding binder and pigment material. Reprinted from [11]. The microcapsules have been coloured brown and the paint matrix yellow for clarification.
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ACCEPTED MANUSCRIPT 3
Controlled release
In this section, the theoretical fundamentals for controlled release will be described and the most important parameters, as visualized in Scheme 5, will be introduced. Focus will be on sustained release which is the dominant release mechanism in coating applications. Although we have previously briefly considered triggered
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release[27, 30, 129, 130], the interested reader is referred to a set of excellent reviews regarding the topics
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targeted or triggered release[131-133].Moreover, the economic and environmental benefits of using ratedetermining release systems in coatings for antifouling purposes will be briefly discussed. The following
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subsections will separately review controlled release from core-shell particles and coatings, respectively. In each subsection, the experimental methodology of the release measurements will be illustrated in detail. The release will be further examined in terms of the physicochemical properties of the core-shell particles and the coating matrix. Finally, the antifouling properties of some functional coatings towards different biofoulers will be
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presented.
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Scheme 5. The figure displays the important parameters for a microencapsulated substance which determine the release rate. These include dimensional and density parameters; the concentration of the active i in the core cCi, of microcapsules in the coating cC which gives the total concentration of active cCi ·cC=ci, the thickness of the coating δP, the radius of the core rc and of the microcapsule rm which gives the thickness shell δS. The kinetic parameters include; the diffusion coefficient in the core DC, in the shell DS and the diffusion coefficient in the paint DP. The concentration gradient of the active is also determined by the core/shell KC/S, shell/coating KS/P and the coating/water KP/W distribution coefficients.
The sustained release of an active from a microcapsule in a coating is controlled by the permeation through the microcapsule barrier (the shell) and the coating matrix. The permeation of the active in the microcapsule barrier and the coating matrix depends on both kinetic and thermodynamic parameters through its diffusion coefficient and its solubility in the barriers. The solubility of the active in, and its interaction with, the microcapsule core and shell materials and the coating matrix will determine its partition between these phases[22, 134-136]. This is described by the distribution coefficients, KiA/B, of the active i between phases A and B (Equation 14, where ciA and ciB are the equilibrium concentrations of i in phase A and phase B, respectively) and we will consider this thermodynamic constant first.
23
ACCEPTED MANUSCRIPT i K A/B
cAi 1 i cBi K B/A
14
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The distribution coefficient can be expressed by the saturation concentration in phase A and the Flory-Huggins interaction parameter between i and “B” (Equation 15) following some minor derivation and implementation of
15
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i ln K A/B ln cAs ,i Bi
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the Flory-Huggins solution theory[123, 124] as described more in detail elsewhere[22].
A higher saturation concentration of i in A and unfavourable B-i interactions (higher χBi) will favour distribution of i in phase A and vice versa, which is a logical and important realization. The Flory-Huggins interaction parameter can be obtained using the Hildebrand interaction parameter as described in Section 2.1.5 (Equations 9-
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13).
The diffusion coefficient on the other hand is a kinetic parameter and is related to the size r of the active (D0=10/r m2/s, r is expressed in Å). For controlled release applications, it is important to make a distinction between
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8
the self-diffusion of the active, D0 and the effective or apparent diffusion D. The notion of an effective diffusion constant is strongly associated with the concept of hindered diffusion[135-138]. As the name suggests, hindered
D
diffusion implies that the diffusion constant will be effectively reduced by a number of parameters as will be
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explained below (Equation 16). Impermeable objects, such as spherulites in semicrystalline polymers or inorganic pigment particles in a coating matrix, will obstruct the diffusion. The obstruction is described by the porosity ε (the cross-sectional area available for diffusion) and the tortuosity τ (the diffusive trajectory relative
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the end-to-end distance in the diffusive direction)[135, 136]. We have chosen to designate this porosity as a macroporosity since the relevant pores sizes of these macropores are significantly larger than the molecular
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dimensions of the actives[38].
V D D0 exp c K B V f
16
On a molecular scale, the active is diffusing in the void, or free volume, of the surrounding matrix, which for most applications consists of polymer chains. The free volume of the matrix Vf, (Equation 17) must be larger than a critical volume for diffusion Vc which is determined by the size of the active. The parameter γ is an overlap factor (1 for most polymers), V is the specific volume of the matrix, V0 is the impermeable volume occupied by the polymer and Vw is the van der Waals volume [38, 139]. This microporosity can also be expressed in terms of a thermal energy of the matrix and a diffusional energy barrier instead of volume-based parameters. The outcome of the expression will still be the same[139]. V f V V0
17
V0 1.3Vw
Interactions between the shell material and the active may not only affect the partition coefficient. Strong and specific interactions; such as coordinate interactions[22, 30] or hydrogen bonds[22, 140], characterized by an equilibrium constant for binding, KeqB, will immobilize a fraction of the active on the shell material[22, 136, 137,
24
ACCEPTED MANUSCRIPT 140]. A shell material with a concentration of cB binding sites will result in an immobilized fraction of actives and only the “free” fraction KB will be available for diffusion according to Equations 16 and 18[136, 137].
K B (1 K Beq cB ) 1
18
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The relationship between these parameters and the release rate is most conveniently explained by assuming
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release under steady state conditions. The release rate ṁ from the microcapsules, which under steady state conditions is equal to the release rate from the coating, is a function of the concentration ci of the active i in the
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core and the total resistance rTOT over the barriers (Equation 19, see Scheme 5). The total resistance (Equation 20) is the sum of the barrier resistances, which depends on the barrier thickness δ, the cross sectional area of the barrier in the direction of the diffusion A (which is very different for the microcapsules and the coating), the effective diffusion coefficient D and the distribution coefficient K between the phases. The permeation P is
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subsequently defined as the product of the diffusion coefficient and the distribution coefficient. It is clear that given the high specific surface area of the microcapsules as compared with the surface area of the coating, the controlled release from a paint system requires rigorous control of the permeation in the core and the shell
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phases of the microcapsule. For non-steady state situations, which are the usual cases, the release must be described by diffusion models. These will be discussed in the following subsections
D
m
ci rTOT
rTOT rx Dxi , K xi , x , Ax x
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x
19
x Ax Dxi K xi
x
x Ax Pxi
20
The importance of controlled release in terms of environmental and economic impact can be explained by a
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simple illustrative example in Fig. 5. Let us consider an antifouling coating containing m∞ amount of biocide. The biocide needs to reach a critical concentration cc, often termed minimum inhibition concentration (MIC) or effective concentration (EC), in a defined volume adjacent to the surface in order to prevent fouling of an arbitrary biofouler. The release of the free biocide from the coating is assumed to obey first order kinetics with
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the rate constant kin which is a function of mtot and the permeability in the coating. The removal of the biocide due to diffusion, degradation, absorption by the biofouler, etc is also assumed to be a first order process with the rate constant kout. It is clear that the majority of the biocide will be wasted during the initial stages of the release. The most efficient release would follow zero order kinetics such that the c(t)=cc over the entire release interval. However, true zero order release kinetic is extremely difficult to achieve[23] and is seldom realized with microencapsulated biocides. Yet, an efficient reduction of the rate constant will still significantly extend the period of efficiency. For example, a reduction of the rate constant kin to 0.1kin for a given mtot and cc=0.01m∞, will prolong this period five times.
25
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Release from core-shell particles
D
3.1
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Fig. 5. Relationship between the surface flux from the coating ṁ(t)= ṁ(t)in and the surface concentration c(t). The solid curves display the surface concentrations c for first order release (black) and for zero-order release (gray). The dashed curves display the corresponding surface flux from the coating. The blue square grid represents the integral of the surface flux and hence the total amount mtot of the biocide. The yellow line grid between the dashed curves for first and zero order surface flux represents the amount of wasted biocide.
This section will first describe the basics for a proper experimental release methodology regarding the release of
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actives (e.g. biocides) from microcapsules to an external aqueous medium. Suitable diffusion models are also discussed as well as their correctness when it comes to model the release in a physically sound way. Thereafter,
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the release results of some illustrative examples will be discussed in terms of the physicochemical properties of the different compartments in the microcapsule: the core, the shell and the outer dispersant layer. In addition, the surface assembly of PEMs on the microcapsule surface (see Section 2.1.4) is presented as a very promising tool for controlling the release of hydrophobic actives. Even though the intended release application usually concerns
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coatings, analysis of the release from a simple microcapsule suspension allows for the evaluation of structurefunction relationship with respect to the core and shell material and to the biocide. Yet, such an assessment requires complete knowledge of the size distribution of the microcapsules as explained below.
3.1.1 Release methodology and diffusion models Controlled release can be evaluated using several different approaches. Regardless of the chosen method, the methodology contains at least three main steps: setup, analysis, and evaluation. The procedures are schematically shown in Scheme 6a. The setup encompasses all laboratory equipment and material that allow for time-dependent release from the microcapsule to some arbitrary external release medium. The analysis involves the procedure for data sampling and determination of the concentration of the active at a given time using a proper analytical instrument. The evaluation is the post-processing work where the release data is put into perspective, e.g. by the use and implementation of a release model. The setup requires knowledge and proper control of the saturation concentration of the active in the release medium as well as its distribution between the microcapsule phase and the release medium. After the microcapsules are formed in the aqueous suspension, as described in Section 2.1, a fraction of the biocide is 26
ACCEPTED MANUSCRIPT released until equilibrium between the concentration of biocide in the microcapsule and in the aqueous phase is reached. This distribution is determined by the distribution coefficient in Equation 14. The magnitude of the distribution coefficients can be estimated by the octanol/water partition coefficients in Chart 1. For hydrophilic actives, prepared using the double emulsion route, the distribution coefficient is close to 1. However, for
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hydrophobic actives, the equilibrium is strongly shifted towards the microcapsules and the water phase is quickly
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saturated with the active. Note that the saturation concentration for hydrophobic actives in water is very low (see Chart 1). Failure to realize the importance of the saturation concentration and the distribution coefficients has led
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to the use of some misconceptions in the literature. The term “yield” is occasionally used to describe the observation that the release of hydrophobic actives reaches a plateau region below 100% released amount of active. However, this “yield” is merely an artefact of the experimental setup where the active reaches the equilibrium concentration in the aqueous medium. If additional water is added, i.e. the microcapsule suspension
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is diluted, further release of the active will occur until equilibrium is once again attained according to the value of the distribution coefficient.
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Scheme 6. a) Experimental methodology for the measurements of the release of an encapsulated active substance from microcapsule in an aqueous microcapsule suspension. The microcapsule suspension is poured into an aqueous solution (occasionally containing solubilizers) and the release of biocide out from the capsules to the surrounding medium will start immediately. Samples are collected over time using a syringe and subsequently pressed through a suitable filter in order to remove the microcapsules, other particles and dust. The concentration is quantitatively analysed by e.g. spectrophotometry. b) The use of semi-permeable dialysis membranes is a common setup for release measurements. The microcapsule suspension is put in a tube where the dialysis membrane has a pore size significantly smaller than the size of the microcapsules. Samples are taken from the surrounding medium, often an aqueous solution, and the concentration of released active substance is analysed. However, drawbacks have been noticed, for instance that the microcapsules aggregate over time due to improper mixing, which will change the release behaviour.
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ACCEPTED MANUSCRIPT In order to achieve complete, or almost complete, release of the active from the microcapsule, a significant amount of water is needed, especially for hydrophobic actives. In practice, our release methodology must always fulfil two criteria; 1), the total amount of the active in the system should be less than 10 % of the corresponding saturation concentration in the release medium. Under this condition, the flux of the active into the microcapsule
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can be neglected. 2) At the equilibrium concentration of the active substance, which is determined by the
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distribution coefficient and the extent of dilution, at least 90 % of the active should be released in the release medium. This allows for obtaining a well-resolved set of experimental data over the entire release interval.
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For hydrophobic biocides, the required dilution is very large and will result in release concentrations which are too low to be accurately quantified using analytical techniques such as spectrophotometric and chromatographic methods. To be able to accomplish a measureable release profile, one must therefore decrease the value of the distribution coefficient. A direct route to alter this coefficient is by the addition of a co-solvent to the aqueous
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release medium. However, co-solvents may generate swelling of the microcapsule shell polymer[141]. A noninvasive, from the microcapsule point of view, alternative is the addition of solubilizers, such as the surfactant sodium dodecyl sulfate (SDS) or cetyltrimethylammonium bromide (CTAB) (see Chart 2), to the release
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medium. Above the critical micelle concentration (CMC), the active is solubilized inside the surfactant micelles, which significantly enhances its solubility[142-145]. Moreover, surfactants are relatively benign with respect to the integrity of the microcapsule in contrast to more hydrophobic co-solvents. In the study of charged
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microcapsules, e.g. microcapsules modified with polyelectrolyte multilayers (see Section 2.1.4), nonionic
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surfactants, such as poly(ethylene glycol)monododeyl ether (Brij® L23), are the proper choice (see Chart 2)[20]. The analysis in terms of data sampling procedure requires a short comment. Since the release from the core-shell
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particles is evaluated in their suspension, the active typically needs to be separated from the particles. A common setup found in the scientific literature is the use of semi-permeable dialysis tubes with pore sizes slightly smaller than the mean size of the microcapsules (see Scheme 6b). However, the dialysis method suffers from some severe drawbacks. Analyses in our laboratory have shown that the equilibrium of the active between
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inner and outer release medium, with respect to dialysis membrane, is not necessarily established during the time interval between two data samplings. Moreover, it is very difficult to obtain proper mixing inside the dialysis tubes. As a consequence, flocculation and aggregation of the microcapsules readily occurs, which strongly affects the release behaviour (See Scheme 6b). In view of the negative aspects using the dialysis tube setup, we have chosen a different approach, which is schematically drawn in Scheme 6a and which in principle can be applied to any system. The release is instigated by diluting the microcapsule suspension according to the two criteria discussed above. For the data sampling, a small sample of the diluted suspension is filtered using a syringe filter to separate the particles from the released active. However, the choice of filter material is of utmost importance since many filters will absorb a significant amount of active. Especially tricky actives are semi-hydrophobic molecules with some surface activity. Such an example is the fungicide OIT[38] (see Chart 1). Only one filter type out of five was sufficiently non-adsorbing for this active substance (see Table 2). If this issue is over-looked, the measured release will be much slower than the true one.
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Polyvinylidene fluoride
98.2 ± 1.1
Polyethersulfone
91.7 ± 1.0
Polyvinylidene fluoride (prefilter: Polyethersulfone)
90.8 ± 0.8
Amphoteric Nylon 6,6
89.2 ± 0.8
Surfactant-free cellulose acetate
50.1 ± 1.3
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Permeability (%)
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Membrane material
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Table 2. The membrane material in different filters evaluated by the research group with respect to the biocide OIT (aq) to find a suitable experimental setup.
Diffusion models
The quantitative analysis of the time-dependent biocide release presented above is tremendously useful.
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Experimental data of high accuracy allows for model evaluation of physicochemical properties that are decisive for the release. Mathematical regression analysis and fitting of experimental data can generate several parameters, out of which the most important is the diffusion coefficient of the active substance within the
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microparticle. Most often it is the apparent, or effective, diffusion coefficient that is calculated (see Section 3) and this parameter permits relevant comparison between different particles, as the diffusion coefficient itself is independent of both size and time.
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There are several extensive reviews concerning modelling of the release of actives from polymeric devices. Most
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of these reviews focus on pharmaceutical applications[146-149] where controlled release is a key concept. Here we often find simplified and empirical approaches to model the release. Many of these models, which are principally based on the work by Weibull, Higuchi and Peppas[150-153], deal with dispersed actives in
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polymeric release reservoirs. The benefit of such models is their simplicity and the possibility to compare various drug devices or release compartments. However, only few physical parameters are included and the models are often valid only for the initial part of the release. Still, in pharmaceutical sciences such models offer a great deal of information considering rather macroscopic release devices where the initial release and initial
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increase of concentration are the important parameters. For sustained release, on the other hand, the long-term release behaviour is crucial. One main goal with sustained release is to prolong the surface flux of the active above a certain critical level[23, 39, 134]. Hence, a proper description also for the extended final part of the release is crucial for improved understanding and prediction. All types of materials used in the release systems described in this review are stationary material. In other words, the materials constituting the core and the polymeric shell are stable and not undergoing erosion or swelling. We therefore have a diffusivity that is constant, with regard to time, in each defined material domain of the microcapsule. The release of active agents from the microcapsules, described in this work, can thus be explained by Fickian diffusion. Regarding the modelling of the experimental release data, it is convenient to express the analysed concentration as a fractional part of released active. The time-dependent released amount m(t) is divided by the total amount of active mtot initially positioned inside the microcapsule. Assuming perfect sink conditions, the complete timedependent release from a microsphere (the most geometrically simple release system, see Scheme 2b) with 29
ACCEPTED MANUSCRIPT homogeneous initial distribution of active substance, is expressed by the following equation derived by Crank[154]
D 2 n 2 m(t ) 6 f ( D, r , t ) 1 2 2 exp t 2 mtot n 1 n r
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21
where D is the apparent diffusion coefficient and r the microsphere radius. In a laboratory study, perfect sink
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conditions are practically never fulfilled and for the long-term part of the release, the deviation from Equation 21 can be significant. The equilibrium distribution, expressed by the distribution constant K, between microsphere and release medium with volumes Vsphere and Vsink, respectively, should therefore be taken into account providing
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a more realistic description using[154]
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Dqn2 m(t ) 1 f ( D, r , t ,Vsink ,Vsphere , K , t ) 1 6 1 exp 2 t 2 mtot 1 n 1 9 9 q n r
,
Vsink VsphereK
22b
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22a
.
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and where qn is the nth positive root of
tan qn
3qn 3 qn2
22c
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For the initial part of the release, Equations 21 and 22 coincide but the deviation is a key factor at longer times. Although only the analytical expressions of the microsphere geometry are shown here, it is always of great importance to include the effect of the equilibrium distribution of the active in the models regardless of internal microcapsule morphology. A second generic parameter that should be attributed to the modelling of release data is the effect of burst release[155]. It is suitable to describe the overall release of the active as a sum of two populations: burst and diffusive release. Compared with the more elaborate expression of the diffusive release, the initial burst fraction can be simply modelled as zero order or first order release[156] with the rate constant kburst. Note that a zero order burst function is discontinuous (Equation 23b) when applied in release modelling. A third common approach for any model describing release from microcapsules is the inclusion of the size polydispersity. The release profile is unmistakably influenced by the microcapsule size and size distribution and to find e.g. the proper diffusion coefficient from regression analysis, the fitting should include this distribution. For most formulation routes of microcapsules described in this review, the radius size distribution P(r) follows the log-normal distribution function[25, 157-159]. 30
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A general release model is given in Equation 23 where the population average of the burst and the diffusive part is sized by the fractions pburst and (1- pburst), respectively. 23a
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m(t ) f ( pburst, kburstD, r , t ) pburst f burst (kburst, t ) 1 pburst f ( D, r , K ,..., t ) P(r )dr mtot
23b
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f burst (kburst, t ) 1 : f burst (kburst, t ) f burst (kburst, t ) f burst (kburst, t ) 1 : 1
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Furthermore, the released active can be affected by e.g. degradation when exposed to the release medium. Since it is the detected amount from the experimental quantitative analysis we relate to as m(t), all chemical reactions in the release medium need to be taken into account. An example of this situation, with corrected m(t) using
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degradation models, has been described by us in a recent publication[38] and is treated in more detail in Section 3.2.1.
