Open access peer-reviewed chapter

Microbial Cellulose — Biosynthesis Mechanisms and Medical Applications

Written By

Wilton R. Lustri, Hélida Gomes de Oliveira Barud, Hernane da Silva Barud, Maristela F. S. Peres, Junkal Gutierrez, Agnieszka Tercjak, Osmir Batista de Oliveira Junior and Sidney José Lima Ribeiro

Submitted: March 26th, 2015 Reviewed: October 19th, 2015 Published: December 9th, 2015

DOI: 10.5772/61797

From the Edited Volume


Edited by Matheus Poletto and Heitor Luiz Ornaghi Junior

Chapter metrics overview

4,069 Chapter Downloads

View Full Metrics


Currently some principles of sustainability, eco-efficiency and green chemistry are guiding the development of a new generation of materials as an alternative to conventional polymers based on petroleum. Then, in the field of biodegradable polymers one of the most promising investigations is focused on the use of microbial cellulose (MC), biocellulose or bacterial cellulose. MC has received substantial interest since it is synthesized from the bacterium Gluconacetobacter genus from a variety of carbon sources such as glucose, fructose, galactose, etc. MC is an interesting emerging biomaterial, with no toxicity, and since its discovery has shown tremendous potential in various fields, because the structural aspect of MC is far superior to those of plant cellulose. Thus, the main focus of the chapter review involves detailed aspects about the biosynthesis and recent advances on microbial production, including mechanism for the biochemistry of the cellulose synthesis, new sources for culture medium, main aspects about static and air-reactor productions and genetic modifications. We also revised microbial cellulose devices for biomedical applications: artificial skin, artificial blood vessels and microvessels, wound dressing of second- or third-degree burn ulcers, scaffolds for tissue engineering, drug delivery systems, dental implants, among others.


  • Microbial cellulose
  • cellulose synthesis
  • medical applications

1. Introduction

Microbial cellulose (MC) presents the same chemical formula as plant cellulose, however with the fibers in nanometer dimensions; confer different properties to MC [1]. The MC is a type of exopolysaccharides composed of glucose monomers bound by glycosidic β (1-4) linkages, with the chemical formula (C6H10O5)n, as can be seen in Figure 1. [2,3]. This biopolymer is produced extracellularly into nanofibers by several genera of bacteria, such as Gluconacetobacter, (formerly Acetobacter), Agrobacterium, Aerobacter, Achromobacter, Azotobacter, Rhizobium, Sarcina, and Salmonella [4].

Figure 1.

Chemical structure of microbial cellulose.

Historical data show that the MC has been used for a long time in the manufacture of a traditional food in the Philippines, known as the coconut cream [5]. Currently, the MC still remains widely utilized as food in various parts of the world however on the other hand aroused great academic and industrial interest due to its unique properties and diverse opportunity of applications. MC produced by Acetobacter xylinum was first reported in 1886 by Brown [6], produced in the presence of oxygen, using glucose as a carbon source.

As proposed by Yamada and colleagues (1997) [7] and subsequently validated by International Journal of Systematic Bacteriology, Acetobacter xylinum was reclassified and scientifically cataloged as Gluconacetobacter xylinus, due to the characteristics phylogeny based on analysis of partial sequences of 16S ribosomal RNA.

This bacterial species belonging to the family Acetobacteriaceae, being morphologically classified as a bacillus Gran-negative, strictly aerobic, no pathogenic which may be found singly arranged, in pairs or in small sets of chain formation of colonies shiny and smooth in Mannitol Agar. Bacteria belonging to this family are able to oxidize fully various carbon sources such as glucose, fructose, galactose, sucrose, mannitol, glycerol, inositol, among others [3,8] and alcohols such as ethanol [7] and is capable of extracellularly producing pulp at temperatures between 25 and 30 °C and pH 3 to 7. The bacterial cellulose may provide arrangements in parallel via hydrogen bonds and to form a tridimensional network. The morphology of the membrane depends directly on the environment and the interface culture medium / air where MC form a thick film, which can be easily manipulated according to the size of the vial used for cultivation [1,8].

Researchers have sought new bacterial strains capable to produce biopolymers with potential industrial application [9]. Although many organisms are capable to produce cellulose, Gluconacetobacter xylinus bacteria are the only known species able to produce cellulose on the industrial scale [10].

In addition G. xylinus, other micro-organisms are considered able to produce cellulose, among others they are Escherichia coli, Salmonella spp. [11] and Pseudomonas spp. [12,13]. The cellulose synthesis genes (MCsA, MCsB, MCsZ and MCsC) of these species were similar to those in G. xylinus [14]. Although these species are also capable to produce the bacterial cellulose, the fact that many of them are potentially pathogenic limits the commercial use of these biopolymers [12,13].

In nature, microorganisms which produce cellulose are usually found in symbiosis with other microorganisms. In the fermentation of Kombucha, for example, Zygosaccharomyces yeasts are used as microorganisms symbionts [15]. The Kombucha, also known as tea fungi and Haipao [16], is produced in China for over 2000 years, with a widely varied yeast population [17,18], whose function is to convert sucrose to organic acids, carbon dioxide and ethanol, the latter being used for the cellulose-producing bacteria for the production of acetaldehyde and acetic acid [19,20].


2. Biochemical and molecular mechanisms of bacterial cellulose biosynthesis by Gluconacetobacter xylinus

Although several species of microorganisms are capable to produce cellulose, G. xylinus is currently considered a model organism for the study these biopolymers [21]. MC biosynthesis consists of a complex process which involves first the polymerization of glucose residues in β1-4-glucan chain [21,22,23], followed by the extracellular secretion of the chains ending the linear arrangement and crystallization of glucan chains through hydrogen bridges and Van der Waals forces hierarchically arranged in strips [22], resulting in formation of a tough three dimensional structure called microfibrils. MC generated by this bacterial species has special characteristics of unidirectional polarity and variable thickness. The crystallization mechanism of the microfibrils in G. xylinum can give rise to two cellulose forms, if the microfibrils is oriented parallel arrangement is synthesized cellulose I, while if the arrangement is antiparallel microfibrils is obtained cellulose II [21]. In G. xylinus, MC synthesis depends on the cycle of pentoses and of the Krebs cycle [1,21,22] that perform respectively the oxidation function carbohydrates and oxidation of organic acids. A particularity G. xylinus is the inability to metabolize glucose anaerobically due to lack of phosphofructokinase-1, an enzyme responsible for catalyzing the reaction of phosphorylation of fructose-6-phosphate, fructose-1,6-bisphosphate, which prevents glycolysis. Thus, MC synthesis by G. xylinus results of a metabolic pool hexose phosphate which is produced directly by phosphorylating exogenous hexose or indirectly by the pentose phosphate pathway and gluconeogenesis. The hexose phosphate conversion of cellulose is direct and does not depend on the intermediate divisions carbon skeleton [1,22]. The conversion of glucose, transported from the external environment into the cytoplasm, is catalyzed by four bacterial enzymes, the glucokinase, which is the enzyme responsible for the phosphorylation of the carbon 6 of glucose, yielding glucose-6-phosphate, the phosphoglucomutase, which catalyzes the reaction isomerization of glucose-6-phosphate to glucose-1-phosphate, the UDPG-pyrophosphorylase (also known as glucose-1-phosphate uridylyltransferase), responsible for synthesis of UDP-glucose (UDPG), and cellulose synthase (CS), responsible for the polymerization of cellulose from UDP-glucose. As previously mentioned, the cellulose synthesis can also occur from endogenous sources, for gluconeogenesis. In G. xylinus, the synthesis from endogenous sources begins with oxaloacetate, into pyruvate by action of the enzyme pyruvate carboxylase. The transformation of the pyruvate in fosfoenolpiruvado, is produced by action of the enzyme phosphoenolpyruvate carboxykinase [1,22,24]. MC synthesis reaction is costly to the cell, consuming about 10% of the ATP generated in bacterial metabolism. Thus, the energy used for the synthesis of CB comes from aerobic metabolism. There are different proposals for the substrate used by the CS. [1,21]. The enzymatic complex of synthesis of cellulose, termed as terminal complex (TC) [25,26], constitute a kind of membrane protein species that, in Gluconacetobacter spp., corresponds to cellulose synthase complex [27].

