Xanthan: Biotechnological Production and Applications

Xanthan: Biotechnological Production and Applications

CHAPTE R 13 Xanthan: Biotechnological Production and Applications Louise C. Candido da Silva*, Brenda N. Targino*, Marianna M. Furtado**, Miriam A. d...

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Xanthan: Biotechnological Production and Applications Louise C. Candido da Silva*, Brenda N. Targino*, Marianna M. Furtado**, Miriam A. de Oliveira Pinto*, Mirian P. Rodarte*, Humberto M. Hungaro* *Federal University of Juiz de Fora, Juiz de Fora, Minas Gerais, Brazil; **University of Campinas, Campinas, São Paulo, Brazil

1 Introduction The global hydrocolloids market is still dominated by polysaccharides extracted from plants and algae (i.e., starch, guar gum, pectin, and alginate). However, polysaccharides produced by microorganisms constitute a growing market and scientific interest in this field is increasing (Freitas et al., 2011; Nwodo et al., 2012). The ability of microorganisms to produce useful compounds has been exploited by humans for centuries. These products can be applied as ingredients in many industries and processes, but they are especially important in the food industry. One example is microbial polysaccharides, which are of an important group of biopolymers synthesized by fungi, yeast, and bacteria. Many bacteria synthesize polysaccharide structures, such as surface lipopolysaccharides, capsular polysaccharides, and exopolysaccharides (EPS) (Becker, 2015). EPS are carbohydrate polymers of high molecular weight that are produced and excreted from bacterial cells. They cover the surface of most cells and play important roles in biological mechanisms, such as the immune response, adhesion, infection, resistance to desiccation, and signal transduction (Bazaka et al., 2011). The composition, structure, biosynthesis, and functional properties of EPS have been extensively studied. Despite the diverse number of bacterial EPS already described, only a few of them have been commercialized as industrial ingredients (i.e., dextran, xanthan, gellan, and curdlan) (Kumar et al., 2007; Poli et al., 2011). Xanthan is currently the most widely accepted commercial bacterial polysaccharide. It is produced on an industrial scale and is used in food, petroleum, and textile, agricultural, and pharmaceutical products. The global xanthan market is constantly expanding and Grand View Research Inc. predicts that it will reach USD 987.7 million in 2020 (Roncˇ evic´ et al., 2017). Microbial Production of Food Ingredients and Additives http://dx.doi.org/10.1016/B978-0-12-811520-6.00013-1


Copyright © 2017 Elsevier Inc. All rights reserved.

386  Chapter 13 Xanthan is produced from bacteria of the genus Xanthomonas, which are important plant pathogens. Its physicochemical characteristics make it an interesting product for encapsulation and film making, and it can also be used as a thickener, viscosifier, stabilizer, emulsifier, and a gelling agent (Faria et al., 2011; Rosalam and England, 2006; Sworn, 2011). Moreover, xanthan has the ability to interact with other polymers and matrix components resulting in the modification of mixture properties. These changes include an increase in viscosity, the formation of strong gels, and an alteration in the charge properties (Sworn, 2011). This biopolymer is also biodegradable, nontoxic, biocompatible, and ecologically safe (Hublik, 2012; Petri, 2015). Xanthan biosynthesis is a complex event that involves the activation of genes located on the gum operon and a high number of enzymes. The process is also energetically demanding. Briefly, the substrate is taken from the culture medium and converted into intermediate nucleotide derivatives, which are then further transformed to monomer units, polymerized into polysaccharide chains, and exported to the extracellular environment (Hublik, 2012). Biotechnological production of xanthan involves preparation of the bacterial inoculum and the substrate, fermentation, cell removal, recovery, and purification of the final product. Xanthomonas campestris is the species most commonly used in the industrial production of this biopolymer; however, other species and pathovars have been evaluated, mainly regarding the growth and production of xanthan from alternative substrates (i.e., agroindustrial waste). Several factors may affect the production yield, molecular structure, and rheological properties of xanthan during fermentation, and among these the culture medium, the physiological state of the bacterial cells, and the conditions for microbial growth are vital (García-Ochoa et al., 2000). Bacterial cells are inactivated or removed from the fermented culture medium by pasteurization or centrifuging, respectively. The addition of organic solvents promotes a reduction of solubility and enhances the precipitation of xanthan. Several downstream processing steps are performed to obtain a final product with desirable qualities. The substrate and recovery are responsible for up to 50% of the costs for producing food-grade xanthan (Palaniraj and Jayaraman, 2011). Further details regarding the structure, properties, and applications of this bacterial biopolymer, as well as microorganism characteristics, biosynthesis, fermentation process conditions, and the challenges to be overcome, are presented in the following sections.

2  Structure and Properties Bacterial EPS represent a wide range of chemical structures containing high molecular weight polysaccharides with a heteropolymeric composition (Kumar et al., 2007). Xanthan is a natural, acidic, extracellular polymer made up of repeated pentasaccharide units (with

Xanthan: Biotechnological Production and Applications   387 molecular weights of 500–2000 kDa), which is produced by phytopathogenic bacteria of the genus Xanthomonas (Dey et al., 2014; Donot et al., 2012; Sworn, 2011). These bacteria can achieve high levels of substrate conversion into EPS because they do not produce significant amounts of other biopolymers (Rehm, 2009). Xanthan is likely produced in response to biotic and abiotic stress factors or as an adaptive response to an extreme environment, and it plays a fundamental role in the survival and pathogenicity of bacterial cells in host plant tissue (Becker, 2015; Donot et al., 2012). This biopolymer is also widely used in several industrial and medical applications because it is natural, nontoxic, biodegradable, rheologically interesting, and less expensive than synthetic polymers (Born et al., 2005; Goswami and Naik, 2014). It is also classified as a generally recognized as safe product by the Food and Drug Administration of the United States, which is fundamental for its application in the food and pharmaceutical industries (Soccol et al., 2013). The molecular structure and conformational state of xanthan are crucial for its rheology, stability, and function (Dario et al., 2011; Renaud et al., 2005). The primary structure is composed of a backbone of β-(1,4)-linked d-glucose units with a trisaccharide side chain on alternate glucose residues. This trisaccharide side chain consists of two mannose units separated by a glucuronic acid (Jansson et al., 1975). The terminal mannose group may carry pyruvate residues and the internal mannose may be acetylated to varying degrees (GarcíaOchoa et al., 2000; Hassler and Doherty, 1990). Xanthan may also be considered as an anionic polyelectrolyte due to its interactions with cations (i.e., Na+, K+, Ca2+, and Mg2+), which are linked to acidic residues in different proportions (Klaic et al., 2016). Moreover, the pyruvate and glucuronic acid groups of the side chain have carboxylic functions that allow chemical modifications to occur in a controlled and specific manner (Roy et al., 2014). Fig. 13.1 shows the primary structure including the sugar residues, the side chain, and the cation binding sites. Xanthan chains are organized into secondary structures, which are aligned with an antiparallel, right-handed, fivefold (5/1) double helix and stabilized by four intramolecular bonds and one intermolecular hydrogen bond (Yui and Ogawa, 1998). This double-stranded helical conformation may undergo order–disorder transitions upon changes in temperature or ionic strength (Bezemer et al., 1993; Jansson et al., 1975; Matsuda et al., 2009). The ordered structure is a rigid single or double-helix strand with persistence length of ∼350 Å, which is formed at low temperature and high ionic strength. The disordered structure is a flexible coiled state with persistence length of ∼50 Å, which occurs at high temperature and low ionic strength (Dario et al., 2011). Order–disorder transitions also depend on the molecular composition of xanthan, particularly with respect to the presence of acetyl or pyruvate groups, which stabilize or destabilize the ordered conformation, respectively (Li and Feke, 2015). Acetyl groups are located closer to the center of the helical structure and tend to stabilize the ordered form, most likely through intramolecular interactions (i.e., hydrogen bonding) between the polymer backbone and the side chains. Conversely, pyruvate groups are located on the

388  Chapter 13

Figure 13.1: Xanthan Primary Structure. Main chain, trisaccharide side chain, and binding sites for cations (M+). From Hublik, G., 2012. Xanthan. In: Moeller, M., Matyjaszewski, K. (Eds.), Polymer Science: A Comprehensive Reference. Elsevier Science & Technology, The Netherlands, pp. 221–229.

periphery of the helical structure and tend to favor the disordered form due to a decrease in electrostatic repulsion when the side chains extend away from the main chain (Viebke, 2005). Therefore a more ordered conformation is obtained by the removal of pyruvate groups, whereas a more disordered conformation is obtained by the removal of acetyl groups. The content of these functional groups in the xanthan molecule can vary with the Xanthomonas strain, medium composition, fermentation conditions, and intentional structural modifications (García-Ochoa et al., 2000; Li and Feke, 2015). The degree of substitution for pyruvate and acetyl groups is usually 30%–40% and 60%–70%, respectively (Thacker et al., 2010). The content of these groups is important because they have a significant influence on the rheological properties of xanthan solutions, especially the viscosity, and also its stability with respect to temperature, solvents, and salts (Kool et al., 2014; Morrison et al., 2004). The presence of salts stabilizes the ordered structure and contributes to the optimal functionality of xanthan, but their effect depends on the concentration and nature of the linked ions (Katzbauer, 1998; Rinaudo, 2001). These ions (especially cations, such as Ca2+, K+, and Na+) can also promote intra- and intermolecular cross-linking and strengthen the structural