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3.1.2 Physicochemical properties of core-shell particles
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In contrast to the microsphere, a microcapsule is defined by its two (or more) compartments. The complete diffusion model of the release from microcapsules includes the partition coefficient between core and polymer
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shell, as well as that between shell and external release medium, and a comprehensive analytical expression has also been presented in the literature[160]. In the following section we present some of our work that describes
The core
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the significance of various parameters related to the release from microcapsules.
The inner core morphology and the distribution of the active in the core will have a profound effect on the release profile[23]. As an example, the only way to achieve zero-order release rate is when the active is distributed in the core as crystals surrounded by a rate determining shell[23]. In addition, the size distribution of the microcapsules has to be perfectly monodisperse. Regarding the magnitude of the release rate, the solubility and the diffusion coefficient of the active in the core are important parameters according to Equation 20. The diffusion coefficient can be dramatically altered by changing the physical state of the core. Nordstierna and Nydén studied the effect of using liquid (dodecane) and solid (octadecane) alkanes as core oils on the release of BHT (see Chart 1). BHT is a pesticide which can be used as an effective antifeedant against pine weevil in the early stages of plant protection. Given the results with accompanied Brownian dynamics simulations, reprinted in Fig. 6, the effective diffusion coefficient was significantly reduced when solid octadecane was used instead of liquid dodecane[22].
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Fig. 6. Fractional release m(t)/mtot of BHT as function of time t from microcapsules with (a) dodecane and (b) octadecane as core medium. Solid line represents the release from octadecane capsules calculated by computer simulation of Brownian motion. Note the different axis scales. Reprinted from[161].
Regarding the solubility, it is clear according to Equations 14, 15, 19 and 20 that a high solubility in the core for the active will reduce its distribution coefficient and hence the permeation. Dowding and Vincent investigated
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the release of 4-nitroanisol (see Chart 1) from polystyrene-based microcapsules with cores of varying polarity. It was demonstrated that the release of the active scaled inversely with its solubility in the core, which is in accordance with theory (Equations 19 and 20). The importance of the active’s solubility is often overlooked.
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There are many examples where the active has a higher solubility in the shell than in the core. Enejder et al. demonstrated that the fungicide IPBC (see Chart 1) was exclusively distributed in the shell of PMMA-based
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microcapsules with a dodecane core[141]. For such systems, the use of core-shell morphology is pointless and
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homogenous microspheres (see Scheme 2b) are the more appropriate choice. The core can also have more unexpected effects on the release. Liquid cores, which are the most common types of cores, may plasticise the shell (see Fig. 7). The plasticising effect from the core will increase the free volume of the polymeric shell (see Equations 16 and 17) and hence the effective diffusion coefficient. An example of
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such a system is PMMA-dodecane. Nordstierna revealed that the release of 4-nitroanisole (see Chart 1) from microcapsules consisting of a dodecane core and a PMMA shell was significantly higher than what was theoretically expected assuming simple diffusion through a homogenous PMMA shell[90].
Fig. 7. The fractional release m(t)/mtot of 4-nitroanisole as a function of time t from microcapsules (open circles) and microspheres (solid squares) with (a) 57mg and (b) 880mg 4-nitroanisole. Lower solid line is the fit of the model for diffusion in microspheres. Middle dashed line is the calculated result when applying the diffusion coefficient value obtained from the microsphere fit but for the microcapsule
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ACCEPTED MANUSCRIPT geometry (open circles). Upper dotted line is the fitting result from microcapsule geometry on the microcapsule data (open circles). Reprinted from[90].
The shell
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The shell is obviously the barrier with the purpose of reducing the release of the active and preventing absorption
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of the external medium. This means that the permeability must be lower in the shell than in the core, which can be achieved by choosing a polymer with a proper polarity. According to Equation 20, the release of the active is
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reduced as the solubility of the active in the shell is reduced. However, this is only valid as long as the solubility of the active is higher in the core than in the shell. If this is not the case, a higher solubility of the active in the shell will result in a reduced release rate[22, 115]. In such cases, the use of core-shell morphology is unnecessary
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and a microsphere morphology should be used (see Scheme 2).
The permeability of an active in the shell can also be reduced by introducing impermeable objects into the shell matrix as previously discussed in Section 3. Romero-Cano and Vincent estimated the effective diffusion constant
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of 4-nitroanisole (see Chart 1) in semi-crystalline PLLA (see Chart 2) to be as low as 10-19 m2/s[162]. Another example of impermeable objects is nanoclays. Given their high aspect ratios, aligned nanoclays may alter the tortuosity of the active in the shell material to a great extent. Fan and coworkers incorporated acidified
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montmorillonite into urea-formaldehyde based microcapsules for self-healing applications[163]. This treatment
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significantly reduced the release of cyclopentadiene. However, compatibility between the nanoclays and the surrounding shell matrix is of paramount importance. We have investigated the release of Disperse Red 13 from PMMA-based microcapsules containing hydrophobically modified montmorillonite[22]. It was found that a high
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reduced release rate.
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fraction of nanoclays impaired the integrity of the PMMA shell matrix resulting in an enhanced rather than
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Fig. 8. The fractional release m(t)/mtot of the fungicide OIT as a function of time t from microspheres formulated by slow evaporation of the volatile solvent () and fast evaporation of the volatile solvent (). Solid lines are fitted models according to Equation 22. The top figure shows the entire release using a logarithmic scale while the first 35 hours of the experiment are presented on a linear time scale in the bottom figure. Reprinted from[38].
The above paragraph deals with the macroporosity discussed in Section 3. However, it is also possible to alter the microporosity. Dowding and Vincent was able to successfully reduce the release of 4-nitroanisole from PS, poly(vinyl phenyl ketone) and poly(2-vinylpyridine) based microcapsules by cross-linking the shell polymer[115] as described in Section 2.1.4. Another, more simple, method to reduce the microporosity is to anneal the shell polymer[38, 88] as is also described in Section 2.1.4. The polymeric matrix of the shell is in many cases in an expanded state due to experimental conditions such as fast evaporation of the volatile solvent. We have recently demonstrated that this is the case when rotary evaporation is used for solvent removal from the microcapsule dispersion[38]. This is in contrast to microcapsules prepared by slow evaporation of the solvent at ambient conditions. As a consequence, the calculated effective diffusion constant of the fungicide OIT (see Chart
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ACCEPTED MANUSCRIPT 1) in PMMA is significantly higher from rotary-evaporation-prepared microcapsules as compared to fume-hoodevaporation-prepared microcapsules (see Fig. 8). In addition, annealing the shell by heating the microcapsule suspension in an autoclave resulted in a significant reduction of the release rate from the rotary-evaporationprepared microcapsules (see Fig. 9). In contrast, this treatment had no effect on the fume-hood-evaporation-
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prepared microcapsules[38] (see Fig. 9). This means that the shell polymer of the latter type of microcapsules is
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already in a conformationally stable state.
Fig. 9. a) The fractional release m(t)/mtot of the fungicide OIT as a function of time t for untreated fume-hood-evaporated microcapsules () and the corresponding heat-treated microcapsules (). b) The release of OIT for untreated rotary-evaporated microcapsules ()and the corresponding heat-treated microcapsules ().[38].
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ACCEPTED MANUSCRIPT The outer dispersant layer and surface modifications In this section we will discuss the outermost layers of the microcapsules and their importance for the release. As mentioned in Section 2.1.3, the dispersant layer; albeit thin, can be viewed as an outer barrier. Before discussing this matter further, it is important to make a point regarding the physicochemical properties of the active, the
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shell and the resulting permeation of the active in the shell. As is clear from Equations 15 and 20, it is beneficial
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for the active to have a low solubility in the shell. This is usually already the case for actives encapsulated in microcapsules prepared via the double emulsion route, since the actives are hydrophilic and the shell polymer
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hydrophobic. However, for microcapsules prepared via the internal phase separation route, the actives are hydrophobic and the shell is semihydrophobic (see Section 2.1.1). This situation is not ideal because hydrophobic actives should require very hydrophilic barriers according to Equations 15 and 20. Yet, knowledge of this problem can be exploited when it comes to the choice of dispersant. We have recently investigated the
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release of the hydrophobic dye Disperse Red 13 (see Chart 1) from PMMA-based microcapsules which have been stabilized by different types of dispersants[27]. The amphiphilic block copolymer dispersants with a polyelectrolyte block (see Chart 2) provides the microcapsules with a highly charged polyelectrolyte brush-like
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surface and this layer is very hydrophilic[25]. It is clear in Fig. 10a that the release rate of the dye is significantly reduced when this block copolymer is used as dispersant as compared to the more conventional uncharged dispersants PMAA and PVA (see reference [27]). Note that the thickness of the block copolymer layer on the
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microcapsule surface is much thinner than the thick gel-like PVA layer (see Fig. 3).
Fig. 10. a) The fractional release m(t)/mtot of the dye Disperse Red 13 as a function of time t from microspheres based on the dispersants PMAA (▼, blue), 600- 4600 (■, black) and 600- 4600 functionalized with 0.5 (●, dark grey), 1.0 (●, grey), 1.5 (●, light grey) and 2.0 (○, white) PDADMAC/PMANa bilayers. The insert magnifies the release from the 600-4600 based microspheres. Redrawn from[27]. b) The
36
ACCEPTED MANUSCRIPT calculated effective diffusion coefficient or permeability of Disperse Red 13 as a function of the PEM thickness on PMMA based microcapsules. The data has been fitted with a hyperbolic function A/(B+δm/C) derived from Equations 19 and 20. Here, Deff(δm=0) is the obtained diffusion coefficient for the bare PMMA surface and the polyelectrolyte brush thickness corresponds to the effective thickness of the collapsed brush. Redrawn from[23].
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The idea of using very hydrophilic barriers against hydrophobic actives can also be implemented when it comes to surface modifications. As mentioned in Section 2.1.4, polyelectrolyte multilayers, PEMs, are very promising
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as such barriers. We have modified PMMA-based microcapsules with PEMs to reduce the release of the
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hydrophobic dye Disperse Red 13 and the fungicide OIT[27, 39] (see Chart 1). As can be seen in Fig. 10, the release rate is profoundly reduced by the PEM barrier and the PEM quickly becomes the rate-determining barrier. For Disperse Red 13, the PEM modification with two polyelectrolyte bilayers results in a 160 times decrease of the effective diffusion constant as compared to the untreated microcapsule[27]. Note that this
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decrease of the release rate is significantly larger compared to the decrease resulting from other modifications mentioned in this review as well as most modifications found in the literature. On the other hand, OIT is much less hydrophobic than Disperse Red 13 and displays surface-active properties[38]. It is therefore not necessarily
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certain that the PEM will have an effect on such a “challenging” molecule. Nonetheless, the effective diffusion constant of OIT is reduced 40 times for microcapsules modified with two polyelectrolyte bilayers as compared to the unmodified microcapsules (see Fig. 11). This emphasizes the universal validity of using highly hydrophilic
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PEM barriers against different types of hydrophobic actives.
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Fig. 11. The fractional release m(t)/mtot of the fungicide OIT as a function of time t from PVA-based () and PEM-modified () microspheres . Filled and unfilled symbols are duplicates. The top figure shows the entire release using a logarithmic scale while the first 25 hours are shown in the bottom figure, on a linear scale.
3.2
Release from coatings
Similar to Section 3.1, this section starts by describing the experimental release methodology with respect to coatings. Regarding the diffusion models, the feature of biocide degradation is also taken into account since the duration of the release experiments relevant for coatings is significantly longer than for microcapsule suspension. Thereafter, some release examples concerning both “free” (not encapsulated) and encapsulated biocides are presented with a focus on the properties of the coating matrix.
3.2.1 Methodology and diffusion models The experimental methodology with respect to the release of actives from coatings contains the same steps as discussed in Section 3.1; setup, analysis, and evaluation. In the studies presented here, the setup is based on an 38
ACCEPTED MANUSCRIPT international standard method for the release of biocides from coatings[164] with some minor modification such as permanent immersion of the coating in water instead of a dipping setup. The entire setup is depicted in Scheme 7.
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Scheme 7. Experimental methodology for release measurements of active substances from coatings. The paint consisting of microcapsules are coated on a grazed polypropylene plate before drying. The coated plate is then placed in a water-filled beaker with lid, and stirred using a shaking table. Samples are collected using an automatic pipette. Since the sample may contain unwanted arbitrary paint species, dust etc. chromatography is used to separate the active substance before quantitative analysis by e.g. spectrophotometry.
For a proper release study, the preparation of the coated substrate is important. The substrate has to allow for thin-film coating and needs to be chemically and physically inert to the coating ingredients. Moreover, the substrate should be physically unaffected by film drying and water contact. In our laboratory we use polypropylene plates as substrate[11, 38, 39]. In order to enhance the adhesion between the substrate and the coating and to prevent coating detachment, the surface is grazed in order to roughen the surface. Moreover, precise control of the surface area and the coating thickness is of utmost importance. This can be achieved by defining the surface area with tape followed by coating with an applicator giving a defined thickness. The paint 39
ACCEPTED MANUSCRIPT is usually dried at slightly elevated temperature (37 °C). The drying time has a profound impact on the release, as discussed in Section 3.2.3. Regarding the experimental methodology, it is important to precisely determine the wet and dry weight of the coating. These parameters will ultimately determine the loading concentration and the
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analysed released fraction.
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According to the international standard method used, the coated plates are placed in a beaker leaning against the beaker wall with the coated surface facing downwards (see Scheme 7). It is important to tightly seal the beaker
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with a top cover in order to prevent evaporation of the release medium. In addition, the stirring (usually realized by shaking) should be gentle in order to prevent release due to mechanical load on the coating. The analysis procedure requires a short comment. In contrast to the situation when the release from
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microcapsules is evaluated (Section 3.1.1), the release medium surrounding the coated plate is a solution since all microcapsules are confined within the coating matrix (see 2.2). This means that the concentration of the active can be analysed without any filtration step. However, other small molecules from the paint will be
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released into the surrounding medium. Therefore, spectrophotometry (UV/Vis), which is normally used for analysis of the release in microcapsule suspension, cannot be directly used due to possible overlapping effects. Instead chromatography-based methods such as HPLC are more suitable. Chromatography in conjunction with
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spectrophotometry can be used to evaluate the concentration as well as possible degradation events. This latter topic is briefly discussed below and in Section 3.2.3. Compared to the previously discussed release studies in
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aqueous microcapsule suspensions, the concentration of active substance is here much lower, giving conditions
Diffusion models
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more closely related to a perfect sink.
The common laboratory procedure when it comes to applying paint on a substrate is to use a paint applicator that
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provides a well-defined thickness of the painted layer, typically between 50 and 200 m. The coating will after the drying procedure form a thin film with a homogeneous distribution of all components (binder, pigment, additives, etc.). Such a film can, from a model perspective, be seen as a homogeneous matrix which enables an experimental quantification of the release by model fitting (see Equation 24). The apparent, or effective, diffusion coefficient D for an active substance migrating through and leaving the coating to a perfect sink can be modelled according to the following function derived by Crank
mt 8 f D, L, t 1 2 mtot
D2 2n 12 1 exp t 2 4 L2 n 0 2n 1
24
where L is the thickness of the coating. This model is often satisfactory despite the perfect sink condition since the concentration of released active substance is a couple of magnitudes lower compared to the situation in Section 3.1. It is important to note that the effective diffusion constant for the coating is a function of all partition coefficients and diffusion constants depicted in Scheme 5. Therefore, this effective diffusion constant can also be viewed as a global permeability for the coating 40
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The measurements of the release from coatings are usually performed over significantly longer time spans than the release measured in microcapsule suspension. As a consequence, potential degradation of the active may influence the measured release. Note that most modern biocides are designed to be chemically or biologically
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degradable in order to prevent unwanted long-term biocidal activity and bioaccumulation in the environment. In
g ( x) m k deg g ( x) dx
25
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d
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order to account for potential degradation, the order m of the degradation reaction must be known (Equation 25):
g(x) is the concentration of non-degraded active that is measured in the sink during time x and x = t - j. The rate constant of degradation is denoted kdeg, and m is the order of the degradation kinetics. The mass change of active is expressed as the fractional parts of the release function (Equation 24) and the degradation (Equation 25). The
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corresponding function that explicitly describes first-order (or rather pseudo-first order) degradation, which is the most common order of degradation kinetic (e.g. hydrolysis), is then given by Equation 26[38]:
m(t ) f ( D, L, j ) d exp kdeg t j dj mtot 0 dj .
26
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We have implemented this modified release function with respect to the degradation of OIT (see Chart 1). This is
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presented in Sections 3.2.3 and 3.3.2.
3.2.2 Solvent-borne vs. water-borne paints The nature of the coating matrix will have a profound effect on the release of the active. It is therefore necessary
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to make a distinction between water-borne and solvent-borne paints. Most water-borne paints consist of polymer lattices dispersed in an aqueous medium. As the paint dries, the space between the particles, i.e. the pores, will be reduced, ultimately resulting in merging of the particles[42, 43]. Any dispersed actives will reside in the pores or be adsorbed on the particle surface. Regarding dispersed microcapsules, the encapsulated active will be
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released into the pores of the coating matrix. The situation is very different for solvent-borne paints. Here, the polymer constituting the coating matrix, i.e. the binder, is dissolved in the solvent. Any dispersed active or microcapsule will be surrounded by the polymer matrix. Therefore, the macroscopic porosity described for the water-borne paints is not relevant for the solvent-borne paints. Instead, the microscopic porosity of the polymeric coating matrix is the important parameter (see Scheme 8).
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Scheme 8. Macroporosity of a latex coating (a) and microporosity of a solvent-borne coating (b) as a function of drying time. a) The porosity of the latex coating will decrease as a function of drying time and is at the initial stages approximately equal to the water content. The porosity ε and tortuosity τ have been defined in Section 3. b) For the solvent-borne paint, the free volume of the matrix Vf will decrease as a function of drying time resulting eventually in a thermally relaxed state. Vf and V0 have been defined in Section 3 and are also discussed in Section 3.1.