One proposed hypothesis is that the UDP-glucose binds to lipids of the plasma membrane [1,21]. Another one considers that the soluble precursor interacts directly with the CS [28]. The CS is a protein complex consisting of three (AxCcSAB, AxCcSC and AxCcSD) or four (AxCcSA, AxCcSB, AxCcSC and AxCcSD) protein subunits encoded by genes exist in an operon chromosomal called MCs.

The two conserved Asp residues (D) invariably are found in loops at the Carboxyl-terminal (C-terminal) ends of predicted strands, a position frequently observed for catalytic residues [21, 39]. The hydrophobic clusters in domain B are more difficult to interpret in terms of secondary structure. AxCcSA and AxCcSAB have a motif consisting of domain, a single conserved residues Asp (D-D-D), presumably important for catalysis, identified along with the conserved sequence motif Gln (Q) Arg-Trp (R-W) in glucotransferases [1, 29]. Through a functional analysis of CS, it appears that the A subunit of this complex with 83kDa, shows catalytic activity. The B subunit of 90kDa, increases the rate of cellulose synthesis by joining a positive allosteric regulator, cyclic diguanosine monophosphate (c-di-GMP). The C subunit (138kDa) and D (17kDa) appear to structural activity. It has been hypothesized that C subunit related to pore formation and extrusion of the cellulose D subunit appears related to the process decrystalization since mutant strains of the gene which encodes the D subunit production are still able to produce cellulose II [23].

In G. xylinus Bis-(3'-5')-cyclic dimeric guanosine monophosphate (c-di-GMP) has been identified as an activator of cellulose biosynthesis [30], considered a second global messenger in bacteria [31]. The free c-di-GMP in the cell is considered to allosterically activate the cellulose synthase BcsA. However, 90% of the cellular c-di-GMP is reversibly bound by the c-di-GMP binding protein BcsB, a membrane protein that is structurally associated with the cellulose synthase [32,33]. It is believed that the spatial proximity is necessary to direct c-di-GMP released from BcsB towards the cellulose synthase. The equilibrium between bound and free c-di-GMP is modulated by the intracellular potassium concentration [34]. The level of free c-di-GMP is regulated by the opposing action of two enzymes, diguanylate cyclase (DGC) that cycles two molecules of GTP under the release of two molecules of PPi, and phosphodiesterase A (PDEA) that degrades c-di-GMP to the inactive GTP under the release of two molecules of PPi, and phosphodiesterase A (PDEA) that degrades c-di-GMP to the inactive 5’-pGpG. G. xylinus has three distinct operons each containing PDEA/DGC pair, which contribute at different levels to the c-di-GMP turnover [35, 36] indicating that cellulose biosynthesis underlies various control mechanisms in G. xylinus. Figure 2 represents a model of the metabolic pathway for the biosynthesis of cellulose by G. xylinus.

Figure 2.

Hypothetical model of the pathway for the biosynthesis of cellulose by G. xylinus from exogenous sources - glucokinase-ATP (1); Phosphoglucomutase (2), glucose-6-phosphate dehydrogenase (3); Phosphoglucoisomerase (4); Fructokinase ATP (5), Aldolase (6); Triosephosphate isomerase (7); Glyceraldehyde 3-phosphate dehydrogenase (8); Phosphoglycerate mutase (9), enolase (10); Pyruvate kinase (11); Pyruvate biphosphate kinase (12), pyruvate dehydrogenase(13); 6-phosphogluconate dehydrogenase (14); Phosphorribulose epimeraase (15); Phosphorribulose isomerase (16); Transaketolase (17); Transaldolase (18); Fructokinase (19); Aldehyde dehydrogenase(20); Alcohol dehydrogenase(21).


3. Cellulose Synthase (CS)

Two bacterial cellulose synthase operons (bcs) [23] were identified in the analyzed genome, but only one (bcsI) is structurally complete. This operon is composed by seven genes encoding for enzymes endo-1,4-beta-glucanases, a homolog CMCax is a putative beta-glucosidase endoglucanase, CPC is a putative homologues four subunits CS: BCSA, BBRC, BCSC and BCSD and beta-glucosidase. Although the three endoglucanases have been identified [37] its exact function in MC biosynthesis is not well understood. Genomic Analysis showed that bcsD gene is conserved as part of the operon and has exactly the same length in all six strains. Interestingly, this gene encodes for CS subunit D, whose function is still speculative, although their crystal structure has been recently resolved.

In cells actively producing cellulose, approximately 50 cellulose-synthesizing multienzyme complexes are organized in a single row along the longitudinal axis of the bacterial rod whereby each complex secretes approx. 12 to 25 glucan chains which assemble into larger microfibrils at the site of synthesis. This so-called linear terminal complex can be visualized by electron microscopy using freeze fracture as 35 Å pores in the outer membrane or as pits when the outer leaflet is fractured away [1,33].

In G. xylinus specie the cellulose synthase complex (BCS) is a multicomponent protein complex encoded in an operon containing at least three genes, bcsA, bcsB, and bcsC, which encodes a transmembrane complex over the cytoplasmic and outer membrane whereby the cellulose synthase (BcsA) and the c-di-GMP binding protein (BcsB) are considered to be localized in the cytoplasmic membrane as shown in Figure 3. [32,33,39]. Cellulose synthesis and transport across the inner bacterial membrane is mediated by a complex of the membrane-integrated catalytic BcsA subunit (green) and the membrane-anchored, periplasmic BcsB domain (blue) and membrane-associated regions BcsB transmembrane anchor (blue). The glycosyltransferase domain is shown brown [40]. BcsC is predicted to form a β-barrel in the outer membrane, preceded by a large periplasmic domain containing tetratricopeptide repeats likely involved in complex assembly [41,42].

The gene corresponding to cellulose synthase BcsA is constituted by a long string that presents between 723-880 amino acid residues, as represented in Figure 3, being the most conserved gene of the operon MCs between species, although the amino-terminus portions (N-terminus) and carboxyl-terminus portions (C-terminus) is not so conserved, since the homology is not restricted to the frequently analyzed D, D, D35Q (R, Q) and RW motif, which spans domains A and B [39].

Figure 3.

Schematic representation of cellulose synthase.

The BcsB protein [Figure 3], related to indirect interaction with c-di-GMP [33] is less well conserved among the species. However, direct comparisons of the MCsB proteins with CelB from A. tumefaciens and R. leguminosarum bv. trifolii revealed significant homology (∼40% similarity) over the entire length of the proteins with several invariable residues. An alanine/proline rich domain is located at the N-terminus region of all proteins except A. aeolicus. One transmembrane domain located at the C-terminus portions has been predicted by various algorithms for all BcsB proteins [39].

The proposed model considers that after the transfer and addition of glycosyl terminal residue, the glucose molecule rotates around acetyl glucan binding to align the channel as shown Figure 4. It is believed that allosteric interactions guide the direction of rotation, causing rotation feature 180⁰ connecting β-1,4 glucan-glucan between the individual glucose units and intramolecular hydrogen bond between oxygen atoms of the hydroxyl groups of the neighboring unit [40,43]. This phenomenon may be sufficient to allow the polymer to move into the channel [Figure 4]. Alternatively, for the translocation of the elongated glucan occurs, replacement is required to UDP-glucose by UDP. Past the channel, induced glucan chain in the BcsA - twist BcssB interface, interaction with BcsB of CBDs (periplasmic carbohydrate binding domains), or aggregation with other glucans may additionally contribute to a unidirectional motion of the polymer [40].

Figure 4.

Proposed model for cellulose synthesis and translocation (adapted). After glycosyl transfer, the newly added Glc could rotate around the acetyl linkage into the plane of the polymer. The rotation direction would be determined by steric interactions and formation of the β-1,4 glucan characteristic intramolecular O3- H••O5 hydrogen bond. The glucan might translocate into the channel during this relaxation. This process would be repeated with a second UDP-Glc but the rotation direction after glycosyl transfer would be in the opposite direction owing to steric constraints. Alternatively, the glucan might not translocate into the channel until UDP is replaced by UDP-Glc. Trp 383 and Cys 318 mark the entrance to the transmembrane channel [40].


4. Cultivation conditions for production of bacterial cellulose

The MC production depends of the appropriate cultivation conditions, which include the composition of the culture medium (synthetic and natural media), temperature, pH and methods agitated or static cultivation. The choice of condition cultivation or another depend on the purpose, once these conditions have significant influence on the properties of structure, physical and mechanical MC.

