Textile Finishing: Recent Developments and Future Trends

By K.L. Mittal and Thomas Bahners

Textiles have been historically and traditionally used to make clothes, but even in ancient times there were technical textiles for making sails, tents, etc. Today, technical textiles are used in various industries for a host of purposes and applications. Recently, there have been exciting developments on various fronts in the textile field to impart novel and innovative functionalities to textiles, e.g., easy-to-clean or dirt-repellent, flame retardancy, anti-bacterial, and fog-harvesting properties, to name a few. Also, textiles for electronics based on graphene, CNTs and other nanomaterials, conductive textiles, textiles for sensor function,   textile-fixed catalysts,  textiles for batteries and energy storage, textiles as substrates for tissue engineering, and textiles for O/W separation have appeared in the literature. All this has been possible through adopting novel ways for finishing textiles, e.g., by appropriate surface modification techniques, and utilizing biomimetic concepts borrowed from nature.

This unique book entitled “Textile Finishing: Recent Developments and Future Trends” is divided into four parts: Part 1: Recent Developments/Current Challenges in Textile Finishing; Part 2: Surface Modification Techniques for Textiles; Part 3: Innovative Functionalities of Textiles; Part 4: Fiber-Reinforced Composites.

The topics covered include: Antimicrobial textile finishes; flame retardant textile finishing; “self-cleaning” or easy-to-clean textiles; metallization of textiles; atmospheric pressure plasma, and uv-based photochemical surface modification of textiles; tunable wettability of textiles; 3D textile structures for fog harvesting; textile-fixed catalysts; medical textiles as substrates for tissue engineering; and fiber-reinforced “green” or “greener” biocomposites and the relevance of fiber/matrix adhesion.

• Language: English
• Publisher: Wiley
• Release date: Aug 22, 2017
• ISBN: 9781119426851

Book preview

Preface

Textiles have been historically and traditionally used to make clothes, but even in ancient times there were technical textiles for making sails, tents, etc. Today technical textiles are used in various industries for a legion of purposes and applications. Recently, there have been exciting developments on various fronts in the textile field to impart novel and innovative functionalities to textiles, e.g., easy-to-clean or dirt-repellent, flame retardancy, anti-bacterial, and fog-harvesting properties, to name just a few. Also textiles for sensor function, textile-fixed catalysts, textiles for batteries and energy storage, textiles as substrates for tissue engineering, and textiles for O/W separation have appeared in the literature. All this has been possible through adopting novel ways for finishing textiles, e.g., by appropriate surface modification techniques, and utilizing biomimetic concepts borrowed from Nature. Nature is a great teacher and we can learn a lot from the creativity of Nature as Nature does not waste time in frivolous activities.

It should be emphasized that by suitable surface modification of any material (including textiles) one can attain the requisite surface characteristics (e.g., adhesion, wetting, superhydrophilicity, superhydrophobicity, omniphobicity, anti-fouling, biocompatibility, to mention just a few) without tampering with the desirable bulk attributes of materials, as a material is surface modified to a depth of about 10nm only. This avenue provides the best of both worlds. Concomitantly, in the last years there has been a flurry of research activity in ameliorating the existing approaches as well as in devising novel, innovative and more efficient ways for surface modification of polymers, including textiles.

Now coming to this book entitled Textile Finishing: Recent Developments and Future Trends it is divided into four parts: Part 1: Recent Developments and Current Challenges in Textile Finishing; Part 2: Surface Modification Techniques for Textiles; Part 3: Innovative Functionalities of Textiles; and Part 4: Fiber-Reinforced Composites. The topics covered include: Antimicrobial textile finishes; flame retardant textile finishing; self-cleaning’ or easy-to-clean textiles; metallization of textiles; wettability characterization in textiles; surface functionalization of textiles by atmospheric pressure plasma (APP); UV-based photochemical surface modification of textiles; tunable wettability of textiles; 3D textile structures for fog harvesting; textile-fixed catalysts and their use in heterogeneous catalysis; medical textiles as substrates for tissue engineering; and fiber-reinforced green or greener" biocomposites and relevance of fiber/matrix adhesion.

This book represents the cumulative wisdom and contribution of many internationally renowned experts actively engaged in various ramifications of textile finishing. Also this unique book containing a wealth of information provides an easily accessible, unified and comprehensive source as well as commentary on the current state-of-the-art in the arena of textile finishing. Apropos, these days the mantras are: nano and green, and wherever applicable and feasible, these two catchwords have been accorded due consideration.

The editors hope that researchers in academia, governmental and other research laboratories and R&D personnel in textile-related industries will find this book of great interest, value and usefulness. It should be of particular interest to those working in polymers, materials science, clothing science and technology, medical field and those seeking novel and innovative characteristics which can be expected in the future. As novel and ameliorated ways are developed for textile finishing, new application vistas will emerge. Also we are quite sanguine that this book will serve as a fountainhead for new research ideas and will facilitate cross-pollination.

Now comes the pleasant task of thanking those who were instrumental in realizing this book. First and foremost, we will like to profusely thank the authors for their sustained interest, enthusiasm, unwavering cooperation and contribution which was sine qua non for materializing this book. Also we very much appreciate the steadfast interest and support of Martin Scrivener (Scrivener Publishing) in this book project and for giving this book a body form.

K.L. Mittal

Hopewell Jct., NY, USA

E-mail: [e-mail]

Thomas Bahners

DTNW gGmbH

Krefeld, Germany

E-mail: [e-mail]

Part 1

RECENT DEVELOPMENTS AND CURRENT CHALLENGES IN TEXTILE FINISHING

Chapter 1

Recent Concepts of Antimicrobial Textile Finishes

Barbara Simončič* and Brigita Tomšič

University of Ljubljana, Faculty of Natural Sciences and Engineering, Department of Textiles, Graphic Arts and Design, Ljubljana, Slovenia

*Corresponding author: [e-mail]

Abstract

The chapter reviews the most important antimicrobial agents for textiles and the mechanisms of their antimicrobial activity. Structures of the leaching and the bound compounds are presented and their modes of antimicrobial functions are discussed. In addition to active antimicrobial agents, the structures of low adhesion compounds and their passive antimicrobial activity are also presented. The importance of dual-action antimicrobial coatings consisting of combined controlled release and biobarrier forming active antimicrobial compounds as well as the active antimicrobial and low adhesion agents is highlighted. Standard microbiological test methods for the determination of the efficiency of antibacterial and antifungal activity of the agents on textiles are described and their pros and cons are discussed. Health and environmental impacts of the antimicrobial compounds are discussed as well as the future trends of their use are indicated.

