Extremophilic Biofilms - ACS Publications - American Chemical Society

Chapter 7

Extremophilic Biofilms: Exploring the Prospects

Downloaded via COLUMBIA UNIV on September 3, 2019 at 10:52:29 (UTC). See https://pubs.acs.org/sharingguidelines for options on how to legitimately share published articles.

Kaustubh Chandrakant Khaire,1 Seema Patel,2 Parmeshwar Vitthal Gavande,3 Vijayan and Suryakant Moholkar,1,4 and Arun Goyal1,3,* 1Center for Energy, Indian Institute of Technology Guwahati,

Guwahati 781039, Assam, India 2Bioinformatics and Medical Informatics Research Center, San Diego State University,

San Diego, California 92182, United States 3Department of Biosciences and Bioengineering, Indian Institute of Technology Guwahati,

Guwahati 781039, Assam, India 4Department of Chemical Engineering, Indian Institute of Technology Guwahati,

Guwahati 781039, Assam, India *E-mail: [email protected].

Prokaryotes are ubiquitous, colonizing even extremely harsh ecological niches. One of the major weapons that enables their survival in the hostile milieu is the self-produced matrix known as a biofilm. The biofilm performs both offensive and defensive roles. Among other functions, it acts as a protective barrier, a reserve nutrient, an osmoprotectant, and a cryoprotectant. Once encased in the biofilm, bacteria are less vulnerable to stressors. Biofilms are formed by a set of genes, expressed upon stressor exposure such as salinity, desiccation, high and low temperatures, pH changes, immune attacks, drugs, heavy metals, and radiation. The virulence role of biofilms has been known for decades. However, the considerable biotechnological potential of the biofilms is now being recognized as well. The biofilm-derived exopolysaccharides (extracellular polymeric substances) have applications in the pharmaceutical, food, and cosmetics industries as stabilizers, coagulants, gelling agents, emulsifiers, and thickeners. The exploitation of a biofilm for industrial application while suppressing its pathogenicity requires a thorough understanding of the biofilm generation mechanism. Extremophilic microbes colonizing hostile habitats also elaborate biofilms that are expected to have noble properties and biotechnological relevance. This chapter reports the valuable insights regarding biofilms from extremophiles and discusses their scopes and some hurdles in the field.

© 2019 American Chemical Society

Rathinam and Sani; Introduction to Biofilm Engineering ACS Symposium Series; American Chemical Society: Washington, DC, 2019.

1. Introduction Groups of microorganisms can stick to each other and to various objects, organism surfaces, or host lumens, forming biofilms. Therefore, the biofilms are dynamic, surface-attached living systems, housing the coordinating microbial communities where the residents are functionally organized. In fact, a biofilm is termed as a city of microbes (1). Biofilm formation has its origin in stress signal acquisition, so the biofilm-encased bacteria are slow growing and slow replicating. Sharing of common goods is the principle in this microcommunity (2). The biofilms are conglomerations of extracellular polymeric substances (EPSs), proteins, extracellular DNA, lipids, surfactants, pigments, and numerous other microbial metabolites (3). The EPS is slimy and acts as a glue, enabling surface adherence of the biofilm (4). The amount of EPS in a biofilm varies, due to the continuous generation and degradation by the resident members of the biofilm (5). Biofilms are like fortresses for bacteria, defending them against all kind of stressors such as salinity, desiccation, pH changes, immune attacks, drugs, heavy metals, and radiation (6, 7). UV radiation has been observed to inactivate nosocomial pathogens, but it is incapable of eliminating biofilms and EPS (8). This indicates the resilience of a biofilm and its importance as an armor for bacteria. In addition, protected by biofilms, bacteria render the human immune surveillance futile and cause pathogenesis (6). There are several instances where a biofilm has led to pathogenic persistence (9), chronic inflammation, implant contamination, and drug resistance (10, 11). In fact, 65% of microbial infections are associated with biofilms (12). Periodontitis (13), cystic fibrosis (14), osteomyelitis, (15), and endocarditis (16) are some of the diseases caused by biofilms. The pathogens that use the biofilm to wreak havoc in the human body include Streptococcus pneumoniae, Porphyromonas gingivalis, Treponema denticola (13), Pseudomonas aeruginosa (14), Staphylococcus aureus (15), Burkholderia cepacia, and Vibrio cholerae (17), among many others. Truly speaking, it would not be wrong to state that all bacterial pathogens use biofilm to establish pathogenesis. The cohesive strength of the biofilm renders it recalcitrant against several antimicrobial agents, which cocoons the pathogen. Apart from pathogenesis, biofilms are associated with biofouling (18). The biofilms clog the membrane systems used for water treatment, food processing, and energy production (19). Scanning electron microscopic studies have revealed that the complex biofilms are composed of multiple types of bacteria. For example, a wound site has biofilms encasing Staph. aureus and Pseudo. aeruginosa. Such mixed-species biofilms undergo compositional change over time. Multispecies biofilms create a conducive environment for resource sharing, resulting in increased biomass retention or carrying capacity (20). The residents of the biofilms forge all kinds of relations between them, depending on the requirement. The bacterium Brevibacillus is sensitive to pyocyanin produced by Pseudo. aeruginosa. In a biofilm, Brevibacillus survived the assault of pyocyanin by taking refuge under a layer of pyocyanin-resistant bacteria Raoultella (21). In contrast, the arrival of an incompatible member can disrupt the biofilm. For example, Streptomyces sp. MC025 inhibited the Staph. aureus biofilm by producing a 2,2-bipyridine: collismycin B (22). Conventionally, biofilms have been regarded as simplified entities, generated by a single or few bacteria. This notion is being dismissed as the large variation in composition and complexity of biofilms is being unraveled. It has been reported that 99% of bacteria exist in biofilms and only 1% in the planktonic state (23). These 1% of bacteria are also not obligate planktonic bacteria but are in a transient state as the conditions are favorable. The phenotypic shift from the planktonic to the biofilm mode of growth involves the switching on of a set of genes. A study of an Escherichia coli model comparing biofilmembedded and suspension cells showed the differential expression of genes related to stress response 142 Rathinam and Sani; Introduction to Biofilm Engineering ACS Symposium Series; American Chemical Society: Washington, DC, 2019.

(e.g., cspABFGI), motility (e.g., flgBCEFH, fliLMQR, and motB), fimbriae (e.g., sfmCHM, fimZ, and csgC), quorum sensing (e.g., ydgG, gadABC, and hdeABD), transport (e.g., gatABC, agaBC, ycjJ, ydf J, phoU, and phnCJKM), sulfur and tryptophan metabolism (e.g., trpLBA, tnaLA, and cysDNCJH), and the extracellular matrix (e.g., wcaBDEC) (24). Biofilm gene expression is regulated by environmental conditions, and the stress genes are switched on during biofilm formation (25). A group of surface proteins are expressed in the first step of biofilm formation. Staph. aureus biofilmassociated protein plays a role in biofilm synthesis (26). Nevertheless, despite the problems associated with biofilms, some of them are used for various beneficial purposes, such as bioleaching, wastewater remediation, and as fuel cells. (7).