The current methods of MC production are static culture [44], submerged fermentation through aerated or agitated cultivation [45], and the airlift bioreactor [46]. Large scale, semi-continuous and continuous fermentation are dominant to meet commercial demand. In all cases, the main objective is to achieve maximum production of MC with optimum form and suitable properties for the application for which it is intended. After all, a wider application of this versatile biopolymer depends on the practical considerations such as the scale-up capability and production costs. G. xylinus has two main operative amphibolic pathways: the pentose phosphate cycle for the oxidation of carbohydrates and the Krebs cycle for the oxidation of organic acids and related compounds [12,21,47,48]. Consequently, several studies have been reported that the composition of the culture medium and the fermentation conditions significantly affect the order structure of cellulose [49,50,51].

Static cultivation is a relatively simple and widely used method of cellulose production. The medium is placed into shallow tray or bottles, inoculated, and cultivated for several days until the cellulose nearly fills the tray. G. xylinus produces a gelatinous MC membrane, which has a denser surface on the side exposed to air, i.e. capable of generating cellulose as an extracellular product on static media (at the air-medium interface) at temperatures between 25 and 30 °C and pH from 4 to 7 [52]. The traditional static culture represents an expensive way of MC production that may hinder its industrial application since the productivity is low and long cultivation time is required. Consequently, authors have proposed new culture system as strategy to increase the MC productivity to a suitable for commercial applications in simple fed-batch [53], in bioreactor for a semi-continuous production [54], in a modified airlift-type bubble column bioreactor [55].

Nutrients required for the growth of these microorganisms are carbon and nitrogen sources, phosphorus, Sulphur, potassium and magnesium salts [56]. Sometimes a complex medium supplying amino acids and vitamins is also used to enhance the cell growth and production [57]. Between the years 1940 to 1960, researchers at the Hebrew University in Jerusalem intensively investigated the biochemistry of simplified production and quantification of cellulose cellulose for produced by G. xylinus. These media, named as HS are widely used nowadays [44]. Typical carbon sources in the production of microbial cellulose include glucose, fructose, sucrose, mannitol, however among others including arabinose, arabitol, citric acid, ethanol, ethylene glycol, diethylene glycol, galactose, glucono lactone, glycerol, inositol, lactose, malic acid, maltose, mannose, methanol, rhamnose, ribose, sorbose, starch, succharide, succinic acid, trehalose, and xylose have been also investigated [58,59,60] to maximize bacterial cellulose production by various Gluconacetobacter strains [8]. The best yield was obtained in fed-batch fermentation, 15.3 g/L in 50 hours of cultivation using glucose as carbon source [61] and other examples of different culture conditions can be found in Table 1. The results shown in Table 1 indicate that, glucose seems to be the best carbon source. It demonstrates the validity of the results that various carbon substrates could be converted to monomer glucose by Gluconacetobacter, followed by polymerization to MC [60]. Among the factors affecting cost, carbon sources play a major role in fermentation [47,62]. Conversion of 60–80% of the utilized carbon source into crude polymer is commonly found in high yielding polysaccharide fermentations. In order to compensate its low sugar conversion yield and to reduce the feedstock cost of MC production, in recent years, MC has been produced by fermenting the hydrolysates of agricultural wastes such as hemicelluloses [63], konjac powder [62], rice bark [64] and waste cotton fabrics [63,65]. An advantage of using agricultural or industrial residual streams as feedstock is the low or no value of the raw material. Several successful efforts have been made to use certain industrial food wastes as growth medium for the MC producer organisms, which is not only a cheap way but also works as a basin for environmental cleaning [51]. Thin stillage (TS) is a wastewater from rice wine distillery rich in carbon sources and organic acids. [66] discovered that TS, when employ to replace distilled water for preparing Hestrin and Schramm medium (the traditional MC production medium), can enhance the MC production 2.5-fold to a concentration of 10.38 g/L with a sugar-MC conversion yield of 57 % (0.57 g MC/g reducing sugar) after 7 days of static cultivation. In 2012, Ha et al. [48] further improved the MC production, 15.28 g/L of MC was obtained after 15 days of cultivation. Yeast extract and peptone are the most commonly used nitrogen sources in MC production as they provide nitrogen and growth factors for Gluconacetobacter strains. Many researchers are trying to find efficient substitutes due to their high cost. Even if various nitrogen sources were added to the HS medium, peptone is found to be the most effective nutrient. However, corn steep liquor (CSL) which produced the second highest production is always chosen as a substitution for the economic viewpoints. Buffering capacity is also important for MC production. Insoluble MC often attaches to pH probe and leads to inaccurate reading [4]. Noro et al. [67] pointed out the buffering capacity of CSL, which could maintain the pH within the optimal range during the production of MC. Jung et al. [68] doubled MC production (from 1.53 to 3.12 g/L) using molasses as carbon source and corn steep liquor as nitrogen source when compared with the results obtained from complex medium. This strategy could not only reduce burden on environment but also achieve the goal of large scale production with low cost. The optimal pH for MC production may vary with carbon source. G. xylinus accumulates gluconic acid at low pH, and a preferred environment for both biomass and MC production can be achieved by shifting pH from 4.0 to 5.5 during cellulose production phase in fed-batch cultures [61]. Under static batch cultivation, the pH of the culture medium decreases due to the respiratory metabolism of G. xylinus, which involves the ethanol oxidation to acetic acid and the glucose conversion into gluconic acid. This fact makes it very important to control the pH within the optimum range for cell growth and cellulose production [48,69]. Gluconacetobacter strains require oxygen as an essential substrate, consequently volumetric oxygen transfer coefficient (kLa) is a key limiting factor in the aerobic fermentation for producing MC. Song et al. [55] investigated the optimum aeration rate for a 50-L spherical type bubble column bioreactor, and it was determined to be 1.0 vvm (30 L/min).

Attempts to enhance MC production by adding different additives in the fermentation medium have been made. The possible mechanisms of these various additives to enhance MC were also proposed such as reduction of the shear force by increasing the viscosity of medium [70]. Different chemical compounds including alcohols [72], glycerol [60,68], organic acids [68], polysaccharides [72] thin stillage from rice wine distillery [66] and thin stillage from beer culture broth [47,53] have been used as additives to the fermentation medium with the aim of increasing MC production.

Microorganism Carbon source Cultivation
Time culture
MC production
(g l-1)
G. xylinus (BRC 5) Glucose Fed-batch 2 15.3 Hwang et al. (1999)[61]
G. xylinus (BPR 2001) Fructose Agitated 3 14.1 Bae et al. (2004) [70]
G. xylinus (KJ1)
Saccharified food wastes airlift-type bioreactor 3 5.6 Song et al. (2009)[55]
A. xylinum (ATCC 700178) CSL-Fru Agitated 5 13.0 Cheng et al. (2011) [72]
Gluconacetobacter sp (F6) Glucose Static 6 4.5 Jahan et al. (2012) [73]
G. xylinus
(MCRC 12334)
TS-Glu Static 7 10.38 Wu et al. (2012)[66]
G. xylinus and Trichoderma ressei Glucose Static 14 6.23 Cavka et al. (2013)[74]
G. xylinus
(PTCC, 1734)
Syrup Static 14 43.5 Moosavi-Nasab
and Yousefi (2011) [75]
G. xylinus
(ATCC 23769)
Glucose Static 15 15.28 Ha and Park (2012)[76]

Table 1.

Bacterial cellulose production under different culture conditions


5. Microbial cellulose for biomedical applications

As previously described, intense research has focused on the use of natural biopolymers in a variety of biomedical materials and devices, including wound dressings, medical implants, drug delivery, vascular grafts, and scaffolds for tissue engineering [78]. Consequently, continual efforts from many researchers, led to novel systems that closely mimic the complex and hierarchical structures inherent to the native tissue are sure to emerge.

In the last decade, several nanocellulose-based materials have been created for a diversity of biomedical applications. Some review articles have highlighted the potential applications of cellulose materials [72,79,80,81,82].

MC represents an interesting emerging nanomaterial, with no toxicity, and since its discovery has shown tremendous potential as an effective biopolymer which offers a wide range of applications, especially the biomedical ones, including the use as biomaterial for artificial skin, artificial blood vessels and microvessels, wound dressing of second- or third-degree burn ulcers and dental implants. Other studies with endothelial, smooth muscle cells and chondrocytes have shown that these cells present good adhesion to bacterial cellulose. [83]

5.1. Pristine MC based biomaterials

One of the main direct applications of MC membranes in biomedical field is related to wound dressing. Fontana et al. [84] were the pioneers in describing the use of bacterial cellulose to replace burned skin. Since then, literature shows a great number of papers related to wound dressing. Cellulose dressings are recommended as a temporary covering for the treatment of wounds, including pressure sores, skin tears, venous stasis, ischemic and diabetic wounds, second-degree burns, skin graft donor sites, traumatic abrasions and lacerations, and biopsy sites by the manufacturers [85].

