WO2007044439A2

WO2007044439A2 - Microbial exopolymers useful for water demineralization

Info

Publication number: WO2007044439A2
Application number: PCT/US2006/038906
Authority: WO
Inventors: Thomas D. Perry
Original assignee: Acillix Incorporated
Priority date: 2005-10-05
Filing date: 2006-10-05
Publication date: 2007-04-19

Abstract

The invention provides methods to control scale formation that utilize microbes and exopolymers and other chemical compounds that they produce. The microbes produce polymers and similarly that control scale formation. The polymers work via a variety of mechanisms depending on their chemical structures. The polymers work by sequestering scale-forming ions, promoting dissolution of existing scale, increasing the solubility of newly formed scale, and inhibiting scale formation on pipes.

Description

MICROBIAL EXOPOLYMERS USEFUL FOR WATER DEMINERALIZATION FIELD OF THE INVENTION

This invention relates to the fields of microbiology, biochemistry and water chemistry. More specifically, this invention relates to the control of minerals in water and mineral scale formation on surfaces in contact with mineral-laden water by providing selected microbes and associated microbial products and methods with which water hardness and scale formation can be controlled, prevented and /or reversed.

1. BACKGROUND OF THE INVENTION

Most fresh water sources provide relatively "hard" water that is high in dissolved minerals such as calcium and magnesium. Heated hard water can form a deposit of calcium and magnesium minerals called "scale". Such scale deposits can contribute to the inefficient operation or failure of water-using appliances and can clog pipes and reduce the efficiency of water flow. Sustained scale formation ultimately requires replacement of the affected pipe or appliance.

The formation of scale in residential and industrial water systems results in reduced heat transfer efficiency, increased pollution and other effects that impose an immense economic burden on industry and on society in general. Many heavy industry applications, and most climate control systems, depend on heat exchanger technology, which suffer particularly elevated energy burdens associated with scale formation. Accordingly, improved scale control abilities would dramatically reduce environmental impacts and fossil fuel energy costs. For example, it is approximated that one millimeter of scale adds 7.5% to energy costs while seven millimeters can increase cost by more than 70% (www.nist.gov). Indeed, the energy-transfer inefficiency imposed by scale results in astronomical amounts of wasted energy and pollution to produce the extra energy required, and ultimately causes a huge economic burden globally. It is estimated that in 2002, the U.S. alone spent $2.9 billion on additional energy generation, $2.5 billion on prevention, treatment and maintenance, and lost $1.4 billion to productivity reduction. Supplying the additional energy resulting from scale-caused energy inefficiencies resulted in the additional production of 5.8 billion tons of green house gases in the U.S., an amount greater than the total green house gas production of the United Kingdom. Additionally, pumps have to pump harder to overcome obstructed flow, materials must be specially engineered and produced, and equipment must be duplicated if production is to remain uninterrupted during maintenance.

Currently accepted scale-inhibition and prevention technologies are limited to chemical and electrochemical water treatments. Chemical water treatments are expensive because of environmental regulations limiting phosphate contamination of wastewater, and recurring chemical costs. Considering this, the market is searching for alternative methods to address this growing issue. For example, ion exchange and electrodialysis have been used to remove calcium and other hard water ions associated with scale formation.

Despite these efforts, there exists a need for technologies capable of controlling scale formation and mineral deposits that are less polluting, less hazardous and/or more cost effective than current technologies.

2. SUMMARY OF THE INVENTION

The invention is based, in part, upon the finding that exopolymers produced by certain bacteria and other microbes can be used to control the mineral hardness in water and formation of scale from mineral-containing water, such as water that is high in calcium and magnesium content and considered "hard".

Accordingly, in one aspect, the invention provides a method of controlling water hardness and scale formation in a water system by contacting the water system with one or more exopolymer-producing microbes under conditions that allow for the production of microbial exopolymer, the microbes producing exopolymer, thereby controlling water hardness and scale formation in the water system.

In another aspect, the invention provides a method of controlling water hardness and scale formation. In this method, one or more exopolymer-producing microbes contacts the water system under conditions that allow for the production of bacterial exopolymer by the bacteria, thereby controlling the water hardness and scale formation in the water system. In one embodiment, the microbe is an exopolymer-producing bacterium. In another embodiment, the microbe is a proteobacterium, such as purple bacteria, chemoautotrophic proteobacteria, or chemoheterotrophic proteobacteria. In other embodiments, the microbe is a fungus. In still other embodiments, the microbe is a lichen. In still other embodiments, the microbe is an alga (e.g., a planktonic alga or a surface biofilm-forming alga). In still other embodiments, there exists a mixture of different types of microorganisms.

In certain embodiments, the exopolymer producing microbe is an alga such as, Volvox aureus, Volvox carteri, Volvox globactor, Volvox dissipatrix, Volvox tertios, Compsopogon coeruleus, Cladophora crispata, Spirogyra rivularis, Enteromorpha micrococca, Eunotia pectinalis, Melosira varian, Stigeoclonium tenue, an Amphora or a Cocconeis species. In still further embodimnets, the EPS producing microbe is an algal surface biofilm-forming alga, such as a Cyanophycota (Cyanobacteria or Blue Green Algae), including Oscillatoria, Lyngbya, Schizothrix, Chroococcus Calothrix; a Chlorophycota (Green Algae) including Ulothrix, Enteromorpha, Spirogyra, Cladophora, Dichotomosiphon, Stigeoclonium, Oedogonium, Mougeotia, Gloeocystis; a Chrornophycota (primarily Diatoms), including Melosira, Ctenophora,

Asterionella, Eunotia, Amphipleura, Cocconeis, Placoneis, Rhoikoneis, Bacillaria, and others; and Rhodophycota (Red Algae) including Compsopogon. In still other embodiments the genera Polysiphonia, Herposiphonia, and Callithamnion are used.

In certain other embodiments, the exopolymer-producing microbe is a salt tolerant organism. In particular embodiments, the salt tolerant organism grows in hard or very hard water. In other embodiments, the very hard water comprises a calcium carbonate concentration of greater than about 180 ppm.

In further embodiments, a Halobacterium, Oscillatoria or Aphanocapsa species is used to produce exopolymer and control water hardness and scale formation. Exemplary Halobacterium species include Halobacterium cutirubrurn Halobacterium denitrificans,Halobacterium distributum, Halobacterium halobium, Halobacterium lacusprofundi, Halobacterium mediterranei, Halobacterium noricense, Halobacterium pharaonis, Halobacterium saccharovorum, Halobacterium salinarium, Halobacterium sodomense, Halobacterium trapanicum, Halobacterium vallismortis and Halobacterium volcanii. Exemplary Oscillatoria species include Oscillatoria simplicissim. Exemplary Aphanocapsa species include Aphanocapsa elachista, Aphanocapsa delicatissima, Aphanocapsa endophytica, Aphanocapsa grevillei, Aphanocapsa pulchra, and Aphanocapsa rivularis.

In yet another aspect, the invention provides a method of controlling water hardness and scale formation in a water system. In this method, one or more exopolymer-producing bacteria contacts the water system under conditions that allow for the production of bacterial exopolymer by the bacteria, thereby controlling the scale formation in the water system.

In certain .embodiments, the exopolymer-producing bacteria is a Proteus species {e.g., Proteus mirabilis). In a particular embodiment, the bacteria is the Proteus mirahilis strain deposited as ATCC #51286. In further embodiments, the exopolymer-producing bacteria is a Bacillus, such as Bacillus cerueus and Bacillus thuringiensis. Ih other embodiments, the Bacillus is G3 or MEX244.1. In yet other embodiments, the Bacillus used in the method of the invention is Bacillus gibsonii, Bacillus pseudalcaliphilus, Bacillus pseudofirmus, Bacillus halodurans, Bacillus subtilis, Bacillus benzoevorans, Bacillus simplex, Bacillus horikoshii, Bacillus cereus, or Bacillus thuringiensis. In further embodiments, the exopolymer-producing bacteria is a Pseudomonas, e.g. Pseudomonas putida or Pseudomonas aeruginosa. In still further embodiments, the exopolymer-producing bacteria is an Azotobacter, e.g., Azotobacter vinelandii, Azotobacter chroococcum ox Azotobacter indicus.

Li particular embodiments of the method of the invention, the exopolymer produced controls water hardness and scale formation by sequestration of ions. These ions include earth metals and minerals, such as sodium, calcium, and magnesium, that are normally occurring in earth surface waters and metals, such as arsenic, from anthropogenic sources. In certain embodiments, the ions include heavy metal ions. In some embodiments, the cation sequestration results in fiocculation and/or precipitation of the exopolymer-mineral complex. Ih other embodiments, the precipitation may lead to coprecipitation of associated anions in from the water column, such as chloride, hydroxide, carbonate, bicarbonate, sulfate, and/or nitrate, as well as heavy metals, such as chromium (VI), arsenic (V) and selenium (VI). Li other embodiments, the exopolymer produced controls scale formation by promoting dissolution of existing scale. Li still other embodiments, the exopolymer produced controls scale formation by increasing the solubility of newly formed scale. Li further embodiments, the exopolymer produced controls scale formation by inhibiting scale formation on a surface of a water system. Li still further embodiments, the exopolymer-producing bacteria further control scale formation by also producing simple acids that dissolve the scale.