2. Steps of Biofilm Formation The biofilm formation involves several stages: (1) biofilm attachment, (2) biofilm formation, and (3) dispersion of biofilms. Psuedo. aeruginosa has been a major model organism used for biofilm formation studies (27). 2.1. Biofilm Attachment Anchoring of bacteria to a substratum by adhesion paves the way for host colonization, quorum sensing, and biofilm formation (28). The surface binding stage has been studied for several microorganisms, and a common pattern has been observed. Initially, the planktonic cells attach to the available surface by physical forces, followed by specific adhesive reactions between the host and the bacterial surface agents (29). The cell appendages, such as polysaccharide chains and protein nanofibers, are the bridges between the bacteria and the substratum (30). The arsenal of adhesins is critical for adherence (31). The soluble adhesin proteins aggregate to form biofilms as observed in Porph. gingivalis (32). Clostridium difficile expresses type IV pili (T4P), which facilitates its biofilm formation (33). The attachment can be reversible or irreversible (27). Cell extremities, such as fimbriae, flagella, and pili, are associated with the reversible surface binding (34). Nonmotile strains of Pseudo. aeruginosa, lacking flagella or motility, showed poor attachment to a poly(vinyl chloride) surface (35). The surface proteins (e.g., LapA, LapD, and SadB) help the hydrophobic interaction between the bacteria and the surface (36). The biofilm formation in Acinetobacter baumannii is mediated by Csu pili, assembled by the chaperone–usher pathway (37). The CsuC–CsuE chaperone–adhesin is instrumental in the bacterial attachment to abiotic surfaces. The ATP-binding cassette transporters (ABC transporters), along with other proteins, help in the reversible to irreversible surface binding process of Pseudomonas fluorescens WCS365. The irreversible binding is associated with secondary intracellular messenger cyclic bis (3′-5′) cyclic dimeric guanosine monophosphate (c-di-GMP) (38). Likewise, another secondary messenger, cyclic adenosine monophosphate (c-AMP), also controls the biofilm synthesis (39). 2.2. Biofilm Formation Once anchored to the biotic or abiotic surface, the bacterial cells begin to form colonies. They gather together, undergo cell division, and create microcolonies. The multiplication is accompanied by EPS production. In the most developed or matured biofilm, the EPS content is more than 90% of the total dry mass. Mycobacterium smegmatis forms a pellicle on a liquid surface, with concurrent 143 Rathinam and Sani; Introduction to Biofilm Engineering ACS Symposium Series; American Chemical Society: Washington, DC, 2019.

division of cells (40). The biofilm structure is dependent on the conditions under which it is formed (e.g., nutrients, pH, and temperature). The biofilm forms are heterogeneous with regional microenvironments, as studied using the Bacillus subtilis model (41). 2.3. Dispersal of Biofilm Biofilm has a carrying capacity, or the number of bacteria it can sustain. Once the biofilm is matured, the crowding and stress buildup reduces its ability to function effectively. Some of the microbial cells break free and leave the matured biofilm. This liberation is the dispersal stage (42). This migration phenomenon is also called “swarming,” which is facilitated by the bacterial-secreted surfactants that reduce the surface tension (43). After detachment from the parental biofilm, bacteria resume the planktonic cell growth cycle. Depending on the niche and the stressors encountered, some bacteria again endeavor to attach to another surface. Therefore, the dispersal is not only the last phase of the biofilm life cycle but also the beginning of the next life cycle. The biofilm formation steps are depicted in Figure 1.

Figure 1. Steps involved in biofilm formation. 2.4. Traits of Biofilm in Different Stages In the Psuedo. aeruginosa model, the planktonic cells were compared with maturation stage biofilm cells, where more than 800 proteins were shown to have a six-fold or higher change in expression level (44). In addition, the bacterial cells in the dispersion stage biofilm were more similar to planktonic bacteria than to maturation stage bacteria (44). In dental plaque bacteria, the matured (2- to 3-week-old) biofilm resisted the disinfecting agent, while the other stages of biofilm were sensitive to the drug treatment. Microarray analysis of the transcriptome of V. cholerae sampled from the three different stages of biofilm showed variations in gene expressions (45). 2.5. Factors Regulating the Fate of Biofilm The stressors inducing biofilm can be diverse. A study found that F-actin, DNA, and Pf1 bacteriophage induce biofilm synthesis in Pseudo. aeruginosa Xen5 (46). Several factors influence each step of the biofilm formation process. The second messenger c-di-GMP acts as a switch of transformation from planktonic cells to the biofilm mode. The enzyme diguanylate cyclase produces c-di-GMP, while phosphodiesterase degrades it (47). Environmental perturbations change the 144 Rathinam and Sani; Introduction to Biofilm Engineering ACS Symposium Series; American Chemical Society: Washington, DC, 2019.

cellular c-di-GMP level. The increased level of c-di-GMP promotes biofilm synthesis, hindering motility. The c-di-GMP signals the LuxR-type regulator VpsT and the NtrC-type regulator VpsR to activate the transcription of vps and rbm genes for EPS and protein component biosynthesis of the biofilm, respectively. It commands the VpsR/VpsT-dependent histone-like nucleoid structuring (HNS) antirepression cascade (48).

3. Types of Extremophiles, Their Biofilms, and EPS The two different prokaryotic domains, bacteria and archaea, can inhabit a variety of extreme environments. Extremophiles are bacteria or archaea capable of thriving in hostile environmental conditions such as radiation, oxidation, desiccation, alkalinity, acidity, salinity, high pressure, high/ low temperature, or in the presence of heavy metal (49). To survive the challenges, extremophiles often live a community life housed in biofilms instead of in a planktonic state. Depending on their extreme condition tolerance, such extremophile bacteria are called thermophiles, hyperthermophiles, psychrophiles, halophiles, alkaliphiles, acidophiles, piezophiles, or radiophiles, as described in Table 1. The extremophiles synthesize various robust molecules like extremozymes (with traits of thermostability, cold adaptability, or salt tolerance), extremolytes (e.g., thermoprotectants, cryoprotectants, etc.), efficient ion efflux channels, pigments, and so on (50–52). They have developed compatible genetic and defense mechanisms through evolution, which allow them to adapt to such extreme conditions (53). Extremophilic biofilms have attracted immense attention for their industrial prospects. Table 1. Classification and Some Examples of Biofilms Producing Extremophiles Environmental Parameter

Type of Extremophile

Growth Conditions

Extremophile

Reference

60–80 °C >80 °C

Thermotoga maritima Geobacillus tepidamans

(58, 57)

Low temperature Pshycrophiles

9

Acidithiobacillus thiooxidans

(72)

Low pH

Acidophiles

pH < 5

Anaerobranca californiensis

(8)

High pressure

Piezophiles

400–600 atm

Shewanella benthica

(82, 101)

Radiation

Radiophlile

Radiation resistant

Deinococcus radiodurans

(87)

High temperature

Thermophiles Hyperthermophiles

3.1. Thermophiles, Their Biofilms, and EPS Microorganisms adapted to higher temperatures (growth temperature 45–75 °C) are called thermophiles, and those adapted to high temperatures (growth temperature ≥80 °C) are called hyperthermophiles (54). The thermophilic, or heat-loving, microbes may belong to both the archaea and bacteria domains (55). Marine and terrestrial hot spring ecosystems are the sources of EPSproducing thermophiles. This type of ecological system is characterized by high temperature, high pressure, and high concentration of toxic elements such as heavy metals and sulfur. In such extreme 145 Rathinam and Sani; Introduction to Biofilm Engineering ACS Symposium Series; American Chemical Society: Washington, DC, 2019.