Xanthan: Biotechnological Production and Applications   389 network. For example, Ca2+ ions form a complex involving two disaccharide units of the main chain and pairs of carboxyl groups on separate helices, promoting their interactions and thus a viscosity increment in the solution (Mohammed et al., 2007). Xanthan is highly soluble in both cold and hot water, likely due to its polyelectrolyte nature, but it is insoluble in most organic solvents. It is also able to increase the solution viscosity, even at small concentrations, and for this reason it is known as “xanthan gum” (Born et al., 2005). This biopolymer shows higher levels of pseudoplastic rheological behavior than most other common hydrocolloids, which means that its viscosity decreases with an increase in shear rate (Hublik, 2012). Conversely, it shows a low degree of nonthixotropicity, which may affect the instantaneous recovery of the initial viscosity of xanthan after shear thinning. The viscosity also depends on temperature, pH, and concentrations of the biopolymer and salts in the solution (García-Ochoa et al., 2000). The xanthan solution generally retains its stability until 70–80°C under favorable pH and salt concentrations. However, higher ionic strength of the solution may increase the thermal stability of this biopolymer due to a shift in the melting point of the helix strand. Likewise, viscosity remains constant at pH 2–12 in ambient conditions, but may reduce at temperatures above 40°C from acidic or alkaline xanthan solutions during longer storage (Hublik, 2012). Commercial xanthan is a tasteless, dry, white to cream-colored powder with an approximate composition of 8%–15% moisture, 7%–12% ash, 0.3%–1% nitrogen, 1.9%–6.0% acetate, 1.0%–5.7% pyruvate, 3.6–14.3 g/L of monovalent salts, 0.085–0.17 g/L of divalent salts, and 13–35 cP (centipoise) viscosity determined at 15.8 s−1, 1 g/L of xanthan, and 25°C for dissolution and measurement (García-Ochoa et al., 2000). Despite the complexity of its structure, it is possible to modify xanthan (and its properties) through controlled and specific processes. Hublik (2012) stated that modifications of xanthan usually occur at three different stages of the production process: during fermentation, during downstream processing, and postproduction. The functional groups that are more susceptible to structural modifications are acetyl and pyruvate. Removal of specific acetyl groups from the xanthan side chains may be useful to further explore the functionality of this biopolymer (Pinto et al., 2011). Acetyl groups are usually modified or removed from the molecule using genetically modified Xanthomonas strains, using controlling conditions in the fermentation process, or through the use of an alkali treatment, whereby the culture broth is heated at pH 9 or above (Becker et al., 1998; Hassler and Doherty, 1990; Hublik, 2012; Kool et al., 2014; Pinto et al., 2011). Use of monovalent (Na+, K+) or divalent (Ca2+) cations during the fermentation process is another method for xanthan structure modification (Hublik, 2012). Xanthan modifications are also possible during the drying process, where higher temperatures increase water loss and the viscosity of solutions with low electrolyte concentrations. However, xanthan solubility in saline solutions will be compromised if the surface of the

390  Chapter 13 particles become excessively dry. The hydration rate in water can be increased if an emulsifier is added to the xanthan particles (Hublik, 2012). Enzymatic degradation can also be used to alter rheological properties of xanthan; for example, removing the terminal β-d-glucuronosyl residue from the molecule also leads to the loss of the mannosyl side-chain terminus, which generates a more viscous product than the original (Becker et al., 1998). Changing the structural and functional properties of xanthan may also be achieved using controlled mechanical degradation under well-defined physical stress. This process saves energy, is environmentally friendly, and is an effective alternative to enzymatic or chemical modification (Eren et al., 2015; Villay et al., 2012); however, changes in the viscosity and storage and loss modulus may be drastic depending on the homogenization pressure applied (Eren et al., 2015). Degradation products (i.e., oligosaccharides) from intact or modified xanthan have also been extensively used in industrial and food applications and they are considered to be value-added products (Li and Feke, 2015). Degradation products of xanthan have become popular because the reduced molecular weight or particle size is often vital to meet the requirements of the application. Several xanthan degradation techniques have been used including biodegradation (Muchová et al., 2009), thermal degradation (Srivastava et al., 2012), chemical degradation (Christensen et al., 1993), and ultrasonic degradation (Li and Feke, 2015). Knowledge of the chemical composition and structure of intact or modified biopolymers is important for us to understand their rheological properties. It also allows us to propose new applications and perform quality control analysis on end products. Fourier transfer infrared spectroscopy (FT-IR) and proton nuclear magnetic resonance (1H-NMR) are widely used in the analysis of carbohydrate polymers, including xanthan (Roy et al., 2014; Tavallaie et al., 2011).

3  Applications of Xanthan The main industrial applications of xanthan are related to its defining characteristics, such as its high viscosity at low concentrations; its pseudoplastic rheological behavior; its resistance to enzymatic degradation; its high stability in a wide range of pH, temperature, and salt solutions; and its synergism with other polymers (Faria et al., 2011; Hublik, 2012). Xanthan’s qualities mean that it is an ideal rheology control agent in aqueous systems and it can act as a thickener, emulsifier, or stabilizer for emulsions and suspensions. However, its industrial applications are not restricted to these purposes. This biopolymer is suitable for the fabrication of biocompatible and inert matrices (Ramasamy et al., 2011; Raschip et al., 2011; Thacker et al., 2010) and its unique rheological properties mean that it can be used in diverse fields, such as food, agriculture, personal care products, or medicines, and in the textile, chemical, and pharmaceutical industries.

Xanthan: Biotechnological Production and Applications   391 The food industry represents the major field of application for xanthan, and it is used in a wide variety of food products including dressings, sauces, baked goods, dairy products, desserts, beverages, and frozen products (Sworn, 2011). Many of these foods require additional texturization, viscosity, flavor release, appearance, and water-control properties, which may be awarded by biopolymers, such as xanthan. Xanthan has a less “gummy mouthfeel” during chewing than other gums due to its pseudoplastic properties in solutions. The content of xanthan in food formulations is usually 0.05–0.7 wt.% (Zhou and Hui, 2014) and it is generally used in combination with guar and locust bean gums (galactomannans) to increase the desired properties. When these polymers are mixed together the viscosity is increased synergistically, becoming greater than the sum of the individual viscosities. Mechanisms have been proposed to explain the interaction between xanthan and galactomannans; however, there are different views regarding the molecular sites of interaction (i.e., side chains or backbone) and the conformational state (i.e., ordered or disordered) of the polymer molecules, which means that this topic is still open for debate (Grisel et al., 2015). When xanthan is applied to food it is subject to significant variations in pH and ionic strength and it can be influenced by the components of the formulation (Freitas et al., 2011). Some components, such as salts, acids, and proteins can influence xanthan due to changes in the charge of the matrix. For instance, salt concentrations can reduced xanthan viscosity. Hydrated xanthan has good salt tolerance and the addition of 20%–30% salt will not adversely affect the viscosity, therefore it is recommended that xanthan be hydrated in water before the addition of salt. Xanthan viscosity is also reduced at pH < 4 due to the conversion of carboxylate groups from ionized to nonionized form (COO− + H+ = COOH), which subsequently suppresses electrostatic repulsion between the xanthan side chains (Rinaudo and Moroni, 2009). However, the stability of xanthan in acidic conditions is not affected with time, which makes this a useful additive for acidic food products, such as salad dressings and vinaigrettes (Sworn, 2011). Dressings containing xanthan also show long-term emulsion stability, constant viscosity over a wide temperature range, better pourability, and enhanced adherence to the salad (Sharma et al., 2006). Polysaccharides and milk proteins are incompatible and their mixing may result in phase separation; despite this, dairy products, such as puddings, ice cream, ice milk, sherbet, milk shakes, and acidified yogurts are successfully manufactured using xanthan. These products are typically formulated using xanthan concentrations of 0.1%–0.3% and they often contain other gelling and thickening agents (Sworn, 2011). A combination of xanthan with other hydrocolloids (i.e., carrageenan, carboxymethyl cellulose, and galactomannans) produces ideal ingredients for dairy products because they optimize the viscosity, enhance long-term stability, improve heat transfer during processing, enhance flavor release, inhibit syneresis, stabilize emulsions, and assist in ice-crystal control (Grisel et al., 2015; Heyman et al., 2014; Martínez-Padilla et al., 2015; Rosalam and England, 2006; Sharma et al., 2006).