MC based wound dressings are commonly available on the market nowadays, for example: BioFill®, Bioprocess®, XCell® and Gengiflex® (for periodontal diseases reconstruction [86]. The biomembrane BioFill® was one of the first commercial product that fulfills the main prerequisites of an ideal wound dressing, including: low cost, good adherence to the wound, water vapor permeability, elasticity, transparency, durability, it constitutes a physical barrier for bacteria, is hemostatic, it presents easy handling and application with minimum exchanges. BioFill® effectiveness has been proven in more than 300 cases in accelerating the healing process, pain relief, etc. [86,87,88,89].

Despite the analgesic mechanism of action of these dressings has not been fully elucidated, some authors suggest that the healing mechanism involves the capture of ions by means of cellulose hydrogen bonds, or the nano MC 3-D network mimics the skin surface creating optimal conditions for healing or regeneration [87,88,89].

It is important to point out that MC wound dressing clearly shortened the time to heal or wound closure over standard care when applied to non-healing lower extremity ulcers, as observed by many researchers [88,89,90]. As can be seen in Figure 5, novel applications of wet MC as wound dressing in the treatment partial thickness burns were applied by Czaja et al. [88,89] presenting excellent results suggesting that MC as a wound dressing promotes a favorable moist environment for a fast wound cleansing, and consequently for rapid healing.

Figure 5.

Bacterial cellulose dressing applied on wounded torso and face. (Reprinted with permission from Czaja, W. K. et al. (2007). The future prospects of microbial cellulose in biomedical applications, Biomacromolecules, Vol.8, No.1, pp. 4. Copyright (2007) American Chemical Society).

Other interesting biomedical applications for MC films have emerged. A Brazilian research group had designed and patented a device to manufacture MC contact lens for therapy in cases of regeneration of cornea [91].

In terms of vascular applications, a German group created BASYC® (Bacterial Cellulose Synthetised), consisting of a tubular biomaterial for applications in microsurgery of arteries and veias [92] and [93] applied vascular stents in animals arteries. Bodin et al. [94] obtained MC tubes by modifying the fermentation process of Acetobacter xylinum on top of silicon tubes, as shown in Figure 6.

Figure 6.

MC tubes presenting different sizes and shapes for applications: a) Mc tubes showing different inner diameters: 1.5mm, 2.4mm, 3.0mm, 4.0 mm and 6.0 mm. b) Branched MC tube fermented on a branched silicone tube. (Reprinted with permission from Bodin et al., 2007 Biotechnology and Bioengineering, Vol. 97, No. 2, June 1, 2007 [94])

Lately, Nimeskern et al. [95] designed and fabricated an ear-shaped pristine MC prototype material applying a Magnetic Resonance Imaging (MRI) scanning technique. This study was extremely important to confirm that MC is a promising tissue engineering material with appropriate mechanical properties for ear cartilage replacement. Thereby, it may be used to create patient-specific ear shapes.

5.2. Microbial cellulose nanocomposites for biomedical applications

Beyond the direct uses, MC can be widely and effectively utilized as either functional reinforcements or excellent matrices due to excellent mechanical properties and biocompatibility which allows it to be engineered in various forms from nano to macro scale. Thus, MC based nanocomposites can be manipulated to improve their properties and/or functionalities becoming one of the reasons that makes MC so exceptional material for biomedical applications.

In relation to biomaterial applications for wound dressing and skin tissue repair several MC based nanobiocomposites were fabricated. Here are some examples: membranes of MC/collagen [96], MC/gelatin [97], MC/ aloe vera films [98], MC/alginate for temporary dressing material [99].

Further, MC composite with kaolin was proved as short-term and long term wound healing materials [100].

Freeze-drying techniques allowed the preparation MC/poly(ethylene glycol) (PEG) composites of by immersing wet MC pellicle in PEG aqueous solution [101]. This same technique was applied by some of us [102] to obtain MC/ silk fibroin (SF) sponge scaffolds. In vitro tests proved non-cytotoxic or genotoxic character of these nanobiocomposites. SEM images revealed a greater number of fibroblast cells (L929 cell line) attached at the MC/SF:50% scaffold surface if compare with the surface of pure MC, suggesting that the presence of fibroin improved cell attachment as is possible to see in Figure 7. This could be related to the SF amino acid sequence that act as cell receptors facilitating cell adhesion and growth. Consequently, MC/SF:50% scaffolds configured an excellent option in bioengineering depicting its potential for tissue regeneration and cultivation of cells on nanobiocomposites.

Figure 7.

To test the hypothesis that the addition of silk fibroin to cellulose scaffolds increases cell adhesion (48 h), L-929 cells were seeded in MC and MC/SF scaffolds. SEM images of the cells attached to MC (a) and MC/SF (b) scaffolds surface; cross-section SEM images of MC (c) and MC/SF (d) evidenced that the cells did not migrate into the scaffolds. (Reprinted with permission from Oliveira Barud et al., 2015, Carbohydrate Polymers, Vol 128, April, 2015 [102]).

Generally, a scaffold provides a foundation for cell attachment, and several materials have been tested as scaffolds to support growth of cells. The need for bio-mimicking scaffolds has led to the exploration of MC as a scaffold material. There is an increased interest in developing adipose tissue as an in vitro model for adipose biology and metabolic disease, and to this end, 2D and 3D porous scaffolds of bacterial nanocellulose and alginate were prepared recently [103].

Bäckdahl et al. [104] also developed MC scaffolds with controlled microporosity by placing paraffin wax and starch particles during culture and removing these particles once the cultivation process was finished. The MC scaffolds were then seeded with smooth muscle cells for investigating the potential tissue engineered blood vessel application.

A variety of surface functionalization through biosynthetic or chemical modification was also investigated. Various approaches to the preparation of functional MC-based nanocomposites by incorporating different guest substrates including small molecules, inorganic nanoparticles or nanowires, and polymers on the surfaces of MC nanofibers are exemplified which can improve the functionality of MC nanomaterials and expand its potential application in the biomedical fields.

Nanocomposites were obtained by the association of nanoparticles presenting antimicrobial activities, including silver nanoparticles [105,106,107,108,109,110]. Additionally, Barud et al. [111] also prepared MC/propol membranes that presented good antimicrobial activities to be used as wound dressing material.

In recent years, several controlled release systems based on nanocellulose material for various pharmaceutical applications have been also investigated to delivery Tetracycline [112], benzalkonium chloride [113], topical release of lidocaine [114] and release of proteins with serum albumin [115].

With respect to bone regeneration in defects of rat tibiae, MC-hydroxyapatite (MCHA) nanobiocomposite were prepared to evaluate the biological properties and performance of the material [116]. The MCHA membranes were effective for bone regeneration and accelerated new bone formation. In addition, reabsorption of the membranes was slow, suggesting that this composite takes time to be completely reabsorbed.


6. Conclusion

Microbial cellulose is a natural renewable polymer synthesized from the bacterium Gluconacetobacter xylinus that is the only known species capable to produce cellulose on an industrial scale. In an appropriate culture medium the bacteria secretes about 50-80 cellulose microfibrils from 3.0 to 3.5 mm thick, free of lignin and hemicellulose, which aggregate themselves to form strips arranged in a 3-D hierarchical network. Besides that MC configures one of the most promising investigations in the field of biodegradable polymers. Due to this uniform structure and morphology MC is endowed with unique characteristics such as high purity, high crystallinity and remarkable mechanical properties, good chemical stability, high water holding capacity featuring it as a completely biocompatible polymer. Despite its high water content, MC shows a good mechanical performance and it can be produced in almost any shape due to its high moldability during formation. MC is an interesting emerging biomaterial, with no toxicity, and since its discovery has shown tremendous potential as an effective biopolymer in various fields, because the structural aspect of MC is far superior to those of plant cellulose. Thus, this chapter reviewed involved detailed aspects about the biosynthesis and recent advances on microbial production, including mechanism for the biochemistry of the cellulose synthesis, new sources for culture medium, main aspects about static and air-reactor productions and genetic modifications. We also revised and presented a great number of different MC based materials that were designed for biomedical applications (dressings, scaffolds, drug delivery systems), among others. Additionally, we hope that this book chapter may aggregate high quality information and may be a benchmark to intensify greater interest of the scientific community in microbial cellulose and related devices and also inspire the development of new materials in this field.