ecosystems, EPS serves an essential role in bacterial survival by offering osmotic tolerance (56). For example, thermophilic bacilli such as Geobacillus thermodenitrificans, Geobacillus tepidamans, Bacillus thermoantarcticus, and Bacillus licheniformis are important EPS producers, which are isolated from hot marine shallow vents. A highly thermostable EPS, which can hydrolyze only at 280 °C, is produced by a bacterium G. tepidamans residing in a terrestrial hot spring (57). Thermotoga maritima is an example of anaerobic, extremely thermophilic fermentative bacteria, with the ability to produce a significant number of biofilms (58). 3.2. Psychrophiles, Their Biofilms, and EPS The psychrophilic bacteria can grow and replicate in cold environments at temperatures 15 °C or lower. Fresh water, oceans, polar soils, glaciers, and permafrost are habitats of psychrophiles. These cold-hardy microbes are adapted to endure extended periods of cryobiosis, a metabolic state of life entered into in response to freezing temperatures (59). Like microbes facing other stressors, the microbial residents of the cold niches are covered by EPS. The marine bacterium Colwellia psychrerythraea elaborates EPS to cope with the limitations of a low-temperature habitat (60). Pseudoalteromonas sp., from the Gamma Proteobacteria family and isolated from marine sediments, was reported to produce EPS as well (61). The EPS of psychrophile, Pseudoalteromonas sp. is likely to be acting as a cryoprotectant for both the microorganism and its enzymes. For example, Pseudomonas sp. ID1 from a marine sediment of Antarctica secretes EPS made of glucose, galactose, and fucose, which protects the bacterial strain from the cold stress (62). Col. psychrerythraea EPS also has cryoprotectant properties. (60). Pseudoalteromonas sp. SM9913-derived EPS can enhance the stability of the cold-adapted protease by preventing its autolysis. It can also bind to metal ions such as Fe2+, Zn2+, Cu2+, and Co2+ (63). EPS produced by psychrophile Pseudoalteromonas sp. has been reported to be highly relevant for metals and radionuclide biosorption (64). 3.3. Halophiles, Their Biofilms, and EPS Hypersaline environments such as salterns, soda lakes, and salt sediments have higher salt concentration than seawater. These hostile niches are also inhabited by bacteria. For the protection of cell membrane integrity from the saline assault and tonicity breakdown from osmotic pressure, halophilic bacteria have EPS capsules encasing them (65). EPS produced by halophiles are of great industrial importance. Halomonas is a well-studied halophilic, biofilm-producing genus. Halomonas eurihalina, a moderately halophilic member, produces an anionic EPS with industrial applications (66). EPS produced by H. eurihalina H96 contained high percentages of sulfate and uronic acid, which has potential application as a jellifying agent (67). It also has the potential to be used for water treatment and detoxification. Halomonas maura S30, a halophilic bacterium, produces an anionic EPS mauran, which holds the capacity to capture heavy metals and exerts an immunomodulator effect (68). The halophilic archaea include Halococcus, Halobacterium, Haloarcula, Haloferax, and Natronococcus (69). 3.4. Acidophiles, Their Biofilms, and EPS The microbes that can grow optimally at pH 3 or less are called acidophilic microorganisms (70). They can survive such extreme low pH by maintaining a large pH gradient to keep the cytoplasmic pH near neutral. Acidophilic chemolithotrophic microorganisms growing in solfataric fields and 146 Rathinam and Sani; Introduction to Biofilm Engineering ACS Symposium Series; American Chemical Society: Washington, DC, 2019.

sulfuric pools have various biotechnological applications, such as bioleaching and bio-oxidation for metals recovery. Pel EPS and its role in biofilm formation by sulfur-oxidizing species Acidithiobacillus thiooxidans has been studied (71). Moderately thermophilic acidophile Sulfobacillus thermosulfidooxidans uses its biofilm for pyrite leaching (72). Snottites are extremely acidic biofilms, formed in hydrogen sulfide–rich caves. A study site, Acquasanta snottites in Italy, was found populated by the sulfide oxidizers such as Acid. thiooxidans and Ferroplasma sp. (73). 3.5. Alkaliphiles, Their Biofilms, and EPS Alkaline habitats are found across the world. Microbial sulfate reduction, ammonification, and leached silicate minerals in water bodies are the drivers of the elevated pH. Some significant soda lakes include Mono Lake (USA), Owens Lake (USA), Magadi lake (Africa), Lonar soda lake (India), and Lake Doroninskoe (Russia) (74–76). Their pH levels can be 10 or higher. Several haloalkaliphilic anaerobic community members, such as Spirochaeta sp., Tindallia sp., and Desulfonatronum sp., can thrive at such high alkaline pH. Chemolithotrophic alkaliphiles have been used for bioleaching of metals such as iron, copper, gold, and uranium. Anaerobranca californiensis and Alkaliphilus metalliredigens strains were found to reduce iron at alkaline pH (77). Bacillus arsenicoselenatis, sp. nov., and Bacillus selenitireducens, sp. nov., use oxyanions of selenium and arsenic for respiration (78). Bacillus subterraneus, sp. nov., reduces iron and manganese (79). Alkaliphilic biofilms are being investigated for nanoparticles production. The biological synthesis of silver nanoparticles by the alkaliphilic actinobacterium Nocardiopsis valliformis OT1 strain has been reported (80). 3.6. Barophiles, Their Biofilms, and EPS Piezophilic or barophilic microbes are capable of growing at high hydrostatic pressure (81). Obligate piezophilic bacteria include Shewanella benthica (82), Psychromonas hadalis (83), and Moritella japonica (84) among others. Biotechnological applications of piezophiles are poorly explored. Piezophilic bacteria and archaea play a critical role in the deep ocean carbon cycle (85). The proposed applications for barophilic biofilms may include the disposal of waste in deep sea, novel bioactive compound production, and as biocatalysts for high-pressure reactors. 3.7. Radiophiles, Their Biofilms, and EPS Radiophiles, or radioisotope-tolerant microbes, possess the ability to resist or sustain ionizing radiations in a vegetative state. Some of the best-known representatives of radiophiles are Deinococcus and Thermococcus (86). The use of Deinococcus radiodurans in managing toxic metals such as mercury and halogenated organics is being investigated (87). Its prospect in radioactive-contaminated groundwater management has also been reported (88). The biofilm of Deino. radiodurans R1 played a role in the bioremediation of uranium, cobalt, and nickel (88). Deinococcus geothermalis, with its glycoconjugates, forms thin biofilms in paper machines (89). It resists dehydration, biocidal chemicals, and other extreme conditions (90). In a study by Peltola et al., it was found capable of reducing iron, uranium, and chromium at high temperatures (90). The genera Knoellia, Lysobacter, Nocardioides, Paracoccus, Pontibacter, Rufibacter, and Microvirga are other radiation-resistant bacteria. The clustering of extremophiles is largely overlapping. A thermophile can be an acidophile (e.g,. Thiobacillus caldus or Alicyclobacillus), a phychrophile can be a barophile (e.g. Alteromonas, Shewanella, Photobacterium, Colwellia, or Moritella), a halophile can be an alkaliphile (e.g., Halomonas 147 Rathinam and Sani; Introduction to Biofilm Engineering ACS Symposium Series; American Chemical Society: Washington, DC, 2019.

