Table of Contents Author Guidelines Submit a Manuscript
International Journal of Polymer Science
Volume 2016 (2016), Article ID 2062360, 12 pages
http://dx.doi.org/10.1155/2016/2062360
Review Article

Alginate Biosynthesis in Azotobacter vinelandii: Overview of Molecular Mechanisms in Connection with the Oxygen Availability

Escuela de Ingeniería Bioquímica, Pontificia Universidad Católica de Valparaíso, Avenida Brasil 2147, Casilla, 4059 Valparaíso, Chile

Received 28 August 2015; Revised 11 January 2016; Accepted 7 February 2016

Academic Editor: Mukund Adsul

Copyright © 2016 Ivette Pacheco-Leyva et al. This is an open access article distributed under the Creative Commons Attribution License, which permits unrestricted use, distribution, and reproduction in any medium, provided the original work is properly cited.

Abstract

The Gram-negative bacterium Azotobacter vinelandii can synthetize the biopolymer alginate that has material properties appropriate for plenty of applications in industry as well as in medicine. In order to settle the foundation for improving alginate production without compromising its quality, a better understanding of the polymer biosynthesis and the mechanism of regulation during fermentation processes is necessary. This knowledge is crucial for the development of novel production strategies. Here, we highlight the key aspects of alginate biosynthesis that can lead to producing an alginate with specific material properties with particular focus on the role of oxygen availability linked with the molecular mechanisms involved in the alginate production.

1. Introduction

Increasing research on the mechanisms of synthesis and biochemical properties of biopolymers, such as polysaccharides, led to improving production process and new applications in diverse areas, mainly in food and pharmaceutical industries [1]. One of the main advantages for the use of biopolymers is its degradability, making them a renewable product option. However, the high costs of biopolymer production are still a major drawback for a widespread industrial application [2].

A particular linear polysaccharide with broad growing interest is alginate, which is a structural component of the brown marine algae and the cell wall of bacteria belonging to the Pseudomonas and Azotobacter genera [35]. The properties of alginates in solution largely depend on four factors: (a) its monomer chemical composition (β-D-mannuronic acid (M-residues) and its epimer, α-L-guluronic acid (G-residues)); (b) the sequence pattern of the monomers; (c) the molecular weight (MW) of the resulting polysaccharide chain; and (d) modifications of the polymer (acetylation degree) [6, 7].

However, algal alginates are complex mixtures containing polysaccharides with a wide range of MW and ratios of M : G. Hence alginates with specific defined M : G ratios or a constant range of MW cannot be easily obtained from particular algae species, due to intrinsic environmental culture conditions, thus limiting their use in the pharmaceutical and chemical industries (more details in Section 2). For this reason, the bioprocesses research area has become interested in developing strategies to produce alginates with particular molecular characteristics through microbial alginate production. In contrast to algal alginates, microbial alginates present exclusive M-residue acetylation, controlled M : G ratios, and specific MW under specific growth conditions [810]. A nonpathogenic bacterium able to produce alginate with high production yields in bioreactors is Azotobacter vinelandii. Yet, the complex regulatory pathways controlling the alginate biosynthesis and material properties in response to external environmental clues remain still unknown, despite some efforts in trying to gain new insights into gene expression patterns under different culturing conditions in A. vinelandii cultures [9, 1113].

In this review, we present an up-to-date biosynthetic overview of microbial alginate biosynthesis from Azotobacter vinelandii, and the perspectives for production process improvement based on a better understanding on the molecular mechanism underlying polymer biosynthesis in relationship with the oxygen availability during the fermentation process.

2. Alginate Structure, Chemical Structure, and Applications

Over the past 40 years, a growing interest in the use of alginate has been observed including different areas, ranging from genetics to pharmaceutics (Figure 1).

Figure 1: (a) Number of publications indexed in Scopus database (August 2015), using keyword alginate (−) in title, abstract, or keywords. (b) Percentage of the word alginate distributed in different subject areas.

Alginate has been placed as the second biopolymer derived from seaweeds with greater demand in the hydrocolloids’ industry [14]. Currently, the only economic way to obtain commercial alginate used for most applications is through its extraction from marine algae, the cost of which ranges between US 2 and 20/kg, and with a total market value of around US 339 million [14]. Furthermore, alginates of very high purity are used in the pharmaceutical industry where they are sold for up to US 3,200/kg.

Since alginate is a biodegradable and a biocompatible polysaccharide, it presents a panoply of food, pharmaceutical, and biotechnological applications (Figure 1(b)). In the food and pharmaceutical industries, alginate is mainly used as a stabilizing, thickening, or gel-film-forming agent [6, 1517], Table 1; in medicine it is used as wound healing material [18], as part of medical treatments [19, 20], or as dietary fiber supplements [21, 22]. Alginate showed potential beneficial physiological effects in the gastrointestinal tract [23]. Moreover, hydrogel-alginates are being investigated in biotechnology as drug delivery agents, as cell encapsulation material, and as scaffold material in tissue engineering [24].

Table 1: Summary of biotechnological and pharmaceutical applications of alginates based on their molecular weights.

Alginate is the main structural component of brown marine algae (Laminaria and Macrocystis) representing about 32% of dry biomass [25], consisting in variable amounts of M-, G-, and MG-residues, linked by 1→4 glycosidic bonds [7]. On the other hand, alginates produced by bacteria are submitted to esterification with O-acetyl groups at the O-2 and/or O-3 of the M-residues [26], where the majority of the M-residues are mono-O-acetylated, and infrequently with 2,3-di-acetylated [27] (Figure 2). Because the monomeric chemical structure of bacterial alginate and the sequence length determine the mechanical properties of the alginates, one of the aims of different investigations is the possibility of manipulating the composition alginates for specific applications have been intensively investigated [28, 29].

Figure 2: Representation of the chemical structure from acetylated alginates produced by Azotobacter vinelandii bacterium [28]. Mannuronic (M) and guluronic (G) acid residues are represented in the alginate chain.

The obligate aerobe bacterium Azotobacter vinelandii produces alginate that acts as a diffusion barrier for nutrients and oxygen [30, 31]. It was reported as a bacterium with a highest respiratory rate [32], implying that it adjusts oxygen consumption rates in order to maintain low levels of cytoplasmic oxygen and in this way permitting the oxygen-sensitive enzymes to be active, like nitrogenase, which is responsible for fixing nitrogen [30, 32].

A. vinelandii under limitation of carbon source or by induction forms cysts that are more resistant to desiccation and is mainly composed of alginate [33, 34]. It also accumulates the intracellular polyester poly-β-hydroxybutyrate (PHB) as a reserve carbon and energy source [35, 36].

Consequently, an increased knowledge about the molecular mechanism involved in alginate biosynthesis will be crucial for the development of novel strategies to improve the production of alginates with defined characteristics tailored for specific applications.

3. The Biosynthetic-Secretory Route of Alginate Production in Azotobacter vinelandii

Microbial polysaccharides have distinct biological functions, as intracellular storage, as envelope, or as extracellular polymers [37]. Microbial alginate is an extracellular polysaccharide as xanthan, cellulose, and sphingan, among others, and they differ in their biosynthetic pathways routes (recently reviewed in Schmid et al. 2015 [37]). Moreover, alginate is secreted trough a secretion system shared among the Gram-negative bacteria [38].

The alginate biosynthesis in bacteria Azotobacter results from a complex regulatory network of proteins, similar to Pseudomonas genera [6, 28, 39].