Li certain embodiments of the method of the invention, the water system includes a water pipe, a heat-transfer system, and/or a boiler. Li particular embodiments, the water system is a heat exchanger, such as a single-pass heat exchanger, a multi-pass heat exchanger, a regenerative heat exchanger, a non-regenerative heat exchanger, a tube heat exchanger, a shell heat exchanger, a plate heat exchanger, a parallel-flow heat exchanger, a cross-flow heat exchanger or a counter-flow heat exchanger. Li further embodiments, the water system of the method of the invention is a waste-water treatment facility, a power-generation facility, a pulp and paper processing plant, a petrochemical refinery, or a metal refinery.

Li further embodiments of the method of the invention, the water system includes one or more chambers in series or parallel, and each of these one or more chambers contains exopolymer-producing bacterial populations. Li particular embodiments, the microbes or bacteria are retained within the chamber(s) by a high- volume filter. Li other embodiments, the bacterial are retained within the one or more chambers by a tangential filter. Li further embodiments, one or more of the high-volume filters retain the bacterial populations but allow bacterial exopolymers and simple acids to pass through the chamber. In other embodiments, one or more of the chambers may be bounded by high- volume filters that retain the bacterial exopolymer. In other embodiments, the water system further includes an outlet system for the chamber for removing calcium-saturated bacterial exopolymer (e.g., as accumulated in the chamber and/or on a high- volume filter that retains the bacterial exopolymer).

In yet other embodiments of the method of the invention, the water system further provides for a means of supplying the one or more exopolymer-producing bacteria with one or more nutrients. In one embodiment, the nutrients supplied include an organic carbon source and a nitrogen source. In other embodiments, the nutrients include a carbon source, a nitrogen source, a phosphorous source and/or micronutrients. Li another embodiment, the apparatus is transparent to sunlight to provide energy for growth of photosynthetic organisms, such as algae or cyanobacteria, contained within the apparatus.

In another aspect, the invention provides a method of controlling water hardness and scale formation in a water system. In this method, an isolated or purified microbial exopolymer is provided to the system in such a manner that the water is in contact with the microbial exopolymer, which controls water hardness and scale-formation in the water system. In one embodiment, the microbial exopolymer is provided in a form that is at least about 50% pure (i.e., free of contaminating substances on a w/w basis). In other embodiments the microbial exopolymer is provided in a form that is at least 75% pure. In still other embodiments the microbial exopolymer is provided in a form that is at least 90% pure.

In still another important aspect, the invention provides an isolated, cation-chelating algal exopolymer having a molecular weight of greater than about 20,000 Daltons. In some embodiments, the isolated, cation-chelating algal exopolymer has a molecular weight that is greater than about 40,000 Daltons. In further embodiments, the isolated, cation-chelating algal exopolymer has a molecular weight that is is greater than about 60,000 Daltons. In particular embodiments, the isolated, cation-chelating algal exopolymer has a molecular weight greater than about 100,000 Daltons.

In another aspect, the invention provides an isolated, cation-chelating bacterial exopolymer having a molecular weight of greater than about 167,000 Daltons. In general, the isolated, cation-chelating bacterial exopolymer compositions of the invention have few or no carboxylate-containing glycosyl residues and effect cation chelation by conformationally and configurationally positioned arrays of electron pair donating groups. In certain embodiments, the isolated, cation-chelating, bacterial exopolymer is at least 50% pure {i.e., 50% free of contaminating substances on a w/w basis). In particular embodiments, the isolated, cation- chelating, bacterial exopolymer is at least 75%, at least 90%, at least 95%, or at least 99% pure.

In further embodiments, the isolated, cation-chelating bacterial exopolymer composition of the invention has a mole percent glycosyl composition of 3% xylose, about 5% arabinose, about 10% galactose, and about 30% mannose. Li still further embodiments, the isolated, cation- chelating bacterial exopolymer compositions have a calcium-binding capacity of about one cation per 8 glycosyl residues.

In a further aspect, the invention provides an isolated, cation-chelating, microbial exopolymer that is produced by a process which comprises providing one or more exopolymer- producing microbes with nutrients sufficient to cause the microbes to produce the cation- chelating microbial exopolymer, and isolating the exopolymer so produced. In certain embodiments, the isolated, cation-chelating, microbial exopolymer is produced by a microbe that is an exopolymer-producing bacteria. In other embodiments, the isolated, cation-chelating, microbial exopolymer of the invention is produced by a microbe that is a proteobacteria, such as purple bacteria, chemoautotrophic proteobacteria, or chemoheterotrophic proteobacteria. In certain embodiments, the exopolymer-producing microbe is algae, such as Volvox aureus, Volvox carteri, Volvox globactor, Volvox dissipatrix, Volvox tertios, Compsopogon coeruleus, Cladophora crispata, Spirogyra rivularis, Enteromorpha micrococca, Eunotia pectinalis, Melosira varian, Stigeoclonium tenue, Amphora or a Cocconeis species. In still further embodiments, the EPS producing microbe is an algal surface biofilm-forming alga, such as a Cyanophycota (Cyanobacteria or Blue Green Algae), including Oscillatoria, Lyngbya, Schizothrix, Chroococcus Calothrix; a Chlorophycota (Green Algae), including Ulothrix, Enteromorpha, Spirogyra, Cladophora, Dichotomosiphon, Stigeoclonium, Oedogoniurn, Mougeotia, Gloeocystis; a Chromophycota (primarily Diatoms), including Melosira, Ctenophora, Asterionella, Eunotia, Amphipleura, Cocconeis, Placoneis, Rhoikoneis, Bacillaria, and others; and Rhodophycota (Red Algae), including Compsopogon. In still other embodiments the genera Polysiphonia, Herposiphonia, and Callithamnion are used. Li still other embodiments, the isolated cation-chelating microbial exopolymer of the invention is produced by a microbe that is a fungus or a lichen. In still other embodiments, the isolated, cation-chelating, microbial exopolymer of the invention is produced by a microbe that is an alga.

In still further embodiments, the EPS producing microbe is a Halobacterium, Oscillatoria or Aphanocapsa. Exemplary Halobacterium species include Halobacterium cutirubrum Halobacterium denitrificans,Halobacterium distributum, Halobacterium halobium, Halobacterium lacusprofundi, Halobacterium mediterranei, Halobacterium noricense, Halobacterium pharaonis, Halobacterium saccharovorum, Halobacterium salinarium, Halobacterium sodomense, Halobacterium trapanicum, Halobacterium vallismortis and Halobacterium volcanii. Exemplary Oscillatoria species include Oscillatoria simplicissim. Exemplary Aphanocapsa species include Aphanocapsa elachista, Aphanocapsa delicatissima, Aphanocapsa endophytica, Aphanocapsa grevillei, Aphanocapsa pulchra, and Aphanocapsa rivularis.

In another aspect, the invention provides an isolated, cation-chelating, bacterial exopolymer that is produced by a process comprising providing one or more exopolymer- producing bacteria with nutrients sufficient to cause the bacteria to produce the cation-chelating bacterial exopolymer, and then isolating the exopolymer so produced. In certain embodiments, the exopolymer compositions of the invention are produced by the exopolymer-producing bacteria Proteus mirabilis. In a particular embodiment, this bacterium is the Proteus mirabilis strain deposited as ATCC #51286. hi other embodiments, the isolated, cation-chelating, bacterial exopolymer compositions of the invention are produced by bacteria of the genus Bacillus. In particular embodiments, the Bacillus species for use in producing the isolated bacterial exopolymers of the invention include Bacillus cerueus and Bacillus thuringiensis. In. other embodiments, the isolated cation-chelating bacterial exopolymers are produced by the Bacillus isolates G3 or MEX244.1. hi other embodiments, the isolated cation-chelating bacterial exopolymers are produced by a Bacillus that is Bacillus gibsonii, Bacillus pseudalcaliphilus, Bacillus pseudofirmus, Bacillus halodurans, Bacillus subtilis, Bacillus benzoevorans, Bacillus simplex, Bacillus horikoshii, Bacillus cereus, or Bacillus thuringiensis. In further embodiments, the isolated, cation-chelating, bacterial exopolymers are produced by bacteria of the genus Pseudomonas, e.g., Pseudomonas putida or Pseudomonas aeruginosa. In still further embodiments, the isolated, cation-chelating, bacterial exopolymers are produced by bacteria of the genus Azotobacter, e.g., Azotobacter vinelandii, Azotobacter chroococcum or Azotobacter indicus.