  1. 1. Donini ÍAN, De Salvi DTB, Fukumoto FK, Lustri WR, Barud HS, Marchetto R, Messaddeq Y, Ribeiro SJL. Biossíntese e recentes avanços na produção de celulose bacteriana. Eclet. Quím. 2010; 35:165-173. DOI: 10.1590/S0100-46702010000400021
  2. 2. Phisalaphong M, Jatupaiboon N. Biosynthesis and characterization of bacteria cellulose–chitosan film. Carbohydr. Polym. 2008;78:482-488. DOI: 10.1016/j.carbpol.2008.04.004
  3. 3. Sheykhnazari S, Tabarsa T, Ashori A, Shakeri A, Golalipour M. Bacterial synthesized cellulose nanofibers. Effects of growth times and culture mediums on the structural characteristics. Carbohydr. Polym. 2011;86:1187-1191. DOI: 10.1016/j.carbpol.2011.06.011.
  4. 4. Shoda M. Sugano Y. Recent advances in bacterial cellulose production. Biotechnol. Bioprocess Eng. 2005;10:1-8. DOI: 10.1007/BF02931175
  5. 5. Lapuz MM, Gallardo EG, Palo MA. The nata organism-cultural requirements, characteristics and identity. Philipp. J. Sci. 1967;96:91-108.
  6. 6. Brown AJ. On an acetic ferment which forms cellulose. J. Chem. Soc. Trans. 1886;49:432-439. DOI: 10.1039/CT8864900432
  7. 7. Yamada Y, Hoshino K, Ishikawa T. The phylogeny of acetic acid bacteria based on the partial sequences of 16S ribosomal RNA: The elevation of the subgenus Gluconoacetobacter to the generic level. Biosci. Biotechnol. Biochem. 1997;6:1244-1251. DOI: 10.1271/bbb.61.1244
  8. 8. Ruka DR, Simon GP, Dean KM. Altering the growth conditions of Gluconacetobacter xylinus to maximize the yield of bacterial cellulose. Carbohydr. Polym. 2012;89:613-622. DOI: 10.1016/j.carbpol.2012.03.059
  9. 9. Ashtaputre AA, Shah AK. Studies on a viscous, gel-forming exopolysaccharide from Sphingomonas paucimobilis GS1. Appl. Environ. Microbiol. 1995;61:1159-1162.
  10. 10. Klemm D, Heublein B, Fink HP, Bohn A. Cellulose: fascinating biopolymer and sustainable raw material. Angewandte Chemie. 2005;44:3358-3393. DOI: 10.1002/anie.200460587
  11. 11. Zogaj X, Nimtz M, Rohde M, Bokranz W, Römling U. The multicellular morphotypes of Salmonella typhimurium and Escherichia coli produce cellulose as the second component of the extracellular matrix. Mol. Microbiol. 2001;39:1452-63. DOI: 10.1046/j.1365-2958.2001.02337.x
  12. 12. Sutherland IW. Microbial polysaccharides from Gram negative bacteria. Int. Dairy. J. 2001;11:663-674. DOI: 10.1016/S0958-6946(01)00112-1
  13. 13. Souza DM, Garcia-Cruz CH. Produção fermentativa de polissacarídeos extracelulares por bactérias. Semina: Ciências Agrárias. 2004;25:331-340. DOI: 10.5433/1679-0359.2004v25n4p331
  14. 14. Solano C, Garcia B, Valle J, Berasain C, Ghigo JM, Gamazo C, Lasa I. Genetic analysis of Salmonella enteritidis biofilm formation: critical role of cellulose. Mol. Microbiol. 2002;43:793-808. DOI: DOI: 10.1046/j.1365-2958.2002.02802.x
  15. 15. Marsh AJ, O’Sullivan O, Hill C, Ross RP, Cotter PD. Sequence-based analysis of the bacterial and fungal compositions of multiple kombucha (tea fungus) samples. Food Microbiol. 2014;38:171-178. DOI: 10.1016/
  16. 16. Liu CH, Hsu WH, Lee FL, Liao CC. The isolation and identification of microbes from a fermented tea beverage, Haipao, and their interactions during Haipao fermentation. Food Microbiol. 1996;13:407-415. DOI: 10.1006/fmic.1996.0047
  17. 17. Mayser P, Fromme S, Leitzmann C, Grunder K. The yeast spectrum of the ‘tea fungus kombucha’. Mocoses. 1995;38:289-295.
  18. 18. Teoh AL, Heard G, Cox J. Yeast ecology of kombucha fermentation. Int. J. Food Microbiol. 2004;95:119-126. DOI: 10.1016/j.ijfoodmicro.2003.12.020
  19. 19. Greenwalt CJ, Ledford RA, Steinkraus KH. Determination and characterization of the antimicrobial activity of the fermented tea kombucha. Lebensm. Wiss. Technol. 1998;31:291-296. DOI: 10.1006/fstl.1997.0354
  20. 20. Dufresne C, Farnworth E. Tea, Kombucha, and health: a review. Food Res. Int. 2000;33:409-421. DOI: 10.1016/S0963-9969(00)00067-3
  21. 21. Chávez-Pacheco JL, Martínez-Yee S, Contreras-Zentella M, Escamilla-Marván E, Celulosa bacteriana en gluconacetobacter xylinum: biosíntesis y aplicaciones Tip Revista Especializada en Ciencias Químico-Biológicas. 2004;7:18-25.
  22. 22. Ross P, Mayer R, Benziman M. Cellulose biosynthesis and function in bacteria. Microbiol. Rev. 1991;55:35-58.
  23. 23. Saxena IM, Kudlicka K, Okuda K, Brown RM Jr. Characterization of genes in the cellulose-synthesizing operon (acs operon) of Acetobacter xylinum: implications for cellulose crystallization. J. Bacteriol. 1994;176:5735-5752.
  24. 24. Krystynowicz A, Koziolkiewicz M, Wiktorowska-Jezierska A, Bielecki S. Klemenska E, Masny A, Płucienniczak A. Molecular basis of cellulose biosynthesis disappearance in submerged culture of Acetobacter xylinum. Acta Biochim. Pol. 2005;52:691-698.
  25. 25. Brown Jr RM. Cellulose microfibril assembly and orientation: recent developments. J. Cell. Sci. Suppl. 1985;2:13–32.
  26. 26. Okuda K. Structure and phylogeny of cell coverings. J. Plant. Res. 2002;115:283-288. DOI: 10.1007/s10265-002-0034-x
  27. 27. Sunagawa N, Fujiwara T, Yoda T, Kawano S, Satoh Y, Yao M, Tajima K, Dairi T. Cellulose complementing factor (Ccp) is a new member of the cellulose synthase complex (terminal complex) in Acetobacter xylinum. J. Biosci. Bioeng. 2013;115:607-612. DOI: 10.1016/j.jbiosc.2012.12.021
  28. 28. Brown Jr RM, Saxena IM. Cellulose biosynthesis: A model for understanding the assembly of biopolymers. Plant Physiol. Biochem. 2000;38:57-67. DOI: 10.1016/S0981-9428(00)00168-6
  29. 29. Kawano S, Yasutake Y, Tajima K, Satoh Y, Yao M, Tanaka I, Munekata M. Crystallization and preliminary crystallographic analysis of the cellulose biosynthesis-related protein CMCax from Acetobacter xylinum. Acta Crystallogr. Sect. F Struct. Biol. Crystall. Commun. 2005;61:252-254. DOI: 10.1107/S174430910500206X
  30. 30. Ross P, Weinhouse H, Aloni Y, Michaeli D, Weinberger-Ohana P, Mayer R, Braun S, de Vroom E, van der Marel GA, van Boom JH, Benziman M. Regulation of cellulose synthesis in Acetobacter xylinum by cyclic diguanylic acid Nature 1987;325:279-281. DOI: 10.1038/325279a0
  31. 31. Römling U, Gomelsky M, Galperin MY. C-di-GMP: the dawning of a novel bacterial signalling system. Mol. Microbiol. 2005;57:629-639. DOI: 10.1111/j.1365-2958.2005.04697.x
  32. 32. Kimura S, Chen HP, Saxena IM, Brown JrRM, Itoh T. Localization of c-di-GMP-binding protein with the linear terminal complexes of Acetobacter xylinum. J. Bacteriol. 2001;183:5668-5674. DOI: 10.1128/JB.183.19.5668-5674.2001