392  Chapter 13 Xanthan interaction with starch can improve the freeze–thaw stability, inhibit retrogradation, and stabilize the starch solution (Arocus et al., 2009; Sworn, 2011). In the bakery industry, xanthan is used to improve water binding, texture, volume, and air incorporation. It can also be used as an ingredient replacement in specific products (i.e., reduced-calorie baked goods and gluten-free breads) without affecting the appearance or taste (Sharma et al., 2006). It is used as a thickener and suspension stabilizer in beverages, contributing to the product appearance and texture, as well as maintaining fruit pulp fibers and insoluble ingredients in suspension (Palaniraj and Jayaraman, 2011). Polysaccharides may also modify the perception and release of volatile and nonvolatile flavor compounds in food and, consequently, determine the acceptance of these products. Xanthan has been evaluated for its capability to mask the astringency of phenolic compounds, allowing the development of antioxidant-rich products without the concomitant taste problems (Troszynska et al., 2010). Besides its extensive applications in the food industry, xanthan has also been used in personal care products (i.e., toothpaste, shampoos, creams, and lotions) to obtain the right consistency, improve the flow, suspend insoluble ingredients, and promote a stable, rich, and creamy lather (Rosalam and England, 2006). Xanthan is also added in agricultural chemical formulations, such as fungicides, herbicides, and insecticides to improve the suspension of actives and to control their spreading and adhesiveness to the plant surface (Flickinger and Draw, 1999). It is also applied in the petroleum industry, especially during drilling, fracturing, pipeline cleaning, workover, and completion (Palaniraj and Jayaraman, 2011). Xanthan is also widely used in human health products and services. It is used as a thickener, emulsifier, or stabilizer in ophthalmic, oral, and topical formulations; as a controlled release agent in tablets; and as a binder in colon-specific drug delivery systems (Ramasamy et al., 2011). These biomedical applications are possible because xanthan is biodegradable and biocompatible in vivo and it is able to form networks (alone or in combination with other polymers) allowing its application as a drug carrier (Petri, 2015). Table 13.1 shows the main applications of xanthan and its function in different areas.

4  Biotechnological Production 4.1 Xanthomonas The genus Xanthomonas belongs to the Xanthomonadaceae family, which resides at the gamma subdivision of Proteobacteria, and encompasses an important ubiquitous group of bacteria that are pathogenic to plants (Buttner and Bonas, 2010). This name derives from Greek, “xanthos” meaning “yellow” and “monas” meaning “entity,” probably because of the pigmentation of the bacterial colonies during growth (Ryan et al., 2011). Xanthomonas was created by Dowson (1939) following a proposal by Burkholder in 1930 for the separation

Xanthan: Biotechnological Production and Applications   393 Table 13.1: Applications of xanthan and its function in different areas. Applications

Usage (%)



Food Salad dressings


Sharma et al. (2006)

Bakery products Beverages

0.05–0.3 0.05–0.2

Prepared foods Soups, sauces, and gravies

0.1–0.3 0.05–0.5

Dairy products


Provides easy pourability, good cling, and forms suspensions Binds water and improves texture Enhances mouthfeel and suspends fruit pulp Stabilizes and avoids syneresis Prevents separation and is thermostable Inhibits syneresis and stabilizes emulsions

Meat products


Binds water and inhibits syneresis

Personal care Toothpaste


Rosalam and England (2006)

Creams and lotions




Provides easy pumpability and gives good stand on Stabilizes emulsions and provides a creamy consistency Controls rheology and suspends insolubles

Industrial Agricultural chemicals


Flickinger and Draw (1999)



Polishes Water-based paints

0.2–0.7 0.1–0.3

Textile and carpet printing




Paper industry


Ceramic glazes Oil drilling

0.3–0.5 0.1–0.4

Enhanced oil recovery


Suspends active ingredients and controls drift and cling Provides pH stability and extends contact time Suspends abrasive components Controls rheology and penetration Improves processing and controls color migration Controls rheology and penetration Aids in suspension and controls rheology Effectively suspends solids Provides good stability against salt and temperature Functions as a mobility control agent

Pharmaceutical Suspensions and emulsions




Sharma et al. (2006)

Sharma et al. (2006) Rosalam and England (2006)

Katzbauer (1998) Byong (1996)

Provides excellent stability and good flow Retards drug release

Source: Palaniraj, A., Jayaraman, V., 2011. Production, recovery and applications of xanthan gum by Xanthomonas campestris. J. Food Eng. 106 (1), 1–12.

394  Chapter 13 of a group of plant pathogens, which were until then assigned to the (now extinct) genus Phytomonas (Sharma et al., 2014). Bacteria were classified according to phenotypic, biochemical, morphological, and pathogenicity characteristics at the time (Simões et al., 2007). This genus has been the subject of numerous taxonomic and phylogenetic studies and reclassifications based on phenotypic and molecular analysis. Most species were characterized at the infrasubspecific level into pathovars based on their distinctive pathogenicity, and individual species can contain multiple pathovars (Giblot-Ducray et al., 2009; Parkinson et al., 2007). These pathovars are pathogenic variants that may infect diverse plant hosts and exhibit different patterns of plant colonization. The Xanthomonas genus is currently comprised of 29 species and 6 subspecies (Fig. 13.2), which may be divided into several pathovars (Euzéby, 2016; Ryan et al., 2011). Members of this genus are short Gram-negative rods of linear shape, which are generally 0.4–0.7 µm wide and 0.8–2 µm long. The bacteria exist singularly or in pairs, and they are motile due to a single flagellum (∼1.7–3.0 µm long). These bacteria have a GC content of 63.3–69.7 mol.%. They are catalase positive, urease, oxidase negative (or weakly positive), nondenitrifying or nitrate reducers, and they can produce H2S but not indole or acetoin. Their growth is inhibited by 6% NaCl; 30% glucose; 0.01% lead acetate, methyl green, or thionin; and by 0.1% (and usually by 0.02%) triphenyl tetrazolium chloride (Saddler and Bradbury, 2015). Proteins are readily digested by these bacteria and some species are able to hydrolyze cellulose, pectin, starch, and Tween 80 (Sharma et al., 2014). They are chemoorganotrophic, able to use various carbohydrates and salts of organic acids as their sole carbon source, and strictly aerobic (as they have respiratory metabolism with oxygen as the terminal electron acceptor) (Saddler and Bradbury, 2015). The main fatty acids found in cells of this genus are 9-methyl decanoic acid (C11:0 iso), 3-hydroxy-9-methyl decanoic acid (C11:0 iso 3OH), and 3-hydroxy-11-methyl dodecanoic acid (C13:0 iso 3OH). This composition of fatty acids serves as a useful criterion to differentiate Xanthomonas from other bacteria (Swings and Civerolo, 1993). The optimal growth temperature for Xanthomonas is 20–30°C depending on the species, with the minimum temperature for growth being >4°C and the maximum being 27.5–39°C (Saddler and Bradbury, 2015). These bacteria ideally grow at pH 6.5–7.5, but anything less than pH 4.5 inhibits growth (Swings and Civerolo, 1993). Xanthomonas metabolize glucose using the Entner–Doudoroff pathway in conjunction with the tricarboxylic acid (TCA) cycle pathway (Palaniraj and Jayaraman, 2011). The pentose phosphate pathway may also be used but this only accounts for a small portion (8%–16%) of the total glucose consumed (García-Ochoa et al., 2000). The glyoxylate cycle may also be used for substrate catabolism and energy production (Petri, 2015). Most Xanthomonas species can grow in chemically defined medium containing minerals, ammonium, nitrogen, a suitable carbon source (i.e., glucose), and amino acids (usually glutamate

Figure 13.2: Unrooted Neighbor-Joining Phylogenetic Tree of Xanthomonas Species Based on 16S rRNA Gene Sequences. Alignment of sequences was performed using CLUSTALW. Bootstrap values are shown at the branch points (based on 1000 replications). Sequence accession numbers for each strain are given in parentheses (genbank data).