In further embodiments, the exopolymers of the invention have a molecular weight of greater than about 167,000 Daltons. Ih particular embodiments, the exopolymers have a purity of at least 50% {i.e., 50% free of contaminating substances on a w/w basis). In other embodiments, the exopolymers produced are about 75%, about 90%, about 95% or about 99% pure. These isolated,

exopolymer compositions of the invention have few or no carboxylate-containing glycosyl residues and effect cation chelation by conformationally and configurationally positioned arrays of electron pair donating groups. In further embodiments, the exopolymer compositions produced have a mole percent glycosyl composition of at least about 3% xylose, at least about 5% arabinose, at least about 10% galactose, and at least about 30% mannose. Li still further embodiments, the exopolymer compositions produced have a calcium-binding capacity of about one cation per 8 glycosyl residues.

In another aspect, the invention provides methods of producing an isolated, cation- chelating, microbial exopolymer comprising providing one or more exopolymer-producing microbes with nutrients sufficient to cause the microbes to produce the cation-chelating, microbial exopolymer, and then isolating the exopolymer so produced. In certain embodiments, the microbe is an exopolymer-producing bacteria. Li other embodiments, the microbe is a proteobacterium. Li certain embodiments, the proteobacteria is purple bacteria, chemoautotrophic proteobacteria, or chemoheterotrophic proteobacteria. Li other embodiments of this method of the invention, the microbe is a fungus. Li still other embodiments, the microbe is a lichen. Li still other embodiments, the microbe is an alga.

Li yet another aspect, the invention provides a method of producing an isolated, cation- chelating, bacterial exopolymer comprising providing one or more exopolymer-producing bacteria with nutrients sufficient to cause the bacteria to produce the cation-chelating, bacterial exopolymer, and then isolating the exopolymer so produced. Li certain embodiments, the exopolymer-producing bacteria provided is Proteus mirabilis. Li a particular embodiment, the bacteria is the strain deposited as ATCC #51286. Li other embodiments, the exopolymer- producing bacteria provided is a Bacillus. Li particular embodiments, the Bacillus provided is Bacillus cerueus and/or Bacillus thuringiensis. In still other embodiments, the Bacillus provided is G3 and/or MEX244.1. In other embodiments, the Bacillus provided is Bacillus gibsonii, Bacillus pseudalcaliphilus, Bacillus pseudofirmus, Bacillus halodurans, Bacillus subtilis, Bacillus benzoevorans, Bacillus simplex, Bacillus horikoshii, Bacillus cereus, or Bacillus thuringiensis. In further embodiments, the exopolymer-producing bacteria provided is a Pseudomonas, e.g., Pseudomonas putida or Pseudomonas aeruginosa. In still further embodiments, the exopolymer-producing bacteria provided is m\ Azotobacter, e.g., Azotobacter vinelandii, Azotobacter chroococcum or Azotobacter indicus.

In an additional aspect, the invention provides an apparatus for processing water to control water hardness and scale formation. The apparatus includes at least one chamber having at least one input opening for receiving the water from a water source, and at least one output opening for discharging treated water from the chamber; a source of cation-chelating, exopolymer-producing microbe disposed within the chamber such that at least a portion of the water passing through the chamber is in fluid communication with the microbe; and at least one filter for filtering the treated water. In some embodiments, the chamber of the apparatus retains the microbe, but not the cation-chelated microbial exopolymer. In certain embodiments, the chamber of the apparatus retains both the microbe and the cation-chelated microbial exopolymer. hi particular embodiments, the chamber of the apparatus retains neither the microbe nor the cation-chelated microbial exopolymer.

In further embodiments, the microbe disposed within the chamber is exopolymer- producing bacteria, hi certain embodiments, the disposed microbe is proteobacteria. hi particular embodiments, the bacteria is purple bacteria, chemoautotrophic proteobacteria, or chemoheterotrophic proteobacteria. hi yet other embodiments, the microbe disposed within the chamber of the apparatus is a fungus, hi further embodiments, the microbe disposed within the chamber of the apparatus is a lichen, hi further embodiments, the microbe disposed within the chamber of the apparatus is an alga.

hi a further aspect, the invention provides an apparatus for processing water to control scale formation. The apparatus comprises at least one chamber having at least one input opening for receiving the water from a water source, and at least one output opening for discharging treated water from the chamber; a source of cation-chelating, exopolymer-producing bacteria disposed within the chamber such that at least a portion of the water passing through the chamber is in fluid communication with the bacteria; and at least one filter for filtering the treated water. In certain embodiments, the filter of the apparatus has a pore size of not more than about 0.2 μm. In particular embodiments, the filter of the apparatus is a 0.2 μm membrane filter, a 5 kD membrane filter, or a combination thereof.

In particular embodiments, the bacteria disposed within the chamber includes exopolymer-producing bacteria. In further embodiments, the source of bacteria includes a bacterial growth matrix. In additional embodiments, the source of bacteria includes a removable cartridge containing polymer-producing bacteria. Li yet other embodiments, the apparatus further includes at least one additional chamber having bacteria disposed within, for subsequently processing the treated water. In further embodiments, the apparatus includes at least one pressure sensor/flow regulator for controlling the water flowing through the chamber.

In other embodiments, the source of bacteria disposed within the chamber provides one or more exopolymer-producing bacteria that includes a Bacillus. In certain embodiments, the Bacillus is Bacillus cerueus and/or Bacillus thuringiensis. In other embodiments, the Bacillus is Bacillus gibsonii, Bacillus pseudalcaliphilu, Bacillus pseudofirmus, Bacillus halodurans, Bacillus subtilis, Bacillus benzoevorans, Bacillus simplex, Bacillus horikoshii, Bacillus cereus, or Bacillus thuringiensis. Ih still further embodiments, the bacteria is Proteus mirabilis. Li a particular embodiment, the invention provides the strain deposited as ATCC #51286). Li other embodiments, the invention provides bacteria that is G3 and/or MEX244.1. Li further embodiments, the bacteria bacteria is of the genus Pseudomonas, e.g., Pseudomonas putida or Pseudomonas aeruginosa. Li still further embodiments, the bacteria is of the genus Azotobacter, e.g., Azotobacter vinelandii, Azotobacter chroococcum or Azotobacter indicus.

Li still further embodiments, the apparatus includes a second output opening in the chamber for removal of cation-saturated bacterial exopolymer. Li particular embodiments, the chamber is just the first of multiple treatment chambers and the cation chelating, exopolymer- producing bacteria facilitate calcium removal within this first chamber. Li further embodiments, the apparatus also includes a second chamber that facilitates the formation of less stable forms of scale (mineral deposits). Li other embodiments, the apparatus includes a second chamber that facilitates the dissolving of scale (mineral deposits). Ih another embodiment, the apparatus comprises a first chamber that facilitates the control of scale by sequestering scale-forming ions, as well as a second chamber that facilitates the control of scale formation by promoting the formation of less stable forms of scale formation, and a third chamber that facilitates the control of scale formation by dissolving scale, as well as a fourth chamber that facilitates the control of scale formation by inhibiting new scale formation.

In another embodiment, the apparatus is part of a reverse osmosis system and/or a zero level discharge reverse osmosis system.

In a further aspect, the invention provides a method of using the cation-chelated microbial exopolymer-containing waste product of a water treatment system by utilizing the cation-chelated microbial exopolymer-containing waste product of the method of the invention as feed for animals (including livestock).

In yet another aspect, the invention provides a method of using the cation-chelated microbial exopolymer-containing waste product of a water treatment system as a source of energy by utilizing the cation-chelated microbial exopolymer-containing waste product of the method of the invention as a biofuel source.

In still another aspect, the invention provides a method of using the cation-chelated microbial exopolymer-containing waste product of a water treatment system as a useful land management material by utilizing the cation-chelated microbial exopolymer-containing waste product of the method of the invention as landfill.

3. BRIEF DESCRIPTION OF THE FIGURES

FIG. 1 A is a diagrammatic representation of an apparatus for cation removal. The apparatus includes two tangential filters, a bacterial source, an input and output site.

FIG. 1 B is a diagrammatic representation of an apparatus for polymer release into a water system. The apparatus includes one tangential filter, a bacterial source, an input and output site.

FIG. 2 is a schematic representation of a phylogenetic tree indicating related bacterial strains.

FIG. 3 is a graphic representation of a heat flux titration curve by hydration for G3 EPS, MEX244.1EPS, P. mimbilis EPS and humic acid.

FIG. 4 is a graphic representation of a heat flux titration curve by a combination of hydration and Ca2⁺ for G3 EPS, MEX244.1EPS, P. mirabilis EPS and humic acid.

FIG. 5 is a graphic representation of a heat flux titration curve in the presence OfCaCO₃ for G3 EPS, MEX244.1EPS, P. mimbilis EPS and humic acid.

FIG. 6 A is a representation of a micrograph of calcite precipitation in the absence of EPS.

FIG. 6 B is a representation of a micrograph of calcite precipitation in EPS from Proteus mirabilis.

FIG. 6 C is a representation of a micrograph of calcite precipitation in EPS from G3.

FIG. 6 D is a representation of a micrograph of calcite precipitation in EPS from MEX244.1.