All of the steps involved in the conversion of central sugar metabolites into the alginate precursor in A. vinelandii have been previously identified and characterized [6, 40]. The alginate precursor, GDP-mannuronic acid, is synthesized from fructose-6-phosphate to mannose-6-phosphate by the bifunctional enzyme phosphomannose isomerase (PMI)/guanosine-diphosphomannose pyrophosphorylase (GMP), designated as AlgA, encoded by the algA gene. A phosphomannomutase (AlgC) directly converts the mannose-6-phosphate into mannose-1-phosphate, which is in turn converted into GDP-mannose by the AlgA enzyme. GDP-mannose is oxidized to GDP-mannuronic acid by GDP-mannose dehydrogenase (AlgD, encoded by algD gene). Because the intracellular levels of GDP-mannose are high and because it is used in different pathways, it has been proposed as the limiting step of alginate biosynthesis in P. aeruginosa [41].

After the production of the polymer precursor GDP-mannuronic acid precursor, its polymerization and transport across the cytoplasmic membrane is carried out by proteins presumably integrating a cytoplasmic membrane complex (polymerase complex). The core of the polymerase complex is composed of the glycosyltransferase Alg8 protein and Alg44 protein [4244]. Furthermore, the protein AlgK is thought to stabilize the polymerase complex, by interacting with Alg44 [43]. Highlighting the important role of this protein, alginate polymerization does not occur in the absence of algK [42, 45].

The polymannuronate polysaccharide resulting from polymerization and then translocation to the A. vinelandii periplasm is composed of M-residues, which can then be further modified during its passage across the periplasm [43]. These modifications consist in acetylation, epimerization, and degradation of the M-residues. More specifically, the polymannuronic molecule undergoes an O-acetylase modification, which is catalyzed by an acetylase enzymatic complex composed of AlgI, AlgV (AlgJ in P. aeruginosa), AlgF, and AlgX proteins [4648]. While M-residue O-acetylation does not occur frequently in alginate, some may be acetylated. O-acetylated M-residues will therefore be protected from epimerization [26], because only nonacetylated M-residues can be epimerized to G-residues by the AlgG epimerase [42], so alginates with a relatively high degree of acetylation display a lower degree of epimerization [27].

Alginate depolymerization occurs at the 4-O-glycosidic bond via β-elimination, by alginate lyases which have been the subject of a recent review [28]. The Azotobacter vinelandii genome encodes six enzymes with alginate lyase activity [31]: the alginate lyase AlgL [49], the bifunctional mannuronan C-5 epimerase and alginate lyase AlgE7 [50], the three AlyA(1–3) lyases [51], and an exolyase, AlyB, that is still uncharacterized [28].

Some of the nonacetylated M-residues are then epimerized to G-residues by the bifunctional AlgG epimerase, which converts poly(β-D-mannuronate) to α-L-guluronate. In P. aeruginosa, AlgG is also part of the periplasmic protein complex that serves as a scaffold for leading the newly formed alginate polymer through the periplasmic space to the outer membrane secretin AlgE porin (AlgJ in A. vinelandii) [52]. A scaffold complex helps to transport the recently modified polysaccharide throughout the periplasm towards AlgE before secretion to the extracellular milieu. This complex is thought to be composed of AlgG, AlgK, and AlgX proteins and possibly AlgL [40, 42, 43, 52]. The exported polysaccharide could be then epimerized by seven extracellular Ca2+-dependent epimerases (AlgE1–7) [53]. Based on these evidences, Figure 3 shows a schematic representation of the alginate biosynthetic steps in A. vinelandii.

Figure 3: Schematic representation of the alginate biosynthetic steps in Azotobacter vinelandii, from evidence-based protein-protein interaction in P. aeruginosa [28, 42, 43]. The biosynthetic alginate pathway is represented as two complementary stages: on the left, the synthesis of the substrate precursor (GDP-mannuronic acid) and its following polymerization, including transfer from cytoplasm; on the right, the modification (periplasmic and extracellular) of the nascent polymer, as well as the export through the outer membrane of the polymer.

4. Genetic Regulation of Alginate Biosynthesis in Azotobacter vinelandii

In Azotobacter vinelandii the alginate biosynthetic gene cluster is arranged as an operon (Figure 4), containing genes coding for enzymes involved in the synthesis of the alginate precursors, as well as those involved in its polymerization, degradation, acetylation, epimerization, and secretion. The availability of the complete genome sequence of A. vinelandii [31] also contributes to the better knowledge of this organism.

Figure 4: Genetic structure genes involved in alginate biosynthesis and modification in Azotobacter vinelandii. Gene operon for alginate biosynthesis algD-A, and algC gene is transcribed separately; alyA13 and alyB alginate lyases encoding genes, and algE17 the epimerases genes.

Several promoters controlling alginate gene cluster transcription have been described: algDp1 ( promoter), algDp2 (AlgU dependent promoter), and algDp3 promoters, all located upstream of algD [54, 55], alg8p promoter, upstream of alg8 [44], and a promoter for sigma 70 located upstream of algG [49]. In addition, two putative promoters algCp1 and algCp2 are situated upstream of algC gene (Figure 4) [56].

The alginate biosynthetic gene cluster expression is controlled by algUmucABCD gene cluster, where algU encodes the alternative sigma factor (AlgU), essential for alginate production [57]. Moreover, AlgU is responsible for transcription driven by the algCp1 and algDp2 promoters (Figure 5), but it does not control the algL or the algA genes, as described for P. aeruginosa [55].

Figure 5: Regulation of alginate biosynthetic genes in A. vinelandii (modified according to reference [40]). Promoters are indicated as banners; mRNAs are indicated as dotted boxes; solid lines indicate the reported mechanism of regulation, and dashed lines indicate unknown mechanism of gene regulation; arrows indicate positive regulation and T-shaped bars indicate negative regulation. OM: outer membrane; PG: peptidoglycan; IM: inner membrane. See text for a more detailed description.

The MucA and MucC proteins negatively regulate alginate production, acting as anti- factors [54]. MucA represses AlgU protein activity, thus suppressing algD transcription from the algDp2 promoter. In contrast, algU gene transcription is autoregulated by AlgU interaction and activation of its promoter locus (algUp2) (Figure 5) [54].

Additionally, expression of the algD promoters is controlled by the global two-component system GacS/GacA, which is conserved among Gram-negative bacteria [58]. The GacS/GacA system controls alginate biosynthesis [58], where GacS controls the expression of algD from its three promoters [58]. Accordingly, mutations in gacS and gacA significantly reduce the algD transcript levels [58]. GacA not only is a positive regulator of the biosynthesis of alginate and PHB [58] but also regulates alginate biosynthesis through activation of the small regulatory RNAs, Rsm (rsmZ1 and rsmZ2). These RNAs interact with the rsmA protein, which binds algD mRNA and thus acts as a transcriptional repressor [59]. The A. vinelandii genome encodes nine small RNAs belonging to the Rsm posttranscriptional regulatory system (rsmZ17 and rsmY1-2) (Figure 5) [59].

Despite the great efforts to understand the alginate biosynthetic gene regulation, little is known about how cultivation conditions could modify gene transcription in A. vinelandii.

5. Alginate Production in Azotobacter vinelandii Cultures: The Balance of Alg8 and AlgL by Oxygen Availability

The glycosyltransferase Alg8 protein belongs to the glycosyltransferase type II family and is localized in the inner cell membrane [78]. The glycosyltransferase type II enzyme family catalyzes the transfer of glycosyl residues to an acceptor molecule, during biosynthesis of polysaccharides, such as the cellulose or chitin synthase [79].

In both Azotobacter vinelandii and Pseudomonas aeruginosa the alg8 gene encodes the Alg8 protein [44]. In P. aeruginosa it has been demonstrated that by adding additional copies of alg8 it is possible to increase alginate production by at least 10 times [80], suggesting that this protein might be involved in a rate-limiting step of alginate production. As a consequence, the possibility of manipulating Alg8 protein levels in A. vinelandii may be a valuable approach for increased alginate production, although this has not being done so far. The attempts to reach high Alg8 protein levels were by manipulating the alg8 gene expression via culture conditions. However, it is important to note that alginate production in A. vinelandii is a multienzymatic and complex process.