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We have investigated the release of the antifouling agent medetomidine (see Chart 1) from both solvent- and water-borne coatings[11]. Both free and encapsulated medetomidine in PMMA-based microcapsules were investigated (see Fig. 12). As can be seen, the release rate is significantly reduced for both paint systems when the active is encapsulated. However, we must note that the effect of the encapsulation on the release rate for a given type of active in a given type of microcapsule can vary significantly. In some cases, the effect of encapsulation is almost absent. Such an example is IPBC (see Chart 1) encapsulated in PMMA-based microcapsules. Whether the encapsulation will have an effect on the release rate or not depends on whether the microcapsule is the rate determining barrier for release according to Equations 19 and 20. However, it is clear from Fig. 12 that the coating matrix has a more profound effect on the release than the encapsulation. As expected, the water-borne coating, containing macroscopic pores, displays a much higher release rate of medetomidine as compared to the solvent-borne coating containing microscopic pores. Note that the observed secondary effect of the encapsulation is not general but depends on the type of microcapsule system used. In Section 3.3.2, we will present a microcapsule system which is virtually independent on the coating matrix.
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Fig. 12. The fractional release m(t)/mtot of medetomidine as a function of time t, on (a) lin–lin scale and (b) log–log scale, from () waterborne paint with free biocide, () water-borne paint with encapsulated biocide, () solvent-borne paint with free biocide, and () solventborne paint with encapsulated biocide. Intersecting lines are solely for clarification. Reprinted from [11].
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3.2.3 Drying time and macroscopic porosity The former section described the importance of porosity of the coating for water- and solvent-borne paints. It is therefore obvious that the drying time of the paint; which will significantly affect the matrix porosity (see Scheme 8), will have a profound effect on the release rate of the active. We have investigated the release of the fungicide OIT from water-borne varnishes, which have been dried at elevated temperatures (37 ºC), as a function of the drying time[38]. Even at this elevated temperature, the complete drying process takes approximately one month. As can be seen in Fig. 13, the drying time has a profound effect on the release rate of OIT. Note also that the degradation of OIT becomes important for these long-term release measurements in contrast to the measurements presented in Section 3.1.2. For the coating which had been dried for seven days, the microencapsulation of OIT results in a twofold reduction of the calculated effective diffusion (or rather permeation) coefficient (Equation 26) as compared to the freely dispersed OIT. However, the effect of altering the macroscopic porosity of the coating matrix is more important than the effect of encapsulation (see Fig. 13). Yet, it is important to note that drying times at elevated temperatures for one month are practically unrealistic, which means that this simple type of encapsulation is still useful.
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Controlled release and antifouling
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Fig. 13. The fractional release m(t)/mtot of the fungicide OIT as a function of time t from varnish coatings. (a) The first 25 hours. (B) The entire experimental time. () Free OIT after 7 days drying with decay fitting (•••) and without decay fitting (—). () Encapsulated OIT after 7 days drying with decay fitting (∙∙∙) and without decay fitting (- - -). () Free OIT after 28 days drying with decay fitting (•••) and without decay fitting (—). () Encapsulated OIT after 28 days drying with decay-fitting (∙∙∙) and without decay fitting (- - -). Reprinted from [38].
In the following subsections, the antifouling and antimicrobial properties of some coatings are presented.
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Emphasis is given to the exceptional antifouling property of biocides with high efficacy and narrow target specificity, in this case medetomidine. We also present the benefit of using strongly rate-determining release systems, such as the PEM-modified microcapsules (see Sections 2.1.4 and 3.1.2), for controlled release of
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hydrophobic biocides, and thereby controlling and prolonging the antifouling/antimicrobial function.
3.3.1 Use of non-hazardous and efficient biocides As mention in Section 2, a promising and, more importantly, feasible antifouling approach is to use a cocktail of biocides with a narrow target specificity towards different types of biofoulers (see Fig. 1). One such biocide, which has been comprehensively investigated in the Marine Paint research programme is medetomidine (see Chart 1). It was shown that medetomidine; which is a α2-adrenoceptor agonist and used as a veterinary sedative, activated the octopamine receptor in barnacles[165]. The barnacle is one of the most problematic biofoulers. Normally, the barnacle settles on a surface in its cyprid larval stage followed by metamorphosis to the adult stage. The octopamine receptor activation of medetomidine increases the swimming activity of the cyprid larvae, preventing it from settling[165]. Medetomidine has been shown to have low toxicity towards several non-target organisms[75, 78-81], and was recently recommended for EU approval by the UK Health and Safety Executive (HSE).
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ACCEPTED MANUSCRIPT The antifouling property of medetomidine was tested by simply mixing the free biocide in a solvent-borne paint which was applied on PMMA plates. After four months immersion in the sea the fouling was assessed. It was found that even though the biocide was not encapsulated, proper antifouling was realized at as low concentrations in the coating as 0.1 wt% (see Fig. 14). As a comparison, the amount of copper used in marine
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paints as biocide is usually as high as 40 wt%.
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Fig. 14. Effect of medetomidine formulated in a solvent-borne paint on barnacle fouling. The percentages given are added amount of medetomidine in the paint formulation.
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Optimized biocide cocktails
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In the Marine Paint project that was mentioned in the Introduction, it became obvious that medetomidine was efficacious only to barnacles, and that a fully protective coating needed to contain a mixture of antifoulants. Experimental work on a set of important fouling organisms and antifouling compounds[66, 166] generated data
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that were used to make a mixture efficacy analysis in silico for all possible combinations of antifoulants. That generated “recipes” for a large number of combinations, all equally effective in controlling the model organisms (barnacles, bryozoans, sea squirts, sea lettuce[166], and multi-species periphyton communities[67]). These
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antifoulant mixtures were then evaluated and ranked based on their estimated environmental risks using Equation 26 to estimate the combined risk for each of the mixtures where ci is the concentration, PECi is the predicted environmental concentration and PNECi is the predicted no effect concentration of biocide i. There was a 100-fold difference between the most and the least risky mixture. An example of the antifouling effect of a coating formulated with such an optimized mixture of biocides in terms of efficacy and environmental low risk is shown in Fig. 15. n
Risk ci i 1
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Already when these ideas emerged it was obvious that conventional paint formulations might fall short and microencapsulation and nanoparticle approaches were therefore explored. It was anticipated that with these paint technologies it might be possible to finetune formulations for certain environments or when fouling organisms acquire tolerance.
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Fig. 15. Marine biofouling on coatings containing (a) no biocides, (b) an optimized mixture of DCOIT, copper, and medetomidine and (c) a tenth of the optimized mixture concentration. Photos by Åsa Arrhenius and Anneli Hilvarsson. The biocide cocktail was optimized from a a set of conventional biocides; copper pyrithione, irgarol, copper, triphenylborane pyridine, DCOIT and medetomidine[17, 167]. The risk for the cocktail containing DCOIT (11 %), copper (89%) and medetomidine (0.035%) was more than ten times lower as compared with conventional copper-based paints and the amount of copper needed for antifouling protection was reduced 24 times.
3.3.2 Use of rate-determining release systems So far in this review, alteration of the coating matrix has been more important than the effect of encapsulating the active with respect to the release. Regarding the release of hydrophobic actives from microcapsules in suspension, the PEM-modified microcapsules displayed especially low release rates. We have therefore investigated the release of the fungicide OIT, encapsulated in PEM-modified microcapsules, from water-borne acrylate paints as a function of drying time[39]. Similar to the varnish example discussed in Section 3.2.3, the release rate of OIT, which has been freely dispersed in the paint, depends strongly on the drying time (see Fig. 16). In contrast, the release rate of OIT; which is encapsulated in PEM-modified microcapsules, is more or less
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rate-determining barrier according to Equations 19 and 20.
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Fig. 16. The fractional release m(t)/mtot of OIT from a latex paint as a function of time t. In a), the first 300 hours of the experiment are displayed and the data have been fitted according to the diffusion models in Equations 24-26. In b), the long-term release is displayed which includes the degradation of the biocide. Freely dispersed OIT is marked with ■ and □, and encapsulated OIT with and. Filled and unfilled symbols are duplicates. The different drying times are marked with, black (one day), blue (seven days) and red (33 days). The fitting for the curve corresponding to free OIT dried for one week was poor and has consequently been excluded. Reprinted from[39].
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The antifouling properties of the above-described coatings against Aspergillus niger was investigated in an accompanying study[39]. The fungus Aspergillus niger, also known as black mold, is a common biofouler on
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exterior walls[9, 13, 39] (see Fig. 1). The antifungal effect of OIT relies on its ability to diffuse over the cell wall of the fungi. In the cytoplasm, the electron deficient sulfur of the isothiazolinone reacts with the thiols in proteins leading to impairment of their enzymatic functions and eventually to cell death[168]. OIT is therefore a broadspectrum biocide of the conventional type and not a selective biocide such as medetomidine. Nonetheless, it is
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clear from Fig. 17 that the encapsulation of OIT in PEM-modified microcapsules profoundly prolonged the antifouling effect of the coating[39]. The minimum surface concentration required for an antifouling effect is determined by the surface flux of active from the coating (the derivative of the release curves in Fig. 16) which is schematically exemplified in Fig. 5 and shown in Fig. 17. Above the critical surface flux of OIT in Fig. 17, fouling of Aspergillus niger is prevented whereas the protection is lost below the critical surface flux.
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Fig. 17. a) Surface flux ṁ(t)/A as a function of time t calculated from the diffusion-modelled release in Figure 15. The colour schemes and symbols are presented in (b). b) Antifouling effect for different coatings assessed by the growth of Aspergillus niger (see Fig. 1). The coatings have been formulated with free (F) and encapsulated OIT in PEM-modified microcapsules (M). A coating containing no addition of OIT was used as a blank reference surface (B). In addition, the drying time for these three types of formulated coatings have been varied (one, seven and 33 days). The plates were removed from the release medium after a specific time which is indicated by the symbols (x-axis) in (a) and their antifouling ability was subsequently assessed. The grey horizontal line in (a) illustrates the critical surface flux needed to reach the critical surface concentration (see Section 3 and Fig. 5).
4
Conclusions
Microencapsulation provides many benefits for coatings containing biocides. The microcapsule protects the biocide from degradation and enables the use of a cocktail of different types of biocide in the coating. Most importantly, the encapsulation allows for proper control of the release and thereby prolongs the duration of the biocidal effect and reduces the waste of biocides. This is advantageous with respect to both economic and environmental perspectives. Hydrophobic biocides are preferably encapsulated using the internal phase separation route whereas the double emulsion route is most suitable for hydrophilic biocides. The intention is always to maximize the solubility of the biocide in the core and minimize its solubility in the shell or shells. The resulting release can consequently be controlled by the choice of core and shell materials. Even the choice of
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shown that addition of a PEM, in which the biocide has very low solubility, on the microcapsule surface is a very
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promising way of reducing the release rate of hydrophobic biocides. When such microcapsules are mixed into a coating formulation, the release is virtually independent of the coating matrix, which is a unique feature. This
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Acknowledgements
The authors are grateful to Alberta Mok, Alireza Movahedi, Anna Ananievskaia, Asvad A. Gabul-Zada, Gia
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Kourouklidou, Hermann Uhr, Kurt Löfgren, Matias Nordin and Ye Li for laboratory, analytical and administrative work regarding microencapsulation and controlled release. The authors are also grateful to all coworkers in the Marine Paint Research Programme and in the innovation companies Capeco and I-Tech.
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We also acknowledge funding from Mistra (the Swedish Foundation for Environmental Research), the Swedish
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Bouillot, Langmuir 21 (2005) 5278. M. Andersson Trojer, Y. Li, M. Wallin, K. Holmberg, M. Nyden, J. Colloid Interface Sci. 409 (2013) 8. E. Donath, G.B. Sukhorukov, F. Caruso, S.A. Davis, H. Mohwald, Angew. Chem., Int. Ed. 37 (1998) 2202. G.B. Sukhorukov, E. Donath, S. Davis, H. Lichtenfeld, F. Caruso, V.I. Popov, H. Mohwald, Polym. Adv. Technol. 9 (1998) 759. G.B. Sukhorukov, E. Donath, H. Lichtenfeld, E. Knippel, M. Knippel, A. Budde, H. Mohwald, Colloids Surf., A 137 (1998) 253. S. Moya, E. Donath, G.B. Sukhorukov, M. Auch, H. Baeumler, H. Lichtenfeld, H. Moehwald, Macromolecules 33 (2000) 4538. L. Wang, M. Schoenhoff, H. Moehwald, J. Phys. Chem. B 106 (2002) 9135. G. Decher, J.B. Schlenoff, Multilayer Thin Films: Sequential Assembly of Nanocomposite Materials. Wiley, Weinheim, 2003. J.M.G. Cowie, Polymers: Chemistry & Physics of Modern Materials. 2nd ed., CRC Press, 1991. D.J. Shaw, Introduction to Colloid and Surface Chemistry. 4th Ed. 1992. B. Karger, L.R. Snyder, C. Eon, J. Chromatogr. 125 (1976) 71. B.L. Karger, L.R. Snyder, C. Eon, Anal. Chem. 50 (1978) 2126. S. Freitas, H.P. Merkle, B. Gander, J. Controlled Release 102 (2005) 313. H. Sah, J. Controlled Release 47 (1997) 233. M. Andersson, O. Hansson, L. Oehrstroem, A. Idstroem, M. Nyden, Colloid Polym. Sci. 289 (2011) 1361. M. Andersson Trojer, A. Maartensson, M. Nyden, Vib. Spectrosc. 61 (2012) 38. A.P. Esser-Kahn, S.A. Odom, N.R. Sottos, S.R. White, J.S. Moore, Macromolecules (Washington, DC, U. S.) 44 (2011) 5539. D. Tarn, C.E. Ashley, M. Xue, E.C. Carnes, J.I. Zink, C.J. Brinker, Acc. Chem. Res. 46 (2013) 792. J.A. Zasadzinski, B. Wong, N. Forbes, G. Braun, G. Wu, Curr. Opin. Colloid Interface Sci. 16 (2011) 203. H. Lewis D, R. Cowsar D, in: Controlled Release Pesticides; ACS, 1977, p 1. J.H. Richards, in: J. Comyn, (Ed.) Polymer Permeability; Elsevier, London, 1985, p 217. E. Rogers C, in: H.B. Scher, (Ed.) Controlled Release Pesticides; ACS, Washingon, 1977, p 17. P.N. Bartlett, J.W. Gardner, Philos. Trans. R. Soc. London, Ser. A 354 (1996) 35. E. Rogers C, in: R. Paul D, F.W. Harris, (Eds.), Controlled Release Polymeric Formulations; ACS, Washingon, 1976, p 15. J. Crank, G.S. Park, Diffusion in Polymers. Academic Press, London, 1968. S. Wang, Microporous Mesoporous Mater. 117 (2008) 1. A. Enejder, F. Svedberg, L. Nordstierna, M. Nyden, Proc. SPIE 7903 (2011) 79030S/1. M. Alauddin, R.E. Verrall, J. Phys. Chem. 88 (1984) 5725.
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Graphical abstract
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ACCEPTED MANUSCRIPT Highlights The encapsulation strategy depends on the physicochemical properties of the active
The important parameters with respect to a proper release methodology are described
The release is reviewed in terms of the properties of the microcapsule
Suitable theoretical diffusion models are presented and discussed
Controlled release and antifouling properties of coatings are presented
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S0001-8686(14)00198-5 doi: 10.1016/j.cis.2014.06.003 CIS 1450
To appear in:
Advances in Colloid and Interface Science
Received date: Revised date: Accepted date:
2 February 2014 6 June 2014 6 June 2014
Please cite this article as: Trojer Markus Andersson, Nordstierna Lars, Bergek Jonatan, Blanck Hans, Holmberg Krister, Nyd´en Magnus, Use of microcapsules as controlled release devices for coatings, Advances in Colloid and Interface Science (2014), doi: 10.1016/j.cis.2014.06.003
This is a PDF file of an unedited manuscript that has been accepted for publication. As a service to our customers we are providing this early version of the manuscript. The manuscript will undergo copyediting, typesetting, and review of the resulting proof before it is published in its final form. Please note that during the production process errors may be discovered which could affect the content, and all legal disclaimers that apply to the journal pertain.
ACCEPTED MANUSCRIPT Use of microcapsules as controlled release devices for coatings Markus Andersson Trojer,*a,b Lars Nordstierna,a Jonatan Bergek,a Hans Blanck, c Krister Holmberga and
Department of Chemical and Biological Engineering, Applied Surface Chemistry, Chalmers University of
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Magnus Nydéna,d
Technology, Göteborg, Sweden.
Department of Colloid Chemistry, Max Planck Institute of Colloids and Interfaces, Potsdam, Germany. E-mail:
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[email protected]
Department of Biological and Environmental Sciences, University of Gothenburg, Göteborg, Sweden
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Ian Wark Research Institute, University of South Australia, Mawson Lakes Campus, Adelaide, South Australia
5095, Australia *E-mail: [email protected]
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Abstract
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Biofouling of surfaces is a considerable problem in many industrial sectors and for the public community in general. The problem is usually approached by the use of functional coatings and most such antifouling coatings
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rely on the effect of biocides. However, a substantial drawback is the poor control over the release of the biocide as well as its degradation in the paint. Encapsulation of the biocides in microcapsules is a promising approach that may overcome some of the problems associated with the more traditional ways of incorporating the antifouling agent into the formulation. In this review, we summarize more than a decade of microcapsule
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research from our lab as well as from other groups working on this topic. Focus will be on two coacervationbased encapsulation techniques; the internal phase separation method and the double emulsion method, which together enable the encapsulation of a broad spectrum of biocides with different physicochemical properties. The release of the biocide from core-shell particles and from encapsulated biocides in coatings is treated in detail. The release behaviour is interpreted in terms of the physicochemical properties of the core-shell particle and the coating matrix. In addition, special attention is given to the experimental release methodology and the implementation of proper diffusion models to describe the release. At the end of the review examples of antifouling properties of some coatings against common biofoulers are presented.