  33. 33. Mayer R, Ross P, Weinhouse H, Amikam D, Volman G, Ohana P, Calhoon RD, Wong HC, Emerick AW, Benziman M. Polypeptide composition of bacterial cyclic diguanylic acid-dependent cellulose synthase and the occurrence of immunologically crossreacting proteins in higher plants, Proc. Natl. Acad. Sci. USA. 1991;88:5472-5476.
  34. 34. Weinhouse H, Sapir S, Amikam D, Shilo Y, Volman G, Ohana P, Benziman M. C-di-GMP-binding protein, a new factor regulating cellulose synthesis in Acetobacter xylinum. FEBS Lett. 1997;416:207-211. DOI: 10.1016/S0014-5793(97)01202-7
  35. 35. Tal R, Wong HC, Calhoon R, Gelfand D, Fear AL, Volman G, Mayer R, Ross P, Amikam D, Weinhouse H, Cohen A, Sapir S, Ohana P, Benziman M, Three cdg operons control cellular turnover of cyclic di-GMP in Acetobacter xylinum: Genetic organization and occurrence of conserved domains in isoenzymes. J. Bacteriol. 1998;180:4416-4425.
  36. 36. Fujiwara T, Komoda K, Sakurai N, Tajima K, Tanaka I, Yao M. The c-di-GMP recognition mechanism of the PilZ domain of bacterial cellulose synthase subunit A. Biochem. Biophys. Res. Commun. 2013;431:802-807. DOI: 10.1016/j.bbrc.2012.12.103
  37. 37. Koo HM, Song SH, Pyun YR, Kim YS. Evidence that a beta-1,4-endoglucanase secreted by Acetobacter xylinum plays an essential role for the formation of cellulose fiber. Biosci. Biotechnol. Biochem. 1998;62:2257-2259. DOI: 10.1271/bbb.62.2257
  38. 38. Legnani C, Vilani C, Calil VL, Barud HS, Quirino WG, Achete CA, Ribeiro SJL, Cremona M. Bacterial cellulose membrane as flexible substrate for organic light emitting devices. Thin Solid Films 2008;517:1016-1020. DOI: 10.1016/j.tsf.2008.06.011
  39. 39. Römling U. Molecular biology of cellulose production in bacteria research in microbiology. 2002;153:205-212.
  40. 40. Morgan JLW, Strumillo J, Zimmer J. Crystallographic snapshot of celulose synthesis and membrane translocation. Nature. 2013;493:181-187. DOI: 10.1038/nature11744
  41. 41. Keiski CL, Harwich M, Jain S, Neculai AM, Yip P, Robinson H, Whitney JC, Riley L, Burrows LL, Ohman DE, Howell PL. AlgK is a TPR-containing protein and the periplasmic component of a novel exopolysaccharide secretin. Structure 2010;18:265-273. DOI: 10.1016/j.str.2009.11.015
  42. 42. Omadjela O, Narahari A, Strumillo J, Mélida H, Mazur O, Bulone V, Zimmer J. MCsA and MCsB form the catalytically active core of bacterial cellulose synthase sufficient for in vitro cellulose synthesis. Proc. Natl. Acad. Sci. USA. 2013;110:17856-17861. DOI: 10.1073/pnas.1314063110
  43. 43. Nishiyama Y, Langan P, Chanzy H. Crystal structure and hydrogen-bonding systemin cellulose Ib from synchrotron X-ray and neutron fiber diffraction. J. Am. Chem. Soc. 2002;124:9074-9082. DOI: 10.1021/ja0257319
  44. 44. Hestrin S, Schramm M. Synthesis of cellulose by Acetobacter xylinum. II. Preparation of freeze-dried cells capable of polymerizing glucose to cellulose. Biochem. J. 1954;58:345-352.
  45. 45. Toyosaki H, Naritomi T, Seto A, Matsuoka M, Tsuchida T, Yoshinaga F. Screening of bacterial cellulose-producing Acetobacter strains suitable for agitated culture. Biosci. Biotechnol. Biochem. 1995;59:1498-1502. DOI: 10.1271/bbb.59.1498
  46. 46. Cheng HP, Wang PM, Chen JW, Wu WT. Cultivation of Acetobacter xylinum for bacterial cellulose production in a modified airlift reactor. Biotechnol. Appl. Biochem. 2002;35:125-32. No tiene
  47. 47. Bielecki S, Krystynowicz A, Turkiewicz M, Kalinowska H. Bacterial Cellulose. Biopolymers Online. Wiley-VCH Verlag GmbH & Co. KGaA 2005. DOI: 10.1002/3527600035.bpol5003
  48. 48. Ha JH, Shah N, Ul-Islam M, Khan T, Park JK. Bacterial cellulose production from a single sugar a-linked glucuronic acid-based oligosaccharide. Process. Biochem. 2011;46:1717-1723. DOI: 10.1016/j.procbio.2011.05.024
  49. 49. Watanabe K, Tabuchi M, Ishikawa A, Takemura H, Tsuchida T, Morinaga Y, Yoshinaga F. Acetobacter xylinum mutant with high cellulose productivity and an ordered structure. Biosci. Biotechnol. Biochem. 1998;62:1290-1292. DOI:10.1271/bbb.62.1290
  50. 50. Duta FP, França FP, Lopes LMA. Optimization of culture for exopolysaccharide production in Rhizobium sp. using the response surface method. Electron. J. Biotechnol. 2006;9: 391-399. DOI: 10.2225
  51. 51. Khan T, Khan S, Park JK. Simple fed-batch cultivation strategy for the enhance production of a single-sugar glucoronic acid-based oligosaccharides by a cellulose-producing Gluconacetobacter hansenii strain. Biotechnol. Bioprocess. Eng. 2008;13:240-247. DOI: 10.1007/s12257-007-0212-8
  52. 52. Iguchi M, Yamanaka S, Budhiono A. Bacterial cellulose - a masterpiece of nature’s arts. J. Mater. Sci. 2000;35:261-270. DOI: 10.1023/A:1004775229149
  53. 53. Shezad O, Khan S, Khan T, Park JK. Physicochemical and mechanical characterization of bacterial cellulose produced with an excellent productivity in static conditionsusing a simple fed-batch cultivation strategy. Carbohydr. Polym. 2010;82:173-180. DOI: 10.1016/j.carbpol.2010.04.052
  54. 54. Kralisch D, Hessler N, Klemm D, Erdmann R, Schmidt W. White biotechnology for cellulose manufacturing - The HoLiR concept. Biotechnol. Bioeng. 2010;105:740-747. DOI: 10.1002/bit.22579
  55. 55. Song H-J, Li H, Seo J-H, Kim M-J, Kim S-J. Pilot-scale production of bacterial cellulose by a spherical type bubble column bioreactor using saccharified food wastes. Korean J. Chem. Eng. 2010;26:141-146. DOI: 10.1007/s11814-009-0022-0
  56. 56. Cherian BM, Leão AL, Souza SF, Oliveira GM, Costa LMM, Brandão CVS, Narine SS. Bacterial nanocelulose for medical implants. In: Thomas, S., Visakh, P.M., Mathew, A.P. editors. Advances in natural polymers. Berlin Heidelberg: Springer, 2013. DOI: 10.1007/978-3-642-20940-6_10
  57. 57. Chawla PR, Bajaj IB, Survase SA, Singhal RS. Microbial cellulose: Fermentative production and applications. Food Technol. Biotechnol. 2009;47:107-124.
  58. 58. El-Saied H, El-Diwany AI, Basta AH, Atwa NA, El-Ghwas DE. Production and characterization of economical bacterial cellulose. BioResources. 2008;3:1196-1217.
  59. 59. Hutchens SA, Benson RS, Evans BR, O'Neill HM, Rawn CJ Biomimetic synthesis of calcium-deficient hydroxyapatite in a natural hydrogel. Biomaterials. 2006;27:4661-4670. DOI: 10.1016/j.biomaterials.2006.04.032
  60. 60. Mikkelsen D, Flanagan BM, Dykes GA, Gidley MJ. Influence of different carbon sources on bacterial cellulose production by Gluconacetobacter xylinus strain ATCC 53524. J. Appl. Microbiol. 2009;107:576-583. DOI: 10.1111/j.1365-2672.2009.04226.x