Moreover, the Alg44 protein acts as link between Alg8 and the AlgJ alginate exporter protein [42, 43]. Since Alg44 has a c-di-GMP intracellular binding domain, it was suggested that this protein presents a regulatory role [81]; although the c-di-GMP levels might not have an impact neither on Alg44 stability nor on its localization, it still seems to be required for the activation of Alg8 [42, 43].

Interestingly, in A. vinelandii batch, cultures controlling the dissolved oxygen tension (DOT) at 1% present higher levels of alg8 and alg44 gene expression, when compared with control cultures (5% DOT) [9]; the authors suggested that this behavior can in turn enhance the MW of the alginate produced under low DOT conditions. Moreover, in continuous cultures under non-nitrogen-fixation conditions at different agitation rates (300, 500, and 700 rpm) and different sucrose concentration in the feed medium, the highest alginate MW (obtained at 500 rpm) is correlated with the highest alg8 expression [12], suggesting that alg8 gene expression can be modulated by not only oxygen availability but also carbon source feed rate, as well. The oxygen availability here is perceived as the amount of oxygen needed for full oxidation of carbon source, taking into account the oxygen transfer rate as well as the DOT level in cultures [82]. Meanwhile, in chemostat cultures under nitrogen-fixation conditions, operated at a dilution rate of 0.07 h−1, expression of both alg44 and alg8 was affected by changes in agitation rate (400, 500, and 800 rpm), implying that the activity of both genes could be controlled by oxygen availability [13]. Although the highest alginate MW was obtained at 500 rpm, this was not correlated with higher alg8 gene expression, which was obtained at 800 rpm. The differences between the two-chemostat culture conditions might be explained by the activation of the nitrogenase protection machinery (non-nitrogen-fixation versus fixation), where the higher alginate MW have directly linked to the alg8 gene expression under nonfixing conditions. This notion agrees with the fact that nitrogenase activity protects cells from oxygen, thus fostering alginate production [30, 83]. Other possible explanation given is that the culture condition might activate the genes coding for alginate lyases, further discussed in this review. However, more studies are needed, especially those involving gene expression and proteomics profiles during A. vinelandii cultures in order to have a better insight of alginate polymerization step.

A possible link among the low specific oxygen uptake rate (), the MW of the alginate synthesized, and alg8 gene expression was found [11]. This work suggests that when the value increases by double, the MW of alginate decreases (about 1.6 times), while alg8 relative expression decreases around sixfold. Moreover, in cultures carried out in continuous mode operated at dilution rate 0.08 h−1, when the value was 2.2 mmol g−1 h−1, both the alginate MW and alg8 gene expression levels were higher than those obtained in cultures in which the value was double [11]. The same correlation between low value and highest alginate MW was reported [12], where a slight increment of 1 in the lead to a reduction in the MW of the alginate produced by A. vinelandii (from 1200 to 500 kDa). Furthermore, in this condition, the lyase-encoding gene algL increased its expression by threefold while alg8 expression decreased by ninefold. Interestingly, for values below 2 mmol g−1 h−1 [12] or exceeding 5 mmol g−1 h−1 [9, 13], the changes in the alginate MW were not correlated with alg8 or algL gene expression levels. Table 2 summarizes the major changes observed on both the alginate MW and gene expression levels, during the small increment values over the specific oxygen uptake rate of A. vinelandii cultures.

Table 2: Molecular weight of alginate and relative gene expression of alg8 and algL with respect to the variations.

Furthermore, the Azotobacter vinelandii genome encodes six enzymes with alginate lyase activity [31]: the alginate lyase AlgL [49], the bifunctional mannuronan C-5 epimerase and alginate lyase AlgE7 [50], and the three AlyA(1–3) lyases [51].

The AlyA1, AlyA2, and AlyA3 belong to the PL7 polysaccharide lyase family, containing an alginate lyase module, linked to three calcium-binding modules [28, 51]. AlyA1 and AlyA2 are more likely to be periplasmic (AlyA1, UniProtKB-M9YEJ6; AlyA2, UniProtKB-C1DHI8) whereas the AlyA3 protein has secreted signal C-terminal domain (AlyA3, UniProtKB-C1DQS5), which is needed for efficient germination in A. vinelandii [51]. In chemostat cultures, conducted at dilution rate of 0.07 h−1 with agitation of 500 rpm, highest alginate MW was reported [13]. In this condition, an increment in the agitation rate (from 400 to 600 rpm) leads to an increment in the lyase-encoding genes alyA1, algL, and alyA2 by twofold.

The algGXLIVFA operon encodes the AlgL protein responsible for the periplasmic alginate lyase activity in A. vinelandii. Disruption of the algL gene generated a strain that overproduces alginate, suggesting that this enzyme is important for alginate biosynthesis [84]. Furthermore, the increase in algL expression was not correlated with a decrease in alginate MW in chemostat cultures [12]. However, algL gene expression pattern could also be affected by the (manipulated by changes in the agitation rate) in chemostat. Supporting this observation, chemostat cultures also showed an increase in algL gene expression (around eightfold) together with higher MW alginate production [11, 12]. By using an A. vinelandii mutant strain carrying algL::WGm nonpolar mutation [84] and culturing under 3% of DOT, no alterations were found in alginate lyase activity in culture broth comparing with the wild-type strain. However, alginates with a high MW were obtained [85], suggesting that the lower MW of the alginate correlates with the higher alginate lyase AlgL activity.

In A. vinelandii ATCC 9046 strain cultures carried out at 1 and 5% DOT, the expression of higher alginate lyase genes (algL, alyA1, alyA2, alyA3, and algE7) correlated with the lower DOT and with the higher MW alginate production [9]. In these conditions (1% DOT), the intracellular and extracellular lyase activities were lower, comparing with cultures grown at 5% DOT, suggesting that dissolved oxygen affected the activity of the alginate lyases and/or their gene expression. However, the alginate lyase activity (intracellular and extracellular) seemed to be associated with the exponential phase of the cultures, where, in the ATCC strain cultured, the maximum of alginate lyase activity was found in the prestationary phase and dropping in the stationary phase [9, 85].

As stated previously (Table 2), in cultures with between 2 mmol g−1 h−1 and 5 mmol g−1 h−1 [9, 1113], the activity of intracellular lyases, namely, AlgL, presented a basal level which was not correlated with a rise in their gene transcriptional levels [9]. This behavior per se may explain the observed rise in alginate MW (Table 2). Even though these observations indicate that dissolved oxygen affects intracellular as well as extracellular alginate lyase activities, it is possible that different alginate lyases could be expressed at different physiological states, as suggested by the study of AlyE3, which is essential for the efficient cyst germination in A. vinelandii [51].

It is important to note that although the AlgL is localized in the periplasm, it has an N-terminal secretion signal (AlgL, UniProtKB-O5219), suggesting that AlgL secretion can occur in response to diverse environmental stimuli (i.e., oxygen concentration). This notion is supported by the observation that AlgL extracellular activity is highly dependent on the dissolved oxygen and that the role of alginate lyase is restricted to a postpolymerization step [9, 85]. Similarly, the alginate lyase AlyA3 also presents extracellular activity, whereas AlyA1 and AlyA2 appear to be periplasmic [51]. These data strongly suggest that alginate lyase expression and extracellular activity occur in response to dissolved oxygen concentrations. Therefore, a detailed analysis of dynamic variations in expression levels and in enzymatic activity throughout the culture is warranted to understand more deeply the alginate polymerization process.

In summary, current evidence indicates that when values of vary between 2 and 5 mmol g−1 h−1 in cultures of A. vinelandii, a rise in expression of algL together with a decrease in expression of alg8 correlates with a decrease in alginate MW (Table 2). As such, this range of could be a target in the development of strategies to manipulate the characteristics of alginates.