Keywords Antifouling, Marine paint, Exterior wall paint, Sustained release, Dyes, Diffusion
Contents of paper
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ACCEPTED MANUSCRIPT Abstract ................................................................................................................................................................... 1 Keywords ................................................................................................................................................................ 1 Contents of paper .................................................................................................................................................... 1 Introduction ..................................................................................................................................................... 3
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Formulation ..................................................................................................................................................... 6
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Microencapsulation ............................................................................................................................. 10
2.1.1
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Internal phase separation ................................................................................................................. 13
The spreading conditions .......................................................................................................................... 13 Predicting morphology using the van Oss formalism ............................................................................... 14 Double emulsion ............................................................................................................................. 16
2.1.3
Choice of dispersant ........................................................................................................................ 17
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Internal phase separation ........................................................................................................................... 18 Double emulsion ....................................................................................................................................... 18 Post-encapsulation modifications .................................................................................................... 19
2.1.5
Solvent choice and environmentally friendly approaches ............................................................... 19
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Microcapsules in paints ....................................................................................................................... 20
3.1
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Controlled release ......................................................................................................................................... 23 Release from core-shell particles ......................................................................................................... 26
3.1.1
Release methodology and diffusion models .................................................................................... 26
Diffusion models ....................................................................................................................................... 29 3.1.2
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Physicochemical properties of core-shell particles ......................................................................... 31
The core..................................................................................................................................................... 31 The shell .................................................................................................................................................... 33 The outer dispersant layer and surface modifications ............................................................................... 36 3.2
Release from coatings .......................................................................................................................... 38
3.2.1
Methodology and diffusion models ................................................................................................. 38
Diffusion models ....................................................................................................................................... 40 3.2.2
Solvent-borne vs. water-borne paints .............................................................................................. 41
3.2.3
Drying time and macroscopic porosity............................................................................................ 43
3.3 3.3.1
Controlled release and antifouling ....................................................................................................... 44 Use of non-hazardous and efficient biocides ................................................................................... 44
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Use of rate-determining release systems ......................................................................................... 46
Conclusions ................................................................................................................................................... 48
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Acknowledgements ............................................................................................................................................... 49
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References ............................................................................................................................................................. 49
Introduction
Coatings , such as paints, varnishes, and lacquers , are used for a large variety of functions and can be divided
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into decorative, protective and functional coatings[1-3]. Functional and smart coatings; including self-healing, self-cleaning, antimicrobial/antifouling[3, 4], lubricating, self-polishing (SPC paints)[5], flame-retardant and piezo-electric coatings, are becoming increasingly important in a variety of industrial sectors[1]. Antifouling and
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antimicrobial coatings are probably the most widespread examples of functional or smart coatings. Fouling of organisms on surfaces, sometimes called biofouling, is a huge problem in many industrial applications such as wastewater plants, implants and wound care products. The perhaps most prominent examples are biofouling on
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ship hulls[6] and exterior walls (see Fig. 1)[7-9] and this review will focus on these applications. Regarding house façades, the biofouling (algae, moulds, lichens, mosses, etc.) results in discolouration and potential
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damage of the substrate[9-13]. The marine biofouling on ship hulls (microalgae, seaweeds, mussels, barnacles, etc.)[14, 15] will result in an increased friction drag leading to increased fuel consumption and therefore
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pollution of the environment[4, 6, 15-18]. As an example, a ship without an antifouling coating may increase its
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fuel consumption by 40 %[5, 17, 19].
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Fig. 1. Examples of potential fouling organisms on house façades and aquatic surfaces. The A. niger image is reprinted from references[20, 21]. The exterior wall image is reprinted from[9]. The biofouling on exterior walls is not only an esthetic issue. Mold growth on houses results in a phenomenon called sick building syndrome. The release of mycotoxins and allergenic bioaerosols (e.g. spore fragments) is connected to the development of respiratory diseases and is consequently a public health problem[9, 13]. Regarding marine biofouling, a layer of slime will increase the fuel consumption by 1-2 %. Soft foulers such as seaweeds, hydroids or bryozans result in a further increase of the fuel consumption to 10%. Shell-forming organisms such as mussels and barnacles, called hard foulers, have such a large effect on the skin friction drag that the fuel consumption increases with 40 %[19].
The most common way to prevent fouling is to incorporate antifouling biocides (antifoulants) in the paint. Regarding marine paints, these biocides are usually termed booster biocides[4, 5, 15] indicating that they improve the performance of the usually copper-based paint. A major problem with the antifouling paints that are on the market today, in addition to the use of poisonous and potentially bioaccumulating biocides, is the poor control of the release of the antifoulants. Many of these coatings suffer from premature leakage or degradation of the biocides. Too fast release of the biocide will result in a loss of the coating’s antifouling function before the end of its intended life time. The desired lifetimes of façade coatings and of marine paints are at least 10 and 2 years, respectively, and achieving release profiles that cover such long times is a very demanding task[2, 22]. One approach to reach the required life times is to use high amounts of biocides in the coating but this may not be economically, environmentally, practically, or legally acceptable. Practically, the use of high concentrations of biocides in the formulation may result in macroscopic phase separation in the film.
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ACCEPTED MANUSCRIPT A very promising solution to many of the biocide-related problems mentioned above, which are discussed more in detail in Section 2, is microencapsulation of the biocides[22, 23]. Microencapsulation involves encapsulation of the biocide in miniature reservoirs called microcapsules or core-shell particles. It is worth mentioning that microcapsules have also found use in other types of functional coatings, including self-healing paints[1, 24].
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Figure 2 shows electron microscopy images and bright field microscopy photographies of various microcapsules.
Fig. 2. Different types of microcapsules based on poly(methyl methacrylate) (PMMA) and polystyrene (PS). a) SEM image of regular microcapsules of core-shell type based on PMMA with a dodecane core[25]. The core and the shell have been coloured in order to better display the internal morphology. b) SEM image of a PMMA-based microcapsule cut with a focused ion beam in order to display the multicore morphology[26]. The core and the shell have been coloured in order to better display the internal morphology c) Bright field microscopy image of a microsphere suspension dyed with Disperse Red 13[27]. d) Bright field microscopy image of microcapsules based on PS with an aqueous gelatine core which has been dyed with the natural dye betanin[28]. e) Bright field microscopy image of microcapsules based on a PMMA shell and a PS core[29]. The PS core has been dyed with β-carotene and the PMMA shell with Methylene blue.
In our group, which has been a part of the interdisciplinary research programme “Marine Paint”[17, 30] and the innovation company “Capeco” [10, 31], we have investigated antifouling and antimicrobial coatings[10, 11, 30, 32-39] for a decade and implemented microcapsule technology[10, 11, 22, 23, 25, 27, 31, 38-41]. It should be noted that a lot of antifouling research is focused on non-biocidal approaches[2, 18, 24, 42-59]. Some recent innovations regarding such biocide-free antifouling approaches comprises the use of anaerobic biofilms[54] and enzymatic peroxide formation[45, 48, 55-58]. However, approaches based on the release of biocides are still the most common and it is actually also a common strategy to combat fouling in biological systems[4, 60, 61]. In
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physicochemical properties of the antifouling biocide is discussed. Furthermore, ways to control the release of
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the biocides by manipulating the physicochemical properties of the microcapsules are investigated and the resulting antifouling properties of the coatings are presented. A special focus will be given to the experimental
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methodology for controlled release measurements and to the subsequent implementation of diffusion models to describe the release. This is important because there are many examples of questionable setups for release experiments and of very pragmatic empirical release models that may not give reliable results that can be treated
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in a scientific way.
In this section, the background to the problems associated with common antifouling formulations will be presented. These problems will be described in detail and the benefits of microencapsulation as a solution to the problems will be discussed. The following subsections will be devoted to the details of the encapsulation
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strategies implemented by us and others for use in antifouling coatings. The controlled release and the
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antifouling assessment of the release systems will be treated in Section 3. The typical practice when it comes to formulating biocides in coatings is to simply disperse them molecularly in the wet paint (see Scheme 1a)[4, 15, 16]. As mentioned in the Introduction, this approach is often not
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satisfactory because the antifouling function usually becomes quite short-lived. The biocidal action is over long before the desired life time of the coating, due to premature release or degradation of the active substance[16, 17]. Most biocides are small molecules and hence subject to fast self-diffusion in the coating matrix[11, 22, 36].
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Therefore, this simple approach requires the use of very high concentrations of the biocide in order to maintain the antifouling function for a long time (see Section 3 for a theoretical argumentation)[7]. The use of excessive amounts of biocide may pollute the environment, and it may also lead to technical problems such as macroscopic phase separation in the paint film if the biocide is incompatible with the binder-solvent system used[11]. This is valid for most biocides except for biocides which are present in the coating as pigments (inorganic crystals) such as copper (CuO or Cu2O). Pigment based biocides are released very slowly; however, the release profile is significantly influenced by the paint system. Yet, this is outside the scope of this review and the interested reader is referred to reviews which are solely focused on antifouling paints[15, 16].
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Scheme 1. An antifouling biocide can be introduced into a paint formulation by molecular dispersion (left) which is the most common approach, by immobilization to a large component in the formulation such as the binder (SPC paints) or a pigment (middle), or by encapsulation (right).
The issue of incompatibility can also prevent the use of different types of biocides in a paint formulation. Most fouling problems are multifaceted and involve a spectrum of different fouling organisms (see Fig. 1) with
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varying sensitivity to biocides[17, 62]. The use of very potent biocides with a broad efficacy towards fouling organisms will also affect non-target organisms[4, 16, 63, 64]. In the past, organometallic compounds were commonly used as antifoulants, with tributyl tin oxide, TBTO, being particularly popular. These biocides were
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very efficient in preventing fouling but, as it turned out, also very harmful to the local ecosystems. Such antifouling agents were banned by the governmental authorities starting in the late 1980’s and the beginning of
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the 2000’s and were phased out from the market during a relatively long transition period. As a response to the new regulations, considerable research efforts were made to implement and evaluate alternative antifoulants with
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better environmental profile. As a summary of the efforts made during this time period one may conclude that it seems very unlikely to find one single biocide that can replace TBTO and similar efficient compounds and at the same time be sufficiently benign to the environment. Instead, there is now a general consensus that an efficient and environmentally friendly antifouling coating requires a cocktail of complementary biocides with different
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efficacy profiles and with a rapid and efficient deactivation in the environment [17, 65, 66]. The biocides that are relevant for this review are presented in Chart 1 and are categorized according to their physicochemical properties. However, it is also common to categorize biocides according to their biocidal mechanism, efficacy profile, target organisms, etc. Triclosan and triclocarban, are antimicrobial compounds used as preservatives in personal care products. Also silver has antimicrobial effects and is presently used to make antimicrobial surfaces. Bronopol is used as a desinfectant in hospitals, but also in industry to avoid bacterial growth, e.g. in paper mills and cooling towers. Clotrimazole[67] is used to treat fungal infections in man and animals, while propiconazole is mainly used in agriculture for plant protection. IPBC is a fungicide used to protect painted surfaces from growth of fungi. The metals copper, zinc and silver, and the metalloid arsenic have broad-spectrum effects on micro-organisms, plants and animals, and have been used for wood preservation. Copper and zinc are also used as antifouling agents with effects on a variety of organisms. The hydrophobic isothiazolinone DCOIT is an antifouling compound effective against bacteria and algae and some animals. Irgarol and medetomidine are antifouling compounds with a narrow efficacy profile, targeting algal photosynthesis and the settling of barnacle larvae, respectively. DINCH is a new plasticizer often used together with antimicrobial agents.
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Chart 1. Biocides and dyes discussed in this review. The logarithm of the octanol-water partition coefficient (logKow values) and the water solubility are given for each substance, as obtained from Chemical databases[68-70].
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ACCEPTED MANUSCRIPT The problem of premature release has been approached by immobilization of the biocide on a more or less immobile object such as binders and pigments[30]. In more advanced paint systems, such as self-polishing marine coatings (SPC-paints)[4, 5], the biocide is attached to the binder by a covalent linkage (see Scheme 1) and the release rate is controlled by the rate of hydrolysis of the linkage. The release kinetics for such systems is
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of zeroth order or pseudo-zeroth order since the concentration of unhydrolyzed ester bonds in the paint and of
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water in the sea is constant. The resulting constant release rate is optimal with respect to environmental and economical aspects as discussed more in detail in Section 3. Note that the self-polishing hydrolysis itself is a
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mechanical antifouling mechanism. This strategy is, due to practical reasons (synthesis procedure, compatibility and solubility issues), only feasible for one or a few biocides per paint system and therefore only applicable to very potent and hence potentially environmentally hazardous biocides. The most well-known example is the selfpolishing coating with tributyl tin covalently attached, a SPC-TBT antifouling paint. This paint system, usually
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with added copper pigment, was very efficient and dominated the market before the ban as discussed above[66, 71-73]. To meet the new environmental regulations, a number of tin-free SPC paints have been developed and the organotin group is in these paints replaced by for example copper, silicon or zinc compounds [4, 5].
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Although these tin-free side groups possess some biocidal properties, the antifouling function is far from sufficiently efficient and booster biocides are therefore required. Note that these booster biocides are molecularly dispersed in the paint and hence subjected to fast release of non-zeroth order kinetics as mentioned earlier in this
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section and in the introduction. Moreover, the selection of suitable biocides is, at least in Europe, restricted due to the Biocidal Products Regulations which are controlled by the European Chemical Agency. Most used
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antifouling biocides are therefore conventional approved agrochemicals and efforts to introduce new antifouling 3.3.1).
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compounds are rare and limited to extraordinarily promising new biocides such as medetomidine (see Section In addition to covalent linkages, biocides may be immobilized by physical interactions[35-37, 74]. We have investigated immobilization; using coordination chemistry, of biocides containing a suitable ligand, with a special focus on the biocide medetomidine (see Chart 1), on common pigments (usually metal oxides
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nanoparticles) used in paints [30, 36]. However, this approach is also limited to a restricted number of biocides, which must contain a suitable ligand[30] and is outside the scope of this review. Microencapsulation can solve both the problem of premature release and the issue of compatibility when using several biocides in the formulation[17]. The release rate of the actives from the coating is determined by the slow release from the microcapsules to the surrounding coating matrix, and the encapsulated biocide is protected from degradation[11, 38, 39] (see Scheme 1 and Section 3.2). Moreover, compatibility is ensured and macroscopic phase separation prevented by the use of microcapsule shell materials that are compatible with the binder used in the formulation. Note that the only interface of importance between the paint and the formulated biocide is that between the microcapsule surface and the surrounding paint matrix. This allows for the use of carefully selected cocktails of biocides[65], containing biocides with high specificity[75-77] towards target organisms and low toxicity towards non-target organisms[75, 78-81], such as medetomidine (see Chart 1 and Section 3.3.2),
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ACCEPTED MANUSCRIPT 2.1
Microencapsulation
The term “microcapsules” covers a broad range of colloidal particles of different sizes, morphologies and chemical compositions (see Scheme 2). Sub-micron particles are typically referred to as nanoparticles. Likewise, the encapsulation is typically termed nanoencapsulation for nanocapsules whereas microencapsulation usually
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regardless of the size and morphology of the corresponding colloidal particle.
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refers to microcapsules. However, from here on, we will use the terms microcapsules and microencapsulation
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Scheme 2. Microcapsule morphologies; a) core-shell, b) microsphere (monolithic/matrix), c) multicore-shell and d) core-multishell
There are numerous methods available to encapsulate biocides, many of which have recently been reviewed by
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us[23]. The choice of encapsulation method depends on the intended release profile and on the physicochemical properties of the biocide (polarity, size, charge, surface activity etc.). This interrelationship is theoretically
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discussed in detail in Section 3 and experimental examples are provided in Section 3.1.2. As illustrated in Chart 1, antimicrobial actives cover a very broad spectrum of chemicals with different physicochemical properties. In
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addition to biocides, we have also investigated the encapsulation of a variety of dyes (see Chart 1). The use of dyes with different physicochemical properties is an effective way to study the controlled release with UVvis[27] or fluorescence spectroscopy. In addition, the distribution and the partition of molecules with given physicochemical properties, into the different compartments of the microcapsule, can be investigated using
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bright field or fluorescence microscopy techniques (see Fig. 2c-e and Chart 1)[28, 29]. We have mainly investigated coacervation-based methods including the internal phase separation which is suitable for encapsulation of hydrophobic actives, and multiple emulsion routes which are suitable for encapsulation of hydrophilic actives. For both methods the coacervation is induced by solvent evaporation from an emulsion (see Scheme 3), and the barrier material is typically polymeric.
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Scheme 3. Encapsulation process in an aqueous continuous medium via the internal phase separation route and the double emulsion route. Reprinted from[22] .
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These methods are popular for a number of reasons. The multitude of polymers available with different physicochemical properties allows for the tailoring of the release according to need, and any core material can in principle be encapsulated[22]. Various types of core and shell materials with different physicochemical properties investigated by us are presented in Chart 2.The experimental procedure is straight-forward and
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feasible for industrial scale up[22, 23] which is discussed in more detail in Section 2.2. The resulting microcapsules contain no by-products such as monomers or initiators which can be present in microcapsules prepared using polymerisation techniques[23]. Even though the two coacervation processes are methodologically similar (see Scheme 3), the interfacial processes involved in the encapsulation are phenomenologically very different (see Sections 2.1.1 and 2.1.2). The theoretical details of these coacervation methods have recently been reviewed by us[22] and we will in this contribution focus on the results obtained in our group including some very recent innovations.
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ACCEPTED MANUSCRIPT
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Chart 2. Polymers used for shells, oils/polymers used for cores and polymers and surfactants used as dispersants. Here, the polyesters (PCL, PLLA and PLGA) and the cellulosics (CA and CTA) belong to the class biopolymers. PS and PMMA are considered commodity plastics whereas PU is an engineering/high performance plastic. The advantage (+) and the drawbacks (-), unrelated to the physicochemical properties, regarding the choice of a specific polymer class is also provided. Solubility parameters (δ [MPa1/2) are given for the shell polymers[82] and the logarithm of the octanol-water partition coefficient (logKow values) and the water solubility are given for each substance, as obtained from Chemical databases[68-70].
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ACCEPTED MANUSCRIPT 2.1.1 Internal phase separation The internal phase separation method was developed by the Vincent group in Bristol in the late 1990’s [83] and has since then been implemented and expanded by a number of research groups[12, 25, 26, 40, 84-87]. Both the
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core and the shell material are dissolved in a water insoluble and volatile solvent or solvent mixture (see Scheme
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3). As the solvent evaporates, the core and the shell materials phase separate inside the emulsion droplet (see Scheme 3). Since we are considering encapsulation in aqueous dispersion, which is the most appropriate system
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for most applications, the core material including the biocide needs to be more hydrophobic than the shell material. However, this simple requirement is not sufficient for ensuring proper microencapsulation as discussed in detail in the following paragraphs. It should be noted that this method has some advantages over the double emulsion route. The shell of the resulting microcapsule is homogenous[83, 88] with an easily controllable shell
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thickness. Moreover, the particles are relatively monodisperse[89] with almost 100% encapsulated active substance[88, 90]. Even though the discussion so far concerns microcapsules in aqueous dispersion, it should be noted that it is possible to produce microcapsules with aqueous cores using the internal phase separation method.
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Atkin and Vincent developed a method based on W/O emulsion systems, or rather acetone/O emulsion systems, for the fabrication of polymeric core-shell particles with aqueous cores[91]. However, as mentioned above, most applications require the microcapsules to be dispersed in aqueous medium. Yet, this can be accomplished by
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extracting the microcapsules from the oil phase to an aqueous phase using a proper choice of surfactants[91]. The choice of core oil can sometimes be restricted. Some oils that are good solvents for the biocide will also dissolve the shell polymer and subsequently hinder the shell solidification. Examples of such solvents are the
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aromatic hydrocarbons mesitylene and toluene, which cannot be used in combination with hydrophobic and semihydrophobic shell polymers (see Chart 2). A way to circumvent this is to use polymers of similar chemical nature as core material. The resulting microcapsule is consequently consisting of both a polymeric core and a polymeric shell. The practically non-existing entropy of mixing two polymers ensures segregation of the
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polymeric core and the polymeric shell and therefore microcapsule formation. As an example, we have investigated the use of polystyrene cores as replacement for aromatic oils (see Fig. 2e[29]. It is however important to realize that the use of a solid instead of a liquid core will have a profound effect on the diffusivity of the biocide as discussed in Section 3.1.2.