  61. 61. Hwang JW, Yang YK, Hwang JK, Pyun Y and Kim YS. Effects of pH and dissolved oxygen on cellulose production by Acetobacter xylinum BRC5 in agitated culture. J. Biosci. Bioengineer. 1999;88:183-188. DOI: 1016/S1389-1723(99)80199-6
  62. 62. Hong F, Qiu K. An alternative carbon sources from konjac powder for enhancing production of bacterial cellulose in static cultures by a model strain Acetobacter aceti subsp. xylinus ATCC23770. Carbohydr. Polym. 2008;72:545. DOI: 10.1016/j.carbpol.2007.09.015
  63. 63. Hong B, Xue G, Weng L, Guo X. Pretreatment of moso bamboo with dilute phosphoric acid. BioResources. 2012;7:4902-4913.
  64. 64. Goelzer FDE, Faria-Tischer PCS, Vitorino JC, Sierakowaki MR, Tischer CA. Production and characterization of nanospheres of bacterial cellulose from Acetobacter xylinum from processed rice bark. Mater. Sci. Enginner.: C. 2009;29:546-551. DOI: 10.1016/j.msec.2008.10.013
  65. 65. Kuo CH, Lin PJ, Lee CK. Enzymatic saccharification of dissolution pretreated wast cellulosic fabrics for bacterial cellulose production by Gluconacetobacter xylinus. J. Chem. Technol. Biotechnol. 2010;85: 1346-1352. DOI: 10.1002/jctb.2439
  66. 66. Wu JM, Liu RH. Thin stillage supplementation greatly enhances bacterial cellulose production by Gluconacetobacter xylinus. Carbohydr. Polym. 2012;90:116-121. DOI: 10.1016/j.carbpol.2012.05.003
  67. 67. Noro N, Sugano Y, Shoda M. Utilization of the buffering capacity of corn steep liquor in bacterial cellulose production by Acetobacter xylinum. Appl. Microbiol. Biotechnol. 2004;64:199-205. DOI: 10.1007/s00253-003-1457-6
  68. 68. Jung HI, Jeong JH, Lee OM, Park GT, Kim KK, Park HC, Lee SM, Kim YG, Son HJ. Influence of glycerol on production and structural-physical properties of cellulose from Acetobacter sp. V6 cultured in shake flasks. Bioresour. Technol. 2010;101:3602-3608. DOI: 10.1016/j.biortech.2009.12.111
  69. 69. Kongruang S. Bacterial cellulose production by Acetobacter xylinum strains from agricultural waste products. Appl. Biochem. Biotechnol. 2008;148:245-256. DOI: 10.1007/s12010-007-8119-6
  70. 70. Bae S, Sugano Y, Shoda M. Improvement of bacterial cellulose production by addition of agar in a jar fermentor. J. Biosci. Bioeng. 2004;97:33-38. DOI: 10.1016/S1389-1723(04)70162-0
  71. 71. Lu XK, Shen XY. Solubility of bacteria cellulose in zinc chlorid aqueous solution. Carbohydr. Polym. 2011;86:239-244. DOI: 10.1016/j.carbpol.2011.04.042
  72. 72. Shah N, Ha JH, Park JK. Effect of reactor surface on production of bacterial cellulose and water soluble oligosaccharides by Gluconacetobacter hansenii PJK, Biotechnol. Bioproc. Engineer. 2010;15:110-118. DOI: 10.1007/s12257-009-3064-6
  73. 73. Cheng KC, Catchmark JM, Demirci A. Effects of CMC addition on bacterial cellulose production in a biofilm reactor and its paper sheets analysis. Biomacromol. 2011;12:730-736. DOI: 10.1021/bm101363t
  74. 74. Jahan F, Kumar V, Rawat G, Saxena RK. Production of microbial cellulose by a bacterium isolated. Appl. Biochem. Biotechnol. 2012;167:1157-1171. DOI: 10.1007/s12010-012-9595-x
  75. 75. Cavka A, Guo X, Tang SJ, Winestrand S, Jönsson LJ and Hong F. Production of bacterial cellulose and enzyme from waste fiber sludge. Biotechnol. Biofuels. 2013;6:25. DOI: 10.1186/1754-6834-6-25.
  76. 76. Moosavi-Nasab M, Yousefi A. Biotechnological production of cellulose by Gluconacetobacter xylinus from agricultural waste. Iranian J. Biotechnol. 2011;9:94-101.
  77. 77. Ha JH, Park JK. Improvement of bacterial cellulose production in Acetobacter xylinum using byproduct produced by Gluconacetobacter hansenii. Korean J. Chem. Eng. 2012;29:563-566. DOI: 10.1007/s11814-011-0224-0
  78. 78. Reis RL, Neves NM, Mano JF, Gomes ME, Marques AP, Azevedo HS. Natural-based polymers for biomedical applications; CRC: Boca Raton, FL, 2008.
  79. 79. Klemm D, Kramer F, Moritz S, Lindstrom T, Ankerfors M, Gray D, Dorris A. Angew. Chem. Int. Ed. Nanocelluloses: A new family of nature-based materials. 2011;50:5438-5466. DOI: 10.1002/anie.201001273
  80. 80. Fu L, Zhang Y, Zhang J, Yang G. Bacterial cellulose for skin repair materials, In:, Fazel R, editor. Biomedical engineering-frontiers and challenges Rijika: Croatia: InTech; 2011. p. 249-274. DOI: 10.5772/1019
  81. 81. Fu L, Zhang J, Yang G. Present status and applications of bacterial cellulose-based materials for skin tissue repair. Carbohydr. Polym. 2013;92,1432-1442. DOI: 10.1016/j.carbpol.2012.10.071
  82. 82. Rajwade JM, Paknikar KM, Kumbhar JV. Applications of bacterial cellulose and its composites in biomedicine. Appl. Microbiol. Biotechnol. 2015;99:2491-2511. DOI: 10.1007/s00253-015-6426-3
  83. 83. Ul-Islam M, Khan T, Park J. Water holding and release properties of bacterial cellulose obtained by in situ and ex situ modification. Carbohydr. Polym. 2012;88:596-603. DOI: 10.1016/j.carbpol.2012.01.006
  84. 84. Fontana J, De Souza A, Fontana C, Torriani I, Moreschi J, Gallotti B, de Souza S, Narcisco GP, Bichara JA, Farah LF. Acetobacter cellulose pellicle as a temporary skin substitute. Appl. Biochem. Biotechnol. 1990;24-25:253-264.
  85. 85. Kowalska-Ludwicka K, Cala J, Grobelski B, Sygut D, Jesionek-Kupnicka D, Kolodziejczyk M, Bielecki S, Pasieka Z. Modified bacterial cellulose tubes for regeneration of damaged peripheral nerves. Arch. Med. Sci. 2013;9:527-534. DOI: 10.5114/aoms.2013.33433
  86. 86. BIO FILL PRODUTOS BIOTECNOLÓGICOS (BRASIL). Farah LFX. Process for the preparation of cellulose film, cellulose film produced thereby artificial skin graft and its use. US 4912049, 30 Sept. 1990, 10 June 1986.
  87. 87. Wouk AFPF, Diniz JM, Círio SM, Santos H, Baltazar EL, Acco A. Membrana biológica Biofill – estudo comparativo com outros agentes promotores da cicatrização da pele em suínos: aspectos clínicos, histopatológicos e morfométricos. Arch. Vet. Scienc. 1998;3:31-37.
  88. 88. Czaja W, Krystynowicz A, Bielecki S, Brown Jr RM. Microbial cellulose-the natural power to heal wounds. Biomater. 2006;27:145-151. DOI: 10.1016/j.biomaterials.2005.07.035
  89. 89. Czaja W, Young DJ, Kawecki M, Brown Jr RM. The future prospects of microbial cellulose in biomedical applications. Biomacromol. 2007;8:1-12. DOI: 10.1021/bm060620d
  90. 90. Portal O, Clark WA, Levinson DJ. Microbial cellulose wound dressing in the treatment of nonhealing lower extremity ulcers. Wounds 2009;21:1-3.
  91. 91. TRIGGER PESQUISA & DESENVOLVIMENTOS BIOTECNOLOGICOS LTDA (Brasil). Y. Messaddeq; S. J. L. Ribeiro; W. Thomazini. Contact lens for therapy in cases of regeneration of cornea, has bacterial cellulose base. BR 200603704-A, 22 Ago. 2006, 08 Abr. 2008.