396  Chapter 13 or methionine). They also show satisfactory growth on nutrient agar (peptic digest of animal tissue, sodium chloride, beef extract, yeast extract, and agar at 5, 5, 1.5, 1.5, and 15 g/L, respectively) with or without yeast extract supplementation, GYCA medium (glucose, yeast extract, CaCO3, and agar at 10, 5, 30, and 20 g/L, respectively), GPPYA medium (glucose, proteose peptone, yeast extract, and agar at 10, 5, 5, and 20 g/L, respectively), and YM agar (glucose, peptone, yeast extract, and agar at 20, 5, 3, and 17 g/L, respectively) (García-Ochoa et al., 2000; Saddler and Bradbury, 2015). Other culture media containing starch may be used for the general growth of Xanthomonas. These are usually fortified with antimicrobials, such as cephalexin, kasugamycin, chlorothalonil, gentamycin, brilliant cresyl blue, methyl green, and methyl violet to make them selective for Xanthomonas; for more information see Sharma et al. (2014). Features that distinguish Xanthomonas from other related bacteria (i.e., Pseudomonas and Enterobacteriaceae species) include their ability to hydrolyze starch and their independence from using asparagine as a source of carbon and nitrogen (Saddler and Bradbury, 2015; Sharma et al., 2014). Culture media for xanthan production have been widely studied and these may have different characteristics to those intended for bacterial growth. Growth rates of Xanthomonas vary widely across the genus; some strains grow quickly, producing visible colonies in 24–36 h at 25°C, while other strains take 2–3 days. Colonies are usually yellow, smooth, butyrous, and mucoid or viscid (Bradbury, 1984). The yellow color is due to the presence of a membrane-bound pigment “xanthomonadin,” which may protect the bacteria from photobiological damage (Rajagopal et al., 1997). The viscous consistency is provided by xanthan, which is produced by most Xanthomonas strains (Saddler and Bradbury, 2015). This EPS has important biological roles in survival and the ability of Xanthomonas members to colonize a diverse number of ecological niches (Chan and Goodwin, 1999). The adhesiveness, anionic structure, and water-retention capacity of xanthan allows microorganisms to adsorb to biological surfaces, protect itself against desiccation and hydrophobic molecules, concentrate nutrients, and immobilize toxic elements (Coplin and Cook, 1990). This biopolymer can also mask bacteria therefore preventing recognition and attack from plant defense responses. It is also related to the biofilm formation in some species (Buttner and Bonas, 2010). As previously mentioned, xanthan is one of the most important commercial ingredients produced by microorganisms (Petri, 2015). Among Xanthomonas, X. campestris is the most studied species and is most commonly used for xanthan production (Sherley and Priyadharshini, 2015). This species includes several pathovars that cause diseases, mainly within the Cruciferae family (Saddler and Bradbury, 2015). Commercial xanthan is produced by X. campestris pathovar (pv) campestris, particularly the NRRL B-1459 strain (Hublik, 2012); however, other species and pathovars have also been investigated and used for the production of xanthan, including X. campestris pv. mangiferaeindicae, X. campestris pv. arracaciae, X. campestris pv. pruni, Xanthomonas axonopodis pv. manihotis, Xanthomonas melonis, and Xanthomonas arboricola pv. pruni (Borges et al., 2009; Moreira et al., 2001; Rottava et al., 2009).

Xanthan: Biotechnological Production and Applications   397

4.2 Biosynthesis EPS biosynthesis is a multistep process encompassing the synthesis of precursors and monomers, the assimilation of these repeating units, and export of the biopolymer. Each of these steps depends on the activation of several genes and enzymes. The biosynthetic mechanisms responsible for the assimilation and exportation of EPS in Gram-negative bacteria are classified as the Wzx/Wzy-dependent pathway, the ATP-binding cassette transporter-dependent pathway, and the synthase-dependent pathway (Becker, 2015; Schmid et al., 2015). Xanthan biosynthesis is characterized as a Wzx/Wzy-dependent mechanism (Jacobs et al., 2012). In this pathway, assimilation of the heteropolysaccharides from individual repeating units occurs on an undecaprenyl phosphate lipid anchor (e.g., polyisoprenol) on the inner membrane. Several glycosyltransferases (GTs) are responsible for catalyzing this reaction. The Wzx protein (or flippase) then shifts the forming polysaccharide through the cytoplasmic membrane and the Wzy protein performs polymerization once the protein has reached the periplasmic space (Islam and Lam, 2014; Schmid et al., 2015). The genes responsible for producing enzymes involved in xanthan synthesis (e.g., GTs and flippase) in Xanthomonas sp. are clustered in a 16 Kb region with 12 open reading frames termed gumB to gumM (Becker, 2015; Becker et al., 1998; Vorhölter et al., 2008). The biosynthetic pathway of xanthan has been described by several authors, with X. campestris pv. campestris frequently being used as the model of choice (Fig. 13.3). Although the pathway has not been fully elucidated, the current consensus is that it consists of five distinct steps: (I) substrate uptake; (II) conversion of the substrate into nucleotide derivatives; (III) assimilation of monosaccharide subunits; (IV) elongation of the polysaccharide chain (polymerization); and (V) exportation of xanthan to the outer environment (Alkhateeb et al., 2016; Donot et al., 2012; Freitas et al., 2011; Ielpi et al., 1993; Schmid et al., 2015; Vorhölter et al., 2008). The stages of the synthesis process are described in more detail: 1. In the first step the carbohydrate source (preferentially glucose) is internalized from the external environment using specific membrane transporters (Freitas et al., 2011). Nonmodified glucose enters the cells by diffusion, active transport, or as gluconate (via a glucose dehydrogenase mechanism) (Iliev and Ivanova, 2002). 2. Two main mechanisms occur during the sugar–nucleotide activation phase (i.e., production of nucleotide derivatives): direct phosphorylation by glucokinase, or periplasmic oxidation (Schatschneider et al., 2014). During glycolysis, the pentose phosphate cycle and the Entner–Doudoroff pathway are responsible for metabolizing glucose to pyruvate, before pyruvate enters into the TCA cycle (Vorhölter et al., 2008). Pyruvate can also be phosphorylated and converted into nucleotide sugars by a hexose monophosphate pathway (not shown in Fig. 13.3) (Ielpi et al., 1993). Glucose-6phosphate dehydrogenase is specific for carbohydrate metabolism in Xanthomonas sp., and is responsible for producing the nucleotide precursor sugars (NDP-sugars) (i.e., UDP-

Figure 13.3: Biosynthetic Pathway of Xanthan in X. campestris. (I) Substrate uptake; (II) glucose conversion into nucleotide derivatives; (III) assembly of oligosaccharide subunits; (IV) polymerization; and (V) exportation of the EPS to the outer environment. The transfer of glucosyl-1-phosphate (from UDP-glucose) to the polyprenol carrier is catalyzed by GumD. GumM, GumH, GumK, and GumI give sequence to the xanthan repeat pentasaccharides unit assembly. GumL pyruvyltransferase and the GumF and GumG acetyltransferases are responsible for the added nonsugar decorations to the mannose residues. GumJ translocates the assembled pentasaccharide unit to the periplasmatic face of the cell membrane. GumE catalyzes the subunits polymerization. The polysaccharide is then exported out of the cell, potentially catalyzed by a complex formed by GumB and GumC. Ac, Acetyl group; Ac-CoA, acetyl-CoA; Glc, glucosyl group; GlcA, glucuronyl group; Man, mannosyl group; PEP, phosphoenolpyruvate; Pyr, pyruvyl group.

Xanthan: Biotechnological Production and Applications   399 glucose, UDP-glucuronate, and GDP-mannose) required to construct the pentasaccharide repeating unit in xanthan (Becker and Vorhölter, 2009; Becker et al., 1998; Freitas et al., 2011; Iliev and Ivanova, 2002). The genes xanA and xanB are responsible for encoding the enzymes involved in the synthesis of these NDP-sugars, which occurs in a branched pathway (Vorhölter et al., 2008). 3. During the assimilation of the monosaccharide subunits, pentasaccharide repeating units are constructed from glucose, mannose, and glucuronic acid in a molar ratio of 2:2:1 (glucose–glucose–mannose–glucuronate–mannose) (Harding et al., 1993; Vorhölter et al., 2008). The synthesis of these repeating units requires GT enzymes encoded by the GumD, GumM, GumH, GumK, and GumI genes (Ielpi et al., 1993; Vorhölter et al., 2008). The repeating units are synthesized by the sequential transfer of monosaccharides from NDP-sugars by GTs. These monosaccharides are connected through a phosphate anchor linkage to a polyisoprenol lipid carrier located in the inner membrane of the cell (Becker and Pühler, 1998; Freitas et al., 2011; Ielpi et al., 1993). This lipid carrier is a long-chain phosphate ester and isoprenoide alcohol, which provides an anchor to the extracellular membrane, organizes the formation of the carbohydrate chain, and stimulates the transport of this chain through the cell membrane (Donot et al., 2012). In order to synthetize monosaccharides repeating units, a UDP-glucose molecule must transfer glycosyl-1-phosphate to the polyisoprenol phosphate transporter. The sequential transfer of d-mannose from GDP-mannose and d-glucuronic acid from UDP-glucuronic acid will complete the assembly of the lipid-linked pentasaccharide unit (Rosalam and England, 2006). Pyruvate and acetyl groups (donated by phosphoenolpyruvate and acetyl-CoA, respectively) are added to the mannose residues in the repeating unit by the pyruvyltransferase (GumL) and the acetyltransferases (GumF and GumG) (Becker, 2015; Becker et al., 1998; Freitas et al., 2011; Ielpi et al., 1993). Pyruvate is added to the terminal mannose and acetyl groups attach to the internal mannose (Donot et al., 2012). In the next step, the flippase (GumJ, the equivalent of the Wzx protein) flips the lipid carrier with the repeating unit from the cytoplasmic face of the inner membrane to the periplasmic face (Becker, 2015; Schmid et al., 2015). 4. Once polyprenol-linked pentasaccharide repeating units are at the periplasmic face of the membrane, polymerization can take place through the actions of the polymerase, Wzy. Studies have demonstrated that the gumE gene product is responsible for the transfer of immature xanthan polymers to the newly translocated repeating units (Becker, 2015; Freitas et al., 2011; Ielpi et al., 1993; Vorhölter et al., 2008). 5. EPS is then exported to the cell surface (Becker, 2015; Schmid et al., 2015). Interaction between GumB and GumC proteins is necessary to complete xanthan polymerization and trigger exportation. GumC is a polysaccharide copolymerase located at the inner membrane of the cell, while GumB is an outer membrane polysaccharide export protein located in the subcellular membrane fraction (Bianco et al., 2014; Schmid et al., 2015). GumC and GumB can form pores at the external

400  Chapter 13 membrane, which allows exportation of mature xanthan (Vorhölter et al., 2008). Further understanding of this process will depend on in-depth characterization of the GumB and GumC proteins, since the complete mechanism and function of the enzymes involved in this step are not fully elucidated (Bianco et al., 2014; Donot et al., 2012). Genomic data and genetic engineering are tools that can be used to identify these genes and the enzymes encoded by them, thus aiding in our understanding of the xanthan biosynthesis process. Genetic approaches along with research into the influence of production factors and substrate sources (addressed in the next chapters) are tools that can help to improve the desirable characteristics, such as the yield, chemical structure, and rheological proprieties of xanthan.