Keywords: Antimicrobial activity, textiles, finishing, mechanisms of action, chemical structures, low-adhesion compounds

1.1 Introduction

The development of effective antimicrobial protection of textile substrates has enabled the expansion of the use of textile products in various industrial sectors, including protective and technical textiles, pharmacy, medicine, transport, tourism, agriculture, and food [1]. It includes protection against all types of microorganisms, i.e., bacteria (antibacterial), viruses (antiviral), fungi (antifungal) and protozoa (antiprotozoal). It may be intended for the protection of the users or the textile fibers. The former protects people against pathogenic and odor-causing microorganisms, which can lead to health and hygiene problems. The latter protects textile substrates against adverse textile aesthetic changes, such as colored stains and discoloration of textiles and biodegradation due to molding and rotting, which results in the reduction of breakage strength, elongation and elasticity and can lead to reduced use value of textiles [1–3].

Microorganisms can be adsorbed onto the textile substrate from the surroundings or colonize on the fibers that are in direct contact with the skin. Natural and synthetic fibers are of organic origin and, as such, represent a culture medium for the growth and development of microorganisms, which reproduce uncontrollably under favorable conditions including moisture, oxygen, heat and dirt. Whereas bacteria are primarily present on synthetic fibers, fungi are present only to a minor extent; natural cellulose, wool and silk fibers can provide excellent conditions for the growth of bacteria, fungi and algae [4–7].

As for the microorganisms present on textile fibers, those that cause various diseases and infections and those that are involved in the process of biodegradation of textile fibers are the most important. The bacterial species Staphylococcus epidermidis and Corynebacterium are the main causes of body and clothing odor [8]. The bacterial species Proteus mirabilis, the fungi Candida albicans and Epidermophyton floccosum, and fungi of the genus Trichophyton may cause skin irritation and infections [8]. On the textile fibers, the pathogenic Gram-positive bacterium Staphylococcus aureus, which is one of the major causes of community-acquired and hospital-acquired infections, can also be found [9]. The most active microorganisms in the process of biodegradation of textile fibers are fungi from the genera Aspergillus Chaetomium, Microsporum, Myrothecium and Penicillium, which cause enzymatic decomposition of both natural and synthetic fibers [6]. The most important bacteria that cause biodegradation of textile fibers are from Bacillus, Pseudomonas and Cellulomonas species [6].

Protection against microorganisms can be achieved by chemical modification of textiles with antimicrobial agents, which prevents or inhibits the growth of microorganisms and subsequent biodegradation of the fibers. There is a variety of classical and modern antimicrobials on the market, which differ in chemical structure, mode of application, antimicrobial action, effectiveness, durability to washing, impact on people and the environment, and price [1–3, 7, 9]. The effectiveness of antimicrobial agents is directly influenced by various factors; of these factors, the chemical structure and concentration of the agent, its antimicrobial mechanism, the type of microorganisms present, the chemical and morphological properties of the textile substrate, and environmental conditions including temperature, pH, the presence of moisture are the most important [3]. An effective, ecologically safe antimicrobial agent should have the following characteristics: efficiency against a broad spectrum of microorganisms, effectiveness at low concentrations and low contact time in the whole lifecycle of the textile product, colorlessness and odorlessness, washing durability, resistance to UV radiation, compatibility with other finishing agents and auxiliaries, preservation of the mechanical and physical properties of textiles, application with the use of standard equipment, economical use, nontoxicity to humans at the concentrations used and environmental friendliness [3]. Although many research works have considered the synthesis of novel antimicrobial agents, none of them fully and simultaneously meets all of the abovementioned characteristics. Accordingly, this research topic still offers many challenges and poses different toxicological and ecological problems and questions for researchers.

1.2 Antimicrobial Agents

Antimicrobial agents for textiles can be classified in several ways, the most frequently used are in terms of chemical structure, origin, concentration, efficiency, mechanism and spectrum of activity, and purpose of the antimicrobial textiles [1–3, 7, 10].

The concentration of the active substance in the antimicrobial agent is of prime importance for its antimicrobial activity [10]. It is found that a minimum inhibitory concentration (MIC) is required for biostatic activity and that the biocidal activity is achieved only if the minimum biocidal concentration (MBC) is exceeded. Whereas the biostats inhibit the growth of microorganisms, the biocides kill the microorganisms. The concentration directly influences the efficiency of the antimicrobial agent and should not be below the MIC, which ensures the resistance of microorganisms to the antimicrobial agent. Biocidal or biostatic activity of the antimicrobial agent is also influenced by the microorganisms because the toxicity of the agent to a particular microorganism can vary [11]. Namely, some antimicrobial agents seem to be more effective against bacteria than fungi or against Gram-positive than Gram-negative bacteria. There are antimicrobial agents that act on a wide range of microorganisms and those with a very limited spectrum of action. Accordingly, the concentration and the mode of action of the antimicrobial agent are directly correlated.

1.2.1 Mechanisms of Antimicrobial Activity

Antimicrobial agents used for textile protection utilize two mechanisms of antimicrobial activity:

a controlled-release (Figure 1.1a) and

a barrier formation mechanism (Figure 1.1b).

Figure 1.1 Schematic presentation of the controlled-release of the antimicrobial agent particles (depicted as grey spots) from the textile surface to surrounding where they kill the microorganisms (a) and the formation of polymer film by the antimicrobial agent on textile surface which acts as a barrier for microorganisms (b).

The controlled-release mechanism is characteristic of the leaching antimicrobial agents [1–3, 7, 10]. The majority of these agents are physically incorporated into the textile fibers, and their antimicrobial activity is attributed to the gradual and persistent release of the agents from the textile into their surroundings in the presence of moisture, where they act as a poison to the antimicrobials. There are also antimicrobial agents that are chemically bonded to the textile fibers, but their antimicrobial activity is due to the controlled-release of the active substance. Because these agents can be regenerated in an appropriate medium to restore their antimicrobial activity, their antimicrobial mechanism is also considered as the regeneration model [7]. Related to the antimicrobial mechanism, there are some important disadvantages in the application of the leaching antimicrobials. A release of the agent from the textile surface to the surrounding results in a decrease in the concentration of the active substance, which eventually falls below the limit of effectiveness. In addition to the deactivation of the antimicrobial agent, this can induce microorganisms to become resistant to it. If the textiles are used in contact with skin, the released antimicrobial agent can kill the beneficial bacteria of the skin microbiota, causing skin irritation. Furthermore, physically bonded leaching antimicrobials are not resistant to washing but are gradually removed from the textiles via repetitive laundering. This may cause serious environmental problems. To prolong the antimicrobial activity, the microencapsulation process has been established in the production of the leaching agent in which the active substance is entrapped in microcapsules from which its release can be controlled [12–14].