  92. 92. Klemm D, Schumann D, Udhardt U, Marsch S. Bacterial synthesized cellulose - artificial blood vessels for microsurgery. Progress Polym. Sci. 2001;26:1561-1603. DOI: 10.1016/S0079-6700(01)00021-1
  93. 93. Wippermann J, Schumann D, Klemm D, Kosmehl H, Satehi-Gelani S, Wahlers T. Preliminary results of small arterial substitute performed with a new cylindrical biomaterial composed of bacterial cellulose. Eur J Vasc Endovasc. 2009; 37(5): 592-96. DOI: 10.1016/j.ejvs.2009.01.007
  94. 94. Bodin A, Backdahl H, Fink H, Gustafsson L, Risberg B, Gatenholm P. Influence of cultivation conditions on mechanical and morphological properties of bacterial cellulose tubes. Biotechnol. Bioeng. 2007;97:425-434. DOI: 10.1002/bit.21314
  95. 95. Nimeskern L, Avila HM, Sundberg J, Gatenholm P, Mueller R, Stok KS. Mechanical evaluation of bacterial nanocellulose as an implant material for ear cartilage replacement. J. Mech. Behav. Biomed. Mater. 2013;22:12-21. DOI: 10.1016/j.jmbbm.2013.03.005
  96. 96. Cai ZJ, Yang G. Bacterial cellulose/collagen composite: Characterization and first evaluation of cytocompatibility. J. Appl. Polym. Sci. 2011;1205:2938-2944. DOI: 10.1002/app.33318
  97. 97. Nakayama A, Kakugo A, Gong JP, Osada Y, Takai M, Erata T, Kawano S. High mechanical strength double–network hydrogel with bacterial cellulose. Adv. Funct. Mater. 2004;14:1124-1128. DOI: 10.1002/adfm.200305197
  98. 98. Saibuatong OA, Phisalaphong M. Novo aloe vera–bacterial cellulose composite film from biosynthesis. Carbohydr. Polym. 2010;79,455-460.
  99. 99. Chiaoprakobkij N, Sanchavanakit N, Subbalekha K, Pavasant P, Phisalaphong M. Characterization and biocompatibility of bacterial cellulose/alginate composite sponges with human keratinocytes and gingival fibroblasts. Carbohydr. Polym. 2011;853:548-553. DOI: 10.1016/j.carbpol.2011.03.011
  100. 100. Wanna D, Alam C, Toivola DM, Alam P. Bacterial cellulose-kaolin nanocomposites for application as biomedical wound healing materials. Adv. Nat. Sci. Nanosci. Nanotechnol. 2013;4:045002. DOI: 10.1088/2043-6262/4/4/045002.
  101. 101. Cai Z, Kim J. Bacterial cellulose/poly (ethylene glycol) composite: Characterization and first evaluation of biocompatibility. Cellulose. 2010;17:83-91. DOI: 10.1007/s10570-009-9362-5
  102. 102. Oliveira Barud HG, Barud HS, Cavicchioli M, Amaral TS, Oliveira Junior OB, Santos DM, Oliveira Almeida Petersen AL, Celes F, Borges VM, de Oliveira CI, de Oliveira PF, Furtado RA, Tavares DC, Ribeiro SJL. Preparation and characterization of a bacterial cellulose/silk fibroin sponge scaffold for tissue regeneration. Carbohydr.Polym. 2015;128:41-51. DOI: 10.1016/j.carbpol.2015.04.007
  103. 103. Krontiras P, Gatenholm P, Hagg D. Adipogenic differentiation of stem cells in three- dimensional porous bacterial nanocellulose scaffolds. J. Biomed. Mater. Res. Part B 2015;103B:195-203. DOI: 10.1002/jbm.b.33198
  104. 104. Bäckdahl H., Helenius G., Bodin A., Nannmark U., Johansson B.R., Risberg B., et al. Mechanical properties of bacterial cellulose and interactions with smooth muscle cells. Biomaterials. 2006; 27(9): 2141-9
  105. 105. Barud HS, Barrios C, Regiane T, Marques RFC, Verelst M, Dexpert-Ghys J, Messaddeq Y, Ribeiro SJL. Self-supported silver nanoparticles containing bacterial cellulose membranes. Mater. Sci. Engineer. C. 2007;1:01-02. DOI: 10.1016/j.msec.2007.05.001
  106. 106. Barud HS, Barrios CE, Regiane T, Marques RFC, Verelst M, Dexpert-Ghys J, Messaddeq Y, Ribeiro SJL. Self-supported silver nanoparticles containing bacterial cellulose membranes. Mater. Sci. Engineer. C: Biomimetic Mater., Sensors and Systems. 2008; 28:515-518.
  107. 107. Barud HS, Regiani T, Marques RFC, Lustri WR, Messaddeq Y, Ribeiro SJL. Antimicrobial bacterial cellulose-silver nanoparticles composite membranes. J. Nanomater. 2011;721631:1-8. DOI: 10.1155/2011/721631
  108. 108. Maria LCS, Santos ALC, Oliveira PC, BARUD, Barud HS, Ribeiro SJL, Messaddeq Y. Synthesis and characterization of silver nanoparticles impregnated into bacterial cellulose. Mater. Lett. 2009;63:797-799. DOI: 10.1016/j.matlet.2009.01.007
  109. 109. Maria LCS, Santos ALC, Oliveira PC, Valle ASS, Barud HS, Messaddeq Y, Ribeiro SJL. Preparation and antibacterial activity of silver nanoparticles impregnated in bacterial cellulose. Polímeros. 2010;20:72-77. DOI: 10.1590/S0104-14282010005000001
  110. 110. Maneerung T, Tokura S, Rujiravanit R. Impregnation of silver nanoparticles into bacterial cellulose for antimicrobial wound dressing. Carbohydr. Polym. 2008;72:43-51. DOI: 10.1016/j.carbpol.2007.07. 025
  111. 111. Barud HS, de Araújo Júnior AM, Saska S, Mestieri LB, Campos JADB, De Freitas RM, Ferreira NU, Nascimento AP, Miguel FG, Vaz MMOLL, Barizon EA, Marquele-Oliveira F, Gaspar AMM, Ribeiro S JL, Berretta AA. Antimicrobial brazilian propolis (EPP-AF) containing biocellulose membranes as promising biomaterial for skin wound healing. evidence-based complementary and alternative medicine. 2013;703024:1-10. DOI: 10.1155/2013/703024
  112. 112. Stoica-Guzun A, Stroescu M, Tache F, Zaharescu T, Grosu E. Effect of electron beam irradiation on bacterial cellulose membranes used as transdermal drug delivery systems. Nucl. Instrum. Methods B 2007;265:434-438. DOI:10.1016/j.nimb.2007.09.036
  113. 113. Wei B, Yang GA, Hong F. Preparation and evaluation of a kind of bacterial cellulose dry films with antibacterial properties. Carbohydr. Polym. 2011;84:533-538. DOI: 10.1016/j.carbpol.2010.12.017
  114. 114. Trovatti E, Silva NHCS, Duarte IF, Rosado CF, Almeida IF, Costa P, Freire CSR, Silvestre AJD, Neto CP. Biocellulose membranes as supports for dermal release of lidocaine. Biomacromol. 2011;12:4162-4168. DOI: 10.1021/bm201303r
  115. 115. Müller A, Ni Z, Hessler N, Wesarg F, Müller FA, Kralisch D, Fischer D. The biopolymer bacterial nanocellulose as drug delivery system: investigation of drug loading and release using the model protein albumin. J Pharm Sci 2013, 102, 579-592. DOI: 10.1002/jps.23385
  116. 116. Saska S, Barud HS, Gaspar AMM, Marchetto R, Ribeiro SJL, Messaddeq Y. Bacterial cellulose-hydroxyapatite nanocomposites for bone regeneration. Int. J. Biomater. 2011;175362:1-8. DOI: 10.1155/2011/175362

Written By

Wilton R. Lustri, Hélida Gomes de Oliveira Barud, Hernane da Silva Barud, Maristela F. S. Peres, Junkal Gutierrez, Agnieszka Tercjak, Osmir Batista de Oliveira Junior and Sidney José Lima Ribeiro

Submitted: March 26th, 2015 Reviewed: October 19th, 2015 Published: December 9th, 2015