4.3  Factors Influencing Production Xanthan production is a complex bioprocess modulated by a multitude of factors (GarcíaOchoa et al., 2000). Variations in the operational conditions during fermentation or downstream processing may influence not only the production yield but also the chemical structure and the rheological properties of the biopolymer (Barua et al., 2016; Borges and Vedruscolo, 2008; Casas et al., 2000; Khosravi-Darani et al., 2011; Lopes et al., 2015). Media composition, temperature, pH, agitation, air flow rate, duration of fermentation, bioreactor type, operational mode (batch or continuous), recovery, purification, and drying are all factors that can influence (directly or indirectly) xanthan production (Borges and Vedruscolo, 2008; de Mello Luvielmo et al., 2016; Faria et al., 2011; GarcíaOchoa et al., 2000; Lopes et al., 2015; Rehm, 2010). The strain of Xanthomonas also plays a key role in consumption of the substrate, production yield, and the chemical composition of xanthan (i.e., the monosaccharide sequence, condensation linkages, and acetyl and pyruvate group incorporation) (Becker, 2015; Becker et al., 1998; GarcíaOchoa et al., 2000; Ielpi et al., 1993). All of these characteristics can influence the quality and application of the biopolymer. Production of xanthan which is of low quality and minimal yield may be attributable to a lack of functional groups in the substrate’s molecular structure, nutritional deficiency of the culture medium, an excess of nonreacted compounds, or the formation of additional by-products. The physiological state of the cells may also cause alterations in the side-chain structures, the molecular weight, and the yield of xanthan, but it does not influence the primary backbone structure (Rosalam and England, 2006). Xanthan biosynthesis exhibits secondary metabolite fermentation kinetics in two different phases: in the first phase the cells grow rapidly but produce little xanthan (trophophase), while in the second phase (idiophase) the cell growth reduces drastically and large amounts of xanthan are produced (Donot et al., 2012; Shu and Yang, 1990). As they are distinct metabolic phases the optimal conditions may vary between them. A two-stage process

Xanthan: Biotechnological Production and Applications   401 that uses two separate sets of conditions to optimize cell growth and xanthan production, respectively, could improve the fermentation process (Shu and Yang, 1990). The economic importance of xanthan to many industrial processes has stimulated research into how to obtain a product with appropriate characteristics for each specific use. Table 13.2 shows some of these recent studies in which xanthan production was evaluated in different fermentative conditions. 4.3.1  Culture medium The composition of the culture medium that provides the nutrients for microbial growth plays an important role in the biosynthesis, molecular structure, and production yield of xanthan (Rosalam and England, 2006). The minimum nutritional requirements for the biosynthesis of xanthan are the sources of carbon, nitrogen, and micronutrients (i.e., phosphate, potassium, magnesium, iron, and calcium) (Brandão et al., 2010; Casas et al., 2000; García-Ochoa et al., 2000; Hublik, 2012; Khosravi-Darani et al., 2009; Lopes et al., 2015). The culture medium represents 20%– 30% of the total cost of the process, therefore optimization of its type and the concentration of its components (mainly the carbon source) can reduce the costs and lead to production of a higher quality biopolymer (Borges and Vedruscolo, 2008; Casas et al., 2000; García-Ochoa et al., 1998; KhosraviDarani et al., 2011). Carbon sources commonly used in the production of this biopolymer are carbohydrates, such as glucose, sucrose, and starch. In an attempt to reduce production costs, many researchers have begun to assess alternative carbon sources for xanthan production. Examples of these alternative sources and the amount of xanthan obtained (g/L) include sugar beet molasses (53), residue from apple juice (45), sugar cane molasses (17.1), cassava wastewater (13.8), cocoa dry pods (4.9), cheese whey (16.4), and date palm juice (24.5) (Brandão et al., 2010; Diniz et al., 2012; Druzian and Plagiarini, 2007; Gilani et al., 2011; Kalogiannis et al., 2003; Niknezhad et al., 2015; Salah et al., 2010). However, glucose and sucrose are still the best carbon sources in terms of yield and quality of the final product (Rosalam and England, 2006). Despite research developments in this field, much of the knowledge about the flow of carbon through the central metabolic pathways in Xanthomonas is specific to glucose. Thus the use of alternative sources demands further research in this regard. EPS biosynthesis is generally favored and regulated by an excess of carbon, concomitant with limitation of another nutrient, such as nitrogen (Freitas et al., 2011; Quinlan, 1986). However, higher levels of carbon may inhibit cell growth through substrate excess or catabolic repression. Glucose concentrations of 30–40 g/kg prevent this effect and maintain xanthan production, but it has been reported that glucose (and other carbon sources) concentrations above 50 g/L may inhibit the growth of Xanthomonas (Niknezhad et al., 2015). The type of carbohydrate may also affect the amount of xanthan produced, most likely due to the synthesis of enzymes and transporters and activation of specific

Table 13.2: Xanthan production by several Xanthomonas strains in different fermentations conditions. Strain

Fermentation Parameters

Medium Composition


X. campestris NRRL B-1459 (parent strain) X. campestris CCTCC M2015714 (mutant from parent strain)

Substrate (g/L) Inoculum (%) Temperature (°C) pH Aeration (vvm) Stirring rate (rpm) Time (h)

Sucrose, glycerol, crude glycerol 10 30 7a 0.4 600 90

Carbon source Fish peptone Yeast extract NaNO3 FeSO4 MgSO4 KH2PO4 K2HPO4 Antifoam

10–80 3 1 0.8 0.01 2.5 2 3.5 0.5

X. campestris pv. manihoti 280-95

Substrate (g/L) Inoculum (%) Temperature (°C) pH Aeration (vvm) Stirring rate (rpm) Time (h)

Sucrose 10 28.5 7–12a 1.5 500 120

Sucrose MgSO4·7H2O KH2PO4 H3BO3 (NH4)2SO4 FeCl3 CaCl2·2H2O ZnSO4 Citric acid

50.0 0.20 5.0 0.006 2.0 0.0024 0.002 0.002 2.0

Recovery Method

Main Results


Only the mutant strain (M2015714) grew and produced xanthan with glycerol or crude glycerol as the sole carbon source. Xanthan production (g/L) for B-1459 and M2015714 with sucrose (40 g/L) was 23.2 and 23.1, respectively. Xanthan production (g/L) for M2015714 with glycerol (40 g/L) was 11.0 and with crude glycerol (40 g/L) was 7.9. M2015714 xanthan had a smaller molecular weight, higher transparency, and lower viscosity compared to the commercial sample. Centrifugation Xanthan production was enand precipitahanced (from 9.4 to 16.5 g/L) tion using etha- by a 24 h alkali stress process. nol 92.6 degree However, alkali stress negativeGL (4:1) ly influences viscosity and this converted the native polygonlike structure of xanthan to a star-like form.

Wang et al. (2016)

Centrifugation and precipitation using cold ethanol (3:1)

de Mello Luvielmo et al. (2016)

Strain 23 wild species of Xanthomonas campestris (isolated from different leaves and fruits)

Fermentation Parameters Substrate (g/L) Inoculum (%) Temperature (°C) pH Aeration (vvm) Stirring rate (rpm) Time (h)

Xanthomonas campes- Substrate (g/L) Inoculum (%) tris LRELP-1 Temperature (°C) pH Aeration (vvm) Stirring rate (rpm) Time (h)

Maltose, glucose, sucrose 10 28–30 7–7.5b — 110–150 96

KWH 10 30 7.00 1 300 120

Medium Composition Carbon source NH4H2PO4 K2HPO4 MgSO4·7H2O

(g/L) 50.0 1.5 2.5 0.1



Recovery Method Centrifugation and precipitation using chilled alcohol 96% (3:1)

Centrifugation and precipitation using 95% ethanol

Main Results Among the 23 tested strains, 16 isolates (70%) were positive for xanthan production. XStl, isolated from the Citrus macroptera plant, was the best gum producer (18.29 g/L). The best yield was achieved using sucrose as the carbon source, pH 7, 28°C, and speed rotation of 150 rpm. FTIR studies suggested that XStl was suitable for large-scale xanthan production Successful batch fermentation production of xanthan gum using KWH as the sole substrate. Maximum xanthan production, reducing sugar conversion, and utilization rates were 11.73 g/L, 67.07%, and 94.82%, respectively. The logistic and Luedeking–Piret kinetic models are capable of describing the batch fermentation process. KWH xanthan had thermal stability equivalent to the commercial xanthan.