The spreading conditions So far, only the most fundamental aspects of the internal phase separation have been considered. A key element with respect to the final particle morphology is the interfacial tensions between the core, the shell and the continuous medium and in particular their spreading coefficients (see Equation 1). We will in the following discussion, for convenience, consider an aqueous dispersion of core-shell particles. This theory, based on spreading coefficients, was developed by Torza and Mazon[92] and later expanded by Loxley and Vincent[83] to predict the morphology of the core-shell particles obtained.
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ACCEPTED MANUSCRIPT Core-shell formation requires that the shell-forming polymer (index p) wets the core material (index o) in the aqueous phase (index w), i.e. spreads between the core and the water (see Equation 1). The spreading coefficient[93] Sp is defined by the interfacial tensions γij between phases i and j as;
Sp Gpc Gpa ow pw op
1
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ΔGcp and ΔGap are the work of cohesion and adhesion, respectively. If Sp>0, the polymer will spread between the
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water and the oil. The spreading coefficients for the oil and the water phases may be derived in a similar manner. It may be assumed that γow>γop which leads to three possible sets of spreading conditions[83, 92];
So<0;
Sw<0;
So<0;
Sw>0;
Sp>0,
2
Sp<0,
3
Sp<0,
4
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Sw<0;
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So<0;
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The core-shell morphology can only be obtained if the conditions in Equation 2 are satisfied. Equation 3 will generate so called acorn particles [83] and Equation 4 will result in separate oil and polymer droplets. The formation of acorn particles is not necessarily an unsuccessful encapsulation. It is also a popular route to produce so called Janus particles[94]. However, this is outside the scope of this review and will not be further treated. If
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no core material, often termed non-volatile-non-solvent, is added during the encapsulation, a homogenous
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polymer particle or microsphere[90, 95] will form and the spreading conditions have no meaning. The different types of morphologies and outcomes from the encapsulation process are depicted in Scheme 4. Since the morphology is highly dependent on the interfacial tensions between the three phases, it is clear that the choice of
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dispersant will be of paramount importance for the end result. This is treated in Section 2.1.3.
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Scheme 4. Morphology of a two-phase droplet system as a function of spreading coefficients; a) engulfed core (core-shell particle), b) partially engulfed core (acorn particle) and c) droplet separation.
Predicting morphology using the van Oss formalism The correct way to predict the microcapsule morphology is to experimentally measure all interfacial tensions between the core phase, the shell polymer and the aqueous dispersant solution using optical[25, 26] or force tensiometry[25, 26, 83]. However, this experimental procedure is rather time-consuming and arduous if the intention is merely to find a suitable core shell pair for encapsulation. The interfacial tension between the solid polymer and the liquid phases is determined by measuring the contact angle in air or in a liquid medium and
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ACCEPTED MANUSCRIPT subsequently applying Young’s equation. This requires that the surface free energy of the polymer is known. If not, the surface free energy of the polymer has to be determined by measuring the contact angle of a set of wellcharacterized liquids on the polymer surface and subsequently applying theoretical approaches, such as the Fowkes[96] or the van Oss-Chaudhury-Good formalism (see below)[97, 98]. For microcapsules that consist of a
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polymeric core as well as a polymeric shell, measuring the interfacial tension between two polymers is difficult
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and is usually performed on polymer melts.
A more straight-forward route to predict the microcapsule morphology is to theoretically calculate the interfacial
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tensions using theoretical approaches such as those mentioned earlier. The interfacial energy is a result of intermolecular forces and can therefore be separated into additive contributions (ion-ion, ion-dipole, Keesom, Debey, London dispersion)[97] in a similar manner as for the bulk[99] (see Section 2.1.5). The most simple representation is the separation of the interfacial energy into a dispersive part γd and a polar part γp as proposed
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by Fowkes[96]. However, it has been demonstrated that the Debey and Keesom contributions to the polar term γp are very small[97, 98]. Moreover, the polar contribution is usually overestimated and neglects the more complicated but important contribution from H-bonding[97, 98]. One popular approach is the van Oss-
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Chaudhury-Good formalism (Equations 5-8), which takes asymmetrical interactions; such as hydrogen-bonding, into account [26, 97, 98]. The apolar γLW contribution is expressed as the total Lifshitz or van der Waals force and the polar term γAB accounts only for Lewis acid-base interaction. The polar term is expressed by an acid
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component γ+ and a base component γ- (see Equation 6)[97].
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i iLW iAB
5 6
iAB 2 i i
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The interfacial tension components can now be expressed by the geometric mean of the usually known surface tension components (Equations 7 and 8) and combined according to Equation 5 to give the total interfacial
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tension.
ijLW
LW i
LW j
ijAB 2 i j
i
2
j
7 8
Clearly, γijLW will always be positive. However, γijAB may attain negative values if i is a strong Lewis base and j is a strong Lewis acid or vice versa[26, 97]. The surface energy components; γLW, γ+ and γ- , can for most common polymers and solvents be found in the literature and the interfacial tensions are conveniently calculated from Equations 7 and 8. It is important to note that this simplistic approach does not take biocide or dispersant effects into account, which can be profound as discussed in Section 2.1.3. The calculated spreading coefficients based on the van Oss approach for the system water-PMMA-alkane (dodecane or hexadecane) are very similar to the experimentally measured counterparts which corroborates the validity of the approach (see Table 1). The more simple Fowkes approach does also predict the correct morphology; however, the values of the coefficients deviate significantly. Moreover, the effect of the dispersant can be significant and for very surface active ones, the deviation from the experimentally or modelled values is large. The acorn morphology is predicted for surfactant-stabilized systems (such as the SDS-based system in Table 1). However, the PVA-stabilized system displays core-shell morphology although acorn morphology is predicted. It should be noted that PVA-stabilized PMMA-based microcapsules occasionally 15
ACCEPTED MANUSCRIPT exhibit an uneven surface with protrusion and indentations which might be an indication of the partially fulfilled wetting conditions. Moreover, certain conditions may result in multi- rather than single core-shell morphology. Regarding the TC4-PDADMAC system, the multiple core droplets are stabilized by the surfactant TC4. The sorbitan monooleate system is a W/O emulsion and the fast evaporation of the solvent acetone freezes the initial
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multicore morphology (see Scheme 3) in the solidified PMMA matrix before the coalescence of the water
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droplets is completed. Note that the prediction does not distinguish between single core-shell and multicore-shell morphology, although attempts have been made to correlate this with the magnitude of Sp[100]. Overall, the
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predicted morphologies agree well with the observed ones in Table 1. Moreover, the theoretical van Oss approach will provide an insight into whether core-shell formation is at all conceivable for a given core-shell pair and has been used by us to predict the morphology of microcapsules with polystyrene cores[29] (see Fig. 2e).
So
Sw
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Table 1. Calculated spreading coefficients for the system water-PMMA-alkane (dodecane or hexadecane) containing (or not containing) different types of dispersants. The pure systems (not containing dispersants) in the table include experimentally measured values as well as theoretically predicted ones according to Equations 5-8. All dispersant-based systems are prepared from O/W emulsions where the dispersant is dissolved in the aqueous phase. The exceptions are TC4-PDADMAC and sorbitan monooleate. Regarding TC4-PDADMAC, the dispersant is residing in the PMMA phase (TC4) and is dissolved in water (PDADMAC) respectively. The sorbitan monooleate system is prepared from a W/O emulsion where the dispersant is dissolved in the alkane. For this system, the spreading coefficients have been estimated from interfacial tensions obtained from the literature.
Sp
Morphology
Predicted
-43.5
Van Oss[29]
-34.1
Fowkes[29]
-23.8
-60.5
26.5
Core-shell
-68.2
31.0
Core-shell
-78.8
16.6
Core-shell
Dispersants SDS[83]
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Experimental[25, 26]
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Pure system
Observed
-9.80
-3.60
-19.4
Acorn
Acorn
-17.4
-25.8
-11.8
Acorn
Core-shell
-31.1
-39.7
1.90
Core-shell
Core-shell
600-4600[25]
-45.4
-29.3
28.4
Core-shell
Core-shell
TC4-PDADMAC[26]
-59.7
-43.7
50.8
Core-shell
Multicore-shell
Sorbitan monooleate[91]
-6.96
-37.8
3.84
Core-shell
Multicore-shell
PVA[83]
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PMAA[25, 83]
2.1.2 Double emulsion The double emulsion route may, in contrast to the internal phase separation route, produce particles with aqueous cores directly in an aqueous suspension (see Scheme 3). The most common type of double emulsion for encapsulation applications is W1/O/W2[22, 101-104], a system that has been extensively reviewed by Garti and coworkers[102, 104-108]. The most explored procedure to produce double emulsions is a two-step homogenization method[109] which shares many similarities with the internal phase separation method (see Scheme 3)[28, 102]. However, the corresponding pre-emulsion is in this case a W/O emulsion formed under high shear[28, 101, 102]. The aqueous phase contains the active substance and the oil phase contains the shell forming polymer and a volatile solvent. The prepared W1/O pre-emulsion is subsequently dispersed in the 16
ACCEPTED MANUSCRIPT continuous W2 phase under moderate shearing and slow evaporation of the solvent in order to solidify the oil phase that forms the shell. This procedure usually results in quite large, polydisperse and multicore-shell particles (see Scheme 2c and Fig. 2d)[28, 101]. However if multicore-shell particles suffice, it is preferable to attain as small inner droplets as possible in order to slow down the coalescence kinetics[101]. The water droplets
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of microemulsions are therefore suitable as inner phases. The microcapsule-forming dispersed phase may also
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consist of a dispersed mesophase, such as cubosomes (dispersed cubic liquid crystals), swelled liposomes (dispersed lamellar phases), etc.[101, 102]. Another way to reduce the kinetics of the segregation of the inner
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droplets is to add thickeners in order to increase the viscosity of the inner phase (see Chart 2 and Fig. 2d)[28]. Modern techniques such as membrane and in particular microfluidic devices[110] enable the formation of monodispersed droplets with controllable inner morphology[111]. Yet, the particles are still relatively large and the shell is occasionally inhomogeneous. For a more detailed investigation of the properties, technical aspects
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and recent developments of microcapsules prepared using microfluidics, the interested reader is referred to a recent review by Datta [112]. However, in contrast to the two-step homogenization method, the microfluidics
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method is as of yet unsuitable for mass production of microcapsules.
2.1.3 Choice of dispersant
The main function of the dispersant is to facilitate the emulsification and to provide colloidal stability of the
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core-shell particle suspension. Polymeric stabilizers are the primary choice for core-shell particle production
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using the internal phase separation or the double emulsion route as will be explained below. The polymer dispersant can provide colloidal stability through steric, electrostatic or depletion stabilization[113]. In many
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cases (PVA or PMAA stabilization), the latter mechanism is predominant and a large excess of dispersants is required to prevent irreversible coalescence[40, 114]. The adsorbed dispersant layer is usually very thin in comparison with the core-shell particle (see Fig. 3) but may nonetheless be viewed as an additional outer shell. This implies that the choice of dispersant type should have a
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large impact on the release rate which is indeed the case[113] as discussed in Section 3.1.2.
Fig. 3. Adsorption isotherms given by the thickness of the adsorbed layer δ as a function of the concentration c of the dispersants PVA (▲), PMAA (▼) and 600-4600 (■) on PMMA surfaces as measured using QCM-D[27].
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ACCEPTED MANUSCRIPT Internal phase separation Normally, the use of surfactants as dispersants during microencapsulation using the internal phase separation method prevents the formation of core-shell particles[22]. From Equations 1-4 it is clear that the interfacial tension γow must remain high for polymer spreading and subsequent core-shell particle formation to occur. Small
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surfactants will significantly reduce γow and this is the reason why the use of surfactants as dispersants prevents
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core-shell particle formation. As an example, Loxley and Vincent studied the effect of using the water-soluble surfactants cetyltrimethylammonium bromide (CTAB) and sodium dodecyl sulfate (SDS) as dispersants which
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resulted in partially engulfed oil cores termed acorn particles[83]. However, contrary to the current knowledge, we have observed that very hydrophobic surfactants can be used for the purpose.
The oil-soluble surfactant TC4
(see Chart 2) enabled core-shell formation when used in combination with the polycation rather than single core-shell (see Fig. 2b)[25, 26].
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poly(diallyldimethylammonium chloride) (PDADMAC)[26]. Yet, the observed morphology was multicore-shell As previously mentioned, the appropriate choice of dispersants are polymeric stabilizers[40, 83]. The ideal surface activity of the polymeric dispersant is a compromise between low surface pressure and stabilization of
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the colloidal surface[40]. We have investigated novel types of ionic dispersant which provided PMMA-based core-shell particles with strong electrostatic stabilization, avoiding the use of excess dispersant in the bulk. These included poly(methacrylic acid) (PMAA) treated with base after adsorption[114], the hydrophobic anionic
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surfactant TC4 in combination with PDADMAC[25, 26] and charged amphiphilic block copolymers of PMMA-
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b-PMANa type (see Chart 2)[40]. In particular the latter dispersant type provided the core-shell particle suspensions with long-term stability by a combination of electrostatic and steric stabilization[25, 40]. This was achieved by using a block copolymer with a hydrophobic block of the same monomer composition as the shell-
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forming polymer[40]. We have shown that the hydrophobic block is dissolved in the polymer phase during the emulsification and becomes amalgamated with the shell as the volatile solvent evaporates[40].
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Double emulsion
In contrast to the internal phase separation route, the core-shell formation from double emulsions is not relying on the spreading of a polymer coacervate per se. This broadens the choice of combinations of shell forming polymers, dispersants and volatile solvents to be used. Nevertheless, double emulsions have a limited shelf life , even more so than regular emulsions[102]. Therefore, successful core-shell formation requires proper stabilization of the W1/O and O/W2 interfaces. Typically, two sets of dispersants are needed; an oil soluble one to stabilize the W1/O interface and a water soluble one to stabilize the O/W 2 interface. Once again, polymeric stabilizers are more suitable than small surfactants[28, 102] but for different reasons compared with the internal phase separation case. Small surfactants have a tendency to migrate from one interface to the other while amphiphilic polymers (graft or block copolymers) tend to adsorb more “irreversibly” at the interface. Therefore, ethylene oxide-propylene oxide block copolymers of Pluronic type are common choices. A very promising type of block copolymers for these purposes are diblock polymers with a large hydrophobic block of the same type as the shell forming polymer and a small hydrophilic polyethylene oxide block (see Chart 2). For example, the inner gelatinous water droplets of the PS based microcapsules in Fig. 2d are stabilised by the P(S-b-EO) blockcopolymer shown in Chart 2. The hydrophobic block will amalgamate with the shell during the evaporation stage
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ACCEPTED MANUSCRIPT as described above[40] and the poly(ethylene oxide) block allows for solvation in organic solvents in contrast to hydrophilic polyelectrolyte blocks[28]. In addition, the polymeric film at the interface prevents coalescence of droplets and migration of water from W 1 to W2.
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2.1.4 Post-encapsulation modifications
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The core-shell particles can, after the encapsulation process, be modified in order to tailor their release
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behaviour. There are many techniques available for post-encapsulation modifications, and in this section we will consider ways to affect the microporosity of shell. We will also discuss surface modifications that are able to introduce additional barriers (shells) on the microcapsule.
The microporosity (or free volume) of the shell can be reduced by annealing or cross-linking[22]. The Vincent
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group investigated ways to cross-link PS and poly(2-vinylpyridine) based microcapsules by simply mixing monomers and initiators in the oil phase[115]. The shell of the microcapsule was cross-linked after the encapsulation by exposing the particle suspension to UV-radiation or heat. The alternate method “annealing” is
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perhaps a more convenient route to reduce the microporosity. Annealing implies relaxing the polymer chains from a kinetically arrested state to a more thermodynamically stable state. It has been shown by the Vincent group, and our group [38, 88], that the encapsulation procedure can result in microcapsules with an expanded shell structure. However, this can be counteracted by heating the microcapsule suspension above the glass
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transition temperature of the shell-forming polymer followed by slow cooling.
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Regarding the surface assembly of additional barriers (shells) on the microcapsule surface, we have investigated the adsorption of polyelectrolyte multilayers (PEMs)[27, 39, 116], as well as of lipid bilayers[116]. The PEMs
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are assembled using the well-known Layer-by-Layer (LbL) technique that for colloidal systems, was developed by the Möhwald group in the late 1990’s[117-119]. The lipid bilayer is assembled by spreading charged liposomes on a microcapsule surface of opposite charge[120, 121]. Both surface modifications require a stable and highly charged surface. The surface charges can be introduced by using an intrinsically charged colloid[25,
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40, 114, 122] or by chemical means[122]. We have investigated the use of an amphiphilic ionic dispersant in order to introduce a high and stable surface charge density as described in Section 2.1.3. The PEM and the lipid bilayer represent two very different barriers in terms of physicochemical properties. The PEM is hydrophilic, highly charged and relatively swollen with solvent. On the other hand, the lipid bilayer is very hydrophobic and; for saturated symmetrical hydrocarbon chains longer than ca 14-16 carbons, also highly dense and crystalline[22, 23].
2.1.5 Solvent choice and environmentally friendly approaches The properties of the volatile solvent used in solvent evaporation-based methods and in particular the internal phase separation method must fulfil certain requirements, which has been reviewed by us in detail elsewhere[12, 22]. Perhaps, the most obvious requirement is that the solvent must dissolve the shell forming polymer and the core. The prediction of solubility of a polymer in a solvent can be done using the Hildebrand’s solubility parameter concept. The Hildebrand solubility parameter δ is based on the cohesive energy density E/V which employs the semi-empirical theorem, like dissolves like (Equation 9-11)[123, 124]. The solubility parameter is
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ACCEPTED MANUSCRIPT related to the interaction parameter χ according to Equation 10 where V0 is the molar volume (assumed to be the same for both components).
EV
9
ij V0 RT ij
10
ijH i j
11
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2
Clearly, substances with similar cohesive energies, i.e. similar intermolecular interactions strengths, will be
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mutually soluble as χ→0. This is valid for non-polar systems but does not account for asymmetric interactions in strongly polar or hydrogen-bonding systems. Here, the extended Hildebrand approach may be used (Equation 12). The solubility parameter is separated into additive components of intermolecular forces (Equation 13) in a similar fashion as for the surface tension (see Section 2.1.1); e.g. apolar, Lewis acid and Lewis base components
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(with indexes “LW”, “+” and “-”)[125, 126]. Moreover, the extended Hildebrand approach allows for negative values of χ. Note the similarity of Equations 12 and 13 with the van Oss approach.
2 2
ijEH iLW jLW
LW 2 i
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i2
2
i
j
i
j
i i
12 13
The most common solvents used as volatile solvents are chlorinated compounds such as dichloromethane or
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chloroform. These chlorinated compounds can dissolve most hydrophobic and semihydrophobic polymers and
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possess some Lewis acid properties[97]. For more hydrophilic polymers such as cellulose acetate (see Chart 2), acetone can be added to the oil phase to increase its polarity and add some additional Lewis base properties[28].