campisalis, Salinicoccus alkaliphilus, or Oceanobacillus oncorhynchi), and so on. Thermoacidophilic archaea include Sulfolobus and Thermococcus. Extremophiles are masters at stress adaptation. The suite of adaptations can protect an extremophile against multiple stressors. Such extremophiles adapted to multiple stresses are termed as polyextremophiles (91). Apart from the above groups of extremophiles, other microbes adapted to adverse conditions include oligotrophic (those that grow in nutritionally limited environments), endolithic (those that grow within rock or within pores of mineral grains), and xerophilic (those that grow in dry conditions, with low water availability) (91). Extremophiles produce hydrocarbons, lipids, proteins, EPSs, and polyhydroxyalkanoates, which can be used to generate bioethanol, biobutanol, biodiesel, biohydrogen, and biogas (92). The extremophiles have shown potential use in fuel cells, electrolytic cells, desalination cells, and electrosynthesis (93). Halophiles and thermophiles demonstrate interesting electrocatalytic performance in microbial electrochemical systems, which can be exploited for treating industrial waste streams (94). Thermophiles have the potential to be used as electrocatalysts for bioelectrochemical applications (95). Their biofilms are expected to be a crucial factor in many applications. In fact, electroactive biofilms play a role in electron transfer, improving the microbial electrocatalytic applications (96). Figure 2 presents a schematic diagram of various types of biofilmproducing extremophiles and their advantages and disadvantages.

Figure 2. Extremophiles and the triggers for their biofilm formation.

4. Scope and Hurdles in Commercial EPS Production by Extremophiles The EPSs of extremophiles are generally made up of monosaccharides and some noncarbohydrate substituents such as phosphate, succinate, pyruvate, and acetate. Due to wide variations in the composition of the EPSs, they have diverse biotechnological applications. They have immense use and prospects especially in the pharmaceutical and food sectors. Some industrially important EPSs have been listed in Table 2. Thermophilic bacteria such as Lactobacillus delbrueckii subsp. bulgaricus, Lactobacillus acidophilus, Lactobacillus delbrueckii subsp. Lactis, Streptococcus macedonicus, and Streptococcus thermophilus are efficient EPS producers. Different species of Leuconostoc, Pseudomonas, Alcaligenes, and Xanthomonas produce dextran, gellan, curdlan, xanthan, mauran, and so on. Dextran is produced by lactic acid 148 Rathinam and Sani; Introduction to Biofilm Engineering ACS Symposium Series; American Chemical Society: Washington, DC, 2019.

bacteria of the genera Leuconostoc, Pediococcus, and Weissella, among others (97). Gellan is synthesized by nonpathogenic bacterium Pediococcus elodea or Sphingomonas elodea (98). Its mucoadhesive and hygroscopic properties have found use in nasal, ophthalmic, oral, and other formulations (98). Curdlan is synthesized by alkali-tolerant pathogenic bacterium Alcaligenes faecalis or Agrobacterium radiobacter (99). Curdlan is used as a stabilizer, formulation aid, surface modifier, or thickener (99). The plant pathogen Xanthomonas campestris pv. campestris produces the EPS xanthan (100). It has several applications in the food and nonfood sectors (100). The halophiles H. maura produce an EPS mauran, which is used for viscosifying solutions (68). Table 2. Type of EPS Produced by Extremophiles and Their Applications EPS

Extremophile

Type of Extremophile Applications

Reference

Mauran

Halomonas maura

Halophile

Viscosifying solutions

(68)

Glucan

Geobacillus tepidamans

Thermophile

Viscosifying agent

(57)

Gellan

Sphingomonas elodea

Alkaliphile

Nasal, ophthalmic, oral, and other formulations

(98)

Xanthan Aphanothece halophytica (like) gum

Halotolerant

Food additives

(98)

Curdlan

Alkalitolerant

Stabilizer, formulation aid, surface modifier, or thickener

(99)

Agrobacterium radiobacter

4.1. Factors Affecting the Commercial Production of Extremophilic EPSs Fermentation is the technology of choice for commercial production of EPSs. The selection of the right bacterial candidate, the optimization of growth medium, and other parameters can enhance EPS production. Microbial strain, media composition, the type of nitrogen and carbon sources, trace elements, oxygenation, rotation, and so on impact the composition, structure, and thickness of the EPS. Pedio. elodea, X. campestris, and Rhizobium meliloti enhance EPS synthesis in the nitrogen-limiting conditions (101). Propionibacterium acidipropionici showed improved growth and EPS synthesis when the yeast extract content in the culture media was increased (102). The fermentation process for EPS production involves an expensive culture medium. Therefore, the cheaper alternatives ought to be explored. The dairy industry discard whey is found to be a promising substrate for EPS biosynthesis by thermophiles such as L. delbrueckii subsp. bulgaricus and Strep. thermophilus (103).

5. Future Perspectives Stress in any form induces the biofilm formation. All bacteria encounter stress, and biofilms are universally produced by them. If an analogy can be offered, biofilms are like the mucus of higher organisms (104). The broad purpose of both the mucus and the biofilm is to entangle invaders and protect one’s own cells or tissues. However, in the biofilm, the strains involved, metabolite composition, viscosity, and amount produced are highly variable. Stressors induce biofilm formation. High temperature, cold, pressure, oxygen tension, pH, salinity, shear stress, radiation, hypoxia, and immune attack are some of the factors driving biofilm formation. Biofilm production 149 Rathinam and Sani; Introduction to Biofilm Engineering ACS Symposium Series; American Chemical Society: Washington, DC, 2019.

is a necessity, but excess or persistent production is a hurdle. Biofilm is a necessary evil. Excess biofilm production is not favored by bacteria in a low-stress condition. Biofilm formation requires physiological changes, alternate gene expression, and the activation of a set of enzymes, which consumes energy and reduces metabolic activity and growth rate. The biofilm production is enhanced under life-threatening conditions. For example, Col. psychrerythraea 34H, under the temperature range –8 to –14 °C, under high pressure (400 and 600 atm), and under salinity (10%–100%) increased the EPS production dramatically (105). As mentioned, excess biofilm production is not a choice, but a necessity for the bacteria and archaea. Thus, to ensure the consistent biofilm production from bacteria for biotechnological applications, the bacteria have to be kept under stress with the appropriate challenges based on the desired biofilm to be harvested. Exploiting the extremophiles for biofilm and other novel component harvesting necessitates the creation of the exact milieu of the microbes. The biochemical adaptations that the extremophiles develop to survive may be lost if the favorable conditions prevail (106). Biofilm production by bacteria concurs with antibiotics and pigments secretion. A stress signal induces the production of these protective weapons. Deinococcus species, elaborating biofilm, have carotenoid pigments to avoid radiation (86). Extremophile-elaborated proteins and enzymes are very stable. Among other factors, the cochaperonin proteins, iron-sulfur-cluster proteins, DNA-binding proteins, and enzymes have been found to maintain their integrity in adverse conditions, which is attributed to their very slow unfolding, among other features. Extremozymes such as protease, glycosyl hydrolase, lipase, and DNA polymerase have been characterized from extremophiles isolated from diverse hostile niches. Biofilm’s barrier function and the EPS are assumed to be contributive in restoring the protein stability. Archaea are considered to be obligate extremophiles, but the mechanism of their biofilm formation is elusive. The quorum sensing and other complementary mechanisms need to be understood for the enhancement of EPS production. Although less is known about eukaryotic extremophilic biofilms as compared to their prokaryotic counterparts, certain eukaryotes have been found to reside in extremophilic habitats and produce EPSs as well. It can be exemplified by halophilic microalga Dunaliella salina, which is known to produce EPS under salt stress (107). This microalgal EPS was used to produce emulsions, and its chemical stability under halophilic conditions was comparable to the bacterial EPSs (107). Overall, the idea of utilizing extremophilic biofilm for diverse applications is interesting, as so far, the thrust has been mostly on biofilm inhibition strategies. Biofilms are dynamic and complex, so their characterizations and modifications over spatial and temporal dimensions ought to be conducted. Desirable EPSs might be obtained by culturing and coaxing cooperative microbes through the right ingredients and conditions. Innovative technologies are being used to enhance biofilm production from various extremophiles. The atmospheric pressure cold plasma treatment induced the thermophile Geobacillus sp. WSUCF1 to enhance biofilm formation (108). Omics approaches are efficient in detecting the signature of biological samples. In that regard, omics profiling can characterize biofilms and offer insights on the factors regulating their matrix compositions. The omics such as genomics, proteomics, transcriptomics, metabolomics, and metagenomics paired with bioinformatics have elucidated the community structure and shed light on the adaptive behavior of different bacterial members (109). These new-age tools are proving important for understanding complex biofilms. For example, metagenomics and metaproteomics have enabled the analyses of air–water interface biofilm samples from ships (110). Researchers concluded that the biofilms comprised 6000 taxa and were well represented by phototrophic bacteria (110). 150 Rathinam and Sani; Introduction to Biofilm Engineering ACS Symposium Series; American Chemical Society: Washington, DC, 2019.