FIG. 7 A is a representation of a micrograph of calcite precipitation in the presence of curdlan.

FIG. 7 B is a representation of a micrograph of calcite precipitation in the presence of lichenan. FIG. 7 C is a representation of a micrograph of calcite precipitation in the presence in the presence of humic acid.

Fig. 8 is a graphic representation of the decrease in conductivity, associated with the removal of water-borne minerals, by serial passage of an artificial seawater through an alginic acid preparation.

Fig. 9 is a schematic representation of a treatment system using microorganisms to dimineralize water. Fig. 10 is a schematic representation of a 50 gallon (100 GPM) cartridge system for use in the microbial demineralization treatment system.

Fig. 11 is a schematic representation of a series of cartridge systems assembled onto a single skid.

Fig. 12 is a schematic representation of a reverse osmosis water treatment system incorporating a microbial exopolymer demineralization system of the invention to treat the reverse osmosis reject water, and produce reject water with reduced mineral ion content and nontoxic byproducts including microbes and salt-polymer complex.

4. DETAILED DESCRIPTION OF THE INVENTION

All patents, patent applications and publications cited herein are hereby incorporated by reference in their entirety. The disclosures of these publications in their entireties are thereby included with this application in order to more fully describe the state of the art as known to those skilled therein as of the date of the invention described and claimed herein.

All scientific literature, patents and patent applications (including priority applications U.S. Serial No. 60/723701, filed October 5, 2005, and U.S. Serial No. 60/845040, filed September 15, 2006) referred to herein establish the knowledge that is available to those with skill in the art, and are formally incorporated by reference herein. The invention provides compositions and associated methods and apparatus for the use of exopolymer-producing microbes to control water hardness of mineral content and scale formation. The following detailed description of the elements and exemplary embodiments of the invention are provided in support of the claimed invention summarized above.

4.1 General

Water is being used at an alarming rate due to population growth and industrial expansion. At the same time, the world's fresh water supply is shrinking as a result of pollution and the draining of underground aquifers. The economics are simple but alarming: water demand has tripled in the past 30 years while the population growth has grown by only 50%. Li 1995, 436 million people in 29 countries lived in places where the water supply was scarce or under stress. China, for example, needs and plans to build 375 wastewater treatment facilities by 2009. Given current trends, by 2035, the World Bank estimates that 3 billion people - one third of the world's estimated population - will not have access to adequate water.

On the demand side, the urgent need to build wastewater treatment systems in developing countries, the tighter regulatory requirements, and the increased water usage by industrial economies, are all considered long-term drivers (Goldman Sachs, "The Essentials of Water Investing," 15 June 2005). For this reason, water is often referred to as the "petroleum for the next century.

As the supply and demand imbalances for water become more severe, water treatment technologies are expected to become increasingly critical (Goldman Sachs, "The Essentials of Water Investing," 15 June 2005). From the process of drawing water from surface reservoirs and underground aquifers, through its eventual discharge, water undergoes specific treatment processes during its life cycle. Water treatment includes various systems for filtration and disinfection to increase the purity and clarity of the water supply to make it suitable for residential, commercial, and industrial uses. The World Bank has estimated that by 2007, investments of between $400 - $600 billion will be required to meet the demand for fresh water (http://www.twst. com/conferences/water_december/water_december .html).

Current demineralization treatments, including reverse osmosis (RO), are energy intensive, inefficient, and result in production of costly and environmentally unfriendly effluent reject water streams. Engineers at various power plants in the US maintain that the RO process efficiency is highly variable (depending mostly on water salinity) leaving behind effluent that costs $6 per 1000 gallons to dispose of. Additionally, as environmental regulation becomes more stringent in the US, it becomes more difficult to obtain permits to discharge high salinity effluent reject water streams. Calpine Corporation, an independent power producer in the United States, has permit problems associated with the disposal of its effluent reject water at 10 out of its 93 power plants. Currently post-RO effluent reject water is handled in one of three ways. It is transferred to sewage systems for a fixed price, disposed of in evaporation ponds, or discharged back into water systems such as oceans or rivers. No economical and non-polluting demineralization technique has been developed that addresses the problems associated with effluent disposal.

In the United States, thermoelectric power generation capacity is projected to increase dramatically by 2025. Thermoelectric power plants use large quantities of water, and the fact that western and southeast United States are already facing water availability issues poses a serious problem. Further, the United States will see the largest increase in population in these same areas, exacerbating this problem. Undoubtedly, power generators will compete more with other water users. Additionally, the Clean Water Act 316(b) mandates that in the United States, the location, design, construction, and capacity of cooling water intake structures reflect the best technology available for minimizing adverse environmental impact.

In general, the instant invention provides for various applications for controlling water hardness and scale formation in industrial, commercial and residential settings, into a cost- effective and more environmentally safe process. The invention includes methods of controlling water hardness and scale formation, methods of manipulating bacterial sources to control scale formation, methods of using exopolymers, which includes exopolysaccharides (EPS), to control water hardness and scale formation, and methods of producing EPS and acids for such use. The polymers work via a variety of mechanisms which include sequestering water-borne minerals and metals, which maybe scale-forming ions, promoting dissolution of existing scale, increasing the solubility of newly formed scale, and inhibiting scale formation on pipes.

The technology includes a device with a single or multiple chambers in series or parallel. Each chamber and/or the system of chambers contains a single or multi-species microbial population. Each chamber and/or the system of chambers may be inoculated by isolated species or be allowed to self-colonize by microorganisms naturally occurring in the water column. Hard water, which may include reverse osmosis reject water, from the treated system will be allowed to flow through the chambers. The "hardness" of the water is related to its concentration of minerals and metals, which is often standardized against calcium carbonate concentration, and is herein characterized into these five groups: soft (calcium carbonate concentration ranging < 17.1 mg/1), slightly hard (calcium carbonate concentration > 17.1 mg/1 but < 60), moderately hard (calcium carbonate concentration > 60 mg/1 but < 120), hard (calcium carbonate concentration > 120 mg/1 but < 180) and very hard (calcium carbonate concentration > 180 mg/1) see Table 1.

Table 1 : Classification of Water Hardness

The fluid, flow provides a fresh source of nutrients and acts to distribute the active chemicals to the system. High- volume filters retain the bacteria within the apparatus while allowing acids and polymers to flow into the system. These chemicals work to control water hardness and scale formation in the aforementioned means. Optimally, in the chamber containing the bacteria sequestering scale-forming ions, polymers do not flow out of the chamber and there exists a method for removal of saturated polymer from the system. The chambers may further contain physical growth matrix to enhance polymer production. The device is attached within the closed-loop or on the input to the water within a heating or industrial boiler system.

An advantage of the technology is that the active acids and polymers are obtained from a perpetually renewing source, the actively metabolizing bacteria, contained within the filter, which limits fixed costs for purchasing chemicals, improves worker safety because they do not need to handle caustic chemicals, and reduces environmental pollution. However, it is possible that the bacteria contained within the filtration apparatus may not have enough nutrients to produce sufficient quantities of active chemicals. In this case, the bacteria contained within the apparatus may be supplied with additional nutrients or another manifestation of the technology is to harvest the polymers and administer them to the system similar to current chemical treatments or to include scale-inhibiting polymers into novel coating materials.

Additionally, the microorganisms may include photosynthetic microorganisms, including algae and cyanobacteria. Ia this case, the apparatus will have exterior or exterior components that are transparent to sunlight. Also, the apparatus may contain reflective surfaces to maximize microbial exposure to sunlight.

4.2 Definitions

As used herein, "microbes" refers to microorganisms which produce exopolysaccharides, including, but not limited bacteria, algae, fungi and lichens. As used herein, "hard water" refers to water containing salt ions such as calcium and magnesium. The relative "strength" of the hard water is defined in Table 1.

As used herein the term "exopolymer" refers to secreted polysaccharides

(exopolysaccarides) that are produced by bacteria and other microbes. As such, exopolymers are biopolymers that are secreted by a microbe into the environment (i.e. external to the microbe) and are frequently found as components of biofilms. Such biofilms are complex aggregations of microorganisms marked by the excretion of a protective and adhesive matrix. Biofilms are produced by bacteria to anchor them and protect them from environmental conditions. Biofilms are also often characterized by surface attachment, structural heterogeneity, genetic diversity, complex community interactions, and an extracellular matrix of polymeric substances.

In general, as used herein, the term "biopolymer" refers to a polymer found in nature. Starch, proteins and peptides, and DNA and RNA are all examples of biopolymers, in which the monomer units, respectively, are sugars, amino acids, and nucleic acids. The exact chemical composition and the sequence in which these units are arranged is called the polymer's primary structure. Many biopolymers spontaneously "fold" into characteristic shapes, which determine their biological functions and depend in a complicated way on their primary structures. Structural biology is the study of the shapes of biopolymers.