A barrier formation mechanism is characteristic of so-called bound antimicrobial agents [2, 10]. These include unique chemical structures that enable chemical binding of the agents to the surface of the textile fibers from where they do not release or leach but act as a barrier to control microorganisms that make contact with the fibers. Chemical binding of the agent to the textile surface can be possible if there are enough reactive functional groups in the agent and in the fibers to form chemical bonds. They have significant advantages over the leaching agents. Because of their non-leaching properties, the concentration of the active substance does not decrease during their operation, which results in a very small probability for microorganisms to develop resistance to them. The barrier formation enables only the microorganisms that are adsorbed onto the textile surface to be killed, so the agents cannot cross the skin barrier and irritate the skin. Bound antimicrobials are much more resistant to repeated laundering compared to leaching agents. Along with these advantages, however, these agents also demonstrate an important weakness. Namely, despite the presence of the barrier on the fiber surface, these can be deactivated by deadly microorganisms, the adsorption of dirt or the neutralization of positive charges owing to complex formation between the cationic antimicrobial group of the biobarrier and the anionic detergent during laundering. Additionally, the bio-barrier can be gradually removed from the fiber surface by abrasion.

1.2.2 Structures of Antimicrobial Agents

1.2.2.1 Leaching Antimicrobial Agents

The most important controlled-release antimicrobial agents used for the chemical modification of textiles include halogenated phenols, cationic surfactants known as quaternary ammonium (QAS) and phosphonium (QPS) salts, zinc pyrithione, polybiguanides, N-halamines, nanoparticles of noble metals and metal oxides, and natural plant-based bioactive substances. The mode of the antimicrobial action of these agents is directly influenced by their chemical structures.

Among halogenated phenols, triclosan, 5-chloro-2-(2,4-dichlorophenoxy)phenol (Figure 1.2) and its derivatives [15–18] are the most widely used biocides. Triclosan has a broad spectrum of activity against many Gram-positive and Gram-negative bacteria at low concentrations. The antimicrobial activity of triclosan is based on its binding to the bacterial enzymes, blocking the active sites of the enzymes responsible for the fatty acid synthesis, which is essential to building the bacterial cell membrane (20).

Figure 1.2 Halogenated phenol, 5-chloro-2-(2,4-dichlorophenoxy) phenol (triclosan).

QASs and QPSs include cationic surface active agents with an amphiphilic molecular structure consisting of the polar cationic ammonium or phosphonium group and nonpolar alkyl or perfluoroalkyl group (Figure 1.3) [20–25]. The antimicrobial activity of QAS depends on the length of the alkyl chain, the presence of the perfluorinated group and the number of cationic ammonium groups in the molecule. QASs are important biocides that attack a broad spectrum of bacteria, but they are less effective with fungi. The antimicrobial function arises from attractive interactions between the cationic ammonium group of the QAS and the negatively charged cell membrane of the bacteria, resulting in the QAS–microorganism complex formation. This, in turn, causes interruption of all essential functions of the cell membrane and thus the interruption of protein activity. QASs also affect bacterial DNA, thereby causing a loss of multiplication ability. If the long alkyl chain is bonded to the cationic ammonium in the structure of the QAS, in addition to a polar interaction between the cationic nitrogen of the ammonium group and the bacterial cell membrane, a nonpolar interaction with the hydrophobic alkyl chain also occurs. This enables penetration of the hydrophobic group into the microorganism, allowing the alkylammonium group in contact with the cell membrane to physically interrupt all key functions of the microorganism cell [27–29].

Figure 1.3 Cationic surfactants: (a) alkyl-3.4-dichlorobenzyl-dimethyl ammonium chloride (b) diquaternary perfluoroalkyl dimethyl ammonium salt and (c) methyltriphenyl phosphonium salt.

Among QPSs, methyltriphenyl phosphonium salt is the most important [30]. It has been reported that quaternary phosphonium salt is more stable than quaternary ammonium salt as well as it exhibits a higher activity by 2 orders of magnitude than the polymeric quaternary ammonium salt with the same structure except the cationic part [31].

N-halamines include the unique heterocyclic organic structures with the antimicrobial chlorinated amine, amide or imide groups (Figure 1.4) [32–38]. Vinyl [32, 33] or trialkoxysilane [34–38] group in the molecule enables N-halamines to be covalently bonded to the textile fibers. N-halamines are biocides with a broad spectrum of activity against bacteria, fungi and viruses. Irrespective of their chemical bonding to the textile substrate, their antimicrobial activity is in accordance with the controlled-release mechanism. Namely, the antimicrobial activity of N-halamines based on the electrophilic substitution of Cl in the N-Cl bond with H in the presence of water results in release of Cl+ ions with antimicrobial properties. Cl+ ions can bind to acceptor regions on microorganisms, which hinders enzymatic and metabolic processes, leading to the destruction of the microorganisms [39]. Because the N-H bond formed in the substitution reaction has no antimicrobial properties, regeneration of the N-Cl bond with further exposure of the agent to dilute sodium hypochlorite is needed to regenerate its antimicrobial activity (Figure 1.5).

Figure 1.4 N-halamines: (a) 3-allyl-5,5-dimethylimidazolidine-2,4-dione [32], (b) 3-(30-triethoxysilylpropyl)-7,7,9,9-tetramethyl-1,3,8-triazaspiro[4.5]-decane-2,4-dione [35], (c) N-(3-triethoxysilylpropyl)-N′-phenylbutanediamide [34].

Figure 1.5 Halogenation of the amide group of imidazolidine-2,4-dione (a) and its antimicrobial activity with the release of the Cl+ cation (b) [39].

Polybiguanides are linear polymeric polycationic amines soluble in water, which consist of a hydrophobic hydrocarbon backbone with the incorporated multiple cationic biguanide repeat units [40–46]. For the textile application, the most important polybiguanide is poly(hexamethylenebiguanide) (PHMB) hydrochloride with an average of 11–15 biguanide units separated by hexamethylene chains (Figure 1.6) [40, 41]. The terminating end groups can be amine, guanide, or cyanoguanide groups. Besides being used as a finishing agent, PHMB hydrochloride has already been incorporated into the antimicrobial electrospun nanofibers [47, 48] as well as covalently bonded into the fiber polymer structure via a combination of copolymerization and a wet-blend-spinning method to prepare nonleaching acrylic fibers with permanent antibacterial activity [49]. The antimicrobial mechanism of PHMB is considered to be similar to that of QAS. Both cationic and hydrophobic features of PHMB suggest that it interacts with microbial cell membrane through electrostatic and hydrophobic interactions. It was demonstrated that electrostatic interactions are a dominant factor [50]. It was proposed that the interaction of PHMB and bacterial membrane phospholipids results in cell membrane disruption.

Figure 1.6 Poly(hexamethylenebiguanide).

Zinc pyrithione (Figure 1.7) is an organic-metal complex with a broad spectrum of antibacterial and antifungal activity [51–54]. It is believed that zinc pyrithione utilises a copper toxicity mechanism. Namely, it is assumed that copper present in the environment could replace zinc in the complex, forming copper pyrithione, which delivers copper to the inside of the microorganism cell. Therefore, antimicrobial activity of zinc pyrithione is derived from the ability to increase the intracellular copper level, which inhibits the growth of microorganisms. In the case of fungi, copper damages the iron-sulphur cluster-containing proteins, essential for metabolism [53].