References Barua et al. (2016)

Li et al. (2016)


Table 13.2: Xanthan production by several Xanthomonas strains in different fermentations conditions. (cont.) Strain X. campestris PTCC1473 X. pelargonii PTCC1474

Fermentation Parameters Substrate (g/L) Inoculum (%) Temperature (°C) pH Aeration (vvm) Stirring rate (rpm) Time (h)

Cheese whey 5 28 7b — 250 48

Medium Composition Cheese whey KH2PO4 MgSO4·7H2O (NH4)NO3 Citric acid H3BO3 ZnCl3 FeCl3 CaCl2

(g/L) 20,50, and 80 5, 10, and 15 0.06, 0.12, and 0.182 2 0.006 0.006 0.0024 0.02

Recovery Method Centrifugation and precipitation using 0.1% calcium chloride in isopropanol

Main Results References Successful development of Niknezhad et al. a mathematical model for (2015) xanthan production, based on RSM results. The concentration of the carbon source was the factor that influenced the production of xanthan the most. PTCC1473 xanthan concentration (16.4 g/L) was achieved using 65.2 (cheese whey), 14.8 (KH2PO4), and 1.1 (MgSO4·7H2O) g/L. PTCC1474 xanthan concentration (12.8 g/L) was achieved at 80, 6.7, and 0.8 g/L of cheese whey, KH2PO4, and MgSO4·7H2O, respectively. X. campestris produced a xanthan yield of 0.42 and X. pelargonii produced a yield of 0.27 (g of xanthan/g of lactose)

Strain Xanthomonas campestris mangiferaeindicae 2103

Xanthomonas campestris pv. Manihotis 1182 Xanthomonas campestris pv. campestris 254 Xanthomonas campestris pv. campestris 629

Fermentation Parameters Substrate (g/L) Inoculum (%) Temperature (°C) pH Aeration (vvm) Stirring rate (rpm) Time (h)

Substrate (g/L) Inoculum (%) Temperature (°C) pH Aeration (vvm) Stirring rate (rpm) Time (h)

CGB 20 28 7b 0.5–1.5 300–700 120

SSAE, Sucrose (control) 10 28 7b — 250 120

Medium Composition CGB (NH2)2CO KH2PO4 antifoam

SSAE Sucrose Urea K2HPO4

(g/L) 10, 20, 40, and 60 0.10 1.0 1.0

20–100 20 1 10

Recovery Method Centrifugation and precipitation using 98% ethanol (3:1)

Main Results CGB optimized fermentation at 0.97 vvm and 497.76 rpm resulted in xanthan production of 5.59 g/L, with 255.40 mPa/s of apparent viscosity at 0.5% (m/v). Variations in the stirring rate influenced the production of xanthan gum, biomass concentration, apparent viscosity, and glucose, mannose, and pyruvic acid concentrations. Crude glycerin products have the potential to be used for the efficient and cost-effective production of xanthan. Centrifugation The SSAE nutrient contents, and precipitacarbon (1.337%), and nitrogen tion using etha- (49.74%) were adequate for nol 96 degree cell growth and xanthan bioGL (3:1) synthesis. The highest viscosity (99.96 mPa·s) and yield (2.64 g/L with 2% SSAE) was achieved by Xanthomonas campestris 1182 after 120 h of fermentation. The yield and viscosity obtained from SSAE were superior to sucrose for all tested strains. SSAE extract can be used as an efficient and cost-effective substrate for xanthan production.

References de Jesus Assis et al. (2014)

de Sousa Costa et al. (2014)


Table 13.2: Xanthan production by several Xanthomonas strains in different fermentations conditions. (cont.) Strain X. campestris ATCC 13951 S. elodea ATCC 31461 Paenibacillus elgii B69

Xanthomonas campestris pv. campestris NRRL B-1459

Fermentation Parameters Substrate (g/L) Inoculum (%) Temperature (°C) pH Aeration (vvm) Stirring rate (rpm) Time (h)

Substrate (g/L) Inoculum (%) Temperature (°C) pH Aeration (vvm) Stirring rate (rpm) Time (h)

Glucose 10 30 7b — 250 48

SCB 10 28 7.5a 0.35 750 24

Medium Composition Glucose Yeast extract K2HPO4 MgSO4·7H2O

(g/L) 30 3 2 0.1

Recovery Method Centrifugation and precipitation using ice-cold ethanol (2:1)

SCB Yeast extract NH4NO3 Na2HPO4 KH2PO4 Antifoam

27.0 2.0 0.8 2.5 2.5 0.5

Centrifugation and precipitation using absolute ethanol and KCl 30% (3:1)

Main Results References XC350G155-19 mutant strain Li et al. (2014) was derived from X. campestris ATCC 13951 by repeated exposure to ampicillin. The EPS productivity and viscosity increased with increasing numbers of serial passages and AMP concentrations. The stability test indicated that there was good stability and heritability of the mutation (caused by ampicillin), even after 30 generations of growth in a nonantibiotic medium. FT-IR and 1H-NMR confirmed Faria et al. (2011) that xanthan produced using SCB as the substrate was similar to commercial xantham. The total sugar content was 85.3%, the monosaccharide composition was 43% glucose, 32% mannose, 24% glucuronic acid, in a ratio of 1.79:1.33:1.

Fermentation Strain Parameters Xanthomonas campes- Substrate (g/L) Inoculum (%) tris PTCC1473 Temperature (°C) pH Aeration (vvm) Stirring rate (rpm) Time (h)

Xanthomonas sp. C1 and C9

Substrate (g/L) Inoculum (%) Temperature (°C) pH Aeration (vvm) Stirring rate (rpm) Time (h)

Date extract with NH4NO3 — 28 — — 200 72

Glycerol, sucrose, glycerol:sucrose 10 28 7b — 180 96

Medium Composition Date extract NH4NO3 H3BO3 MgCl2 Na2SO4 H3BO3 ZnO Fe2Cl3·6H2O CaCO3 FeSO4 HCl

Carbon source NH4H2PO4 K2HPO4 H3BO3 (NH4)2SO FeCl3 CaCl2·2H2O ZnSO4

(g/L) 40, 50, and 60 0.2, 0.25, and 0.3 2.1 0.507 4.6 0.006 0.006 0.020 0.020 0.008 0.13 mL

50.0 2.5 5.0 0.006 2.0 0.0024 0.002 0.002

Recovery Method Centrifugation and precipitation using isopropanol (3:1) and NaCl (1 g/L)

Main Results Concentrations of date extract higher than 50 g/L caused a decrease in cell growth and xanthan production. The highest cell growth and xanthan production (11.2 g/L) were obtained using 40 g/L of the carbon source. Nitrogen source concentration did not impact on cell growth but the C/N ratio did interfere with xanthan production. Date extract can be used as an alternative and cost-effective carbon source for xanthan production, since it is capable of supplying the nitrogen needs of the cell. Centrifugation The productivity with glycerol and precipitawas 0.157 for C1 and 0.186 for tion using etha- C9. When glycerol and sucrose nol 92.6 degree were used 0.363 and 0.363 g/L GL (3:1) were obtained for C1 and C9, respectively.

References Khosravi-Darani et al. (2011)

Reis et al. (2010)


Table 13.2: Xanthan production by several Xanthomonas strains in different fermentations conditions. (cont.) Strain Xanthomonas arbicola pv. juglandis (Ceviz 1) X. axonopodis pv. vesicatoria (XCVA3-1) X. axonopodis pv. begonia (Xcb-9) X. axonopodis pv. dieffenbachia (Xad-2) X. campestris NRRL B-1459

Fermentation Parameters Substrate (g/L) Inoculum (%) Temperature (°C) pH Aeration (vvm) Stirring rate (rpm) Time (h)

Glucose 10 30 7a 1 200 72

Medium Composition Glucose Citric acid KH2PO4 MgCl2 Na2SO4 H3BO3 ZnO FeCl3·6H2O CaCO3

(g/L) 40.0 2.1 2.866 0.507 0.089 0.006 0.006 0.020 0.020

Recovery Method Centrifugation and precipitation using isopropanol (2:1)

Main Results References Ceviz 1 showed the highest Gumus et al. gum productivity (8.22 g/L), (2010) followed by Xcb-9 (7.74 g/L), the control bacterial NRRL B-1459 (7.46 g/L), and finally XCVA3-1 (6.40 g/L). The Xad-2 strain did not produce xanthan. The highest viscosity value (428 mPa·s at 1% solution) was obtained from the XCVA3-1 strain.

CGB, Crude glycerin biodiesel; KWH, kitchen waste hydrolysate; RSM, response surface methodology; SCB, sugarcane broth; SSAE, shrimp shell extract. a pH was maintained constant throughout the fermentation process. b pH was only adjusted at the beginning of the fermentation process.