4.3 Microbes

Microbes are micro-organisms usually unicellular and sometimes multicellular in composition. Microbes can be identified throughout nature. Microbes can be helpful in recycling other organisms' remains and waste products, or employed in biotechnology, for brewing, baking and environmental clean-up. Microbes may also be parasitic and result in deleterious infections and diseases. Microbes are found throughout the taxonomic structure and include, but are not limited to, bacteria, fungi, algae and lichens.

Although not limited to, the invention describes the use of microbes, particularly bacteria, and algae, and their products, for water hardness and mineral scale control. Bacteria can be divided into five phyla: Proteobacteria, Cyanobacteria, Gram-Positive Eubacteria, Spirochetes, and Chlamydiae. Proteobacteria, the most diverse group of bacteria, include nitrogen-fixing bacteria in the root nodules of legumes, as well as enteric bacteria that live in the intestinal tract of animals (including E. coli). The nitrogen-fixing bacteria convert inorganic nitrogen in the form of atmospheric N₂ to NH₄ ⁺ (ammonium) and NO₃ ^" (nitrates) in the soil, which can be incorporated into the amino acids and nucleic acids of plants, after being absorbed through their roots. Proteobacteria are categorized in three main subgroups: purple bacteria, chemoautotrophic proteobacteria, and chemoheterotrophic proteobacteria.

Purple bacteria use energy from the sun but extract electrons from substances other than water, and therefore release no oxygen. Most species are strict anaerobes and live in the sediment of ponds and lakes. Purple non-sulfur bacteria are found among the alpha and beta subgroups, including: Rhodospirallales {Rhodospirillum and Rhodopilά), Rhizobiales (Rhodopseudomonas and Rhodobium) and Rhodobacteraceae (Rhodobacter). Purple sulfur bacteria are included among the gamma subgroup, and make up the order Chromatiales.

Chemoautotrophic proteobacteria are free living species. Many are mutualists, including the nitrogen-fixing bacteria. They play roles in the cycles of chemicals within the environment by fixing nitrogen as legume root symbionts, thus contributing to the plant's nutrition and providing organic nitrogen to the environment. Examples of chemoautotropic proteobacteria include Neisseria meningitidis, Neisseria gonorrhoeae, and Bordetella pertussis

Chemoheterotrophic proteobacteria are parasitic. Some, including the enteric bacteria, live in the intestinal tracts of animals. Many of these are facultative anaerobes, able to participate in aerobic or anaerobic respiration. Many are harmless but some, {e.g. Salmonella), are pathogenic. Examples of chemoheterotrophic proteobacteria include Salmonella enterica , Escherichia coli, Vibrio cholerae, and Pseudomonas aeruginos.

Although not limited to any particular bacteria, the bacteria may be one of the following: Bacillus abysseus (ATCC #14409), Bacillus acidocaldarius (ATCC #43030), Bacillus alcalophilus (ATCC #43592), Bacillus apiarius (ATCC #29575), Bacillus capitovalis (ATCC #29318), Bacillus cereus (ATCC #23260), Bacillus circulans (ATCC #13403), Bacillus coagulans (ATCC# 12245), Bacillus colofoetidus (ATCC #11811), Bacillus dendrolimus (ATCC #19266), Bacillus fastidiosus (ATCC #29312), Bacillus firmus (ATCC #14575), Bacillus glucanolyticus (ATCC #49278), Bacillus lentimorbus (ATCC #14707), Bacillus licheniformis (ATCC #21039), Bacillus macerans (ATCC #49035), Bacillus megaterium (ATCC #13402), Bacillus naphthovorans (ATCC #BAA-550), Bacillus natto (ATCC #15245), Bacillus palustris (ATCC #15329), Bacillus pasteuri i(ATCC#11859), Bacillus polymyxa (ATCC #12321),

Bacillus pumilus (ATCC #14884), Bacillus sphaericus (ATCC #12123), Bacillus stearothermophilus (ATCC # 12976), Bacillus sύbmarinus (ATCC #14415), Bacillus subtilis (ATCC #12432), Bacillus sulfasportare (ATCC #39909), Bacillus thermoleovorans (ATCC #43505) or Bacillus thiamine- Iy ticus (ATCC #13023).

Further, the bacteria may be one of the following: Pseudomonas acidovorans (ATCC #15667), Pseudomonas aeruginosa (ATCC #25319), Pseudomonas alcaligenes (ATCC #14909), Pseudomonas aureofaciens (ATCC #17418), Pseudomonas boreopolis (ATCC #15452), Pseudomonas caryophylli (ATCC #11441), Pseudomonas cepacia (ATCC #49709), Pseudomonas chlororaphis (ATCC #13986), Pseudomonas cichorii (ATCC #13455), Pseudomonas citronellolis (ATCC #13674), Pseudomonas creosotensis (ATCC #14582), Pseudomonas dacunhae (ATCC #13261), Pseudomonas delafieldii (ATCC #17506), Pseudomonas denitriflcans (ATCC #13867), Pseudomonas desmolytica (ATCC #15005), Pseudomonas diminuta (ATCC #11568), Pseudomonas echinoides (ATCC #14820), Pseudomonas excibis (ATCC #12293), Pseutomonas fluorescens (ATCC #14150), Pseudomonas fulva (ATCC #14598), Pseudomonas haloplanktis (ATCC #14393), Pseudomonas iodinum (ATCC #15729), Pseudomonas lanceolata (ATCC #14669), Pseudomonas maltophilia (ATCC #17674), Pseudomonas mangiferae-indicae (ATCC #11637), Pseudomonas marginalis (ATCC #13889), Pseudomonas melanogenum (ATCC #14535), Pseudomonas meliae (ATCC #33050), Pseudomonas nactus (ATCC #12294), Pseudomonas natήegens (ATCC #14048), Pseudomonas oxalaticus (ATCC #11451), Pseudomonas papaveris (ATCC #13041), Pseudomonas perlurida (ATCC #15048), Pseudomonas piscicida (ATCC #15251), Pseudomonas primulae (ATCC #13038), Pseudomonas pseudoalcaligenes (ATCC #12815), Pseudomonas putida (ATCC #27393), Pseudomonas reptilivora (ATCC #14039), Pseudomonas rhodos (ATCC #14821), Pseudomonas savastanoi (ATCC #13522), Pseudomonas schuylkilliensis (ATCC #15916), Pseudomonas stutzeri (ATCC #11607), Pseudomonas syringae (ATCC #13396), Pseudomonas syzygii (ATCC #49543), Pseudomonas testosteroni (ATCC #11996), Pseudomonas tolaasii (ATCC #14340), Pseudomonas trifolii (ATCC #14537), or Pseudomonas tuticorinensis (ATCC #12230).

Still further, the bacteria maybe one of the following: Azobacter nigricans (ATCC #35009), Azobacter beijerinckii (ATCC #17087), Azobacter chroococcum (ATCC #4412), Azobacter salinestris (ATCC #49674), Azobacter vinelandii (ATCC #12837). Algae comprise several different groups of living organisms usually found in wet places or water bodies. They capture light energy through photosynthesis, converting inorganic substances into simple sugars with the captured energy. Algae were traditionally regarded as simple plants, and some are closely related to the higher plants. Others appear to represent different protist groups, alongside other organisms that are traditionally considered more animal- like (protozoa).

Algae range from single-celled organisms to multi-cellular organisms, some with fairly complex differentiated forms and some are called seaweeds. All lack leaves, roots, flowers, and other organ structures that characterize higher plants. They are distinguished from other protozoa in that they are photoautotrophic, although this is not a hard and fast distinction as some groups may contain members that are mixotrophic, deriving energy both from photosynthesis as well as through the uptake of organic carbon either by osmotrophy, myzotrophy, or phagotrophy. Some unicellular algae rely entirely on external energy sources and have reduced or lost their photosynthetic apparatus.

All algae have photosynthetic machinery ultimately derived from the cyanobacteria, and produce oxygen as a by-product of photosynthesis, unlike other, non-cyanobacterial photosynthetic bacteria. Algae are common in terrestrial as well as aquatic environments, but usually inconspicuous on the land and more common in moist, tropical climates. The various sorts of algae play significant roles in aquatic ecology. Microscopic forms that live suspended in the water column, called phytoplankton, provide the food base for most marine food chains. In very high densities (so-called algal blooms) they may discolor the water and outcompete or poison other life forms. The seaweeds grow mostly in shallow marine waters; some are used as human food or are harvested for useful substances such as agar or fertilizer. The study of algae is called phycology or algology. Examples of EPS producing algae include, but are not limited to, Volvox aureus, Volvox carteri, Volvox globactor, Volvox dissipatrix and Volvox tertios.