Figure 1.7 Organic metal compound, bis(2-pyridylthio)zinc 1,1′-dioxide (zinc pyrithione).

Nanoparticles of noble metals and metal oxides have proved to be highly effective biocides for bacteria as well as fungi. They mainly include silver (Ag) and silver salts (AgNO3, AgCl) [55–57], titanium dioxide (TiO2) [58–60], zinc oxide (ZnO) [60–62] and copper II oxide (CuO) [63]. Their antimicrobial activity is attributed to the controlled release of both metal ions and the nanoparticles (NPs). The efficiency of the latter strongly increases with decreasing size because particles with smaller size showed enhanced activity because of the larger surface area-to-volume ratio and surface reactivity than those of the larger size. It is believed that metal ions and the nanoparticles react simultaneously because a small number of metal ions are released from the NPs as a result of the oxidation of NPs in the presence of water and oxygen. The interaction of NPs with microorganisms directly depends on their physical and chemical properties and the type and physiological state of the microorganism. Both NPs and ions can bind to the surface of microorganisms’ cells because of the electrostatic attractive forces [57, 58, 64]. NPs smaller than 10 nm can also penetrate into the interior of microorganism cells. It is also demonstrated that the NPs produce secondary products that cause cell destruction. To stabilize nanoparticle structure, control the concentration of released nanoparticles or metal ions, prolong the release time and thereby improve the durability and wash resistance of the finish, nanosized metals and metal oxides have been embedded into different polymer matrices created on the fibers surface [65–68] or applied in combination with cross-linking or grafting agents, such as 3-(glycidoxypropyl)trimethoxysilane [69] and aminopropyltriethoxysilane [70].

In textiles, Ag [71–76], TiO2 [70, 76–79] and ZnO [80–86] NPs and their combinations [87–90] have been intensively investigated. According to literature, the mechanisms of the antimicrobial activity of Ag NPs, i.e., nanoscale clusters of metallic silver atoms, Ag⁰, and silver ions, are very similar [91–95]. Both NPs and ions can form intermolecular interactions with the cell membrane of bacteria. Ag NPs penetrate into the interior of microorganism cells, where they bind to the thiol groups of enzymes and nucleic acids. When present in the cell, NPs hinder or deactivate its critical physiological functions, such as cell wall synthesis, membrane transport, and the synthesis of nucleic groups, including deoxyribonucleic and ribonucleic acids and electron transport. Their interactions with the thiol groups of proteins hinder their enzymatic functions. Additionally, the binding of metal ions to the DNA of bacteria causes them to lose their ability to reproduce. In the presence of oxygen, metal ions and NPs may also catalytically accelerate the formation of reactive oxygen species (ROS), which are highly toxic to cells [94, 96]. The formation of ROS is shown in the following reaction:

(1.1)

Although ROSs are normal side products in the process of cell respiration, their production in excessive amounts results in oxidative stress, which causes damage to the lipids, proteins and DNA of microorganisms and consequently destroys the microorganisms’ cells.

TiO2 and ZnO NPs also exhibit excellent antimicrobial activity against Gram-negative and Gram-positive bacteria, fungi, molds and viruses [97–101]. The reason for this is attributed to their oxidative photocatalytic activity, which is a characteristic of semiconductor photocatalysts. Namely, it has been observed that light activation (photocatalysis) of TiO2 and ZnO NPs significantly enhances their antimicrobial activity. The photocatalytic killing of microorganisms can be explained by different mechanisms. It is assumed that the photocatalytic process on TiO2 and ZnO NPs includes the formation of ROS, such as superoxide anions (•O2–), H2O2 and hydroxyl radicals (•OH) as follows [97, 102]:

(1.2)

(1.3)

(1.4)

(1.5)

(1.6)

The photocatalytic process on TiO2 and ZnO NPs also inhibits the fungal growth by affecting cellular functions, which causes deformation in hyphae and distorted conidial heads and shrivels hyphal walls in mycelia. Peroxidation of lipids also induces DNA damage and disruption of the cell membrane morphology [101].

Chitosan (Figure 1.8a) is a partially deacetylated chitin and represents a biodegradable, biocompatible and nontoxic biopolymer with antimicrobial properties [103–105]. The antimicrobial activity of chitosan is directly influenced by its molecular weight, degree of deacetylation, and pH [105, 106]. In general, the antimicrobial activity of chitosan increases with increasing deacetylation degree and decreasing molecular weight and pH. The mechanism of the antimicrobial activity of chitosan is attributed to the cationic amine groups created by their protonation at pH levels lower than the pKa. This also increases the solubility of chitosan in water.

Figure 1.8 Chitosan and its derivatives: (a) chitosan [103], (b) N,N,N-trimethyl chitosan iodide, (c) N-[(2-hydroxy-3-trimethylammonium)propyl]chitosan chloride [103], (d) O-quaternary ammonium salt-chitosan derivative bearing N-methyl-N-R-N, N-bis(2-hydroxyethyl) ammonium bromide (R = alkyl) [109], (e) N,N,N-trimethyl O-(2-hydroxy-3-trimethyammonium propyl) chitosan [110], (f) N-quaternary triphenylphosphonium salt-chitosan derivative [111, 112].

Because the activity of chitosan is limited to the acidic pH, quarternization of chitosan (Figures 1.8 b and c) has been performed to prepare derivatives with cationic ammonium groups [103, 107–110]. This significantly improved aqueous solubility and antimicrobial activity over a broader pH range compared to native chitosan. Furthermore, a series of novel dihydroxy quaternary ammonium salts were successfully grafted onto the free -OH of chitosan to prepare water-soluble O-quaternary ammonium salt-chitosan derivatives. It was found that quaternized chitosan exhibits better antimicrobial properties than chitosan. The antibacterial activity gradually increases with increasing length of the alkyl chain in the alkyl dimethyl ammonium group and in the presence of hydroxyl groups. As an alternative to quaternary chitosan derivatives with quaternary ammonium moiety, chitosan derivatives with cationic lipophilic quaternary phosphonium groups have been synthesized [111–114]. These studies showed that quaternary phosphonium groups have better antimicrobial properties than quaternary ammonium groups.

The mechanism of the antimicrobial action of chitosan is based on the electrostatic attraction between the cationic groups of chitosan and the negatively charged microbial cell membrane, which restrains all essential functions of the microbial cell membrane. Oligomeric chitosan can also penetrate into the cells of microorganisms and prevent the growth of cells by prohibiting the transformation of DNA into RNA. Chitosan seems to be effective against bacteria and fungi, but in the case of bacteria, it is more effective against Gram-positive than Gram-negative bacteria [103–105, 115].