6. Conclusions Biofilm formation is influenced by extreme environmental conditions such as high or low temperatures, pressure, and so on. Extremophilic biofilms are unique compared to their mesophilic counterparts. There is great diversity in extremophilic biofilms in their content and structure. Extremophilic biofilms are rich in novel metabolites, which can be explored for environmental protection and betterment of humankind through the use of omics, bioinformatics, and genetic engineering approaches. Genetic modification of extremophiles, optimization of fermentation, and product recovery procedures can facilitate the harnessing of biofilms. Future research on extremophilic biofilms ought to explore the adaption or construction of microfactories to sustainably empower the food sector, the pharmaceutical industry, bioremediation, and waste management.

References 1. 2.

3.

4. 5. 6. 7. 8. 9. 10. 11. 12. 13. 14. 15.

Watnick, P.; Kolter, R. Biofilm, city of microbes. J. Bacteriol. 2000, 182, 2675–2679. Lee, K. W. K.; Periasamy, S.; Mukherjee, M.; Xie, C.; Kjelleberg, S.; Rice, S. A. Biofilm development and enhanced stress resistance of a model, mixed-species community biofilm. ISME J. 2014, 8, 894–907. Jiao, Y.; Cody, G. D.; Harding, A. K.; Wilmes, P.; Schrenk, M.; Wheeler, K. E.; Thelen, M. P. Characterization of extracellular polymeric substances from acidophilic microbial biofilms. Appl. Environ. Microbiol. 2010, 76, 2916–2922. Flemming, H. C.; Wingender, J. The biofilm matrix. Nat. Rev. Microbiol. 2010, 8, 623–633. Alpkvist, E.; Picioreanu, C.; van Loosdrecht, M. C.; Heyden, A. Three‐dimensional biofilm model with individual cells and continuum EPS matrix. Biotechnol. Bioeng. 2010, 94, 961–979. Percival, S. L.; Malic, S.; Cruz, H.; Williams, D. W. In Biofilms and Veterinary Medicine; Percival, S. L., Ed.; Series on Biofilms 6; Springer-Verlag: Berlin, 2011; Vol. 14, pp 41–68. Pennafirme, S.; Lima, I.; Bitencourt, J. A.; Crapez, M. A. C.; Lopes, R. T. Microbial biofilm study by synchrotron X-ray microscopy. Phys. Chem. 2015, 116, 116–119. Pholawat Tingpej, M. D.; Tiengtip, R. Decontamination efficacy of ultraviolet radiation against biofilms of common nosocomial bacteria. J. Med. Assoc. Thai. 2015, 98, 582–588. Del Pozo, J. L. Biofilm-related disease. Expert Rev. Anti. Infect. Ther. 2018, 16, 51–65. Rabin, N.; Zheng, Y.; Opoku-Temeng, C.; Du, Y.; Bonsu, E.; Sintim, H. O. Agents that inhibit bacterial biofilm formation. Future Med. Chem. 2015, 7, 647–671. Høiby, N.; Bjarnsholt, T.; Givskov, M.; Molin, S.; Ciofu, O. Antibiotic resistance of bacterial biofilms. Int. J. Antimicrob. Agents. 2010, 35, 322–332. Abdel-Aziz, S. M.; Aeron, A. Bacterial biofilm: dispersal and inhibition strategies. Sch. J. Biotechnol. 2010, 1, 1–10. Kuramitsu, H. K.; Chen, W.; Ikegami, A. Biofilm formation by the periodontopathic bacteria Treponema denticola and Porphyromonas gingivalis. J. Periodontol. 2005, 76, 2047–2051. Macia, M. D.; Rojo-Molinero, E.; Oliver, A. Antimicrobial susceptibility testing in biofilmgrowing bacteria. Clin. Microbiol. Infect. 2014, 20, 981–990. Junka, A.; Szymczyk, P.; Ziółkowski, G.; Karuga-Kuzniewska, E.; Smutnicka, D.; Bil-Lula, I.; Bartoszewicz, M.; Mahabady, S.; Sedghizadeh, P. P. Bad to the bone: on in vitro and ex vivo microbial biofilm ability to directly destroy colonized bone surfaces without participation of host immunity or osteoclastogenesis. PLoS One 2017, 12, 1–20. 151 Rathinam and Sani; Introduction to Biofilm Engineering ACS Symposium Series; American Chemical Society: Washington, DC, 2019.