Algae that colonize solid surfaces to form an "algal surface biofilm" may also be used as microbes of the invention. Such "algal surface biofilm" forming algae and their use in an "Algal surface biofilm Water Purification Method" is described in U.S. Patent No. 5,851,398, the contents of which are hereby incorporated by reference in their entirety. In particular, algal surface biofilm species such as Compsopogon coeruleus, Cladophora crispata, Spirogyra rivularis, Enteromorpha micrococca, Eunotia pectinalis, and Melosira varian, Stigeoclonium tenue, among others, may also be used. Other such algae include small pennolean diatoms, particularly Amphora and Cocconeis species. The algal growth of the listed groups is random on the growing surface, and wave action maybe passed across and through the turf to enhance metabolite cellular-ambient water exchange. The use of a screen, such as a plastic screen, as a growing surface provides good results, although other surfaces known in the art can be used. Typically, such a growing surface can be a plastic screen having screen grip dimensions in the range of approximately 0.5 to 5 mm. It should be plain to those familiar with algae that the above-identified genera and species are all attached, as opposed to planktonic, algae.

Further microalgae for growing an algal surface biofilm include the benthic microalgae such as: Cyanophycota (Cyanobacteria or Blue Green Algae), including Oscillatoria, Lyngbya, Schizothrix, Chroococcus Calothrix; Chlorophycota (Green Algae) including Ulothrix, Enteromorpha, Spirogyra, Cladophora, Dichotomosiphon, Stigeoclonium, Oedogonium, Mougeotia, Gloeocystis; Chromophycota (primarily Diatoms), including Melosira, Ctenophora, Asterionella, Eunotia, Amphipleura, Cocconeis, Placoneis, Rhoikoneis, Bacillaria, and others; and Rhodophycota (Red Algae) including Compsopogon. In waters of brackish to higher salinities, such as that of estuaries, coastal waters, or seawater, the genera Polysiphonia, Herposiphonia, and Callithamnion, among others, are also useful. Accordingly, for growing the algal surface biofilm, the present invention utilizes major groups of benthic microalgae. The benthic microalgae for practicing the present invention can be selected from the group consisting of green and blue-green algae for low to moderate saline waters (0-10 ppm) and including red and brown algae for high saline waters. The brown algae include diatoms. Still further, the algae are selected from the group comprising green and red algae for low-to-moderate salinity waters. The microalgae spores for growing such benthic microalgae can be obtained as described in the U.S. Patent Nos. 4,333,263, 4,966,096 and 5,097,795, each of which is hereby incorporated by reference in its entirety.

Still further light-sensitive microorganisms for use in the invention include Halobacterium, Oscillatoria or Aphanocapsa species (see Uma and Subramanian (1990) Proc. Natl. Svmp. Cyanobactt. Nitrog. Fix.. IAPJ, New Delhi. Pages 437-444). The genus Halobacterium consists of several species of archaea with an obligate aerobic metabolism which require an environment with a high concentration of salt; many of their proteins will not function in low-salt environments. They grow on amino acids in their aerobic conditions. Their cell walls are also quite different from those of bacteria, as ordinary lipoprotein membranes fail in high salt concentrations. In shape, they may be either rods or cocci, and in color, either red or purple. They reproduce using binary fission (by constriction), and are motile. Halobacterium grows best in a 37 degree Celsius environment. Halobacterium can be found in the Great Salt Lake, the Dead Sea, Lake Magadi, and any other waters with high salt concentration. Purple Halobacterium species owe their color to bacteriorhodopsin, a light-sensitive protein which provides chemical energy for the cell by using sunlight to pump protons out of the cell. The resulting proton gradient across the cell membrane is used to drive the synthesis of the energy carrier ATP. Thus, when these protons flow back in, they are used in the synthesis of ATP. The bacteriorhodopsin protein is chemically very similar to the light-detecting pigment rhodopsin, found in the vertebrate retina. Exemplary Halobacterium species include Halobacterium cutirubrum Halobacterium denitrificansJHalobacterium distrϊbutum, Halobacterium halobium, Halobacterium lacusprofundi, Halobacterium mediterranei, Halobacterium noricense, Halobacterium pharaonis, Halobacterium saccharovorum, Halobacterium salinarium, Halobacterium sodomense, Halobacterium trapanicum, Halobacterium vallismortis and Halobacterium volcanii. Oscillatoria are Cyanobacteria that form cylindrical or sometimes slightly tapering, unbranched filaments (trichomes), often with a rounded or capitate apical cell. Other cells are discoid, with further developing cross-walls. There are no heterocysts or akinetes. Trichomes leave a thin mucilaginous trail as they glide. This genus is well known for its motility, trichomes being able to glide apparently by means of wave movements of microfibrils, so long as the cells are in contact with a solid substrate. Mucilage is secreted through pores in the cell walls and may help to provide better contact with the substrate surface. Movement has been timed at up to 11 μm per second. Species of Oscillatoria occur in a diverse range of conditions, in damp soil or on dripping rocks, in freshwater, in the sea and in hot springs. Some are tolerant of high levels of organic pollution and some are shade-tolerant and able to survive in water below blooms of green algae. In water they may be benthic or planktonic. O. rubescens is a red species that can form conspicuous red blooms in eutrophicated lakes. Oscillatoria is implicated in irritation of skin and mucous membranes suffered by people swimming off tropical coastlines. Exemplary Oscillatoria species include Oscillatoria simplicissim. Aphanocapsa are Cyanobacteria that take the form of many-celled aggregates of widely spaced cells in a globular mucilage. Like nearly all blue-green algae, this genus may produce lipopolysaccharides capable of causing skin irritation and gastrointestinal distress. Aphanocapsa delicatissima is a dominant component of the summer phytoplankton assemblage in Lake Michigan. Other exemplary Aphanocapsa species include Aphanocapsa elachista, Aphanocapsa endophytica, Aphanocapsa grevillei, Aphanocapsa pulchra, and Aphanocapsa rivularis.

Fungi are a group of organisms that are now thought of as one of the four kingdoms of eukaryotes, the others being animals, plants and protists. Long counted among the plantae they are more recently considered to be more closely related to the animals and grouped together with these in the taxon of opisthokonts. Notable differences from animals include the mostly saprobiontic nutrition of fungi and in many cases the presence of a cell wall. This cell wall comprises chitin as a structural component, which together with their chemo-organo-heterotrophy distinguishes fungi from the photoautotrophic plants. Chitin cell walls and haploid nuclei differentiate them from structurally similar protists such as water molds. Fungi reproduce either sexually or asexually through spores. Both unicellular and multicellular forms exist.

Fungi occur in all environments on the planet and include important decomposers and parasites. Parasitic fungi infect animals, including humans, other mammals, birds, and insects, with consequences varying from mild itching to death. Other parasitic fungi infect plants, causing diseases such as butt rot and making trees more vulnerable to toppling. The vast majority of vascular plants are associated with mutualistic fungi, called mycorrhizae, which assist their roots in absorption of nutrients and water.

Lichens are symbiotic organisms made up by the association of microscopic green algae or cyanobacteria and filamentous fungi. Lichens take the external shape of the fungal partner and hence are named based on the fungus. The fungus most commonly forms the majority of the lichen's bulk, though in filamentous and gelatinous lichens this may not always be the case. Some lichen taxonomists place lichens in their own division, the Mycophycophyta, but this practice ignores the fact that the components may belong to separate lineages. The algal cells contain chlorophyll, permitting them to live in a purely mineral environment by producing their own organic compounds. The fungus protects the alga against drying out and, in some cases, provides it with minerals obtained from the substratum. If a cyanobacterium, such as in Terricolous Lichens, is present this can fix atmospheric nitrogen, complementing the activities of the green alga. Examples of fungi and lichens which produce EPS include, but are not limited to, Acremonium persicinum, Acremonium pullulan, Aspergillus flavipes, Aureobasidium pullulans, Cordyceps militaris, Epicoccum purpurascens, Ganoderma lucidum, Plerutous pulmonarius, Paecilomyces sinclairii, Phanerocheates chrysosporium, Phellinus linteus, Plerutous florida, Sclerotium glucanicum, Rhizobium, Bradyrhizobium, Cordyceps militaris and Volvariella volvacea

AA Bacteria and Bacterial Sources

Bacteria which produce and secrete EPS are described herein. Examples of bacteria which produce exopolysaccharides include, but are not limited to, G3, MEX244.1, Bacillus thuringiensis, Bacillus cereus, Bacillus gibsonii, Bacillus pseudalcaliphilus, Bacillus pseudofirmus, Bacillus halodurans, Bacillus subtilis, Bacillus benzoevorans, Bacillus simplex, Bacillus horikoshii, Burkholderia cepacia, Proteus mirabilis, Lactobacillus delbruecldi, Lactobacillus acidophilus, Lactobacillus fermentum, Streptococcus thermophilus, Pediococcus spp, Leuconostoc mesenteroides, Sinorhizobium meliloti, Staphylococcus aureus, Lactobacillus delbrueckii, Salmonella typhimurium, Salmonella Poona, Salmonella enterica and Lactobacillus reuteri.

The bacteria may be natural isolates that produce the appropriate chemicals. These bacteria can be optimized for different pH levels, nutrient concentrations, temperature ranges, and other variables found within different piping systems. Also, microorganisms may be engineered to hyperexpress the active chemicals. The gene(s) coding for chemical expression may be identified and cloned into other microorganisms with better growth profiles in a given environment. The chemicals themselves, particularly the complex polysaccharides, may have useful novel properties that make them superior to current chemical treatments.