Plant extracts are natural bioactive antimicrobial and antioxidative agents that represent nontoxic, safe, human- and environment-friendly biodegradable compounds [116–118]. They are mainly extracted from the following parts of the plants: Aloe vera leaves [119, 120], tea and olive tree leaves [117, 121, 122], neem (Azadirachta indica) leaves [117, 120], Eucalyptus leaves [117, 123], henna (Lawsonia inermis) leaves [124], pomegranate (Punica granatum) fruit peels [125], turmeric (Curcuma longa) roots [116, 126] and others. The extracted antimicrobial compounds include phenolics (simple phenols, phenolic acids, naphthoquinones, flavonoids, flavones, flavonols, tannins, coumarins), terpenoids, curcumin, essential oils, alkaloids, lectins, polypeptides, polyacetylenes and others [116, 118]. Some of these are presented in Figure 1.9.

Figure 1.9 Plant extracts: (a) tannins, (b) lawsone, (c) flavone, (d) terpenoids, (e) alkaloids, (f) lapachol, (g) curcumin.

Because many of the identified compounds from plants are colored, they are used as natural antimicrobial dyes and pigments for dyeing natural and synthetic fibers. The most important antimicrobial mechanisms of these compounds comprise intercalation into the microorganism cell wall, formation of complex with the cell wall, disruption of the cell membrane, inactivation of the enzymes, and interactions with DNA. Some of the antimicrobial compounds are also insect growth regulators.

Despite all the benefits attributed to bioactive antimicrobial agents, the most important limitation of these compounds for the application to textile substrates is their very low affinity to textile substrates, which makes it difficult to achieve a concentration greater than the MIC [117]. Furthermore, the weak binding of the substances to the fibers does not preserve their durable antimicrobial activity.

1.2.2.2 Bound Antimicrobial Agents

A precondition for chemical binding of antimicrobial compounds to the textile surface is the presence of the reactive groups in the structure that are able to chemically bind to the functional groups of the fibers. The most important bound antimicrobial agents used for the chemical modification of textiles include polymerizable surfactants (surfmers) [128–131], QAS-functional trialkoxysilanes [132–138], reactive dyes with incorporated QAS [139–142] and reactive quaternized chitosan [143–145]. The scenario for the synthesis of these antimicrobial compounds was to bind QAS to the textile fibers covalently and subsequently convert the controlled- release mechanism of QAS to biobarrier formation.

Surfmer monomers (Figure 1.10) include the biocidal QAS group linked to the vinyl, acrylic or methacrylic groups. The presence of the vinyl (ethenyl) group enables polymerisation of the surfmer into the organic polymer consisting of the polyvinyl backbone and the side antimicrobial QAS groups of different structures. To increase the durability of the polymer film, surfmers are copolymerized with glycidyl methacrylate or vinyl methacrylate that act as cross-linking agents and anchors for the polymer film to fiber functional groups. The comonomers should be carefully selected according to the nature of the substrate to which they are applied. The copolymerization of surfmers with glycidyl methacrylate and chemical binding of the copolymer to the cellulose fibers are presented in Figure 1.11.

Figure 1.10 Surfmer, alkyl[2-(acryloyloxy)ethyl]dimethylammonium bromide [128].

Figure 1.11 Copolymerization of surfmer with glycidyl methacrylate and chemical binding of copolymer to the cellulose fibers.

The QAS-functional trialkoxysilanes (Figure 1.12) are a class of hybrid organic–inorganic sol-gel precursors with the chemical structure R′–Si(OR)3 [146, 147], where R’ represents the QAC functional antimicrobial group and OR reactive alkoxide group. The latter reacts readily with water to cause hydrolysis in the presence of a mineral acid or a base as a catalyst (Figure 1.13). The hydrolysis reaction replaces alkoxide groups with hydroxyl groups (–OH), forming the silanol groups (–Si–OH), which produce siloxane bonds (Si–O–Si) in the subsequent condensation reaction [146]. This type of reaction can continue to build a large, silicon-containing polymer network with a three-dimensional structure by the process of polymerisation. Upon precipitation they irreversibly form gel, which, upon drying, affords amorphous xerogel with a porous structure. The xerogel reforms into a crystallized polycondensate during heating at a temperature of 150 °C. In the condensation reaction, the precursor’s silanol (–Si–OH) groups can also react with the hydroxyl (–OH) groups of fibers forming the covalent (Si–O–C) bonds (Figure 1.13). This strongly increases the adhesion of the polymer film to the fibers and the degree of polymer film orientation on the fiber surface.

Figure 1.12 Organofunctional trialkoxysilanes: (a) alkyldimethyl-3(trimethoxysilyl) propylammonium bromide [132], (b) perfluorooctylated quaternary ammonium silane coupling agent [134].

Figure 1.13 Hydrolysis, polycondensation and chemical binding of QAC-functional trialkoxysilane to cellulose fibers.

Covalent binding of the QAS group to the textile fibers can also be achieved via the 2-chloro-4,6-disubstituted-1,3,5-triazine group of the reactive dyes (Figure 1.14) [141]. In this case, the high reactivity of the 2,4,6-trichloro-1,3,5-triazine group is beneficial for the synthesis of the antimicrobial dye. Although the first and the second chlorines of the 2,4,6-trichloro-1,3,5-triazine group are substituted by the dye chromogen and the QAS group, the third chlorine is still available for substitution by the hydroxyl group of the cellulose fibers under appropriate conditions. This reaction is presented in Figure 1.14.

Figure 1.14 Reactive dye with incorporated alkyldimethyl ammonium bromide chloride.

To improve the reactivity of chitosan, functional acrylamidomethyl groups have been introduced into the primary alcohol groups (C-6), which can form covalent bonds with cellulose in alkaline conditions (Figure 1.15) [145]. Furthermore, chitosan can also strongly bind to cellulose fibers via cross-linking agents, including most polycarboxylic acids (1,2,3,4-butantetracarboxylic and citric acids) (Figure 1.16) [148, 149] and derivatives of imidazolidinone [150]. In the presence of a cross-linking agent, hydroxyl groups of chitosan and cellulose can form covalent bonds with carboxyl groups of polycarboxylic acid in an esterification reaction or with hydroxyl groups of imidazolidinone in an etherification reaction, thus leading to the formation of a cross-link between chitosan and cellulose. The chemical binding of chitosan to cellulose fibers can also be achieved by the oxidation of cellulose fibers with potassium periodate under acidic conditions to form aldehyde groups, which react with the fibers (Figure 1.17) [151].

Figure 1.15 Chemical binding of O-acrylamidomethyl-N-[(2-hydroxy-3-trimethylammonium)propyl] chitosan chloride on the cellulose [145].

Figure 1.16 Chemical binding of chitosan on the cellulose via (1,2,3,4-butantetracarboxylic and citric acids) [148] (left) and derivatives of imidazolidinone [150] (right).

Figure 1.17 Chemical bonding of chitosan to pre-oxidized cellulose fibers [151].

1.3 Low Adhesion Agents

In addition to active antimicrobial agents, microbial growth can be hindered or even prevented by the application of the so-called low adhesion agents. These compounds are not active antimicrobials but are able to create a coating on the fibers, which significantly decreases the attractive interactions between the microorganisms and fibers. This phenomenon has been called a passive antimicrobial activity [65]. There are many factors that influence the adhesion of microorganisms to a solid surface [152], among which the surface free energy (SFE) of the bacterial cells and the solid substrate, surface charge, morphology and topography of the solid substrate are the most prominent [153–155].