5.1. Oxygen Sensing Mechanisms in Azotobacter vinelandii

Current evidences demonstrate that the oxygen transfer rate, the dissolved oxygen tension levels, and the oxygen uptake rate affect alginate biosynthesis in A. vinelandii cultures [8, 9, 12, 13, 36, 40, 8689]. Despite the importance of the oxygen and the intrinsic relationship with it, no strong evidence of the molecular mechanism involved in sensing it during A. vinelandii culturing is available, as well as its further downstream mechanism still being lacking. In this section we discuss that oxygen availability during A. vinelandii culturing is a key factor and we suggest a possible mechanism of action.

In A. vinelandii the mechanism involved in sensing oxygen availability remains to be fully investigated. In bacteria, several oxygen sensing mechanisms exist. However they can be clustered in two groups based on how the signal is perceived. One category can interact with external environment while, on the other hand, the second category senses physiological changes resulting from variations in the external environment. Nevertheless, both sensing mechanisms operating together control directly the switch between aerobic and anaerobic metabolism [90]. Among the oxygen sensing mechanism, the FNR, ArcA/B, and ubiquinone-8 (Q8) are well characterized in E. coli [90].

In A. vinelandii the absence of an Fnr-like protein, CydR, overexpressing the β-ketothiolase and acetoacetyl-coA reductase [91], both enzymes catalyze the production of β-hydroxybutyryl-CoA, which is the PHB precursor [40]. It has been demonstrated that low aeration culture conditions in A. vinelandii cultures enhanced the metabolic flux from pyruvate towards acetyl-CoA. This had an influence on the increment on the metabolic flux towards PHB production, concomitantly with the higher alginate production [8], suggesting that the aeration conditions could affect the alginate production, by regulating possible gene targets of CydR. Supporting this observation, batch cultures of A. vinelandii OP mutant strain carried out at 600 rpm showed lowest compared with wild-type strain (ATCC 9046) [92]. The A. vinelandii OP strain contains an insertion element in the algU gene, which in turn represses alginate synthesis [93] and it has been suggested that AlgU is required for cydR gene expression [94].

CydR controls the expression of cydAB operon that encodes a cytochrome bd terminal oxidase, and cydAB gene expression correlates with the NADH:ubiquinone oxidoreductase activity (NDHII) [91]. In A. vinelandii, the Na+-translocating NADH:ubiquinone oxidoreductases (Na+-NQR) are encoded in the nqr operon, and it had been linked to regulating negatively alginate production [95]. Additionally, A. vinelandii genome contains genes linked to NADH:ubiquinone oxidoreductases (NDH), the NDH-II type, and 13 genes encoding subunits of NDH-I type [95]. The NADH oxidation in A. vinelandii is mediated by two NADH:ubiquinone oxidoreductases [96], and the fast NADH oxidation is linked to a fast quinone reduction. The ubiC-A operon in A. vinelandii is responsible for the transcription of the genes necessaries for Q8 biosynthesis [95]. A mutation in the intragenic region ubiA correlates with lower Q8 protein levels, accompanied with an improvement in the alginate production, but all the more, with a higher expression of biosynthetic alginate genes, algD, algC, and algA. Moreover, the Q8 protein seems to be responsible for at least 8% of the respiratory capacity in A. vinelandii, during low and high aeration cultures [95].

Interestingly, in other bacteria as E. coli, the role of quinones as a redox signal for the pathways involved in sensing oxygen and regulation of expression of genes involved in oxidative and fermentative catabolism is well known, specifically the ArcB/A two-component system [9799].

Figure 6 summarizes the plausible regulation of alg genes in A. vinelandii, via a signaling cascade activated by oxygen availability. On one hand, the Na+NQR protein regulates negatively algD and algC gene targets, while the ArcB/A two-component system regulates algD and alg8 gene expression under oxygen availability. When oxygen is limiting, the sensor kinase ArcB autophosphorylates and then transphosphorylates the regulator ArcA, which activates algD, alg8, and alg44 gene expression. The autophosphorylation of ArcB is inhibited at higher oxygen concentrations, by the accumulation of Q8 (oxidized form). In this sense, in A. vinelandii, a tight control of alg genes via a signaling cascade activated by oxygen availability may exist (Figure 6).

Figure 6: Schematic representation of the possible gene regulation mechanism by oxygen in Azotobacter vinelandii. Oxygen availability is depicted in the figure as low O2 (left side) and high O2 (right side). Light red dotted boxes indicate the Na+-translocating NADH:ubiquinone oxidoreductase (Na+NQR) that regulates negatively algD and algC gene targets, although the exact mechanism of algD and algC gene regulation at high O2 by Na+NQR is still unknown. Gray slashed boxes represent the ArcB/A two-component redox sensor: under high oxygen availability, the autophosphorylation of ArcB (B blocks) is inhibited by oxidized quinones (Q8). ArcA (A blocks) in the nonphosphorylated state is unable to bind specifically to algD, alg8, and alg44 gene targets. Low oxygen causes a decrease in the level of oxidized quinones (Q8H2), allowing the autophosphorylation of ArcA. ArcA-P binds specifically to its target sites and coordinates the cellular response to oxygen availability. Arrows indicate positive regulation and T-shaped bars indicate negative regulation. Flag-type boxes indicate genes described in the figure. Question mark indicates unknown gene regulation mechanism. OM: outer membrane; PG: peptidoglycan; IM: inner membrane.

Although recently Flores et al., 2015 [36], discussed mainly the influence of the oxygen on production of alginate during A. vinelandii cultures, not much attention is paid to which molecular pathways are involved during alginate biosynthesis. In our work, we propose a possible mechanism of action of the oxygen availability during A. vinelandii culturing, offering a new path to look at and in this way contributing to the better knowledge of controlling bacterial alginates production.

Despite the enormous efforts in understanding the microbial alginate biosynthesis under defined culture conditions, there is still a way to go. The decoding of the A. vinelandii genome has open the possibility to getting access to new information; however no wide genetic screen studies during alginate production have been reported yet. So, it will be necessarily an improvement in the knowledge of A. vinelandii alginate biosynthesis gene regulation in alginate production processes, in order to generate a tailored and affordable alginate product.

6. Conclusion

In the present review we discuss that oxygen availability during Azotobacter vinelandii cultures might exert a tight control over the expression of alginate-related genes, which will impact the quality of the polysaccharide or will regulate enzymatic activities that modified the nascent alginate chain. Current evidence indicates a prevailing equilibrium in alg8 and algL gene expression, which is being regulated by oxygen availability. This equilibrium will further impact the alginate molecular weight. Accordingly, more information regarding oxygen sensing, transportation, and signaling pathways during specific culture conditions of A. vinelandii will be needed in order to obtain alginates with specific characteristics.

Conflict of Interests

The authors declare that there is no conflict of interests regarding the publication of this paper.

Acknowledgments

This work was supported by a Grant from CONICYT-Chile (Project PCCI40039) and DI-PUCV 037-98. The authors acknowledge Dr. Nuno Rodrigues Dos Santos for his critical comments on the paper.