16. Chung, P. Y.; Toh, Y. S. Anti‐biofilm agents: recent breakthrough against multi‐drug resistant Staphylococcus aureus. Pathog. Dis. 2014, 70, 231–239. 17. Silva, A. J.; Benitez, J. A. Vibrio cholerae biofilms and cholera pathogenesis. PLoS Negl. Trop. Dis. 2016, 10, 1–25. 18. Vanysacker, L.; Boerjan, B.; Declerck, P.; Vankelecom, I. F. Biofouling ecology as a means to better understand membrane biofouling. Appl. Microbiol. Biotechnol. 2014, 98, 8047–8072. 19. Wood, T. L.; Guha, R.; Tang, L.; Geitner, M.; Kumar, M.; Wood, T. K. Living biofoulingresistant membranes as a model for the beneficial use of engineered biofilms. Proc. Natl. Acad. Sci. 2016, 113, E2802–E2811. 20. Brileya, K. A.; Camilleri, L. B.; Zane, G. M.; Wall, J. D.; Fields, M. W. Biofilm growth mode promotes maximum carrying capacity and community stability during product inhibition syntrophy. Front. Microbiol. 2014, 5, 693. 21. Narisawa, N.; Haruta, S.; Arai, H.; Ishii, M.; Igarashi, Y. Coexistence of antibiotic-producing and antibiotic-sensitive bacteria in biofilms is mediated by resistant bacteria. Appl. Environ. Microbiol. 2008, 74, 3887–3894. 22. Lee, J. H.; Kim, E.; Choi, H.; Lee, J. Collismycin C from the micronesian marine bacterium Streptomyces sp. MC025 inhibits Staphylococcus aureus biofilm formation. Mar. Drugs 2017, 15, 387. 23. Abdel-Aziz, S. M.; Aeron, A. Bacterial biofilm: dispersal and inhibition strategies. SAJ Biotechnol. 2014, 1, 1–10. 24. Domka, J.; Lee, J.; Bansal, T.; Wood, T. K. Temporal gene-expression in Escherichia coli K-12 biofilms. Environ. Microbiol. 2007, 9, 332–346. 25. Ren, D.; Bedzyk, L. A.; Thomas, S. M.; Ye, R. W.; Wood, T. K. Gene expression in Escherichia coli biofilms. Appl. Microbiol. Biotechnol. 2004, 64, 515–524. 26. Lasa, I.; Penadés, J. R. Bap: a family of surface proteins involved in biofilm formation. Res. Microbiol. 2006, 157, 99–107. 27. Anuj, S.; Whiley, D. M. In PCR for Clinical Microbiology: An Australian and International Perspective; Schuller, M., Sloots, T. P., James, G. S., Halliday, C. L., Carter, I. W. J., Eds.; Springer Sci. Busi. Med.: Dordrecht, Netherlands, 2010; pp 191–195. 28. Arciola, C. R.; Campoccia, D.; Ravaioli, S.; Montanaro, L. Polysaccharide intercellular adhesin in biofilm: structural and regulatory aspects. Front. Cell. Infect. Microbiol. 2015, 5, 1–10. 29. Love, R. M. Biofilm–substrate interaction: from initial adhesion to complex interactions and biofilm maturity. Endod. Top. 2010, 22, 50–57. 30. Hori, K. In Biofilm and Materials Science; Kanematsu, H., Barry, D. M., Eds.; Springer Int.: Cham, Switzerland, 2015; pp 23–34. 31. Patel, S.; Mathivanan, N.; Goyal, A. Bacterial adhesins, the pathogenic weapons to trick host defense arsenal. Biomed. Pharmacother. 2017, 93, 763–771. 32. Xie, H. Biogenesis and function of Porphyromonas gingivalis outer membrane vesicles. Future Microbiol. 2015, 10, 1517–1527. 33. Maldarelli, G. A.; Piepenbrink, K. H.; Scott, A. J.; Freiberg, J. A.; Song, Y.; Achermann, Y.; von Rosenvinge, E. C. Type IV pili promote early biofilm formation by Clostridium difficile. Pathog. Dis. 2016, 74, 1–6.

152 Rathinam and Sani; Introduction to Biofilm Engineering ACS Symposium Series; American Chemical Society: Washington, DC, 2019.

34. Di Martino, P.; Cafferini, N.; Joly, B.; Darfeuille-Michaud, A. Klebsiella pneumoniae type 3 pili facilitate adherence and biofilm formation on abiotic surfaces. Res. Microbiol. 2013, 154, 9–16. 35. O’toole, G. A.; Kolter, R. Flagellar and twitching motility are necessary for Pseudomonas aeruginosa biofilm development. Mol. Microbiol. 1998, 30, 295–304. 36. Hinsa, S. M.; O’Toole, G. A. Biofilm formation by Pseudomonas fluorescens WCS365: a role for LapD. Microbiol. 2006, 152, 1375–1383. 37. Pakharukova, N.; Tuittila, M.; Paavilainen, S.; Malmi, H.; Parilova, O.; Teneberg, S.; Zavialov, A. V. Structural basis for Acinetobacter baumannii biofilm formation. Proc. Natl. Acad. Sci. 2018, 115, 5558–5563. 38. Ishihara, Y.; Hyodo, M.; Hayakawa, Y.; Kamegaya, T.; Yamada, K.; Okamoto, A.; Ohta, M. Effect of cyclic bis (3′–5′) diguanylic acid and its analogs on bacterial biofilm formation. FEMS Microbiol. Lett. 2009, 301, 193–200. 39. Peng, X.; Zhang, Y.; Bai, G.; Zhou, X.; Wu, H. Cyclic di‐AMP mediates biofilm formation. Mol. Microbiol. 2016, 99, 945–959. 40. Sochorova, Z.; Petráčková, D.; Sitařová, B.; Buriankova, K.; Bezoušková, S.; Benada, O.; Weiser, J. Morphological and proteomic analysis of early stage air-liquid interface biofilm formation in Mycobacterium smegmatis. Microbiol. 2014, 160, 1346–1356. 41. Dervaux, J.; Magniez, J. C.; Libchaber, A. On growth and form of Bacillus subtilis biofilms. Interface Focus 2014, 4, 20130051. 42. Garrett, T. R.; Bhakoo, M.; Zhang, Z. Bacterial adhesion and biofilms on surfaces. Prog. Nat. Sci. 2008, 18, 1049–1056. 43. Connelly, M. B.; Young, G. M.; Sloma, A. Extracellular proteolytic activity plays a central role in swarming motility in Bacillus subtilis. J. Bacteriol. 2004, 186, 4159–4167. 44. Sauer, K.; Camper, A. K.; Ehrlich, G. D.; Costerton, J. W.; Davies, D. G. Pseudomonas aeruginosa displays multiple phenotypes during development as a biofilm. J. Bacterial. 2002, 184, 1140–1154. 45. Moorthy, S.; Watnick, P. I. Identification of novel stage‐specific genetic requirements through whole genome transcription profiling of Vibrio cholerae biofilm development. Mol. Microbiol. 2005, 57, 1623–1635. 46. Bucki, R.; Niemirowicz, K.; Wnorowska, U.; Wątek, M.; Byfield, F. J.; Cruz, K.; Wróblewska, M.; Janmey, P. A. Polyelectrolyte-mediated increase of biofilm mass formation. BMC Microbiol. 2015, 15, 1–10. 47. Massie, J. P.; Reynolds, E. L.; Koestler, B. J.; Cong, J. P.; Agostoni, M.; Waters, C. M. Quantification of high-specificity cyclic diguanylate signaling. Proc. Natl. Acad. Sci. 2012, 109, 12746–12751. 48. Ayala, J. C.; Wang, H.; Silva, A. J.; Benitez, J. A. Repression by H-NS of genes required for the biosynthesis of the Vibrio cholerae biofilm matrix is modulated by the second messenger cyclic diguanylic acid. Mol. Microbiol. 2015, 97, 630–645. 49. Imhoff, J. F. In Modern Topics in the Phototrophic Prokaryotes; Hallenbeck, P. C., Ed.; Environmental and Applied Aspects; Springer Int.: Cham, Switzerland, 2017; Vol. 6, pp 47–85. 50. Suriya, J.; Bharathiraja, S.; Krishnan, M.; Manivasagan, P.; Kim, S. K. In Marine Enzyme Biotechnology: Production and Industrial Application, Part-II Marine Organisms Producing


51. 52. 53. 54. 55. 56.

57.

58. 59. 60.

61.

62. 63.

64.

65. 66.

67.