When studying the influence of SFE on the bacterial growth, it has been found that the SFE difference between the bacterial cells and the solid substrate represents an important indicator of bacterial adhesion [156, 157]. The lower the difference, the higher the degree of bacterial adhesion. According to literature [153], the bacterial adhesion is minimal when the value of SFE of the solid substrate is in the range of 20–30 mJ/m². Furthermore, it has been shown that the particularly low dispersion part (γLW) of SFE has a tremendous effect on reducing the bacterial attachment. Therefore, the application of low adhesion agents should reduce the SFE of the textile fibers, especially in the case of natural cellulose fibers, which are highly hydrophilic because of the bleaching and mercerisation pretreatment processes. However, by tuning the hydrophobicity of the fiber surface, bacterial adhesion can be either promoted or inhibited depending on the properties of bacterial cells, such as hydrophobicity, zeta potential and abundance of fibrils and fimbriae [154]. From these results, it is clear that a mere preparation of hydrophobic fiber surface is not sufficient to obtain the low adhesion phenomenon. The latter could be achieved only on the superhydrophobic surface of the fibers.

A superhydrophobic surface is characterized as a biomimetic self-cleaning surface constructed with a carefully designed chemical composition and topography [158, 159]. It possesses extremely low surface free energy and dual micro- and nano-structural roughness. Because the self-cleaning mechanism has been previously observed in the leaves of the sacred lotus (Nelumbo nucifera), this effect is referred to as the lotus effect [160–163]. Understanding the complementary roles of surface free energy and roughness on natural non-wetting surfaces has led to the development of a number of biomimetic super- and ultra-hydrophobic surfaces that exhibit apparent contact angles with water greater than 150° and are characterised by low contact angle hysteresis, which leads to roll-off angles lower than 10° [164–169]. When these droplets roll off the solid surface, contaminant particles are picked up by the water and carried away as a result of the interdependence among surface roughness, reduced particle adhesion, and water-repellent properties. The surface topography strongly influences the contact angle hysteresis and, consequently, the sliding angle [170–174]. A superhydrophobic surface with the lotus effect follows the Cassie–Baxter theory of wetting [172]. According to this theoretical model, a water droplet does not entirely wet the rough surface but sits on a composite surface made of solid and air. This state is usually called the fakir state. In this state, a droplet interacts only with the top of the roughness peaks and leaves pockets of air between the droplet and the substrate, the observed contact angle is influenced by the area fraction of the droplet that is actually in contact with the surface. In this case, the solid surface is assumed to be a rough composite surface, and the wetting regime is considered as heterogeneous wetting.

An important class of nonpolar, low-energy surface chemical agents represents functional sol-gel precursors that contain various alkyl and perfluoroalkyl organic groups, among which organotrialkoxysilanes and organofunctional trialkoxysilanes (Figure 1.18) [175–183], organofunctional trichlorosilanes [184, 185], organically modified silicates [178, 186], polydimethylsiloxane and polysiloxane derivatives [65, 187, 188], poly(acrylate-g-siloxane) oligomers [165], and polyhedral oligomeric silsesquioxanes (Figure 1.18) [189, 190] are the most prominent. Because degradation of long perfluoroalkyl groups leads to the release of biopersistent perfluorooctanoic acid or perfluorooctane sulfonate into the environment—both of which can cause serious ecological problems because they have been detected in solids, biota and aqueous systems—these agents have already been removed from the market [191]. When oleophobic properties are unnecessary, the superhydrophobic properties of textile fibers can be successfully obtained with the use of non-fluorinated water-repellent precursors.

Figure 1.18 Alkyltrialkoxysilane (a) and perfluoroalkyltrialkoxysilane (b), polyhedral oligomeric silsesquioxanes (c and d).

Besides the surface free energy and roughness, the surface charge of the solid substrate is an important property that affects the adhesion of microorganisms [154]. Whereas the most bacterial cells are negatively charged, a negatively charged solid surface is more resistant to bacterial adhesion than the cationic one. At this point, it should be stressed that the cationic surface with the positively charged functional groups, such as QAS, acts as an active biobarrier forming antimicrobial agent. Recent research showed that the surface charge could be controlled by the films of zwitterionic compounds (Figure 1.19) [155, 192–202] which can effectively reduce or eliminate bacterial and protein adsorption. Among these compounds, zwitterionic polymers represent the functional polymers which include poly(phosphobetaine), poly(sulfobetaine) and poly(carboxybetaine). It is assumed that the low- or anti-adhesion (low- or non-fouling) properties of zwitterionic compounds are derived from their highly hydrophilic charged structure which contains a positive and a negative charge in the same monomer unit enabling them to bind water molecules via electrostatically induced hydration [193]. This results in the formation of a strongly bound water layer which acts as a barrier to prevent adsorption of proteins on the surface. Zwitterionic compounds can be applied to the solid substrates as self-assembled monolayers (Figure 1.20a) [194, 196, 198] or polymeric brushes (Figure 1.20b) [192, 195, 197, 199–202], where the uniformity of surface coating as well as its strong adhesion to the solid substrate are of great importance for achieving durable anti-adhesion properties. Zwitterionic polymers have already been applied to the textile fibers [194, 202], therefore they offer a great potential for the construction of environment-friendly antifouling textile surfaces.

Figure 1.19 Poly(methacryloyloxyethyl phosphorylcholine) (a) [195], poly(sulfobetaine methacrylate) (b) [192, 195], poly(carboxybetaine methacrylate) (c) [192], siloxane sulfopropylbetaine (d) [194], and L-cysteine betaine (e) [198].

Figure 1.20 Formation of anti-adhesion self-assembled monolayers of L-cysteine betaine (a) [198] and poly(phosphobetaine) brush (b) [199, 201] on the activated solid surface which resists bacteria, represents a live bacterium.

1.4 Dual-Action Antimicrobial Agents

To achieve more efficient antimicrobial activity and higher durability of the coating, antimicrobial agents of different chemical structures and mechanisms of action have been used in combination. The so-called dual-action antimicrobial coatings with the combined controlled-release and biobarrier forming action include two-level dual-functional antibacterial coating with both QAS and silver where silver nanoparticles are embedded into the polymer matrix created by Si-QAS (Figure 1.21) [203], chitosansilver [204, 205], chitosan-zinc [206–208] and chitosan-copper [208] bionanocomposites, polyhexamethylene biguanide-functionalized cationic silver nanoparticles [209], titania matrix precipitated by polyhexamethylene biguanide with the incorporated enzyme (Figure 1.22) [210] as well as N-halamine in combination with QAS (Figure 1.23) [211]. A reactive triclosan derivative with the included amine group which can be converted to a N-halamine structure containing nitrogen-halogen covalent bond was also applied to produce antimicrobial cellulose fibers (Figure 1.24) [212].