References

  1. B. H. A. Rehm, “Bacterial polymers: biosynthesis, modifications and applications,” Nature Reviews Microbiology, vol. 8, no. 8, pp. 578–592, 2010. View at Publisher · View at Google Scholar · View at Scopus
  2. S. Bengtsson, A. R. Pisco, M. A. M. Reis, and P. C. Lemos, “Production of polyhydroxyalkanoates from fermented sugar cane molasses by a mixed culture enriched in glycogen accumulating organisms,” Journal of Biotechnology, vol. 145, no. 3, pp. 253–263, 2010. View at Publisher · View at Google Scholar · View at Scopus
  3. F. Clementi, “Alginate production by Azotobacter vinelandii,” Critical Reviews in Biotechnology, vol. 17, no. 4, pp. 327–361, 1997. View at Publisher · View at Google Scholar · View at Scopus
  4. D. E. Pszczola, “Discovering treasures of the deep,” Food Technology, vol. 52, no. 4, pp. 74–80, 1998. View at Google Scholar
  5. I. W. Sutherland, Biotechnology of Microbial Exopolysaccharydes, Cambridge University Press, Cambridge, UK, 1990.
  6. U. Remminghorst and B. H. A. Rehm, “Bacterial alginates: from biosynthesis to applications,” Biotechnology Letters, vol. 28, no. 21, pp. 1701–1712, 2006. View at Publisher · View at Google Scholar · View at Scopus
  7. J. L. Geddie and I. W. Sutherland, “The effect of acetylation on cation binding by algal and bacterial alginates,” Biotechnology and Applied Biochemistry, vol. 20, no. 1, pp. 117–129, 1994. View at Google Scholar · View at Scopus
  8. T. Castillo, E. Heinzle, S. Peifer, K. Schneider, and C. F. Peña M, “Oxygen supply strongly influences metabolic fluxes, the production of poly(3-hydroxybutyrate) and alginate, and the degree of acetylation of alginate in Azotobacter vinelandii,” Process Biochemistry, vol. 48, no. 7, pp. 995–1003, 2013. View at Publisher · View at Google Scholar · View at Scopus
  9. C. Flores, S. Moreno, G. Espín, C. Peña, and E. Galindo, “Expression of alginases and alginate polymerase genes in response to oxygen, and their relationship with the alginate molecular weight in Azotobacter vinelandii,” Enzyme and Microbial Technology, vol. 53, no. 2, pp. 85–91, 2013. View at Publisher · View at Google Scholar · View at Scopus
  10. Ç. Kıvılcımdan Moral, Ö. Doğan, and F. D. Sanin, “Effect of oxygen tension and medium components on monomer distribution of alginate,” Applied Biochemistry and Biotechnology, vol. 176, no. 3, pp. 875–891, 2015. View at Publisher · View at Google Scholar
  11. A. Díaz-Barrera, A. Aguirre, J. Berrios, and F. Acevedo, “Continuous cultures for alginate production by Azotobacter vinelandii growing at different oxygen uptake rates,” Process Biochemistry, vol. 46, no. 9, pp. 1879–1883, 2011. View at Publisher · View at Google Scholar · View at Scopus
  12. A. Díaz-Barrera, E. Soto, and C. Altamirano, “Alginate production and alg8 gene expression by Azotobacter vinelandii in continuous cultures,” Journal of Industrial Microbiology and Biotechnology, vol. 39, no. 4, pp. 613–621, 2012. View at Publisher · View at Google Scholar · View at Scopus
  13. A. Díaz-Barrera, F. Martínez, F. Guevara Pezoa, F. Acevedo, and B. Lin, “Evaluation of gene expression and alginate production in response to oxygen transfer in continuous culture of Azotobacter vinelandii,” PLoS ONE, vol. 9, no. 8, Article ID e105993, 2014. View at Publisher · View at Google Scholar
  14. N. Rhein-Knudsen, M. T. Ale, and A. S. Meyer, “Seaweed hydrocolloid production: an update on enzyme assisted extraction and modification technologies,” Marine Drugs, vol. 13, no. 6, pp. 3340–3359, 2015. View at Publisher · View at Google Scholar
  15. W. Sabra, A.-P. Zeng, and W.-D. Deckwer, “Bacterial alginate: physiology, product quality and process aspects,” Applied Microbiology and Biotechnology, vol. 56, no. 3-4, pp. 315–325, 2001. View at Publisher · View at Google Scholar · View at Scopus
  16. B. H. A. Rehm and S. Valla, “Bacterial alginates: biosynthesis and applications,” Applied Microbiology and Biotechnology, vol. 48, no. 3, pp. 281–288, 1997. View at Publisher · View at Google Scholar · View at Scopus
  17. P. Gacesa, “Bacterial alginate biosynthesis—recent progress and future prospects,” Microbiology, vol. 144, no. 5, pp. 1133–1143, 1998. View at Publisher · View at Google Scholar · View at Scopus
  18. D. Hoefer, J. K. Schnepf, T. R. Hammer, M. Fischer, and C. Marquardt, “Biotechnologically produced microbial alginate dressings show enhanced gel forming capacity compared to commercial alginate dressings of marine origin,” Journal of Materials Science: Materials in Medicine, vol. 26, no. 4, article 162, 2015. View at Publisher · View at Google Scholar
  19. E. Ruvinov and S. Cohen, “Alginate biomaterial for the treatment of myocardial infarction: progress, translational strategies, and clinical outlook,” Advanced Drug Delivery Reviews, vol. 96, pp. 54–76, 2016. View at Publisher · View at Google Scholar
  20. J. Venkatesan, I. Bhatnagar, P. Manivasagan, K.-H. Kang, and S.-K. Kim, “Alginate composites for bone tissue engineering: a review,” International Journal of Biological Macromolecules, vol. 72, pp. 269–281, 2015. View at Publisher · View at Google Scholar · View at Scopus
  21. I. A. Brownlee, A. Allen, J. P. Pearson et al., “Alginate as a source of dietary fiber,” Critical Reviews in Food Science and Nutrition, vol. 45, no. 6, pp. 497–510, 2005. View at Publisher · View at Google Scholar · View at Scopus
  22. M. G. Jensen, M. Kristensen, and A. Astrup, “Effect of alginate supplementation on weight loss in obese subjects completing a 12-wk energy-restricted diet: a randomized controlled trial,” The American Journal of Clinical Nutrition, vol. 96, no. 1, pp. 5–13, 2012. View at Publisher · View at Google Scholar · View at Scopus
  23. P. W. Dettmar, V. Strugala, and J. Craig Richardson, “The key role alginates play in health,” Food Hydrocolloids, vol. 25, no. 2, pp. 263–266, 2011. View at Publisher · View at Google Scholar · View at Scopus
  24. M. Liu, L. Dai, H. Shi, S. Xiong, and C. Zhou, “In vitro evaluation of alginate/halloysite nanotube composite scaffolds for tissue engineering,” Materials Science and Engineering: C, vol. 49, pp. 700–712, 2015. View at Publisher · View at Google Scholar · View at Scopus
  25. N. V. Konda, S. Singh, B. A. Simmons, and D. Klein-Marcuschamer, “An investigation on the economic feasibility of macroalgae as a potential feedstock for biorefineries,” BioEnergy Research, vol. 8, no. 3, pp. 1046–1056, 2015. View at Publisher · View at Google Scholar
  26. I. W. Davidson, I. W. Sutherland, and C. J. Lawson, “Localization of O-acetyl groups of bacterial alginate,” Journal of General Microbiology, vol. 98, no. 2, pp. 603–606, 1977. View at Publisher · View at Google Scholar · View at Scopus
  27. G. Skjåk-Bræk, S. Paoletti, and T. Gianferrara, “Selective acetylation of mannuronic acid residues in calcium alginate gels,” Carbohydrate Research, vol. 185, no. 1, pp. 119–129, 1989. View at Publisher · View at Google Scholar · View at Scopus