Enzymes; Kim, S. K., Toldrá, F., Eds.; Advances in Food and Nutrition Research; Elsevier and Academic Press: Cambridge, MA, 2016; Vol. 79, pp 43–66. Hough, D. W.; Danson, M. J. Extremozymes. Curr. Opin. Chem. Biol. 1999, 3, 39–46. Lentzen, G.; Schwarz, T. Extremolytes: natural compounds from extremophiles for versatile applications. Appl. Microbiol. Biotechnol. 2006, 72, 623–634. Mehta, D.; Satyanarayana, T. Functional genomics of the extremophilic bacteria and archaea. Curr. Dev. Biotechnol. Bioeng. Funct. Genomics Metab. Eng. 2016, 45–78. Luke, K. A.; Higgins, C. L.; Wittung‐Stafshede, P. Thermodynamic stability and folding of proteins from hyperthermophilic organisms. FEBS J. 2007, 274, 4023–4033. Vieille, C.; Zeikus, G. J. Hyperthermophilic enzymes: sources, uses, and molecular mechanisms for thermostability. Microbiol. Mol. Biol. Rev. 2001, 65, 1–43. Chaudhari, R.; Hajoori, M.; Suthar, M.; Desai, S.; Mahavir, B. Isolation, screening and characterization of marine bacteria for exopolysaccharide production. Biosci. Discov. 2017, 8, 643–649. Kambourova, M.; Mandeva, R.; Dimova, D.; Poli, A.; Nicolaus, B.; Tommonaro, G. Production and characterization of a microbial glucan, synthesized by Geobacillus tepidamans V264 isolated from Bulgarian hot spring. Carbohydr. Polym. 2009, 77, 338–343. VanFossen, A. L.; Lewis, D. L.; Nichols, J. D.; Kelly, R. M. Polysaccharide degradation and synthesis by extremely thermophilic anaerobes. Ann. N.Y. Acad. Sci. 2008, 1125, 322–337. Moyer, C. L.; Collins, R. E.; Morita, R. Y. Psychrophiles and psychrotrophs. Ref. Modul. Life Sci. 2017, 1, 1–6. Casillo, A.; Ziaco, M.; Lindner, B.; Parrilli, E.; Schwudke, D.; Holgado, A.; Tutino, M. L. Unusual lipid A from a cold‐adapted bacterium: detailed structural characterization. ChemBioChem 2017, 18, 1845–1854. Roca, C.; Lehmann, M.; Torres, C. A.; Baptista, S.; Gaudêncio, S. P.; Freitas, F.; Reis, M. A. Exopolysaccharide production by a marine Pseudoalteromonas sp. strain isolated from Madeira Archipelago ocean sediments. N. Biotechnol. 2016, 33, 460–466. Carrión, O.; Delgado, L.; Mercade, E. New emulsifying and cryoprotective exopolysaccharide from Antarctic Pseudomonas sp. ID1. Carbohydr. Polym. 2015, 117, 1028–1034. Qin, G.; Zhu, L.; Chen, X.; Wang, P. G.; Zhang, Y. Structural characterization and ecological roles of a novel exopolysaccharide from the deep-sea psychrotolerant bacterium Pseudoalteromonas sp. SM9913. Microbiol. 2007, 153, 1566–1572. Caruso, C.; Rizzo, C.; Mangano, S.; Poli, A.; Di Donato, P.; Nicolaus, B.; Giudice, A. L. Extracellular polymeric substances with metal adsorption capacity produced by Pseudoalteromonas sp. MER144 from Antarctic seawater. Environ. Sci. Pollut. Res. 2018, 25, 4667–4677. Kunte, H. J. Osmoregulation in bacteria: compatible solute accumulation and osmosensing. Env. Chem. 2006, 3, 94–99. Llamas, I.; Sánchez, M. J.; Argandona, M.; Béjar, V.; Quesada, E.; Del Moral, A. Analysis of the genome of the moderate halophile Halomonas eurihalina. Curr. Microbiol. 2002, 45, 233–239. Béjar, V.; Llamas, I.; Calvo, C.; Quesada, E. Characterization of exopolysaccharides produced by 19 halophilic strains of the species Halomonas eurihalina. J. Biotechnol. 1998, 61, 135–141. 154 Rathinam and Sani; Introduction to Biofilm Engineering ACS Symposium Series; American Chemical Society: Washington, DC, 2019.

68. Llamas, I.; Del Moral, A.; Martínez-Checa, F.; Arco, Y.; Arias, S.; Quesada, E. Halomonas maura is a physiologically versatile bacterium of both ecological and biotechnological interest. Antonie Van Leeuwenhoek 2006, 89, 395–403. 69. Falb, M.; Müller, K.; Königsmaier, L.; Oberwinkler, T.; Horn, P.; von Gronau, S.; Gonzalez, O.; Pfeiffer, F.; Bornberg-Bauer, E.; Oesterhelt, D. Metabolism of halophilic archaea. Extremophil. 2008, 12, 177–196. 70. Baker-Austin, C.; Dopson, M. Life in acid: pH homeostasis in acidophiles. Trend. Microbiol. 2007, 15, 165–171. 71. Díaz, M.; Castro, M.; Copaja, S.; Guiliani, N. Biofilm formation by the acidophile bacterium Acidithiobacillus thiooxidans involves c-di-GMP pathway and pel exopolysaccharide. Gen. 2018, 9, 1–15. 72. Li, Q.; Sand, W.; Zhang, R. Enhancement of biofilm formation on pyrite by Sulfobacillus thermosulfidooxidans. Min. 2016, 6, 71. 73. Gorlenko, J. D. S.; Schaperdoth, I.; Macalady, J. L. Metagenomic evidence for sulfide oxidation in extremely acidic cave biofilms. Geomicrobiol. J. 2014, 3, 94–204. 74. Pikuta, E. V.; Detkova, E. N.; Bej, A. K.; Marsic, D.; Hoover, R. B. Anaerobic halo-alkaliphilic bacterial community of athalassic, hypersaline Mono Lake and Owens Lake in California. Proc. SPIE-The Int. Soc. Opt. Eng. 2003, 4859, 130–145. 75. Deshmukh, K. B.; Pathak, A. P.; Karuppayil, M. S. Bacterial diversity of Lonar soda lake of India. Ind. J. Microbiol. 2011, 51, 107–111. 76. Gorlenko, V. M.; Buryukhaev, S. P.; Matyugina, E. B.; Borzenko, S. V.; Namsaraev, Z. B.; Bryantseva, I. A.; Boldareva, E. N.; Sorokin, D. Y.; Namsaraev, B. B. Microbial communities of the stratified soda Lake Doroninskoe (Transbaikal region). Microbiol. 2010, 79, 390–401. 77. Gorlenko, V.; Tsapin, A.; Namsaraev, Z.; Teal, T.; Tourova, T.; Engler, D.; Mielke, R.; Nealson, K. Anaerobranca californiensis sp. nov., an anaerobic, alkalithermophilic, fermentative bacterium isolated from a hot spring on Mono Lake. I. J. Syst. & Evol. Microbial. 2004, 54, 739–743. 78. Blum, J. S.; Oremland, R. S.; Borglin, S. E. Reduction of elemental selenium to selenide: experiments with anoxic sediments and bacteria that respire se-oxyanions. Geomicrobiol. J. 2003, 20, 587–602. 79. Kanso, S.; Greene, A. C.; Patel, B. K. Bacillus subterraneus sp. nov., an iron-and manganesereducing bacterium from a deep subsurface Australian thermal aquifer. I. J. Syst. & Evol. Microbial. 2002, 52, 869–874. 80. Rathod, D.; Golinska, P.; Wypij, M.; Dahm, H.; Rai, M. A new report of Nocardiopsis valliformis strain OT1 from alkaline Lonar crater of India and its use in synthesis of silver nanoparticles with special reference to evaluation of antibacterial activity and cytotoxicity. Med. Microbiol. Immunol. 2016, 205, 435–447. 81. Kato, C.; Bartlett, D. H. The molecular biology of barophilic bacteria. Extremophil. 1997, 1, 111–116. 82. Lauro, F. M.; Chastain, R. A.; Ferriera, S.; Johnson, J.; Yayanos, A. A.; Bartlett, D. H. Draft genome sequence of the deep-sea bacterium Shewanella benthica strain KT99. Genome Announc. 2013, 1, 10–12.