Xanthan: Biotechnological Production and Applications   409 catabolic pathways. For example, high yields of xanthan have been obtained from glucose, sucrose, maltose, and soluble starch (Leela and Sharma, 2000); however, lactose shows low productive when compared with other carbon sources, mainly because the Xanthomonas have limited β-galactosidase activity (Frank and Somkuti, 1979; Fu and Tseng, 1990). The nitrogen source also plays an important role in microbial growth, xanthan production, and the structural characteristics of the xanthan molecule. This element may be added to the culture medium as an organic or inorganic substance, such as ammonium salts (i.e., nitrate, phosphate, sulfate, and chloride), sodium nitrate, urea, yeast extract, peptone, casein hydrolysates, soybean whey, glutamate, or additional amino acids (Palaniraj and Jayaraman, 2011; Souw and Demain, 1979). Conflicts exist as to whether (NH4)2HPO4 or glutamate is the best nitrogen source for the growth of Xanthomonas and xanthan production (Cadmus et al., 1978; García-Ochoa et al., 2000; Souw and Demain, 1979). High concentrations of nitrogen are needed for rapid cell growth; however, the rheological properties of xanthan may be affected when there is an excess of this element (Casas et al., 2000; Sutherland, 1982). Experimental data suggest that excess nitrogen reduces the concentration of pyruvate in the xanthan molecule; while the acetate content is not affected (Borges and Vedruscolo, 2008). The ratio of nitrogen to carbon is more important than the concentration of each element alone. This ratio influences the microbial growth regulation and the production of metabolites of commercial interest (i.e. xanthan). The C:N ratio during the production phase should be lower than the ratio in the growth phase (García-Ochoa et al., 2000; Souw and Demain, 1979). Phosphate is also an important component of the culture medium. Phosphate influences cell growth, buffers the culture medium, and affects xanthan production (Silva et al., 2009). Low phosphate concentrations may affect the composition of xanthan, causing a reduction in the pyruvate content of the polymer structure (Davidson, 1978), while high phosphate concentrations (>50 mM) may inhibit xanthan production (Souw and Demain, 1979; Umashankar et al., 1996). The magnesium composition of the culture medium also influences cell growth and the production and structure of xanthan. Magnesium acts as a cofactor for many enzymes, plays an important role in the activation of sugar uptake systems, and is present in the cell wall and membrane (Niknezhad et al., 2015). 4.3.2 Temperature Temperature is one variable that is directly related to the xanthan production process. An optimal temperature of 28–30°C is chosen for EPS production in most bacterial strains (Giavasis, 2013). A range of temperatures (20–38°C) have been tested in studies that evaluated the influence of this variable on xanthan production (García-Ochoa et al., 2000; Gomashe et al., 2013; Psomas et al., 2007), and most found that 28°C was optimal for cell growth and xanthan yield (Borges and Vedruscolo, 2008; García-Ochoa et al., 2000).

410  Chapter 13 Conversely, Shu and Yang (1990) reported different optimal temperatures for cell growth (24–27°C) and xanthan production (30–33°C). When cells are grown at 25°C there is an increase in the number of acetyl groups, molecular weight, and viscosity (Casas et al., 2000; Giavasis, 2013; Lopes et al., 2015). Since the optimal temperatures for cell growth and xanthan production are different, it may be advantageous to use a two-stage incubation to optimize the fermentation process (Shu and Yang, 1990). 4.3.3 pH pH also has an optimum value for cell growth and EPS accumulation. It is generally assumed that neutral pH is optimal for polysaccharide synthesis and microbial growth during the fermentation process (Barua et al., 2016; Lopes et al., 2015). Some studies have reported that pH 6–8 is suitable for xanthan production, but this production is drastically reduced when the pH drops below 5 (Casas et al., 2000; Esgalhado et al., 1995; Gumus et al., 2010; Sherley and Priyadharshini, 2015). The precise control of pH is only possible in benchtop reactors when a buffer or a base (i.e., NaOH) can be added to the growth medium (Sherley and Priyadharshini, 2015). The pH may decrease during the fermentation process due to the presence of acid groups in the xanthan molecule (García-Ochoa et al., 2000); however, Psomas et al. (2007) observed an increase in pH (7–10) after 72 h using their agitation and temperature conditions. de Mello Luvielmo et al. (2016) evaluated the effect of alkali stress on the yield, viscosity, structure, and ultrastructure of xanthan. They concluded that alkali stress can induce the cells to increase EPS production as a protective mechanism against adverse conditions. However, a lower viscosity was observed when compared to the alkali stress-free gum, regardless of the alkali stress time. The data indicate that there are still issues to be elucidated regarding the effect of pH on the xanthan synthesis. 4.3.4  Stirrer speed and air flow rate Bacterial EPS are produced under suitable aerobic conditions. The aerobic properties of Xanthomonas dictate that high levels of oxygen are necessary to achieve efficient xanthan production (Donot et al., 2012; Freitas et al., 2011). The production rate increases considerably when the oxygen transfer rate is increased (Psomas et al., 2007); however, the viscosity of the culture medium also increases (Borges et al., 2008; Khosravi-Darani et al., 2011; Seviour et al., 2011; Shu and Yang, 1990). This increase in viscosity reduces the aeration rate of the culture medium and reduces the homogeneity and nutrient availability (Palaniraj and Jayaraman, 2011). Therefore an accurate control of the air flow rate and stirrer speed is desirable to avoid stress (Borges and Vedruscolo, 2008). Optimization of mechanical mixing (i.e., using different paddle configurations or altering the stirring rate) can be used to minimize this problem; however, these methods can change

Xanthan: Biotechnological Production and Applications   411 the properties of the biopolymer and the increase in mechanical stress can lead to the cell rupturing (Freitas et al., 2011). Oxygen limitation exists at low stirring rates (100– 300 rpm), but when the rates are increased (above 500 rpm) hydrodynamic stress damages the rheological properties of xanthan (Borges and Vedruscolo, 2008; Casas et al., 2000). Although high stirring rates impair the efficiency of the process, a relatively high agitation speed is essential to fulfill the oxygen requirements in viscous media. Alternatively, it is possible to gradually increase the agitation speed throughout the fermentation process while increasing viscosity (Seviour et al., 2011). 4.3.5  Duration of fermentation process The duration of fermentation is closely related to the cell growth kinetics and the amount of substrate available for bioconversion (Gilani et al., 2011; Letisse et al., 2001). The fermentative process will cease once the cell decline phase is reached or once the substrate is exhausted. Thus it is only possible to maintain xanthan synthesis for long periods of time using continuous culture (Seviour et al., 2011). Experimental data has demonstrated that the fermentation time influences the average molecular weight by changing the degree of pyruvilation and acetylation in the xanthan molecule. Acetyl and pyruvate content of the xanthan molecule and average molecular weight increases with time, regardless of the other operational conditions (Casas et al., 2000; Psomas et al., 2007; Shu and Yang, 1990; Tait et al., 1986). 4.3.6  Operational production mode: batch or continuous Xanthan can be produced through batch or continuous fermentation. Batch-scale fermentation takes ∼80 h and achieves 75%–80% conversion efficiency (carbon source into xanthan) (Rosalam and England, 2006; Vuyst et al., 1987). However, the growing viscosity during the fermentation process affects oxygen and nutrient availability (Sherley and Priyadharshini, 2015). Moreover, the environmental conditions can change throughout the fermentation cycle and could lead to adverse conditions, such as nutrient exhaustion, extreme pH, and toxic products (Rosalam and England, 2006). Continuous fermentation may be used as an alternative to batch fermentation (Vuyst et al., 1987). The culture medium is continuously added to the culture vessel to maintain the optimal conditions and provide a steady supply of nutrients (Rosalam and England, 2006; Sherley and Priyadharshini, 2015). A conversion rate of 60%–70% can be reached in continuous fermentation (Becker et al., 1998). The fermentation time can be reduced and nutrient use by microorganisms can be improved, reducing the overall cost of xanthan production (Vuyst et al., 1987). Nevertheless, the increased risk of contamination and the threat from fast-growing mutants that do not synthesize xanthan are the main challenges to be overcome before this process is used industrially (Becker et al., 1998; Seviour et al., 2011; Sherley and Priyadharshini, 2015; Vuyst et al., 1987).