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Acetone is also the solvent of choice regarding encapsulation of aqueous cores using the internal phase separation method[22, 91]. However, for O/W emulsions it is important to note that acetone will be distributed between the water and the oil phase and therefore evaporate faster than the more hydrophobic solvent. This will affect the point of internal separation and shell solidification.
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For the implementation of solvent evaporation based-methods on an industrial scale the environmental and toxicity aspects of the solvent must also be taken into account. The use of chlorinated solvents in the industry is highly restricted today[12]. Replacement of the chlorinated solvents with more environmentally friendly ones is not easy since they, in addition to the above mentioned property requirements, must be cheap and/or recyclable for industrial large-scale production[127]. However, we have found that ethyl acetate fulfils these requirements[31]. We and others have subsequently used ethyl acetate to prepare microcapsules using the internal phase separation method[11, 12, 31], as well as the double emulsion route[128].
2.2
Microcapsules in paints
The incorporation of microcapsules into a paint system is simple and requires a minor workload and equipment. The microcapsules are formulated into wet paint as an additive, which mean that no specific preceding formulation of the paint is needed. Hence, the addition of microcapsules can be performed at any time prior to the application of the coating and, moreover, commercially available paints can be used. Our standard procedure of microcapsule addition involves two steps.
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ACCEPTED MANUSCRIPT First, water should be removed from the aqueous microcapsule suspension. Section 2.1 describes the formulation of microcapsules and the final suspension contains a microcapsule-to-water ratio of approximately 1:20. Before the addition of the microcapsules to the wet paint, the water content of the microcapsule suspension needs to be significantly decreased, using for instance centrifugation or ultrafiltration. A large amount of the continuous
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solvent phase may cause undesirable effects in the paint and must in some cases be exchanged with a solvent
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which is more compatible with the paint solvent. Completely dry microcapsules should be avoided as the microcapsule agglomeration in the dry state can be difficult to break by mere stirring of a highly viscous paint
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system. However, the agglomeration is strongly dependent on the type of dispersant used in the microcapsule suspension. The use of conventional dispersants such as PVA and PMAA results in irreversible aggregation if the excess dispersant in the water phase is removed. However, microcapsule suspensions prepared using the highly charged dispersants described in Section 2.1.3 are fully dispersible also from a completely dried state
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when all excess dispersant has been removed. For the microcapsule suspension stabilized by PVA or PMAA, a microcapsule-to-water ratio of approximately 1:1 is convenient since it produces a manageable fluid material, which will allow for completely homogenous dispersion of the microcapsules in the wet paint system. Water
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removal from the suspension is, from our experience, a critical stage in the scale-up for industrial production. The microcapsule-to-water ratio should, at industrial quantities in a relatively fast step, increase from approximately 1:20 to 1:1 but where the latter ratio is important not to stretch further to avoid agglomeration.
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Moreover, recycling of the removed aqueous containing dispersant is most probably preferred.
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In the second step, the concentrated microcapsules suspension is added to the wet paint. If a waterborne paint is used, the formulation only requires a simple, but thorough, mechanical stirring of the paint system after addition
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of the manageable fluid material. In the case of solvent-borne paint, the situation is different. The water content of the concentrated suspension might need to be exchanged to a solvent that is more compatible with the solvent of the paint. Moreover, attention should be made with regard to the paint solvent and its influence on the microcapsule material. The various microcapsules presented in this article are all suitable for at least water-borne
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paints and varnishes. Fig. 4 displays electron micrographs of the area inside dried paint matrices revealed by scratching the coating with a thin needle [11]. It is clear that the microcapsules are stable and properly dispersed in both solvent-borne (xylene-based paint in Fig. 4a and b) and latex paint systems (Fig. 4c and d). Further information regarding the application of coatings for laboratory studies is presented in Section 3.2.1.
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Fig. 4. SEM images of solvent-borne (a and b) and water-borne (c and d) paint matrixes containing well-dispersed microcapsules (c and d)[11]. The microcapsules are homogeneously embedded in the coating and their geometry is unaffected by the mechanical stirring during the addition of manageable fluid material of microcapsules to the paint, the drying of the applied paint, and by the presence of the surrounding binder and pigment material. Reprinted from [11]. The microcapsules have been coloured brown and the paint matrix yellow for clarification.
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Controlled release
In this section, the theoretical fundamentals for controlled release will be described and the most important parameters, as visualized in Scheme 5, will be introduced. Focus will be on sustained release which is the dominant release mechanism in coating applications. Although we have previously briefly considered triggered
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release[27, 30, 129, 130], the interested reader is referred to a set of excellent reviews regarding the topics
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targeted or triggered release[131-133].Moreover, the economic and environmental benefits of using ratedetermining release systems in coatings for antifouling purposes will be briefly discussed. The following
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subsections will separately review controlled release from core-shell particles and coatings, respectively. In each subsection, the experimental methodology of the release measurements will be illustrated in detail. The release will be further examined in terms of the physicochemical properties of the core-shell particles and the coating matrix. Finally, the antifouling properties of some functional coatings towards different biofoulers will be
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presented.
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Scheme 5. The figure displays the important parameters for a microencapsulated substance which determine the release rate. These include dimensional and density parameters; the concentration of the active i in the core cCi, of microcapsules in the coating cC which gives the total concentration of active cCi ·cC=ci, the thickness of the coating δP, the radius of the core rc and of the microcapsule rm which gives the thickness shell δS. The kinetic parameters include; the diffusion coefficient in the core DC, in the shell DS and the diffusion coefficient in the paint DP. The concentration gradient of the active is also determined by the core/shell KC/S, shell/coating KS/P and the coating/water KP/W distribution coefficients.
The sustained release of an active from a microcapsule in a coating is controlled by the permeation through the microcapsule barrier (the shell) and the coating matrix. The permeation of the active in the microcapsule barrier and the coating matrix depends on both kinetic and thermodynamic parameters through its diffusion coefficient and its solubility in the barriers. The solubility of the active in, and its interaction with, the microcapsule core and shell materials and the coating matrix will determine its partition between these phases[22, 134-136]. This is described by the distribution coefficients, KiA/B, of the active i between phases A and B (Equation 14, where ciA and ciB are the equilibrium concentrations of i in phase A and phase B, respectively) and we will consider this thermodynamic constant first.
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ACCEPTED MANUSCRIPT i K A/B
cAi 1 i cBi K B/A
14
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The distribution coefficient can be expressed by the saturation concentration in phase A and the Flory-Huggins interaction parameter between i and “B” (Equation 15) following some minor derivation and implementation of
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i ln K A/B ln cAs ,i Bi
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the Flory-Huggins solution theory[123, 124] as described more in detail elsewhere[22].
A higher saturation concentration of i in A and unfavourable B-i interactions (higher χBi) will favour distribution of i in phase A and vice versa, which is a logical and important realization. The Flory-Huggins interaction parameter can be obtained using the Hildebrand interaction parameter as described in Section 2.1.5 (Equations 9-
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13).
The diffusion coefficient on the other hand is a kinetic parameter and is related to the size r of the active (D0=10/r m2/s, r is expressed in Å). For controlled release applications, it is important to make a distinction between
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the self-diffusion of the active, D0 and the effective or apparent diffusion D. The notion of an effective diffusion constant is strongly associated with the concept of hindered diffusion[135-138]. As the name suggests, hindered
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diffusion implies that the diffusion constant will be effectively reduced by a number of parameters as will be
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explained below (Equation 16). Impermeable objects, such as spherulites in semicrystalline polymers or inorganic pigment particles in a coating matrix, will obstruct the diffusion. The obstruction is described by the porosity ε (the cross-sectional area available for diffusion) and the tortuosity τ (the diffusive trajectory relative
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the end-to-end distance in the diffusive direction)[135, 136]. We have chosen to designate this porosity as a macroporosity since the relevant pores sizes of these macropores are significantly larger than the molecular
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dimensions of the actives[38].
V D D0 exp c K B V f
16
On a molecular scale, the active is diffusing in the void, or free volume, of the surrounding matrix, which for most applications consists of polymer chains. The free volume of the matrix Vf, (Equation 17) must be larger than a critical volume for diffusion Vc which is determined by the size of the active. The parameter γ is an overlap factor (1 for most polymers), V is the specific volume of the matrix, V0 is the impermeable volume occupied by the polymer and Vw is the van der Waals volume [38, 139]. This microporosity can also be expressed in terms of a thermal energy of the matrix and a diffusional energy barrier instead of volume-based parameters. The outcome of the expression will still be the same[139]. V f V V0
17
V0 1.3Vw
Interactions between the shell material and the active may not only affect the partition coefficient. Strong and specific interactions; such as coordinate interactions[22, 30] or hydrogen bonds[22, 140], characterized by an equilibrium constant for binding, KeqB, will immobilize a fraction of the active on the shell material[22, 136, 137,
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ACCEPTED MANUSCRIPT 140]. A shell material with a concentration of cB binding sites will result in an immobilized fraction of actives and only the “free” fraction KB will be available for diffusion according to Equations 16 and 18[136, 137].
K B (1 K Beq cB ) 1
18
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The relationship between these parameters and the release rate is most conveniently explained by assuming
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release under steady state conditions. The release rate ṁ from the microcapsules, which under steady state conditions is equal to the release rate from the coating, is a function of the concentration ci of the active i in the
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core and the total resistance rTOT over the barriers (Equation 19, see Scheme 5). The total resistance (Equation 20) is the sum of the barrier resistances, which depends on the barrier thickness δ, the cross sectional area of the barrier in the direction of the diffusion A (which is very different for the microcapsules and the coating), the effective diffusion coefficient D and the distribution coefficient K between the phases. The permeation P is
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subsequently defined as the product of the diffusion coefficient and the distribution coefficient. It is clear that given the high specific surface area of the microcapsules as compared with the surface area of the coating, the controlled release from a paint system requires rigorous control of the permeation in the core and the shell
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phases of the microcapsule. For non-steady state situations, which are the usual cases, the release must be described by diffusion models. These will be discussed in the following subsections
D
m
ci rTOT
rTOT rx Dxi , K xi , x , Ax x
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x
19
x Ax Dxi K xi
x
x Ax Pxi
20
The importance of controlled release in terms of environmental and economic impact can be explained by a
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simple illustrative example in Fig. 5. Let us consider an antifouling coating containing m∞ amount of biocide. The biocide needs to reach a critical concentration cc, often termed minimum inhibition concentration (MIC) or effective concentration (EC), in a defined volume adjacent to the surface in order to prevent fouling of an arbitrary biofouler. The release of the free biocide from the coating is assumed to obey first order kinetics with
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the rate constant kin which is a function of mtot and the permeability in the coating. The removal of the biocide due to diffusion, degradation, absorption by the biofouler, etc is also assumed to be a first order process with the rate constant kout. It is clear that the majority of the biocide will be wasted during the initial stages of the release. The most efficient release would follow zero order kinetics such that the c(t)=cc over the entire release interval. However, true zero order release kinetic is extremely difficult to achieve[23] and is seldom realized with microencapsulated biocides. Yet, an efficient reduction of the rate constant will still significantly extend the period of efficiency. For example, a reduction of the rate constant kin to 0.1kin for a given mtot and cc=0.01m∞, will prolong this period five times.
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Release from core-shell particles
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3.1
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Fig. 5. Relationship between the surface flux from the coating ṁ(t)= ṁ(t)in and the surface concentration c(t). The solid curves display the surface concentrations c for first order release (black) and for zero-order release (gray). The dashed curves display the corresponding surface flux from the coating. The blue square grid represents the integral of the surface flux and hence the total amount mtot of the biocide. The yellow line grid between the dashed curves for first and zero order surface flux represents the amount of wasted biocide.
This section will first describe the basics for a proper experimental release methodology regarding the release of
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actives (e.g. biocides) from microcapsules to an external aqueous medium. Suitable diffusion models are also discussed as well as their correctness when it comes to model the release in a physically sound way. Thereafter,
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the release results of some illustrative examples will be discussed in terms of the physicochemical properties of the different compartments in the microcapsule: the core, the shell and the outer dispersant layer. In addition, the surface assembly of PEMs on the microcapsule surface (see Section 2.1.4) is presented as a very promising tool for controlling the release of hydrophobic actives. Even though the intended release application usually concerns
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coatings, analysis of the release from a simple microcapsule suspension allows for the evaluation of structurefunction relationship with respect to the core and shell material and to the biocide. Yet, such an assessment requires complete knowledge of the size distribution of the microcapsules as explained below.
3.1.1 Release methodology and diffusion models Controlled release can be evaluated using several different approaches. Regardless of the chosen method, the methodology contains at least three main steps: setup, analysis, and evaluation. The procedures are schematically shown in Scheme 6a. The setup encompasses all laboratory equipment and material that allow for time-dependent release from the microcapsule to some arbitrary external release medium. The analysis involves the procedure for data sampling and determination of the concentration of the active at a given time using a proper analytical instrument. The evaluation is the post-processing work where the release data is put into perspective, e.g. by the use and implementation of a release model. The setup requires knowledge and proper control of the saturation concentration of the active in the release medium as well as its distribution between the microcapsule phase and the release medium. After the microcapsules are formed in the aqueous suspension, as described in Section 2.1, a fraction of the biocide is 26
ACCEPTED MANUSCRIPT released until equilibrium between the concentration of biocide in the microcapsule and in the aqueous phase is reached. This distribution is determined by the distribution coefficient in Equation 14. The magnitude of the distribution coefficients can be estimated by the octanol/water partition coefficients in Chart 1. For hydrophilic actives, prepared using the double emulsion route, the distribution coefficient is close to 1. However, for
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hydrophobic actives, the equilibrium is strongly shifted towards the microcapsules and the water phase is quickly
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saturated with the active. Note that the saturation concentration for hydrophobic actives in water is very low (see Chart 1). Failure to realize the importance of the saturation concentration and the distribution coefficients has led
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to the use of some misconceptions in the literature. The term “yield” is occasionally used to describe the observation that the release of hydrophobic actives reaches a plateau region below 100% released amount of active. However, this “yield” is merely an artefact of the experimental setup where the active reaches the equilibrium concentration in the aqueous medium. If additional water is added, i.e. the microcapsule suspension
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is diluted, further release of the active will occur until equilibrium is once again attained according to the value of the distribution coefficient.
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Scheme 6. a) Experimental methodology for the measurements of the release of an encapsulated active substance from microcapsule in an aqueous microcapsule suspension. The microcapsule suspension is poured into an aqueous solution (occasionally containing solubilizers) and the release of biocide out from the capsules to the surrounding medium will start immediately. Samples are collected over time using a syringe and subsequently pressed through a suitable filter in order to remove the microcapsules, other particles and dust. The concentration is quantitatively analysed by e.g. spectrophotometry. b) The use of semi-permeable dialysis membranes is a common setup for release measurements. The microcapsule suspension is put in a tube where the dialysis membrane has a pore size significantly smaller than the size of the microcapsules. Samples are taken from the surrounding medium, often an aqueous solution, and the concentration of released active substance is analysed. However, drawbacks have been noticed, for instance that the microcapsules aggregate over time due to improper mixing, which will change the release behaviour.
27
ACCEPTED MANUSCRIPT In order to achieve complete, or almost complete, release of the active from the microcapsule, a significant amount of water is needed, especially for hydrophobic actives. In practice, our release methodology must always fulfil two criteria; 1), the total amount of the active in the system should be less than 10 % of the corresponding saturation concentration in the release medium. Under this condition, the flux of the active into the microcapsule
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can be neglected. 2) At the equilibrium concentration of the active substance, which is determined by the
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distribution coefficient and the extent of dilution, at least 90 % of the active should be released in the release medium. This allows for obtaining a well-resolved set of experimental data over the entire release interval.
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For hydrophobic biocides, the required dilution is very large and will result in release concentrations which are too low to be accurately quantified using analytical techniques such as spectrophotometric and chromatographic methods. To be able to accomplish a measureable release profile, one must therefore decrease the value of the distribution coefficient. A direct route to alter this coefficient is by the addition of a co-solvent to the aqueous
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release medium. However, co-solvents may generate swelling of the microcapsule shell polymer[141]. A noninvasive, from the microcapsule point of view, alternative is the addition of solubilizers, such as the surfactant sodium dodecyl sulfate (SDS) or cetyltrimethylammonium bromide (CTAB) (see Chart 2), to the release
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medium. Above the critical micelle concentration (CMC), the active is solubilized inside the surfactant micelles, which significantly enhances its solubility[142-145]. Moreover, surfactants are relatively benign with respect to the integrity of the microcapsule in contrast to more hydrophobic co-solvents. In the study of charged
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microcapsules, e.g. microcapsules modified with polyelectrolyte multilayers (see Section 2.1.4), nonionic
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surfactants, such as poly(ethylene glycol)monododeyl ether (Brij® L23), are the proper choice (see Chart 2)[20]. The analysis in terms of data sampling procedure requires a short comment. Since the release from the core-shell
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particles is evaluated in their suspension, the active typically needs to be separated from the particles. A common setup found in the scientific literature is the use of semi-permeable dialysis tubes with pore sizes slightly smaller than the mean size of the microcapsules (see Scheme 6b). However, the dialysis method suffers from some severe drawbacks. Analyses in our laboratory have shown that the equilibrium of the active between
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inner and outer release medium, with respect to dialysis membrane, is not necessarily established during the time interval between two data samplings. Moreover, it is very difficult to obtain proper mixing inside the dialysis tubes. As a consequence, flocculation and aggregation of the microcapsules readily occurs, which strongly affects the release behaviour (See Scheme 6b). In view of the negative aspects using the dialysis tube setup, we have chosen a different approach, which is schematically drawn in Scheme 6a and which in principle can be applied to any system. The release is instigated by diluting the microcapsule suspension according to the two criteria discussed above. For the data sampling, a small sample of the diluted suspension is filtered using a syringe filter to separate the particles from the released active. However, the choice of filter material is of utmost importance since many filters will absorb a significant amount of active. Especially tricky actives are semi-hydrophobic molecules with some surface activity. Such an example is the fungicide OIT[38] (see Chart 1). Only one filter type out of five was sufficiently non-adsorbing for this active substance (see Table 2). If this issue is over-looked, the measured release will be much slower than the true one.
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Polyvinylidene fluoride
98.2 ± 1.1
Polyethersulfone
91.7 ± 1.0
Polyvinylidene fluoride (prefilter: Polyethersulfone)
90.8 ± 0.8
Amphoteric Nylon 6,6
89.2 ± 0.8
Surfactant-free cellulose acetate
50.1 ± 1.3
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Permeability (%)
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Membrane material
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Table 2. The membrane material in different filters evaluated by the research group with respect to the biocide OIT (aq) to find a suitable experimental setup.
Diffusion models
The quantitative analysis of the time-dependent biocide release presented above is tremendously useful.