Other applications of the technology include any application of heat transfer engineering, including residential use, industrial use, waste- water treatment facilities, power-generation facilities.

Alkaliphilic bacteria are often the primary colonizers of fresh limestone surfaces. A freshly exposed calcite mineral surface under aqueous conditions has a pH of 8 to 10 (Horikoshi 1998), which naturally enriches for alkaliphilic or alkalitolerant bacteria. These initial colonizers produce metabolic byproducts, such as EPS. These byproducts may contribute to early dissolution processes during biofilm development (Perry IV, et al. (2004) Env. Sci. Technol. 38: 3040-3046; Perry IV et al. (2005) J. Am. Chem. Soc. 127: 5744-5745). The bacteria have the metabolic ability to produce different EPS depending on growth stage, nutrient conditions, and other environmental factors, which will have different dissolution effects.

Non-polar electron-donating groups are important in stabilizing the EPS-calcium complex. The EPS monosaccharide residues detected in the study are a mixture of five and six membered sugars without reactive moieties that would typically be implicated in reaction with a polar mineral surface, such as carboxylates (Perry TV, Estroff, et al. (in prep) BiogeochemΛ. However, carboxylates are absent on the monosaccharides detected in the harvested EPS. The absence of these moieties indicates that hydration of the polymer, rather than Coulombic interactions, may be the driving force for surface adsorption (Dimova, et al. (2003 Langmuir 19(15): 6097- 6103) and that the role of hydroxyls and linkage esters in binding to a mineral surface are significant (Perry IV, Estroff, et al. (in prep) Biogeochem.^*).

In natural environments, biological macromolecules such as EPS play an underestimated role in dissolution reactions. In terrestrial environments, humic acids are the most abundant organic species and have been demonstrated to play a role in mineral weathering. However, EPS polysaccharides are the second most abundant biopolymer. The observation that EPS have different binding interactions with calcite and that the effect is of similar magnitude to that of humic acid indicates that these polymers are considered when modeling mineral weathering, since EPS is an important contributor to the dissolution of minerals in the environment.

4.5 Bacterial Isolation and Genotyping

The microbial samples are collected from the dark interior of Tomb 25, Athienou Archaeological Project, Malloura, Cyprus. The microorganisms are removed from the stone surface by swabbing using a Q-tip in a sterile solution of saline (0.85% NaCl) and dilute (<0.1%) non-toxic surfactant (Triton X-1000) in deionized water. Collected organisms are released into suspension by vortexing and are enriched for alkaliphilic organisms by inoculating the suspensions on a solid alkaliphilic growth medium modified from (Horikoshi 1998) set to pH 10.5 and allowed to grow at room temperature. Per liter of deionized water, the medium consisted of 10 g dextrose, 7 g NaHCO₃, 1O g polypeptone, 1O g yeast extract, 1 g KH₂PO₄, 0.2 g Mg₂SO₄ '7H₂O, and 20 g agar. The dextrose and NaHCO₃ each are prepared in separate 100 mL flasks to prevent hydrolysis. Each solution is adjusted to the desired pH, autoclaved, and combined after cooling. Pure bacterial cultures are obtained by repeated streaking. Several isolates are screened for their ability to produce polymer at high pH values and one isolate (identified as isolate GS) is chosen for further experimentation. The isolate MEX244.1 is selected from a library of microorganisms collected from the Acropolis at the Maya site at Ek' Balam, Yucatan, Mexico (McNamara et al. (in press) Microb. Ecolog.^'), and enriched under alkaline conditions using a medium containing precipitated calcium carbonate (Di Bonaventura et al. (1999) Geomicrobiol. J. 16: 55-64). A strain of Proteus mirabilis (ATCC #51286) is acquired from the American Type Culture Collection (Manassas, VA).

The selected isolates are identified by 16S rRNA gene sequencing. DNA is extracted using the UltraClean Soil DNA Kit (MoBio Labs, Carlsbad, CA). A portion of the 16S rDNA genes are amplified using the primers 27f and 1492r (Lane 1991) in PCR protocol (Schabereiter- Gurtner et al. (2001) J. Microbiol. Meth. 45: 77-87) carried out in a Robocycler (Stratagene, La Jolla, CA) for 35 cycles under the following conditions: initial denaturation at 94°C for 3 min, followed by 15 cycles of denaturation at 94°C for 1 min, primer annealing at 5O⁰C for 1 min, elongation at 72°C for 2 min with a final extension step at 72°C for 5 min. PCR reactions are conducted in 50 μL volumes and contained 25 pmol of each primer, 0.2 mM of each dNTP, 5.0 μL of 10x PCR buffer (200 mM Tris-HCI, pH 8.4, 500 mM KCl), 2 mM MgCI₂, 2 U of Taq DNA polymerase (Invitrogen, Carlsbad, CA), 4 μL of template DNA from the extractions, and nanopure deionized water (18.3 MΩ cm; Barnstead, Dubuque, IA). The amplified fragments are precipitated using a QIAquick PCR purification kit (Qiagen, Valencia, CA) and resuspended. The fragments served as the template for the sequencing PCR reaction using three primers to obtain complete sequences: 27f, 907r, 1942r (Lane 1991) and a BigDye Terminator kit (Applied Biosystems, Foster City, CA).

16S rRNA gene sequences are edited and assembled using the Sequencher software (Gene Codes) and are checked for quality by manually mapping to a secondary structure of Bacillus cereus 16S rRNA (Cannone, et al. (2002) BioMed. Central. Bioinf. 3(2): doi: 10.1186/1471-2105-3-2.; Cole et al. (2005) Nucl. Acids Res. 1(33): Database Issue:D294-6). Related sequences are identified by blasting against Genbank (Altschul et al. (1997) Nucl. Acids. Res. 25(17): 3389-3402.) and by searching against the RDP (Maidak, et al. (1999 Nucl. Acids Res, 25(109- 110).; Maidak e£ α/. (2001) Nucl. Acids Res. 29: 173-174). Phylogenetic analyses are conducted in PAUP, version 4.ObIO (Swofford, D. L. (1993 J. Gen. Physiol. 102: A9.). Relationships are determined using the Neighbor-joining method with Jukes-Cantor correction and checked for consistency using Parsimony and Maximum Likelihood methods. Almost the full length of the sequences (E. coli positions 98 - 1460) are used in the phylogenetic analyses. For each analysis, bootstrap resampling with minimum evolution method with 1000 replicates is used to test robustness.

4.6 Εxopolysaccharides

Biofilms are a rich source of biological ligands. Mature biofilms on mineral surfaces are temporally, spatially, and taxonomically dynamic communities of microorganisms that can affect dissolution through production of metabolic byproducts. Bacteria, Archaea, algae, fungi, and lichens increase calcite dissolution rates through the production of metabolic byproducts, such as organic and inorganic acids (Christensen et al (1990). Physical and Chemical Properties in Biofilms. Biofilms. W. G. Charaklis and K. C. Marshall. New York, Wiley.; Sand, W. (1997) Mernat. Biodeterioration & Biodegradation 40(2-4): 183-190.; Perry IV et al. (2005) J. Am. Chem. Soc. 127: 5744-5745.). Other complex biologically derived compounds including lipids and phospholipids (Suess, Ε. (1970) Geochim. Cosmochim. Acta 34: 157-168.) Geochim. Cosmochim. Acta 34: 157-168.; Frye, et al (1993) Chem. Geol. 109: 215-226), proteins (Suess, Ε. (1970) Geochim. Cosmochim. Acta 34: 157-168.), humics (Hoch et al. (2000) Geochim. Cosmochim. Acta 64: 61-72.) Geochim. Cosmochim. Acta 64: 61-72.), and other carboxylated compounds (Fredd et al. (1998) J. Coll. Inter. ScL 204: 187-194; Wu et al (2002) Langmuir 18: 6813-6820) inhibit dissolution (Compton et al (1993) J. Coll. Inter. Sci. 158: 439-445; Thomas et al. (1993) Chem. Geol. 109: 227-237).