83. Nogi, Y.; Hosoya, S.; Kato, C.; Horikoshi, K. Psychromonas hadalis sp. nov., a novel piezophilic bacterium isolated from the bottom of the Japan Trench. Int. J. Syst. Evol. Microbiol. 2007, 57, 1360–1364. 84. Fang, J.; Li, C.; Zhang, L.; Davis, T.; Kato, C.; Bartlett, D. H. Hydrogen isotope fractionation in lipid biosynthesis by the piezophilic bacterium Moritella japonica DSK1. Chem. Geol. 2014, 367, 34–38. 85. Fang, J.; Zhang, L.; Li, J.; Kato, C.; Tamburini, C.; Zhang, Y.; Dang, H.; Wang, G.; Wang, F. The POM-DOM piezophilic microorganism continuum (PDPMC)—The role of piezophilic microorganisms in the global ocean carbon cycle. Sci. China Earth Sci. 2015, 58, 106–115. 86. Tian, B.; Hua, Y. Carotenoid biosynthesis in extremophilic Deinococcus thermus bacteria. Trend. Microbiol. 2010, 18, 512–520. 87. Brim, H.; McFarlan, S. C.; Fredrickson, J. K.; Minton, K. W.; Zhai, M.; Wackett, L. P.; Daly, M. J. Engineering Deinococcus radiodurans for metal remediation in radioactive mixed waste environments. Nat. Biotechnol. 2000, 18, 85–90. 88. Appukuttan, D.; Rao, A. S.; Apte, S. K. Engineering of Deinococcus radiodurans R1 for bioprecipitation of uranium from dilute nuclear waste. Appl. & Env. Microbiol. 2006, 72, 7873–7878. 89. Daly, M. J. Engineering radiation-resistant bacteria for environmental biotechnology. Curr. Opin. Biotechnol. 2000, 11, 280–285. 90. Peltola, M.; Neu, T. R.; Raulio, M.; Kolari, M.; Salkinoja‐Salonen, M. S. Architecture of Deinococcus geothermalis biofilms on glass and steel: a lectin study. Environ. Microbiol. 2008, 10, 1752–1759. 91. Gupta, G. N.; Srivastava, S.; Khare, S. K.; Prakash, V. Extremophiles: an overview of microorganism from extreme environment. Int. J. Agric. Environ. Biotechnol. 2014, 7, 371–372. 92. Sani, R. K.; Rathinam, N. K. In Extremophilic Enzymatic Processing of Lignocellulosic Feedstocks to Bioenergy; Sani, R. K., Rathinam, N. K., Eds.; Springer Int.: Cham, Switzerland, 2017; pp 1–30. 93. Sani, R. K.; Rathinam, N. K. In Extremophilic Microbial Processing of Lignocellulosic Feedstocks to Biofuels, Value-Added Products, and Usable Power; Sani, R. K., Rathinam, N. K, Eds..; Springer Int.: Cham, Switzerland, 2018; pp 229–245. 94. Shrestha, N.; Chilkoor, G.; Vemuri, B.; Rathinam, N.; Sani, R. K.; Gadhamshetty, V. Extremophiles for microbial-electrochemistry applications: a critical review. Bioresour. Technol. 2018, 255, 318–330. 95. Rathinam, N. K.; Bibra, M.; Salem, D. R.; Sani, R. K. Thermophiles for biohydrogen production in microbial electrolytic cells. Bioresour. Technol. 2019, 277, 171–178. 96. Angelaalincy, M. J.; Navanietha, K. R.; Shakambari, G.; Ashokkumar, B.; Kathiresan, S.; Varalakshmi, P. Biofilm engineering approaches for improving the performance of microbial fuel cells and bioelectrochemical systems. Front. Ener. Res. 2018, 6, 1–12. 97. Kothari, D.; Das, D.; Patel, S.; Goyal, A. Dextran and food application. In Polysaccharides; Springer Int.: Cham, Switzerland, 2015; pp 1–16. 98. Sworn, G. In Handbook of Hydrocolloids, 2nd ed.; Phillips, G. O., Williams, P. A., Eds.; Woodhead: Cambridge, UK, 2009; pp 186–227.


99. Divyasri, D.; Gunasekar, V.; Benny, I. S.; Ponnusami, V. A review on industrial applications of curdlan. Int. J. Chem. Tech. Res. 2014, 6, 3000–3003. 100. Tavernier, P.; Portais, J. C.; Nava Saucedo, J. E. Exopolysaccharide and poly-βhydroxybutyrate coproduction in two Rhizobium meliloti strains. Appl. Environ. Microbiol. 2016, 63, 21–26. 101. Gorret, N.; Maubois, J. L.; Ghoul, M.; Engasser, J. M. Exopolysaccharide production by Propionibacterium acidi-propionici on milk microfiltrate. J. Appl. Microbiol. 2001, 90, 779–787. 102. Briczinski, E. P.; Roberts, R. F. Production of an exopolysaccharide-containing whey protein concentrate by fermentation of whey. J. Dairy Sci. 2002, 85, 3189–3197. 103. Asgarani, E.; Ghashghaei, T.; Soudi, M. R.; Alimadadi, N. Enterobacterial repetitive intergenic consensus (ERIC) PCR based genetic diversity of Xanthomonas sp. and its relation to xanthan production. Iran. J. Microbiol. 2015, 7, 38–44. 104. Cone, R. A. Barrier properties of mucus. Adv. Drug. Deliv. Rev. 2009, 61, 75–85. 105. Marx, J. G.; Carpenter, S. D.; Deming, J. W. Production of cryoprotectant extracellular polysaccharide substances (EPS) by the marine psychrophilic bacterium Colwellia psychrerythraea strain 34H under extreme conditions. Can. J. Microbiol. 2009, 55, 63–72. 106. Hande, K.; Öner, E. T.; Robert, F. H. D. In Handbook of Carbohydrate Polymers: Development, Properties and Applications; Ryouichi, I., Youto, M., Eds.; Nova Sci.: Colchester, UK, 2010; pp 605–619. 107. Mishra, A.; Jha, B. Isolation and characterization of extracellular polymeric substances from micro-algae Dunaliella salina under salt stress. Bioresour. Technol. 2009, 100, 3382–3386. 108. Rathinam, N. K.; Gorky, B. M.; Rajan, M.; Salem, D.; Sani, R. K. Short term atmospheric pressure cold plasma treatment: a novel strategy for enhancing the substrate utilization in a thermophile, Geobacillus sp. strain WSUCF1. Bioresour. Technol. 2019, 278, 477–480. 109. Jerez, C. A. The use of genomics, proteomics and other OMICS technologies for the global understanding of biomining microorganisms. Hydrometall. 2008, 94, 162–169. 110. Leary, D. H.; Li, R. W.; Hamdan, L. J.; Hervey, W. J.; Lebedev, N.; Wang, Z.; Deschamps, J. R.; Kusterbeck, A. W.; Vora, G. J. Integrated metagenomic and metaproteomic analyses of marine biofilm communities. Biofoul. 2014, 30, 1211–1223.


Extremophilic Biofilms - ACS Publications - American Chemical Society

Recommend Documents