  28. H. Ertesvåg, “Alginate-modifying enzymes: biological roles and biotechnological uses,” Frontiers in Microbiology, vol. 6, no. 523, 2015. View at Publisher · View at Google Scholar
  29. H. Ertesvåg, S. Valla, and G. Skjåk-Bræk, “Enzymatic alginate modification,” in Alginates: Biology and Applications, B. H. A. Rehm, Ed., Microbiology Monographs, pp. 95–115, Springer, Berlin, Germany, 2009. View at Publisher · View at Google Scholar
  30. W. Sabra, A.-P. Zeng, H. Lunsdorf, and W.-D. Deckwer, “Effect of oxygen on formation and structure of Azotobacter vinelandii alginate and its role in protecting nitrogenase,” Applied and Environmental Microbiology, vol. 66, no. 9, pp. 4037–4044, 2000. View at Publisher · View at Google Scholar · View at Scopus
  31. J. C. Setubal, P. dos Santos, B. S. Goldman et al., “Genome sequence of Azotobacter vinelandii, an obligate aerobe specialized to support diverse anaerobic metabolic processes,” Journal of Bacteriology, vol. 191, no. 14, pp. 4534–4545, 2009. View at Publisher · View at Google Scholar · View at Scopus
  32. E. Post, D. Kleiner, and J. Oelze, “Whole cell respiration and nitrogenase activities in Azotobacter vinelandii growing in oxygen controlled continuous culture,” Archives of Microbiology, vol. 134, no. 1, pp. 68–72, 1983. View at Publisher · View at Google Scholar · View at Scopus
  33. H. L. Sadoff, “Encystment and germination in Azotobacter vinelandii,” Bacteriological Reviews, vol. 39, no. 4, pp. 516–539, 1975. View at Google Scholar · View at Scopus
  34. D. Segura, C. Núñez, and G. Espín, “Azotobacter cysts,” in Encyclopedia of Life Sciences, John Wiley & Sons, New York, NY, USA, 2001. View at Publisher · View at Google Scholar
  35. A. Díaz-Barrera and E. Soto, “Biotechnological uses of Azotobacter vinelandii: current state, limits and prospects,” African Journal of Biotechnology, vol. 9, no. 33, pp. 5240–5250, 2010. View at Google Scholar · View at Scopus
  36. C. Flores, A. Díaz-Barrera, F. Martínez, E. Galindo, and C. Peña, “Role of oxygen in the polymerization and de-polymerization of alginate produced by Azotobacter vinelandii,” Journal of Chemical Technology and Biotechnology, vol. 90, no. 3, pp. 356–365, 2015. View at Publisher · View at Google Scholar · View at Scopus
  37. J. Schmid, V. Sieber, and B. Rehm, “Bacterial exopolysaccharides: biosynthesis pathways and engineering strategies,” Frontiers in Microbiology, vol. 6, 2015. View at Publisher · View at Google Scholar
  38. J. C. Whitney and P. L. Howell, “Synthase-dependent exopolysaccharide secretion in Gram-negative bacteria,” Trends in Microbiology, vol. 21, no. 2, pp. 63–72, 2013. View at Publisher · View at Google Scholar · View at Scopus
  39. I. D. Hay, Z. U. Rehman, A. Ghafoor, and B. H. A. Rehm, “Bacterial biosynthesis of alginates,” Journal of Chemical Technology and Biotechnology, vol. 85, no. 6, pp. 752–759, 2010. View at Publisher · View at Google Scholar · View at Scopus
  40. E. Galindo, C. Peña, C. Núñez, D. Segura, and G. Espín, “Molecular and bioengineering strategies to improve alginate and polydydroxyalkanoate production by Azotobacter vinelandii,” Microbial Cell Factories, vol. 6, article 7, 2007. View at Publisher · View at Google Scholar · View at Scopus
  41. P. J. Tatnell, N. J. Russell, and P. Gacesa, “GDP-mannose dehydrogenase is the key regulatory enzyme in alginate biosynthesis in Pseudomonas aeruginosa: evidence from metabolite studies,” Microbiology, vol. 140, no. 7, pp. 1745–1754, 1994. View at Publisher · View at Google Scholar · View at Scopus
  42. Z. U. Rehman, Y. Wang, M. F. Moradali, I. D. Hay, and B. H. A. Rehm, “Insights into the assembly of the alginate biosynthesis machinery in Pseudomonas aeruginosa,” Applied and Environmental Microbiology, vol. 79, no. 10, pp. 3264–3272, 2013. View at Publisher · View at Google Scholar · View at Scopus
  43. M. Fata Moradali, I. Donati, I. M. Sims, S. Ghods, and B. H. Rehm, “Alginate polymerization and modification are linked in Pseudomonas aeruginosa,” mBio, vol. 6, no. 3, Article ID e00453-15, 2015. View at Publisher · View at Google Scholar
  44. H. Mejía-Ruíz, J. Guzmán, S. Moreno, G. Soberón-Chávez, and G. Espín, “The Azotobacter vinelandii alg8 and alg44 genes are essential for alginate synthesis and can be transcribed from an algD-independent promoter,” Gene, vol. 199, no. 1-2, pp. 271–277, 1997. View at Publisher · View at Google Scholar · View at Scopus
  45. H. Mejía-Ruíz, S. Moreno, J. Guzmán et al., “Isolation and characterization of an Azotobacter vinelandii algK mutant,” FEMS Microbiology Letters, vol. 156, no. 1, pp. 101–106, 1997. View at Publisher · View at Google Scholar · View at Scopus
  46. L. M. Riley, J. T. Weadge, P. Baker et al., “Structural and functional characterization of Pseudomonas aeruginosa AlgX: role of Algx in alginate acetylation,” Journal of Biological Chemistry, vol. 288, no. 31, pp. 22299–22314, 2013. View at Publisher · View at Google Scholar · View at Scopus
  47. M. J. Franklin and D. E. Ohman, “Mutant analysis and cellular localization of the AlgI, AlgJ, and AlgF proteins required for O acetylation of alginate in Pseudomonas aeruginosa,” Journal of Bacteriology, vol. 184, no. 11, pp. 3000–3007, 2002. View at Publisher · View at Google Scholar · View at Scopus
  48. P. Baker, T. Ricer, P. J. Moynihan et al., “P. aeruginosa SGNH hydrolase-like proteins AlgJ and AlgX have similar topology but separate and distinct roles in alginate acetylation,” PLoS Pathogens, vol. 10, no. 8, Article ID e1004334, 2014. View at Publisher · View at Google Scholar
  49. A. Vazquez, S. Moreno, J. Guzmán, A. Alvarado, and G. Espín, “Transcriptional organization of the Azotobacter vinelandii algGXLVIFA genes: characterization of algF mutants,” Gene, vol. 232, no. 2, pp. 217–222, 1999. View at Publisher · View at Google Scholar · View at Scopus
  50. B. I. G. Svanem, W. I. Strand, H. Ertesvåg et al., “The catalytic activities of the bifunctional Azotobacter vinelandii mannuronan C-5-epimerase and alginate lyase AlgE7 probably originate from the same active site in the enzyme,” Journal of Biological Chemistry, vol. 276, no. 34, pp. 31542–31550, 2001. View at Publisher · View at Google Scholar · View at Scopus
  51. M. Gimmestad, H. Ertesvåg, T. M. B. Heggeset, O. Aarstad, B. I. G. Svanem, and S. Valla, “Characterization of three new Azotobacter vinelandii alginate lyases, one of which is involved in cyst germination,” Journal of Bacteriology, vol. 191, no. 15, pp. 4845–4853, 2009. View at Publisher · View at Google Scholar · View at Scopus
  52. S. Jain and D. E. Ohman, “Role of an alginate lyase for alginate transport in mucoid Pseudomonas aeruginosa,” Infection and Immunity, vol. 73, no. 10, pp. 6429–6436, 2005. View at Publisher · View at Google Scholar · View at Scopus