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Experimental data of high accuracy allows for model evaluation of physicochemical properties that are decisive for the release. Mathematical regression analysis and fitting of experimental data can generate several parameters, out of which the most important is the diffusion coefficient of the active substance within the
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microparticle. Most often it is the apparent, or effective, diffusion coefficient that is calculated (see Section 3) and this parameter permits relevant comparison between different particles, as the diffusion coefficient itself is independent of both size and time.
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There are several extensive reviews concerning modelling of the release of actives from polymeric devices. Most
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of these reviews focus on pharmaceutical applications[146-149] where controlled release is a key concept. Here we often find simplified and empirical approaches to model the release. Many of these models, which are principally based on the work by Weibull, Higuchi and Peppas[150-153], deal with dispersed actives in
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polymeric release reservoirs. The benefit of such models is their simplicity and the possibility to compare various drug devices or release compartments. However, only few physical parameters are included and the models are often valid only for the initial part of the release. Still, in pharmaceutical sciences such models offer a great deal of information considering rather macroscopic release devices where the initial release and initial
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increase of concentration are the important parameters. For sustained release, on the other hand, the long-term release behaviour is crucial. One main goal with sustained release is to prolong the surface flux of the active above a certain critical level[23, 39, 134]. Hence, a proper description also for the extended final part of the release is crucial for improved understanding and prediction. All types of materials used in the release systems described in this review are stationary material. In other words, the materials constituting the core and the polymeric shell are stable and not undergoing erosion or swelling. We therefore have a diffusivity that is constant, with regard to time, in each defined material domain of the microcapsule. The release of active agents from the microcapsules, described in this work, can thus be explained by Fickian diffusion. Regarding the modelling of the experimental release data, it is convenient to express the analysed concentration as a fractional part of released active. The time-dependent released amount m(t) is divided by the total amount of active mtot initially positioned inside the microcapsule. Assuming perfect sink conditions, the complete timedependent release from a microsphere (the most geometrically simple release system, see Scheme 2b) with 29
ACCEPTED MANUSCRIPT homogeneous initial distribution of active substance, is expressed by the following equation derived by Crank[154]
D 2 n 2 m(t ) 6 f ( D, r , t ) 1 2 2 exp t 2 mtot n 1 n r
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where D is the apparent diffusion coefficient and r the microsphere radius. In a laboratory study, perfect sink
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conditions are practically never fulfilled and for the long-term part of the release, the deviation from Equation 21 can be significant. The equilibrium distribution, expressed by the distribution constant K, between microsphere and release medium with volumes Vsphere and Vsink, respectively, should therefore be taken into account providing
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a more realistic description using[154]
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Dqn2 m(t ) 1 f ( D, r , t ,Vsink ,Vsphere , K , t ) 1 6 1 exp 2 t 2 mtot 1 n 1 9 9 q n r
,
Vsink VsphereK
22b
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D
22a
.
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and where qn is the nth positive root of
tan qn
3qn 3 qn2
22c
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For the initial part of the release, Equations 21 and 22 coincide but the deviation is a key factor at longer times. Although only the analytical expressions of the microsphere geometry are shown here, it is always of great importance to include the effect of the equilibrium distribution of the active in the models regardless of internal microcapsule morphology. A second generic parameter that should be attributed to the modelling of release data is the effect of burst release[155]. It is suitable to describe the overall release of the active as a sum of two populations: burst and diffusive release. Compared with the more elaborate expression of the diffusive release, the initial burst fraction can be simply modelled as zero order or first order release[156] with the rate constant kburst. Note that a zero order burst function is discontinuous (Equation 23b) when applied in release modelling. A third common approach for any model describing release from microcapsules is the inclusion of the size polydispersity. The release profile is unmistakably influenced by the microcapsule size and size distribution and to find e.g. the proper diffusion coefficient from regression analysis, the fitting should include this distribution. For most formulation routes of microcapsules described in this review, the radius size distribution P(r) follows the log-normal distribution function[25, 157-159]. 30
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A general release model is given in Equation 23 where the population average of the burst and the diffusive part is sized by the fractions pburst and (1- pburst), respectively. 23a
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m(t ) f ( pburst, kburstD, r , t ) pburst f burst (kburst, t ) 1 pburst f ( D, r , K ,..., t ) P(r )dr mtot
23b
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f burst (kburst, t ) 1 : f burst (kburst, t ) f burst (kburst, t ) f burst (kburst, t ) 1 : 1
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Furthermore, the released active can be affected by e.g. degradation when exposed to the release medium. Since it is the detected amount from the experimental quantitative analysis we relate to as m(t), all chemical reactions in the release medium need to be taken into account. An example of this situation, with corrected m(t) using
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degradation models, has been described by us in a recent publication[38] and is treated in more detail in Section 3.2.1.
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3.1.2 Physicochemical properties of core-shell particles
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In contrast to the microsphere, a microcapsule is defined by its two (or more) compartments. The complete diffusion model of the release from microcapsules includes the partition coefficient between core and polymer
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shell, as well as that between shell and external release medium, and a comprehensive analytical expression has also been presented in the literature[160]. In the following section we present some of our work that describes
The core
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the significance of various parameters related to the release from microcapsules.
The inner core morphology and the distribution of the active in the core will have a profound effect on the release profile[23]. As an example, the only way to achieve zero-order release rate is when the active is distributed in the core as crystals surrounded by a rate determining shell[23]. In addition, the size distribution of the microcapsules has to be perfectly monodisperse. Regarding the magnitude of the release rate, the solubility and the diffusion coefficient of the active in the core are important parameters according to Equation 20. The diffusion coefficient can be dramatically altered by changing the physical state of the core. Nordstierna and Nydén studied the effect of using liquid (dodecane) and solid (octadecane) alkanes as core oils on the release of BHT (see Chart 1). BHT is a pesticide which can be used as an effective antifeedant against pine weevil in the early stages of plant protection. Given the results with accompanied Brownian dynamics simulations, reprinted in Fig. 6, the effective diffusion coefficient was significantly reduced when solid octadecane was used instead of liquid dodecane[22].
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Fig. 6. Fractional release m(t)/mtot of BHT as function of time t from microcapsules with (a) dodecane and (b) octadecane as core medium. Solid line represents the release from octadecane capsules calculated by computer simulation of Brownian motion. Note the different axis scales. Reprinted from[161].
Regarding the solubility, it is clear according to Equations 14, 15, 19 and 20 that a high solubility in the core for the active will reduce its distribution coefficient and hence the permeation. Dowding and Vincent investigated
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the release of 4-nitroanisol (see Chart 1) from polystyrene-based microcapsules with cores of varying polarity. It was demonstrated that the release of the active scaled inversely with its solubility in the core, which is in accordance with theory (Equations 19 and 20). The importance of the active’s solubility is often overlooked.
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There are many examples where the active has a higher solubility in the shell than in the core. Enejder et al. demonstrated that the fungicide IPBC (see Chart 1) was exclusively distributed in the shell of PMMA-based
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microcapsules with a dodecane core[141]. For such systems, the use of core-shell morphology is pointless and
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homogenous microspheres (see Scheme 2b) are the more appropriate choice. The core can also have more unexpected effects on the release. Liquid cores, which are the most common types of cores, may plasticise the shell (see Fig. 7). The plasticising effect from the core will increase the free volume of the polymeric shell (see Equations 16 and 17) and hence the effective diffusion coefficient. An example of
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such a system is PMMA-dodecane. Nordstierna revealed that the release of 4-nitroanisole (see Chart 1) from microcapsules consisting of a dodecane core and a PMMA shell was significantly higher than what was theoretically expected assuming simple diffusion through a homogenous PMMA shell[90].
Fig. 7. The fractional release m(t)/mtot of 4-nitroanisole as a function of time t from microcapsules (open circles) and microspheres (solid squares) with (a) 57mg and (b) 880mg 4-nitroanisole. Lower solid line is the fit of the model for diffusion in microspheres. Middle dashed line is the calculated result when applying the diffusion coefficient value obtained from the microsphere fit but for the microcapsule
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The shell
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The shell is obviously the barrier with the purpose of reducing the release of the active and preventing absorption
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of the external medium. This means that the permeability must be lower in the shell than in the core, which can be achieved by choosing a polymer with a proper polarity. According to Equation 20, the release of the active is
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reduced as the solubility of the active in the shell is reduced. However, this is only valid as long as the solubility of the active is higher in the core than in the shell. If this is not the case, a higher solubility of the active in the shell will result in a reduced release rate[22, 115]. In such cases, the use of core-shell morphology is unnecessary
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and a microsphere morphology should be used (see Scheme 2).
The permeability of an active in the shell can also be reduced by introducing impermeable objects into the shell matrix as previously discussed in Section 3. Romero-Cano and Vincent estimated the effective diffusion constant
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of 4-nitroanisole (see Chart 1) in semi-crystalline PLLA (see Chart 2) to be as low as 10-19 m2/s[162]. Another example of impermeable objects is nanoclays. Given their high aspect ratios, aligned nanoclays may alter the tortuosity of the active in the shell material to a great extent. Fan and coworkers incorporated acidified
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montmorillonite into urea-formaldehyde based microcapsules for self-healing applications[163]. This treatment
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significantly reduced the release of cyclopentadiene. However, compatibility between the nanoclays and the surrounding shell matrix is of paramount importance. We have investigated the release of Disperse Red 13 from PMMA-based microcapsules containing hydrophobically modified montmorillonite[22]. It was found that a high
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reduced release rate.
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fraction of nanoclays impaired the integrity of the PMMA shell matrix resulting in an enhanced rather than
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Fig. 8. The fractional release m(t)/mtot of the fungicide OIT as a function of time t from microspheres formulated by slow evaporation of the volatile solvent () and fast evaporation of the volatile solvent (). Solid lines are fitted models according to Equation 22. The top figure shows the entire release using a logarithmic scale while the first 35 hours of the experiment are presented on a linear time scale in the bottom figure. Reprinted from[38].
The above paragraph deals with the macroporosity discussed in Section 3. However, it is also possible to alter the microporosity. Dowding and Vincent was able to successfully reduce the release of 4-nitroanisole from PS, poly(vinyl phenyl ketone) and poly(2-vinylpyridine) based microcapsules by cross-linking the shell polymer[115] as described in Section 2.1.4. Another, more simple, method to reduce the microporosity is to anneal the shell polymer[38, 88] as is also described in Section 2.1.4. The polymeric matrix of the shell is in many cases in an expanded state due to experimental conditions such as fast evaporation of the volatile solvent. We have recently demonstrated that this is the case when rotary evaporation is used for solvent removal from the microcapsule dispersion[38]. This is in contrast to microcapsules prepared by slow evaporation of the solvent at ambient conditions. As a consequence, the calculated effective diffusion constant of the fungicide OIT (see Chart
34
ACCEPTED MANUSCRIPT 1) in PMMA is significantly higher from rotary-evaporation-prepared microcapsules as compared to fume-hoodevaporation-prepared microcapsules (see Fig. 8). In addition, annealing the shell by heating the microcapsule suspension in an autoclave resulted in a significant reduction of the release rate from the rotary-evaporationprepared microcapsules (see Fig. 9). In contrast, this treatment had no effect on the fume-hood-evaporation-
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prepared microcapsules[38] (see Fig. 9). This means that the shell polymer of the latter type of microcapsules is
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already in a conformationally stable state.
Fig. 9. a) The fractional release m(t)/mtot of the fungicide OIT as a function of time t for untreated fume-hood-evaporated microcapsules () and the corresponding heat-treated microcapsules (). b) The release of OIT for untreated rotary-evaporated microcapsules ()and the corresponding heat-treated microcapsules ().[38].
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ACCEPTED MANUSCRIPT The outer dispersant layer and surface modifications In this section we will discuss the outermost layers of the microcapsules and their importance for the release. As mentioned in Section 2.1.3, the dispersant layer; albeit thin, can be viewed as an outer barrier. Before discussing this matter further, it is important to make a point regarding the physicochemical properties of the active, the
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shell and the resulting permeation of the active in the shell. As is clear from Equations 15 and 20, it is beneficial
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for the active to have a low solubility in the shell. This is usually already the case for actives encapsulated in microcapsules prepared via the double emulsion route, since the actives are hydrophilic and the shell polymer
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hydrophobic. However, for microcapsules prepared via the internal phase separation route, the actives are hydrophobic and the shell is semihydrophobic (see Section 2.1.1). This situation is not ideal because hydrophobic actives should require very hydrophilic barriers according to Equations 15 and 20. Yet, knowledge of this problem can be exploited when it comes to the choice of dispersant. We have recently investigated the
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release of the hydrophobic dye Disperse Red 13 (see Chart 1) from PMMA-based microcapsules which have been stabilized by different types of dispersants[27]. The amphiphilic block copolymer dispersants with a polyelectrolyte block (see Chart 2) provides the microcapsules with a highly charged polyelectrolyte brush-like
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surface and this layer is very hydrophilic[25]. It is clear in Fig. 10a that the release rate of the dye is significantly reduced when this block copolymer is used as dispersant as compared to the more conventional uncharged dispersants PMAA and PVA (see reference [27]). Note that the thickness of the block copolymer layer on the
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microcapsule surface is much thinner than the thick gel-like PVA layer (see Fig. 3).
Fig. 10. a) The fractional release m(t)/mtot of the dye Disperse Red 13 as a function of time t from microspheres based on the dispersants PMAA (▼, blue), 600- 4600 (■, black) and 600- 4600 functionalized with 0.5 (●, dark grey), 1.0 (●, grey), 1.5 (●, light grey) and 2.0 (○, white) PDADMAC/PMANa bilayers. The insert magnifies the release from the 600-4600 based microspheres. Redrawn from[27]. b) The
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ACCEPTED MANUSCRIPT calculated effective diffusion coefficient or permeability of Disperse Red 13 as a function of the PEM thickness on PMMA based microcapsules. The data has been fitted with a hyperbolic function A/(B+δm/C) derived from Equations 19 and 20. Here, Deff(δm=0) is the obtained diffusion coefficient for the bare PMMA surface and the polyelectrolyte brush thickness corresponds to the effective thickness of the collapsed brush. Redrawn from[23].
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The idea of using very hydrophilic barriers against hydrophobic actives can also be implemented when it comes to surface modifications. As mentioned in Section 2.1.4, polyelectrolyte multilayers, PEMs, are very promising
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as such barriers. We have modified PMMA-based microcapsules with PEMs to reduce the release of the
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hydrophobic dye Disperse Red 13 and the fungicide OIT[27, 39] (see Chart 1). As can be seen in Fig. 10, the release rate is profoundly reduced by the PEM barrier and the PEM quickly becomes the rate-determining barrier. For Disperse Red 13, the PEM modification with two polyelectrolyte bilayers results in a 160 times decrease of the effective diffusion constant as compared to the untreated microcapsule[27]. Note that this
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decrease of the release rate is significantly larger compared to the decrease resulting from other modifications mentioned in this review as well as most modifications found in the literature. On the other hand, OIT is much less hydrophobic than Disperse Red 13 and displays surface-active properties[38]. It is therefore not necessarily
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certain that the PEM will have an effect on such a “challenging” molecule. Nonetheless, the effective diffusion constant of OIT is reduced 40 times for microcapsules modified with two polyelectrolyte bilayers as compared to the unmodified microcapsules (see Fig. 11). This emphasizes the universal validity of using highly hydrophilic
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PEM barriers against different types of hydrophobic actives.
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Fig. 11. The fractional release m(t)/mtot of the fungicide OIT as a function of time t from PVA-based () and PEM-modified () microspheres . Filled and unfilled symbols are duplicates. The top figure shows the entire release using a logarithmic scale while the first 25 hours are shown in the bottom figure, on a linear scale.
3.2
Release from coatings
Similar to Section 3.1, this section starts by describing the experimental release methodology with respect to coatings. Regarding the diffusion models, the feature of biocide degradation is also taken into account since the duration of the release experiments relevant for coatings is significantly longer than for microcapsule suspension. Thereafter, some release examples concerning both “free” (not encapsulated) and encapsulated biocides are presented with a focus on the properties of the coating matrix.
3.2.1 Methodology and diffusion models The experimental methodology with respect to the release of actives from coatings contains the same steps as discussed in Section 3.1; setup, analysis, and evaluation. In the studies presented here, the setup is based on an 38
ACCEPTED MANUSCRIPT international standard method for the release of biocides from coatings[164] with some minor modification such as permanent immersion of the coating in water instead of a dipping setup. The entire setup is depicted in Scheme 7.
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Scheme 7. Experimental methodology for release measurements of active substances from coatings. The paint consisting of microcapsules are coated on a grazed polypropylene plate before drying. The coated plate is then placed in a water-filled beaker with lid, and stirred using a shaking table. Samples are collected using an automatic pipette. Since the sample may contain unwanted arbitrary paint species, dust etc. chromatography is used to separate the active substance before quantitative analysis by e.g. spectrophotometry.
For a proper release study, the preparation of the coated substrate is important. The substrate has to allow for thin-film coating and needs to be chemically and physically inert to the coating ingredients. Moreover, the substrate should be physically unaffected by film drying and water contact. In our laboratory we use polypropylene plates as substrate[11, 38, 39]. In order to enhance the adhesion between the substrate and the coating and to prevent coating detachment, the surface is grazed in order to roughen the surface. Moreover, precise control of the surface area and the coating thickness is of utmost importance. This can be achieved by defining the surface area with tape followed by coating with an applicator giving a defined thickness. The paint 39
ACCEPTED MANUSCRIPT is usually dried at slightly elevated temperature (37 °C). The drying time has a profound impact on the release, as discussed in Section 3.2.3. Regarding the experimental methodology, it is important to precisely determine the wet and dry weight of the coating. These parameters will ultimately determine the loading concentration and the
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analysed released fraction.
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According to the international standard method used, the coated plates are placed in a beaker leaning against the beaker wall with the coated surface facing downwards (see Scheme 7). It is important to tightly seal the beaker
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with a top cover in order to prevent evaporation of the release medium. In addition, the stirring (usually realized by shaking) should be gentle in order to prevent release due to mechanical load on the coating. The analysis procedure requires a short comment. In contrast to the situation when the release from
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microcapsules is evaluated (Section 3.1.1), the release medium surrounding the coated plate is a solution since all microcapsules are confined within the coating matrix (see 2.2). This means that the concentration of the active can be analysed without any filtration step. However, other small molecules from the paint will be
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released into the surrounding medium. Therefore, spectrophotometry (UV/Vis), which is normally used for analysis of the release in microcapsule suspension, cannot be directly used due to possible overlapping effects. Instead chromatography-based methods such as HPLC are more suitable. Chromatography in conjunction with
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spectrophotometry can be used to evaluate the concentration as well as possible degradation events. This latter topic is briefly discussed below and in Section 3.2.3. Compared to the previously discussed release studies in
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aqueous microcapsule suspensions, the concentration of active substance is here much lower, giving conditions
Diffusion models
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more closely related to a perfect sink.