Figure 1.21 Silver nanoparticles embedded into the polymer matrix created by Si-QAS [203].

Figure 1.22 The schematic presentation of the encapsulated enzyme in the precipitated titania matrix created on the polyhexamethylene biguanide [210].

Figure 1.23 N-halamine/quaternary ammonium polysiloxane copolymer, poly[3-(5,5-dimethylhydantoinylpropyl)siloxane-co-3-dimethyldodecylammoniumpropylsiloxane chloride] [211].

Figure 1.24 4-(4-chloro-6-(5-chloro-2-(2,4-dichloro-phenoxy)phenoxy)-1,3,5-triazin-2-ylamino)-benzenesulfonic acid sodium salt [212].

Dual-action antimicrobial coatings have also been created by the combination of active antimicrobial and low adhesion agents. These coatings include functional as well as ‘’smart" polymer films. Functional polymer coatings are formed by combining the low surface free energy agent with a controlled release agent, such as silver [65], by combining the low surface free energy agent with a biobarrier forming agent [213] or by combining the low surface free energy agent with both the controlled-release and biobarrier forming agents (Figure 1.25). ‘’Smart" polymer coatings consist of responsive monolayers which are capable of switching between an attractive and a repellent state [154, 214]. One of the most prominent smart polymer which repeatedly switches between its two equilibrium states, a cationic biobarrier forming N,N-dimethyl-2-morpholinone with the attractive function to kill bacteria under dry conditions and a zwitterionic low adhesion carboxy betaine with the protecting function to release and resist dead bacteria under wet conditions (Figure 1.26) [214].

Figure 1.25 Silver nanoparticles embedded into the polymer matrix created by the mixture of Si-QAS, perfluoroalkyltrialkoxysilane and polyhedral oligomeric silsesquioxanes.

represents silver nanoparticles.

Figure 1.26 Two equilibrium states of the smart polymer: cationic N,N-dimethyl-2-morpholinone which kills bacteria (left) and a zwitterionic carboxy betaine which resists live bacteria and releases dead bacteria (right) [214].

represents a live bacterium.

represents a dead bacterium.

1.5 Evaluation of Antimicrobial Activity of Functionalized Textiles

There are many established standard microbiological test methods for the determination of the efficiency of antimicrobial treatments on textiles [2, 3]. Most of them involve measurement of bacteria or fungi growth in vitro and involve either the quantitative or qualitative measurement of microorganism populations. In Table 1.1, their classification is given.

Table 1.1 Classification of standardized microbiological test methods for the evaluation of antimicrobial activity of textiles.

In general, semi-quantitative methods are easily performed but ensure only a comparative assessment of antimicrobial activity between test (antimicrobial treated) textile sample and control (untreated) textile sample. In contrast, quantitative methods reflect accurate results of the antibacterial activity of the treated textiles, thus enabling their classification among those appropriate for hygiene products or those appropriate for use in medicine. However, quantitative methods of evaluation have two main weaknesses: (i) insufficient correlation between laboratory results and actual conditions in the field because most of them include testing against only a single microorganism strain, whereas effectiveness against different types of microorganisms must be ensured in practical use, and (ii) rather poor reproducibility of test results, thus well-trained laboratory personnel are essential for accurate and repeatable results. Regardless of the test method used, the results of antimicrobial activity are obtained in terms of reduced microorganism growth or fiber damage [2].

Among the test methods for the determination of antimicrobial efficiency, the suitability of the selected one directly depends on the mechanism of action of the antimicrobial agent and the type of the tested microorganism. Therefore, in the following section, the well-established standardized microbiological test methods will be reviewed, divided between those for the determination of antibacterial or antifungal activity.

1.5.1 Standardized Methods for the Determination of Antibacterial Activity

Test methods for the determination of antibacterial activity can be generally divided into two types: those based on the agar-based zone of inhibition, which are semi-quantitative; and bacteria colony counting tests, which ensure quantitative evaluation of antimicrobial activity [2, 3]. The most often used standards for the implementation of the tests based on the agar-based zone of inhibition are EN ISO 20645 [215], SN 195920 [216], AATCC 147 [217] and AATCC 90 [218], whereas EN ISO 20743 [219], AATCC 100 [220], ASTM E2149 – 01 [221] and JIS L 1902 [222] are the standards based on colony counting. Both types of methods prescribe testing of antibacterial activity, mostly against Staphylococcus aureus as a Gram-positive strain and Escherichia coli or Klebsiella pneumoniae as a Gram-negative strain.

The agar-based zone of inhibition test method is relatively quick but is mostly suitable for the determination of antimicrobial activity of leaching antimicrobial finishes; it is rarely used for the evaluation of antimicrobials that act according to the mechanism of biobarrier formation. These types of methods include the following steps: preparation of the selected bacteria inoculum; inoculation of agar and placement of the tested textile sample on the inoculated agar plates such that good contact between the sample and agar is obtained (Figure 1.27). Finally, the assessment of the antibacterial activity is obtained by determination of the size of the inhibition zone—the clean area around the sample (Figures 1.28 and 1.29).

Figure 1.27 Preparation of the selected bacteria inoculum (a), inoculation of agar plates (b) and placement of the tested textile sample (c) as prescribed by the agar-based zone of inhibition test method.

Figure 1.28 Bacteria growth on agar plates covered by control textile sample (a) and test textile sample treated by diffusible antimicrobial agent (b), forming the inhibition zone.

Figure 1.29 Microscopic image of nutrient agar after the removal of the control textile sample (a) and test textile sample treated by diffusible antimicrobial agent (b): 1 – printed marks of the fabric; 2 – bacteria colonies; 3 – inhibition zone.

The higher the extent of the inhibition zone, the greater the infusibility of an antimicrobial agent and thus the more effective the antibacterial agent. At this point, it should be stressed that the extent of the inhibition zone cannot be interpreted as a quantitative evaluation of antibacterial activity but is only recommended as a quick test to evaluate the efficiency of an antibacterial treatment [2]. In the case of a non-leaching antibacterial agent, the level of antibacterial activity is assessed by the degree of bacterial growth in the contact area between the agar and the test sample and its comparison with the bacterial growth obtained in the case of the control textile sample. However, this provides a qualitative result on the efficiency of only biobarrier-based antibacterial treatments, which is relatively subjective.

To obtain a quantitative evaluation of antimicrobial efficiency, test methods based on bacteria colony counting are used. In this case, the quantity of bacteria inoculated onto test and control textile samples is assessed at ‘0’ contact time and after a 18–24 h incubation period or other selected time intervals. Subsequently, enumeration of bacteria is obtained by the intensity of luminescence produced by an enzymatic reaction [adenosine triphosphate (ATP) method] or by visual counting of colonies on the agar plate as culture forming units (CFUs). Based on the amount of ATP or number of CFUs determined at ‘0’ contact time and after the 18–24 h incubation period, the antibacterial activity value is determined (Figure 1.30).