  53. H. Ertesvåg, H. K. Høidal, I. K. Hals, A. Rian, B. Doseth, and S. Valla, “A family of modular type mannuronan C-5-epimerase genes controls alginate structure in Azotobacter vinelandii,” Molecular Microbiology, vol. 16, no. 4, pp. 719–731, 1995. View at Publisher · View at Google Scholar · View at Scopus
  54. C. Núñez, R. León, J. Guzmán, G. Espín, and G. Soberón-Chávez, “Role of Azotobacter vinelandii mucA and mucC gene products in alginate production,” Journal of Bacteriology, vol. 182, no. 23, pp. 6550–6556, 2000. View at Publisher · View at Google Scholar · View at Scopus
  55. L. Lloret, R. Barreto, R. Léon et al., “Genetic analysis of the transcriptional arrangement of Azotobacter vinelandii alginate biosynthetic genes: identification of two independent promoters,” Molecular Microbiology, vol. 21, no. 3, pp. 449–457, 1996. View at Publisher · View at Google Scholar · View at Scopus
  56. G. Gaona, C. Núñez, J. B. Goldberg et al., “Characterization of the Azotobacter vinelandii algC gene involved in alginate and lipopolysaccharide production,” FEMS Microbiology Letters, vol. 238, no. 1, pp. 199–206, 2004. View at Publisher · View at Google Scholar · View at Scopus
  57. S. Moreno, R. Nájera, J. Guzmán, G. Soberón-Chávez, and G. Espín, “Role of alternative σ factor AlgU in encystment of Azotobacter vinelandii,” Journal of Bacteriology, vol. 180, no. 10, pp. 2766–2769, 1998. View at Google Scholar · View at Scopus
  58. M. Castañeda, J. Sánchez, S. Moreno, C. Núñez, and G. Espín, “The global regulators GacA and σS form part of a cascade that controls alginate production in Azotobacter vinelandii,” Journal of Bacteriology, vol. 183, no. 23, pp. 6787–6793, 2001. View at Publisher · View at Google Scholar · View at Scopus
  59. J. Manzo, M. Cocotl-Yañez, T. Tzontecomani et al., “Post-transcriptional regulation of the alginate biosynthetic gene algD by the Gac/Rsm system in Azotobacter vinelandii,” Journal of Molecular Microbiology and Biotechnology, vol. 21, no. 3-4, pp. 147–159, 2012. View at Publisher · View at Google Scholar · View at Scopus
  60. M. A. Azevedo, A. I. Bourbon, A. A. Vicente, and M. A. Cerqueira, “Alginate/chitosan nanoparticles for encapsulation and controlled release of vitamin B2,” International Journal of Biological Macromolecules, vol. 71, pp. 141–146, 2014. View at Publisher · View at Google Scholar · View at Scopus
  61. X. Zhao, B. Li, C. Xue, and L. Sun, “Effect of molecular weight on the antioxidant property of low molecular weight alginate from Laminaria japonica,” Journal of Applied Phycology, vol. 24, no. 2, pp. 295–300, 2012. View at Publisher · View at Google Scholar · View at Scopus
  62. C. A. Bonino, M. D. Krebs, C. D. Saquing et al., “Electrospinning alginate-based nanofibers: from blends to crosslinked low molecular weight alginate-only systems,” Carbohydrate Polymers, vol. 85, no. 1, pp. 111–119, 2011. View at Publisher · View at Google Scholar · View at Scopus
  63. K. Fujiki, H. Matsuyama, and T. Yano, “Protective effect of sodium alginates against bacterial infection in common carp, Cyprinus carpio L.,” Journal of Fish Diseases, vol. 17, no. 4, pp. 349–355, 1994. View at Publisher · View at Google Scholar
  64. T. Kuda, H. Goto, M. Yokoyama, and T. Fujii, “Effects of dietary concentration of laminaran and depolymerised alginate on rat cecal microflora and plasma lipids,” Fisheries Science, vol. 64, no. 4, pp. 589–593, 1998. View at Google Scholar · View at Scopus
  65. T. Kuda, T. Yano, N. Matsuda, and M. Nishizawa, “Inhibitory effects of laminaran and low molecular alginate against the putrefactive compounds produced by intestinal microflora in vitro and in rats,” Food Chemistry, vol. 91, no. 4, pp. 745–749, 2005. View at Publisher · View at Google Scholar · View at Scopus
  66. I. Pajic-Lijakovic, S. Levic, M. Hadnađev et al., “Structural changes of Ca-alginate beads caused by immobilized yeast cell growth,” Biochemical Engineering Journal, vol. 103, pp. 32–38, 2015. View at Publisher · View at Google Scholar
  67. F. E. Vasile, A. M. Romero, M. A. Judis, and M. F. Mazzobre, “Prosopis alba exudate gum as excipient for improving fish oil stability in alginate—chitosan beads,” Food Chemistry, vol. 190, pp. 1093–1101, 2016. View at Publisher · View at Google Scholar
  68. F. Mancini, L. Montanari, D. Peressini, and P. Fantozzi, “Influence of alginate concentration and molecular weight on functional properties of mayonnaise,” LWT—Food Science and Technology, vol. 35, no. 6, pp. 517–525, 2002. View at Publisher · View at Google Scholar · View at Scopus
  69. O. Aizpurua-Olaizola, P. Navarro, A. Vallejo, M. Olivares, N. Etxebarria, and A. Usobiaga, “Microencapsulation and storage stability of polyphenols from Vitis vinifera grape wastes,” Food Chemistry, vol. 190, pp. 614–621, 2016. View at Publisher · View at Google Scholar
  70. W. Cheng, C.-H. Liu, C.-M. Kuo, and J.-C. Chen, “Dietary administration of sodium alginate enhances the immune ability of white shrimp Litopenaeus vannamei and its resistance against Vibrio alginolyticus,” Fish and Shellfish Immunology, vol. 18, no. 1, pp. 1–12, 2005. View at Publisher · View at Google Scholar · View at Scopus
  71. M. D. Wilcox, I. A. Brownlee, J. C. Richardson, P. W. Dettmar, and J. P. Pearson, “The modulation of pancreatic lipase activity by alginates,” Food Chemistry, vol. 146, pp. 479–484, 2014. View at Publisher · View at Google Scholar · View at Scopus
  72. B. An, H. Lee, S. Lee, S. Lee, and J. Choi, “Determining the selectivity of divalent metal cations for the carboxyl group of alginate hydrogel beads during competitive sorption,” Journal of Hazardous Materials, vol. 298, pp. 11–18, 2015. View at Publisher · View at Google Scholar
  73. W. Cheng, R.-T. Tsai, and C.-C. Chang, “Dietary sodium alginate administration enhances Mx gene expression of the tiger grouper, Epinephelus fuscoguttatus receiving poly I:C,” Aquaculture, vol. 324-325, pp. 201–208, 2012. View at Publisher · View at Google Scholar · View at Scopus
  74. S.-T. Chiu, R.-T. Tsai, J.-P. Hsu, C.-H. Liu, and W. Cheng, “Dietary sodium alginate administration to enhance the non-specific immune responses, and disease resistance of the juvenile grouper Epinephelus fuscoguttatus,” Aquaculture, vol. 277, no. 1-2, pp. 66–72, 2008. View at Publisher · View at Google Scholar · View at Scopus
  75. C.-H. Liu, S.-P. Yeh, C.-M. Kuo, W. Cheng, and C.-H. Chou, “The effect of sodium alginate on the immune response of tiger shrimp via dietary administration: activity and gene transcription,” Fish and Shellfish Immunology, vol. 21, no. 4, pp. 442–452, 2006. View at Publisher · View at Google Scholar · View at Scopus
  76. K. Fujiki and T. Yano, “Effects of sodium alginate on the non-specific defence system of the common carp (Cyprinus carpio L.),” Fish and Shellfish Immunology, vol. 7, no. 6, pp. 417–427, 1997. View at Publisher · View at Google Scholar · View at Scopus