The common laboratory procedure when it comes to applying paint on a substrate is to use a paint applicator that
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provides a well-defined thickness of the painted layer, typically between 50 and 200 m. The coating will after the drying procedure form a thin film with a homogeneous distribution of all components (binder, pigment, additives, etc.). Such a film can, from a model perspective, be seen as a homogeneous matrix which enables an experimental quantification of the release by model fitting (see Equation 24). The apparent, or effective, diffusion coefficient D for an active substance migrating through and leaving the coating to a perfect sink can be modelled according to the following function derived by Crank
mt 8 f D, L, t 1 2 mtot
D2 2n 12 1 exp t 2 4 L2 n 0 2n 1
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where L is the thickness of the coating. This model is often satisfactory despite the perfect sink condition since the concentration of released active substance is a couple of magnitudes lower compared to the situation in Section 3.1. It is important to note that the effective diffusion constant for the coating is a function of all partition coefficients and diffusion constants depicted in Scheme 5. Therefore, this effective diffusion constant can also be viewed as a global permeability for the coating 40
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The measurements of the release from coatings are usually performed over significantly longer time spans than the release measured in microcapsule suspension. As a consequence, potential degradation of the active may influence the measured release. Note that most modern biocides are designed to be chemically or biologically
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degradable in order to prevent unwanted long-term biocidal activity and bioaccumulation in the environment. In
g ( x) m k deg g ( x) dx
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order to account for potential degradation, the order m of the degradation reaction must be known (Equation 25):
g(x) is the concentration of non-degraded active that is measured in the sink during time x and x = t - j. The rate constant of degradation is denoted kdeg, and m is the order of the degradation kinetics. The mass change of active is expressed as the fractional parts of the release function (Equation 24) and the degradation (Equation 25). The
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corresponding function that explicitly describes first-order (or rather pseudo-first order) degradation, which is the most common order of degradation kinetic (e.g. hydrolysis), is then given by Equation 26[38]:
m(t ) f ( D, L, j ) d exp kdeg t j dj mtot 0 dj .
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We have implemented this modified release function with respect to the degradation of OIT (see Chart 1). This is
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presented in Sections 3.2.3 and 3.3.2.
3.2.2 Solvent-borne vs. water-borne paints The nature of the coating matrix will have a profound effect on the release of the active. It is therefore necessary
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to make a distinction between water-borne and solvent-borne paints. Most water-borne paints consist of polymer lattices dispersed in an aqueous medium. As the paint dries, the space between the particles, i.e. the pores, will be reduced, ultimately resulting in merging of the particles[42, 43]. Any dispersed actives will reside in the pores or be adsorbed on the particle surface. Regarding dispersed microcapsules, the encapsulated active will be
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released into the pores of the coating matrix. The situation is very different for solvent-borne paints. Here, the polymer constituting the coating matrix, i.e. the binder, is dissolved in the solvent. Any dispersed active or microcapsule will be surrounded by the polymer matrix. Therefore, the macroscopic porosity described for the water-borne paints is not relevant for the solvent-borne paints. Instead, the microscopic porosity of the polymeric coating matrix is the important parameter (see Scheme 8).
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Scheme 8. Macroporosity of a latex coating (a) and microporosity of a solvent-borne coating (b) as a function of drying time. a) The porosity of the latex coating will decrease as a function of drying time and is at the initial stages approximately equal to the water content. The porosity ε and tortuosity τ have been defined in Section 3. b) For the solvent-borne paint, the free volume of the matrix Vf will decrease as a function of drying time resulting eventually in a thermally relaxed state. Vf and V0 have been defined in Section 3 and are also discussed in Section 3.1.
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We have investigated the release of the antifouling agent medetomidine (see Chart 1) from both solvent- and water-borne coatings[11]. Both free and encapsulated medetomidine in PMMA-based microcapsules were investigated (see Fig. 12). As can be seen, the release rate is significantly reduced for both paint systems when the active is encapsulated. However, we must note that the effect of the encapsulation on the release rate for a given type of active in a given type of microcapsule can vary significantly. In some cases, the effect of encapsulation is almost absent. Such an example is IPBC (see Chart 1) encapsulated in PMMA-based microcapsules. Whether the encapsulation will have an effect on the release rate or not depends on whether the microcapsule is the rate determining barrier for release according to Equations 19 and 20. However, it is clear from Fig. 12 that the coating matrix has a more profound effect on the release than the encapsulation. As expected, the water-borne coating, containing macroscopic pores, displays a much higher release rate of medetomidine as compared to the solvent-borne coating containing microscopic pores. Note that the observed secondary effect of the encapsulation is not general but depends on the type of microcapsule system used. In Section 3.3.2, we will present a microcapsule system which is virtually independent on the coating matrix.
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Fig. 12. The fractional release m(t)/mtot of medetomidine as a function of time t, on (a) lin–lin scale and (b) log–log scale, from () waterborne paint with free biocide, () water-borne paint with encapsulated biocide, () solvent-borne paint with free biocide, and () solventborne paint with encapsulated biocide. Intersecting lines are solely for clarification. Reprinted from [11].
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3.2.3 Drying time and macroscopic porosity The former section described the importance of porosity of the coating for water- and solvent-borne paints. It is therefore obvious that the drying time of the paint; which will significantly affect the matrix porosity (see Scheme 8), will have a profound effect on the release rate of the active. We have investigated the release of the fungicide OIT from water-borne varnishes, which have been dried at elevated temperatures (37 ºC), as a function of the drying time[38]. Even at this elevated temperature, the complete drying process takes approximately one month. As can be seen in Fig. 13, the drying time has a profound effect on the release rate of OIT. Note also that the degradation of OIT becomes important for these long-term release measurements in contrast to the measurements presented in Section 3.1.2. For the coating which had been dried for seven days, the microencapsulation of OIT results in a twofold reduction of the calculated effective diffusion (or rather permeation) coefficient (Equation 26) as compared to the freely dispersed OIT. However, the effect of altering the macroscopic porosity of the coating matrix is more important than the effect of encapsulation (see Fig. 13). Yet, it is important to note that drying times at elevated temperatures for one month are practically unrealistic, which means that this simple type of encapsulation is still useful.
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Controlled release and antifouling
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Fig. 13. The fractional release m(t)/mtot of the fungicide OIT as a function of time t from varnish coatings. (a) The first 25 hours. (B) The entire experimental time. () Free OIT after 7 days drying with decay fitting (•••) and without decay fitting (—). () Encapsulated OIT after 7 days drying with decay fitting (∙∙∙) and without decay fitting (- - -). () Free OIT after 28 days drying with decay fitting (•••) and without decay fitting (—). () Encapsulated OIT after 28 days drying with decay-fitting (∙∙∙) and without decay fitting (- - -). Reprinted from [38].
In the following subsections, the antifouling and antimicrobial properties of some coatings are presented.
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Emphasis is given to the exceptional antifouling property of biocides with high efficacy and narrow target specificity, in this case medetomidine. We also present the benefit of using strongly rate-determining release systems, such as the PEM-modified microcapsules (see Sections 2.1.4 and 3.1.2), for controlled release of
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hydrophobic biocides, and thereby controlling and prolonging the antifouling/antimicrobial function.
3.3.1 Use of non-hazardous and efficient biocides As mention in Section 2, a promising and, more importantly, feasible antifouling approach is to use a cocktail of biocides with a narrow target specificity towards different types of biofoulers (see Fig. 1). One such biocide, which has been comprehensively investigated in the Marine Paint research programme is medetomidine (see Chart 1). It was shown that medetomidine; which is a α2-adrenoceptor agonist and used as a veterinary sedative, activated the octopamine receptor in barnacles[165]. The barnacle is one of the most problematic biofoulers. Normally, the barnacle settles on a surface in its cyprid larval stage followed by metamorphosis to the adult stage. The octopamine receptor activation of medetomidine increases the swimming activity of the cyprid larvae, preventing it from settling[165]. Medetomidine has been shown to have low toxicity towards several non-target organisms[75, 78-81], and was recently recommended for EU approval by the UK Health and Safety Executive (HSE).
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ACCEPTED MANUSCRIPT The antifouling property of medetomidine was tested by simply mixing the free biocide in a solvent-borne paint which was applied on PMMA plates. After four months immersion in the sea the fouling was assessed. It was found that even though the biocide was not encapsulated, proper antifouling was realized at as low concentrations in the coating as 0.1 wt% (see Fig. 14). As a comparison, the amount of copper used in marine
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paints as biocide is usually as high as 40 wt%.
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Fig. 14. Effect of medetomidine formulated in a solvent-borne paint on barnacle fouling. The percentages given are added amount of medetomidine in the paint formulation.
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Optimized biocide cocktails
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In the Marine Paint project that was mentioned in the Introduction, it became obvious that medetomidine was efficacious only to barnacles, and that a fully protective coating needed to contain a mixture of antifoulants. Experimental work on a set of important fouling organisms and antifouling compounds[66, 166] generated data
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that were used to make a mixture efficacy analysis in silico for all possible combinations of antifoulants. That generated “recipes” for a large number of combinations, all equally effective in controlling the model organisms (barnacles, bryozoans, sea squirts, sea lettuce[166], and multi-species periphyton communities[67]). These
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antifoulant mixtures were then evaluated and ranked based on their estimated environmental risks using Equation 26 to estimate the combined risk for each of the mixtures where ci is the concentration, PECi is the predicted environmental concentration and PNECi is the predicted no effect concentration of biocide i. There was a 100-fold difference between the most and the least risky mixture. An example of the antifouling effect of a coating formulated with such an optimized mixture of biocides in terms of efficacy and environmental low risk is shown in Fig. 15. n
Risk ci i 1
PEC i PNEC i
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Already when these ideas emerged it was obvious that conventional paint formulations might fall short and microencapsulation and nanoparticle approaches were therefore explored. It was anticipated that with these paint technologies it might be possible to finetune formulations for certain environments or when fouling organisms acquire tolerance.
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Fig. 15. Marine biofouling on coatings containing (a) no biocides, (b) an optimized mixture of DCOIT, copper, and medetomidine and (c) a tenth of the optimized mixture concentration. Photos by Åsa Arrhenius and Anneli Hilvarsson. The biocide cocktail was optimized from a a set of conventional biocides; copper pyrithione, irgarol, copper, triphenylborane pyridine, DCOIT and medetomidine[17, 167]. The risk for the cocktail containing DCOIT (11 %), copper (89%) and medetomidine (0.035%) was more than ten times lower as compared with conventional copper-based paints and the amount of copper needed for antifouling protection was reduced 24 times.
3.3.2 Use of rate-determining release systems So far in this review, alteration of the coating matrix has been more important than the effect of encapsulating the active with respect to the release. Regarding the release of hydrophobic actives from microcapsules in suspension, the PEM-modified microcapsules displayed especially low release rates. We have therefore investigated the release of the fungicide OIT, encapsulated in PEM-modified microcapsules, from water-borne acrylate paints as a function of drying time[39]. Similar to the varnish example discussed in Section 3.2.3, the release rate of OIT, which has been freely dispersed in the paint, depends strongly on the drying time (see Fig. 16). In contrast, the release rate of OIT; which is encapsulated in PEM-modified microcapsules, is more or less
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ACCEPTED MANUSCRIPT independent of the drying time and hence the macroscopic coating porosity (see Fig. 16). Clearly, the PEM is the
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rate-determining barrier according to Equations 19 and 20.
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Fig. 16. The fractional release m(t)/mtot of OIT from a latex paint as a function of time t. In a), the first 300 hours of the experiment are displayed and the data have been fitted according to the diffusion models in Equations 24-26. In b), the long-term release is displayed which includes the degradation of the biocide. Freely dispersed OIT is marked with ■ and □, and encapsulated OIT with and. Filled and unfilled symbols are duplicates. The different drying times are marked with, black (one day), blue (seven days) and red (33 days). The fitting for the curve corresponding to free OIT dried for one week was poor and has consequently been excluded. Reprinted from[39].
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The antifouling properties of the above-described coatings against Aspergillus niger was investigated in an accompanying study[39]. The fungus Aspergillus niger, also known as black mold, is a common biofouler on
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exterior walls[9, 13, 39] (see Fig. 1). The antifungal effect of OIT relies on its ability to diffuse over the cell wall of the fungi. In the cytoplasm, the electron deficient sulfur of the isothiazolinone reacts with the thiols in proteins leading to impairment of their enzymatic functions and eventually to cell death[168]. OIT is therefore a broadspectrum biocide of the conventional type and not a selective biocide such as medetomidine. Nonetheless, it is
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clear from Fig. 17 that the encapsulation of OIT in PEM-modified microcapsules profoundly prolonged the antifouling effect of the coating[39]. The minimum surface concentration required for an antifouling effect is determined by the surface flux of active from the coating (the derivative of the release curves in Fig. 16) which is schematically exemplified in Fig. 5 and shown in Fig. 17. Above the critical surface flux of OIT in Fig. 17, fouling of Aspergillus niger is prevented whereas the protection is lost below the critical surface flux.
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Fig. 17. a) Surface flux ṁ(t)/A as a function of time t calculated from the diffusion-modelled release in Figure 15. The colour schemes and symbols are presented in (b). b) Antifouling effect for different coatings assessed by the growth of Aspergillus niger (see Fig. 1). The coatings have been formulated with free (F) and encapsulated OIT in PEM-modified microcapsules (M). A coating containing no addition of OIT was used as a blank reference surface (B). In addition, the drying time for these three types of formulated coatings have been varied (one, seven and 33 days). The plates were removed from the release medium after a specific time which is indicated by the symbols (x-axis) in (a) and their antifouling ability was subsequently assessed. The grey horizontal line in (a) illustrates the critical surface flux needed to reach the critical surface concentration (see Section 3 and Fig. 5).
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Conclusions
Microencapsulation provides many benefits for coatings containing biocides. The microcapsule protects the biocide from degradation and enables the use of a cocktail of different types of biocide in the coating. Most importantly, the encapsulation allows for proper control of the release and thereby prolongs the duration of the biocidal effect and reduces the waste of biocides. This is advantageous with respect to both economic and environmental perspectives. Hydrophobic biocides are preferably encapsulated using the internal phase separation route whereas the double emulsion route is most suitable for hydrophilic biocides. The intention is always to maximize the solubility of the biocide in the core and minimize its solubility in the shell or shells. The resulting release can consequently be controlled by the choice of core and shell materials. Even the choice of
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ACCEPTED MANUSCRIPT dispersant has a significant effect on the release. Regarding the experimental methodology for release measurements, it is of utmost importance to have knowledge of and control over the saturation concentration of the active in the release medium and the distribution coefficient between the microcapsule and the release medium. The release can be further altered by manipulating the shell porosity and by surface modifications. It is
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shown that addition of a PEM, in which the biocide has very low solubility, on the microcapsule surface is a very
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promising way of reducing the release rate of hydrophobic biocides. When such microcapsules are mixed into a coating formulation, the release is virtually independent of the coating matrix, which is a unique feature. This
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Acknowledgements
The authors are grateful to Alberta Mok, Alireza Movahedi, Anna Ananievskaia, Asvad A. Gabul-Zada, Gia
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Kourouklidou, Hermann Uhr, Kurt Löfgren, Matias Nordin and Ye Li for laboratory, analytical and administrative work regarding microencapsulation and controlled release. The authors are also grateful to all coworkers in the Marine Paint Research Programme and in the innovation companies Capeco and I-Tech.
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We also acknowledge funding from Mistra (the Swedish Foundation for Environmental Research), the Swedish
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S.K. Ghosh, in: S.K. Ghosh, (Ed.) Functional Coatings, 1st ed.; Wiley-VCH, Weinheim, 2006. S. Kiil, D.M. Yebra, Adv. Mar. Antifouling Coat. Technol. (2009) 334. J.R. Rischbieth, F. Marson, Nature (London, U. K.) 192 (1961) 748. I. Omae, Chem. Rev. (Washington, DC, U. S.) 103 (2003) 3431. I. Omae, Kagaku Kogyo 54 (2003) 646. International Maritime Organization, International Convention on the Control of Harmful Anti-fouling Systems on Ships, 2011, [Cited: 2011:11:07]. Available from http://www.imo.org M. Edge, N.S. Allen, D. Turner, J. Robinson, K. Seal, Progress in Organic Coatings 43 (2001) 10. M.A. Shirakawa, R.G. Tavares, C.C. Gaylarde, M.E.S. Taqueda, K. Loh, V.M. John, Science of The Total Environment 408 (2010) 5878. O.C.G. Adan, R.A. Samson, Fundamentals of mold growth in indoor environments and strategies for healthy living. Wageningen Academic Publishers, Wageningen, 2011. L. Nordstierna, J. Borg, M. Andersson Trojer, in: (Ed.)^(Eds.)European coatings congress, 2013. L. Nordstierna, A.A. Abdalla, M. Masuda, G. Skarnemark, M. Nydén, Prog. Org. Coat. 69 (2010) 45. L. Nordstierna, A. Movahedi, M. Nyden, J. Dispersion Sci. Technol. 32 (2011) 310. B. Crook, N.C. Burton, Fungal Biology Reviews 24 (2010) 106. L. Petrone, Advances in Colloid and Interface Science 195–196 (2013) 1. E. Almeida, T.C. Diamantino, O. De Sousa, Prog. Org. Coat. 59 (2007) 2. L.D. Chambers, K.R. Stokes, F.C. Walsh, R.J.K. Wood, Surf. Coat. Technol. 201 (2006) 3642. T. Backhaus, Å. Arrhenius, Marine Paint Final Report 2003-2011, U.o. Gothenburg, Gothenburg, 2012. J.A. Callow, M.E. Callow, Nat Commun 2 (2011) 244. P. Schultz Michael, Biofouling 23 (2007) 331. K. Nielsen, J. Mogensen, M. Johansen, T. Larsen, J. Frisvad, Analytical and Bioanalytical Chemistry 395 (2009) 1225. N. Read, in: K. Mendgen, D.-E. Lesemann, (Eds.), Electron Microscopy of Plant Pathogens; Springer Berlin Heidelberg, 1991, p 17. M. Andersson Trojer, in: P. Somasundaran, (Ed.) Encyclopedia of Surface and Colloid Science; Taylor and Francis, New York, DOI: 10.1081/E-ESCS-120048578, 2013. M. Andersson Trojer, L. Nordstierna, M. Nordin, B.M. Nyden, K. Holmberg, Phys. Chem. Chem. Phys. 15 (2013) 17727. T. Nesterova, K. Dam-Johansen, L.T. Pedersen, S. Kiil, Prog. Org. Coat. 75 (2012) 309. M. Andersson Trojer, Y. Li, C. Abrahamsson, A. Mohamed, J. Eastoe, K. Holmberg, M. Nyden, Soft Matter 9 (2013) 1468.
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[32]
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Graphical abstract
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ACCEPTED MANUSCRIPT Highlights The encapsulation strategy depends on the physicochemical properties of the active
The important parameters with respect to a proper release methodology are described
The release is reviewed in terms of the properties of the microcapsule
Suitable theoretical diffusion models are presented and discussed
Controlled release and antifouling properties of coatings are presented
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