Figure 1.30 The quantity of bacterial colonies of E. coli inoculated onto control (a) and test textile samples (b) after 24 h incubation period.

Among the analytical methods for the quantitative evaluation of antibacterial efficiency, the most recent one is EN ISO 20743 standard. The standard includes procedures for evaluation of antimicrobial activity according to the following methods: Absorption method, Transfer method and Printing method, offering the user to select the most appropriate one based on the intended use of the textile product and its surface properties. In addition, the JIS L 1902 standard includes procedures for the performance of the Absorption method and Printing method, whereas there is no great difference between those covered by EN ISO 20743. The Absorption method, which can be easily compared with AATCC 100, prescribes inoculation of the test bacterial suspension directly onto textile samples (Figure 1.31a). In this case, highly hydrophilic textiles are recommended to obtain complete wetting of the test samples with bacteria inoculum, which ensures good contact of the bacteria strain with the antimicrobial agent on/in the fibers. In the case of samples with hydrophobic surface properties, the Transfer method can be used. In this method, test bacteria are first placed on an agar plate and then transferred onto textile samples, which is achieved by pressing the sample against the inoculated agar for a certain amount of time (Figure 1.31b). Therefore, along with antimicrobial activity, the Transfer method also enables an assessment of passive antimicrobial efficiency of fibers as a consequence of the inhibited adsorption of bacteria on the hydrophobic low-adhesion fiber surface. The last method within EN ISO 20743 standard, the Printing method, is an evaluation method in which test bacteria inoculum is filtered through membrane filter paper and subsequently printed onto textile samples by using a special printing apparatus. In this case, textiles with both hydrophilic or hydrophobic surface properties can be tested.

Figure 1.31 Inoculation of the test bacterial suspension onto textile samples according to Absorption method (a) and Transfer method (b).

All aforementioned methods are performed in static conditions, which are not optimal for the evaluation of antimicrobial efficiency of biobarrier-based antimicrobial agents. Namely, in the static conditions where the bacteria inoculum is poured over the test samples or directly transferred, deactivation of the biobarrier can occur owing to the presence of dead bacteria cells, which can also represent an additional source of food for the remaining bacteria, thus influencing the results of antibacterial efficiency. In this case, the use of ASTM standard method E2149–01 is more suitable. This test method was designed to evaluate the bacterial resistance of the non-leaching antimicrobial treatment and is performed under dynamic contact conditions [221]. Namely, good contact between the test textile sample and bacterial suspension is ensured by their constant agitation during the specified period of time. Such constant shaking allows dead bacterial cells, which first made contact with the biobarrier, to be removed from its surface, thus allowing antimicrobial active sites of the coating to freely interact with the remaining bacterial cells in the inoculum.

1.5.2 Standardized Methods for the Determination of Antifungal Activity

Standardised methods for the antifungal activity assessment of textiles mainly prescribe a visual estimation of fungal growth on textile samples or their de-coloration. Such methods are described in the standards DIN 53931 [223], AATCC 30–2004 [224] or SN 195921 [225]. However, the test procedure for quantitative determination of antifungal activity is given only in ISO 13629 [226, 227]. These test methods involve testing the resistance of the antimicrobial treatment against two types of fungi, Aspergillus niger and Chaetomium globosum. In some cases, where mechanical properties are important, test methods also include determination of the mechanical properties of textile samples before and after burial in soil that contains fungi. The most contemporary standard that describes this procedure is EN ISO 11721 [228, 229].

In general, test methods based on visual estimation of antifungal activity are performed by inoculation of agar plates with the spores’ suspension of the tested fungus strain (Figure 1.32) followed by their incubation for a certain length of time under appropriate conditions. After incubation, the test textile sample and control textile sample are placed on an inoculated culture medium and re-incubated (Figure 1.33). At the end of the incubation period, the textile sample are examined in terms of fungus growth intensity on the surface of the test and control textile samples (Figure 1.34) and fungus growth intensity on culture medium which was in direct contact with both textile samples (Figure 1.35). In the first case, the degree of overgrowth of the sample surface is visually determined, whereas in the second case, the intensity of fungus growth is observed by microscopic observation of the culture medium after removal of the test samples from the agar plate; meanwhile, the intensity of spore formation of the tested fungus is qualitatively estimated.

Figure 1.32 Pre-culture of the tested fungus strain (a) and its spores’ suspension (b), prepared by pouring certain amount of sterilized water containing anionic surfactant on the pre-culture and subsequent agitation and filtration.

Figure 1.33 Inoculation of agar plates with the spores’ suspension of the tested fungus strain (a), placement of the test textile samples on the inoculated culture medium (b) and incubation of agar plates for certain period of time (c).

Figure 1.34 Growth intensity of fungus C. globosum on the surface of the control (a) and test textile samples treated by biobarrier – forming (b) and diffusible antimicrobial agent (c).

Figure 1.35 Microscopic observation of the intensity of spore formation of the fungus C. globosum in the culture medium, after removal of the control textile sample (a – strong overgrowth, extensive spore formation) and tested textile samples treated by biobarrier – forming (b – weak, only mycelium) and diffusible antimicrobial agent (c – clean area, no mycelium) from the agar plate.

It should be stressed that for the study of the antifungal activity of the treated textiles, the selection of an appropriate agar medium is crucial. Namely, for example, the DIN 53931 standard prescribes the use of malt-extract agar enriched by oatmeal of high nutrition value, which can result in intensive overgrowth of the tested fungus, preventing the evaluation of antifungal activity. Accordingly, it was shown that the replacement of this highly nutritious culture medium by synthetic nutrient-poor agar was crucial for successful determination of the antifungal activity of silver-treated cotton fabric [230].

ISO 13629 standard specifies a test method for quantitative determination of antifungal activity by either luminescence or the plate count method. According to the test procedure, test and control textile samples are inoculated with a spore suspension of the reference fungus and incubated for a certain length of time under appropriate conditions. Quantitative determination of the antifungal activity is subsequently carried out by the ATP luminescence method or by the colony plate count method. Based on the hydrophilic/hydrophobic surface properties of the test samples, ISO 13629 comprises two methods—i.e., the absorption method, in which test fungus suspension is inoculated directly onto the textile samples, and the transfer method, in which test fungus is placed on an agar plate and printed onto the textile samples.

An important standardized in vitro method EN ISO 11721 includes testing of antimicrobial activity not solely for a single bacterium or fungus strain but a mixture of different microorganisms. This standard specifies determination of the resistance of chemically pretreated textiles to the action of microorganisms present in soil. Because biodegradation of textiles is mostly related to the decomposition action of fungi, this standard was classified among test methods for the determination of antifungal activity. This standard method is applicable to flat textiles made of cellulose-containing yarns (tents, tarpaulins, webbing and tapes) that typically make contact with soil during use. Implementation of the experiment comprises preparation of the test soil and the