412  Chapter 13

4.4  Recovery of Xanthan The recovery of microbial EPS is a critical step that determines the cost and functional properties of the final product. It is often based on isolation and purification processes developed in plants and seaweeds (Pace and Righelato, 1980). The downstream processing should not degrade xanthan and should include inactivation or removal of bacterial cells, precipitation of the biopolymer, washing, drying, milling, and packing (García-Ochoa et al., 2000; Palaniraj and Jayaraman, 2011). Its main objectives are to concentrate the fermented broth to facilitate transport and storage; to reduce or eliminate contaminants, such as salts, bacterial cells, solid particles, and undesirable enzymes; and to improve the functional proprieties, color, odor, taste, and microbiological stability of the final product (Smith and Pace, 1982). Several methods (i.e., chemical, thermal, and mechanical treatments) have been developed to lyse, inactivate, or remove cells from the fermented broth. Chemical treatments often use hypochlorite or alkali compounds, which can cause depyruvylation of xanthan when applied at elevated pH (García-Ochoa et al., 2000; Palaniraj and Jayaraman, 2011). Thermal treatments (i.e., pasteurization or sterilization) are often the methods of choice for inactivating bacterial cells and enzymes in industrial processes. They also improve xanthan dissolution and reduce the viscosity of the fermented broth, thus facilitating cell removal. However, these treatments should be conducted under appropriate conditions (80–130°C, 10–20 min, pH 6.3–6.9) in order to avoid thermal EPS degradation (García-Ochoa et al., 2000; Palaniraj and Jayaraman, 2011; Smith and Pace, 1982). Xanthan clarification can be carried out by protease-mediated protein degradation prior to pasteurization. The production of cell-free xanthan is achieved by mechanical treatments, such as filtration or centrifugation of the fermented broth (Hublik, 2012; Rosalam and England, 2006). However, these treatments may be affected by the high viscosity of the medium, which should be reduced through dilution or heating (Rosalam and England, 2006). Enzymatic lysis of the bacteria can also be used to solve this problem; however, the enzymes must be removed after they have been used, which increases the recovery process costs (Palaniraj and Jayaraman, 2011). Xanthan must be extracted from the fermented broth once the bacterial cells have been inactivated or removed. One of the most common methods is precipitation, which reduces the solubility of the EPS through the addition of organic solvents, such as ethanol, acetone, and isopropyl alcohol; salts; or a mixture of salts and alcohol (García-Ochoa et al., 2000). These solvents can also be used to wash out impurities including colored components, salts, and cell debris (Palaniraj and Jayaraman, 2011). The quantity of solvent used depends on the nature of the reagent. For example, total xanthan precipitation is possible when two to three volumes of isopropyl alcohol or acetone are added per volume of fermented broth

Xanthan: Biotechnological Production and Applications   413 (Gumus et al., 2010; Rottava et al., 2009; Salah et al., 2010; Silva et al., 2009; Zhang and Chen, 2010); however, more than four volumes of ethanol are required per volume of fermented broth (Borges et al., 2008; de Mello Luvielmo et al., 2016). Using a combination of alcohol and salt is one way to increase the precipitation of xanthan and reduce the volume of reagents required. This occurs due to a reversal of charges effect by ionic bonds between cations of salt and anionic groups of the biopolymer resulting in decreased affinity of the latter for water (García-Ochoa et al., 1993, 2000; Rosalam and England, 2006). The polyvalent cations from calcium, aluminum, and quaternary ammonium salts are more effective for xanthan precipitation than monovalent cations, such as sodium or potassium (Palaniraj and Jayaraman, 2011). The solvents used to precipitate xanthan in the industrial processes may be collected from the solid–liquid separation steps, and recovered by distillation for reuse (Hublik, 2012; Rosalam and England, 2006). Ultrafiltration (UF) is an alternative method for xanthan recovery. This method recovers the biopolymer by passing the fermentation broth through membranes of defined porosity and does not change the rheological properties and molecular weight of xanthan. UF is also considered to be an energy saving, environmentally safe, and cost-effective method (Lo et al., 1996). The precipitated biopolymer should be separated from the fermented broth, washed, and dried to obtain the final purity required. The specific purification methods should be chosen in accordance with the proposed application. For example, if the final product is to be used as a food additive it should be free of bacterial cells and solvents; however, if it is to be used in the textile industry the requirements are less restrictive (García-Ochoa et al., 2000; Palaniraj and Jayaraman, 2011). Xanthan can be obtained by tangential-flow filtration for industrial nonfood applications (Becker et al., 1998). Wet xanthan can be redissolved and washed with a KCl solution to reduce the viscosity. This process can be repeated until the desired quality is achieved. The precipitate may be treated with chemical compounds before the drying process to inactivate cellulases and improve dispersion of the final product (Hublik, 2012; Smith and Pace, 1982). Precipitated xanthan is then dried in batch or continuous dryers, at low temperatures under a vacuum, or with forced circulation of an inert gas to prevent combustion of the organic solvent in the precipitate. The drying conditions should be chosen to avoid chemical and thermal degradation, excessive coloring, and changes in the density, dispersibility, and solubility of xanthan. The dry biopolymer can be milled to the desired mesh size in order to control the dispersibility and dissolution rates (Kumar et al., 2007); however, this step should be conducted at controlled temperatures to avoid excessive heating and the consequent darkening or degradation of the product (García-Ochoa et al., 2000). The final product

414  Chapter 13 should be packaged into a material with low-water permeability once it is hygroscopic and subjected to hydrolytic degradation (Rosalam and England, 2006; Smith and Pace, 1982). Most commercial xanthans have a final moisture content of about 10% (Faria et al., 2011).

5  Industrial Production Following the discovery of dextran in the early 1940s, extensive research on microbial polysaccharides was performed at the Northern Regional Research Laboratories of the US Department of Agriculture in the 1950s. The EPS produced by X. campestris NRRL B-1459 was found to have the most interesting properties (including all those mentioned previously), which allowed it to compete with other natural and synthetic gums. In 1960 the Kelco Company undertook a pilot plant for xanthan production, which became the second microbial polysaccharide to be industrially produced and this biopolymer was commercialized about 10 years later (Kang and Pettitt, 1993; Pace and Righelato, 1980). As previously mentioned, the major substrates used in the production of xanthan are carbon and nitrogen sources. Most commercial xanthan is obtained by the fermentation of glucose or invert sugars (Becker et al., 1998; Faria et al., 2010; Rosalam and England, 2006). Ammonium or nitrate salts, casein hydrolysates, peptone, soybean whey, or yeast extract are frequently used as the nitrogen source (Palaniraj and Jayaraman, 2011). The commercial production of xanthan starts with the preparation of X. campestris inoculum using a suitable culture medium. This stage involves several steps that require a set of reactors ranging from 10 L (for the initial inoculation) to 100,000 L (in the production process) (Rosalam and England, 2006). The fermentation process is conducted in sterile mechanically-agitated reactors, under monitored and controlled conditions. Batch processing is preferred because fewer parameters need to be controlled. The temperature is usually established at 28–32°C and the pH at 6.5–7.5, depending on the selected strain. The aeration rate should be higher than 0.3 (v/v), and the specific power input for agitation should be higher than 1 kW/m3 (Hublik, 2012; Palaniraj and Jayaraman, 2011; Rosalam and England, 2006). The final concentration and productivity of xanthan depends on the oxygen supply. Industrial batch fermentations achieve xanthan productivities of 0.4–0.7 g/L/h and concentrations of 3%–5%. As mentioned previously, one of the major problems in the industrial production of xanthan is the growing viscosity of the culture during fermentation, which affects the availability of oxygen and nutrients (Hublik, 2012). After fermentation, processes for recovering and purifying xanthan are applied. Fig. 13.4 depicts the process for the industrial production of xanthan including the downstream processing steps.

Xanthan: Biotechnological Production and Applications   415

Figure 13.4: An Example of the Xanthan Industrial Process and Its Multiple Downstream Processing Steps. From Rosalam, S., England, R., 2006. Review of xanthan gum production from unmodified starches by Xanthomonas comprestris sp. Enzyme Microb. Technol. 39 (2), 197–207.

6  Prospects and Challenges Xanthan is one of the most important commercial microbial EPS for the food and nonfood industries. The commercial success of this biopolymer is based on its rheological properties, which allow it to be used as a thickening, emulsifying, or stabilizing agent in a wide range of products and processes (i.e., in the food, agrochemical, and cosmetics industries; in personal care products; and in the oil drilling process). Xanthan has been widely studied and our understanding of its biosynthesis, biotechnological production, molecular structure, and rheological properties has expanded. However, several challenges must still be overcome to increase the industrial production and applications of this biopolymer. Substrates and downstream processing techniques are responsible for the high production costs. The use of

416  Chapter 13 agricultural and industrial wastes as a carbon source is an interesting alternative to reduce the production costs; however, the successful conversion of some of these nutrient sources into xanthan is still a challenge. Optimization of the fermentation conditions and mutagenesis or genetic manipulation of Xanthomonas strains to obtain higher yields may help to circumvent this issue. The metabolic engineering and overexpression of certain genes may also lead to the assimilation of substrates not used previously, the control of the structure and rheological properties of xanthan, and an increase in the rate of synthesis. Another important challenge is to optimize the downstream processes in order to successfully remove the microbial cells from the fermentation broth and reduce the use of solvents for the precipitation of xanthan. The use of immobilized cells, ultrafiltration, and the addition of salts in the precipitation have been investigated to address this issue. Modified xanthan and its derivate products have proved to be interesting compounds due to the changes in the structural characteristics and rheological properties achieved with the use of these products. The bacterial polysaccharides market is likely to grow further in the coming years. The inclusion of these new products and their applications will create niches where the traditional hydrocolloids are not able to compete.

Acknowledgments The co-authors thank the financial support of Fundação de Amparo à Pesquisa do Estado de Minas Gerais (FAPEMIG) for their research projects into the Microbial Production of Food Ingredients area.

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