  77. H. Tomida, T. Yasufuku, T. Fujii, Y. Kondo, T. Kai, and M. Anraku, “Polysaccharides as potential antioxidative compounds for extended-release matrix tablets,” Carbohydrate Research, vol. 345, no. 1, pp. 82–86, 2010. View at Publisher · View at Google Scholar · View at Scopus
  78. L. L. Oglesby, S. Jain, and D. E. Ohman, “Membrane topology and roles of Pseudomonas aeruginosa Alg8 and Alg44 in alginate polymerization,” Microbiology, vol. 154, no. 6, pp. 1605–1615, 2008. View at Publisher · View at Google Scholar · View at Scopus
  79. I. M. Saxena, R. M. Jr. Brown, M. Fevre, R. A. Geremia, and B. Henrissat, “Multidomain architecture of β-glycosil tranferases: implications for mechanism of action,” Journal of Bacteriology, vol. 177, no. 6, pp. 1419–1419, 1995. View at Google Scholar
  80. U. Remminghorst and B. H. A. Rehm, “In vitro alginate polymerization and the functional role of Alg8 in alginate production by Pseudomonas aeruginosa,” Applied and Environmental Microbiology, vol. 72, no. 1, pp. 298–305, 2006. View at Publisher · View at Google Scholar · View at Scopus
  81. M. Merighi, V. T. Lee, M. Hyodo, Y. Hayakawa, and S. Lory, “The second messenger bis-(3′-5′)-cyclic-GMP and its PilZ domain-containing receptor Alg44 are required for alginate biosynthesis in Pseudomonas aeruginosa,” Molecular Microbiology, vol. 65, no. 4, pp. 876–895, 2007. View at Publisher · View at Google Scholar · View at Scopus
  82. S. Alexeeva, K. J. Hellingwerf, and M. J. Teixeira de Mattos, “Quantitative assessment of oxygen availability: perceived aerobiosis and its effect on flux distribution in the respiratory chain of Escherichia coli,” Journal of Bacteriology, vol. 184, no. 5, pp. 1402–1406, 2002. View at Publisher · View at Google Scholar · View at Scopus
  83. J. Oelze, “Respiratory protection of nitrogenase in Azotobacter species: Is a widely held hypothesis unequivocally supported by experimental evidence?” FEMS Microbiology Reviews, vol. 24, no. 4, pp. 321–333, 2000. View at Publisher · View at Google Scholar · View at Scopus
  84. M. A. Trujillo-Roldán, S. Moreno, D. Segura, E. Galindo, and G. Espín, “Alginate production by an Azotobacter vinelandii mutant unable to produce alginate lyase,” Applied Microbiology and Biotechnology, vol. 60, no. 6, pp. 733–737, 2003. View at Publisher · View at Google Scholar · View at Scopus
  85. M. A. Trujillo-Roldán, S. Moreno, G. Espín, and E. Galindo, “The roles of oxygen and alginate-lyase in determining the molecular weight of alginate produced by Azotobacter vinelandii,” Applied Microbiology and Biotechnology, vol. 63, no. 6, pp. 742–747, 2004. View at Publisher · View at Google Scholar · View at Scopus
  86. A. Díaz-Barrera, C. Peña, and E. Galindo, “The oxygen transfer rate influences the molecular mass of the alginate produced by Azotobacter vinelandii,” Applied Microbiology and Biotechnology, vol. 76, no. 4, pp. 903–910, 2007. View at Publisher · View at Google Scholar · View at Scopus
  87. A. Díaz-Barrera, P. Silva, R. Ávalos, and F. Acevedo, “Alginate molecular mass produced by Azotobacter vinelandii in response to changes of the O2 transfer rate in chemostat cultures,” Biotechnology Letters, vol. 31, no. 6, pp. 825–829, 2009. View at Publisher · View at Google Scholar · View at Scopus
  88. E. Lozano, E. Galindo, and C. F. Peña, “Oxygen transfer rate during the production of alginate by Azotobacter vinelandii under oxygen-limited and non oxygen-limited conditions,” Microbial Cell Factories, vol. 10, article 13, 2011. View at Publisher · View at Google Scholar · View at Scopus
  89. C. Peña, M. A. Trujillo-Roldán, and E. Galindo, “Influence of dissolved oxygen tension and agitation speed on alginate production and its molecular weight in cultures of Azotobacter vinelandii,” Enzyme and Microbial Technology, vol. 27, no. 6, pp. 390–398, 2000. View at Publisher · View at Google Scholar · View at Scopus
  90. J. Green and M. S. Paget, “Bacterial redox sensors,” Nature Reviews Microbiology, vol. 2, no. 12, pp. 954–966, 2004. View at Publisher · View at Google Scholar · View at Scopus
  91. G. Wu, A. J. G. Moir, G. Sawers, S. Hill, and R. K. Poole, “Biosynthesis of poly-β-hydroxybutyrate (PHB) is controlled by CydR (Fnr) in the obligate aerobe Azotobacter vinelandii,” FEMS Microbiology Letters, vol. 194, no. 2, pp. 215–220, 2001. View at Publisher · View at Google Scholar · View at Scopus
  92. A. Díaz-Barrera, R. Andler, I. Martínez, and C. Peña, “Poly-3-hydroxybutyrate production by Azotobacter vinelandii strains in batch cultures at different oxygen transfer rates,” Journal of Chemical Technology & Biotechnology, 2015. View at Publisher · View at Google Scholar
  93. J. M. Martínez-Salazar, S. Moreno, R. Nájera et al., “Characterization of the genes coding for the putative sigma factor AlgU and its regulators MucA, MucB, MucC, and MucD in Azotobacter vinelandii and evaluation of their roles in alginate biosynthesis,” Journal of Bacteriology, vol. 178, no. 7, pp. 1800–1808, 1996. View at Google Scholar · View at Scopus
  94. R. León and G. Espín, “flhDC, but not fleQ, regulates flagella biogenesis in Azotobacter vinelandii, and is under AlgU and CydR negative control,” Microbiology, vol. 154, no. 6, pp. 1719–1728, 2008. View at Publisher · View at Google Scholar · View at Scopus
  95. C. Núñez, A. V. Bogachev, G. Guzmán, I. Tello, J. Guzmán, and G. Espín, “The Na+-translocating NADH: ubiquinone oxidoreductase of Azotobacter vinelandii negatively regulates alginate synthesis,” Microbiology, vol. 155, no. 1, pp. 249–256, 2009. View at Publisher · View at Google Scholar · View at Scopus
  96. Y. V. Bertsova, A. V. Bogachev, and V. P. Skulachev, “Non-coupled NADH: ubiquinone oxidoreductase of Azotobacter vinelandii is required for diazotrophic growth at high oxygen concentrations,” Journal of Bacteriology, vol. 183, no. 23, pp. 6869–6874, 2001. View at Publisher · View at Google Scholar · View at Scopus
  97. M. Bekker, S. Alexeeva, W. Laan, G. Sawers, J. T. De Mattos, and K. Hellingwerf, “The ArcBA two-component system of Escherichia coli is regulated by the redox state of both the ubiquinone and the menaquinone pool,” Journal of Bacteriology, vol. 192, no. 3, pp. 746–754, 2010. View at Publisher · View at Google Scholar · View at Scopus
  98. D. Georgellis, O. Kwon, and E. C. C. Lin, “Quinones as the redox signal for the Arc two-component system of bacteria,” Science, vol. 292, no. 5525, pp. 2314–2316, 2001. View at Publisher · View at Google Scholar · View at Scopus
  99. R. Malpica, G. R. Peña Sandoval, C. Rodríguez, B. Franco, and D. Georgellis, “Signaling by the Arc two-component system provides a link between the redox state of the quinone pool and gene expression,” Antioxidants and Redox Signaling, vol. 8, no. 5-6, pp. 781–795, 2006. View at Publisher · View at Google Scholar · View at Scopus