Bacteria, Archaea, cyanobacteria, algae, fungi, and lichens stimulate calcite dissolution through production of metabolic by-products, including organic acids and exopolysaccharides (EPS) (Christensen, et al (1990). Physical and Chemical Properties in Biofilms. Biofilms. W. G. Charaklis and K. C. Marshall. New York, Wiley.; Perry et al. (2005) Biodeterioration of stone. In: Scientific Examination of Art: Modern Techniques in Conservartion and Analysis. Nat. Acad. Sci., Washington, DC, pp. 72-84. The exudates are produced by microorganisms in biofilms, which are heterogeneous communities of microorganisms attached to the stone surface in an anchoring matrix of excreted EPS (Costerton et al. (1999) ScL 284(5418): 1318-1322). The composition of EPS is genotypically, phenotypically, and environmentally regulated. Its chemical structure varies by microorganism, growth stage, nutrient abundance, and other environmental stimuli (Christensen, et al (1990). Physical and Chemical Properties in Biofilms. Biofilms. W. G. Charaklis and K. C. Marshall. New York, Wiley.). The EPS is generally comprised of a variety of sugars, including uronic acids, and often contains functional groups (such as carboxylic acids) that can interact with mineral ions. The specific dissolution effects of EPS depend on mineral type, ligand functionality, acidic moieties, and pH. EPS can impact mineral weathering by a variety of mechanisms (Barker et al. (1997) J. F. Banfield and K. H. Nealson. Washington, D.C., Minearlogical Soc. of America. 35: 391-428.) and has been demonstrated to either accelerate or retard dissolution rate, although the precise mechanisms and reactions are not well understood (Welch, et al (1994) Geomicrobiol. J. 12: 227- 238; Banfield et al. (1999). Proc. Natl. Acad. Sci. (USA) 96(7): 3404-3411 : Flemming. et al. (2001) Water Sci. Technol 43(6): 9-16).

Exopolysaccharides (EPS) are a particularly important class of biological ligand, which biofilm bacteria produce in large amounts. EPS bind minerals and affect the dissolution rate. EPS are generally composed of a variety of sugars often containing functional groups (such as carboxylic acids) that can interact with mineral ions, such as iron or aluminum (Barker, et al. (1996) Chem. Geol. 132: 55-69). The chemical structure of EPS is genotypically, phenotypically, and environmentally regulated. It varies by microorganism, growth stage, nutrient abundance, and various environmental stimuli (Christensen et al. (1990). Physical and Chemical Properties in Biofilms. Biofilms. W. G. Charaklis and K. C. Marshall. New York, Wiley.). EPS binds to minerals with different strengths, and the complex nature of the EPS- mineral interaction arises from the detailed chemical compositions of EPS and mineral surfaces. The specific dissolution effects of EPS depend on mineral type, ligand functionality, acidic moieties, and pH.

EPS polymers are complex macromolecules and difficult to study. These polymers are built up from monomelic units, and the study of these simpler units provides some important insights. For example, although literature reports on the effects of simple organic polydentate ligands on calcite dissolution are sparse, it appears that the distinctive dissolution effects of ligands depend strongly on their chemistry. Chelating agents, including polyaspartic acid (PASP), ethylenediamine tetraacetic acid (EDTA), diethylenetriaminepentaacetic acid (DTPA), and 1,2-cyclohexanediaminetetraacetic acid (CTDA), increase the dissolution rate (Fredd et al. (1998) J. Coll. Inter. Sci. 204: 187-194; Wu et al (2002) Langmuir 18: 6813-6820.). Many biologically derived compounds, however, inhibit dissolution, including lipids and phospholipids (Suess, E. (1970) Geochim. Cosmochim. Acta 34: 157-168.; Frye, et al. (1993) Chem. Geol. 109: 215-226), proteins (Suess, E. (1970) Geochim. Cosmochim. Acta 34: 157-168.), humics (Hoch et al. (2000) Geochim. Cosmochim. Acta 64: 61-72.), and other carboxylated compounds (Fredd et al. (1998) J. Coll. Inter. Sci. 204: 187-194; Wu et al (2002) Langmuir 18: 6813- 6820.), though the effect varies widely with chemical identity and aqueous conditions (Compton et al (1993) J. Coll. Inter. Sci. 158: 439-445; Thomas et al. (1993) Chem. Geol. 109: 227-237). Polysaccharides, the dominant EPS constituent, have been observed to bind to cations on mineral surfaces (carbonates and silicates), both inhibiting and promoting dissolution (Thomas et al. (1993) Chem. Geol. 109: 227-237; Welch, et al (1994) Geomicrobiol. J. 12: 227- 238). However, no previous studies investigating the effect of polysaccharides on calcite dissolution have been reported.

4.7 Mechanism of Action of Exopolvsaccharides

EPS acts by several mechanisms (Barker et al. (1997) J. F. Banfϊeld and K. H. Nealson. Washington, D. C, Minearlog. Soc. Am. 35: 391-428.), such as by decreasing aqueous saturation through secondary precipitation or by chelating dissolution-inhibiting ions (Perry IV et al. (2005) J. Am. Chem. Soc. 127: 5744-5745). Different types of EPS have been observed to either accelerate or slow mineral dissolution rates (Thomas et al. (1993) Chem. Geol. 109: 227-237; Welch, et al (1994) Geomicrobiol. J. 12: 227- 238). However, how these mechanisms and reactions precisely work is not well established (Welch, et al (1994) Geomicrobiol. J. 12: 227- 238; Banfield et α/. (1999). Proc. Natl. Acad. Sci. OJSA) 96(7): 3404-3411 ; Flemming, et al (2001) Water Sci. Technol. 43(6): 9-16). A model, commercially available microbial polysaccharide produced by ubiquitous microorganisms accelerates calcite dissolution by a crystallographically specific, surface-chelation mechanism (Perry IV, et al. (2004) Env. Sci. Technol. 38: 3040-3046). Quantification of EPS-mineral interaction is, however, challenging due to the difficulty of retrieving EPS from natural microbial isolates and the chemical complexity of the interactions.

Alkaliphilic bacteria are often the primary colonizers of fresh limestone surfaces. A freshly exposed calcite mineral surface under aqueous conditions has a pH of 8 to 10 (Horikoshi, K. (1998). Extremophiles: Microbial Life in Extreme Environments. K. Horikoshi and W. D. Grant. New York, Wiley-Liss: 155-180), which naturally enriches for alkaliphilic or alkalitolerant bacteria. These initial colonizers produce metabolic byproducts, such as EPS. These byproducts may contribute to early dissolution processes during biofilm development (Perry IV₅ et al. (2004) Env. Sci. Technol. 38: 3040-3046; Perry IV et al. (2005) J. Am. Chem. Soc. 127: 5744-5745.). The bacteria have the metabolic ability to produce different EPS depending on growth stage, nutrient conditions, and other environmental factors, which have different dissolution effects.

Non-polar electron-donating groups are important in stabilizing the EPS-calcium complex. The EPS monosaccharide residues detected are a mixture of five and six membered sugars without reactive moieties that would typically be implicated in reaction with a polar mineral surface, such as carboxylates (Perry IV, Estroff, et al. (in prep) Biogeochem.). However, carboxylates are absent on the monosaccharides detected in the harvested EPS. The absence of these moieties indicates that hydration of the polymer, rather than Coulombic interactions, may be the driving force for surface adsorption (Dimova et al. (2003). Langmuir 19(15): 6097- 6103.) and that the role of hydroxyls and linkage esters in binding to a mineral surface are significant (Perry IV, Estroff, et al. (in prep) Biogeochem.).

In natural environments, biological macromolecules such as EPS may play an underestimated role in dissolution reactions. In terrestrial environments, humic acids are the most abundant organic species and have been demonstrated to play a role in mineral weathering. However, EPS polysaccharides are the second most abundant biopolymer. The observation that EPS can have different binding interactions with calcite and that the effect can be of similar magnitude to that of humic acid indicates that these polymers should be considered when modeling mineral weathering, since EPS may be an important contributor to the dissolution of minerals in the environment.

4.8 Algal surface biofilm Growth

Algal surface biofilm growth is achieved in an aqueous environment by providing any suitable vacant area in which spores may settle. The algal surface biofilm may be grown in a trough or floway. The water entering the floway may come from any source of water. The water can contain one or more undesirable elements, such as calcium and other scale-forming minerals.

The first colonizations of an algal surface biofilm are typically microscopic diatoms or blue green algae (cyanobacteria) which are then rapidly dominated by the turf species. In accordance with the present invention, the harvesting of such turfs may occur before the turf species are overgrown in turn by the larger macroalgae. The harvest interval may range from about one to about four weeks. This harvest timing keeps production rates at a high level and minimizes predation by grazing microorganisms. The rate of harvesting is dependent on nutrient levels, light levels, temperature and surge action, and perhaps other floway operating conditions.

Regrowth of the algal surface biofilm is facilitated if the newly harvested surface is sufficiently coarse to allow the filamentous base of the algae to remain following harvesting. Alternately, all algae is removed and the growing surface "seeded" with new algal spores. This process is valuable in special cases, for example, when necessary for micrograzer control, after a shut down of either the floway, or of the source generating the water, for repairs, power loss, etc.

Using screens as the growing surface, harvesting is accomplished by simply scraping the surface or, in the context of artificial growing techniques, the screen is set up for removal from the floway for harvesting. Vacuum harvesting techniques greatly decrease labor. In addition to the use of screens, other growing surfaces comprise, for example, any rough surface on which algae can grow.

As used herein, the term "algal surface biofilm" and its derivatives refers to a colony of attached microalgae and/or smaller macroalgae and/or spores of the microalgae or smaller macroalgae. The term "microalgae" refers to algae that are generally smaller than approximately 2 centimeters in height or length. Examples of such algae may be found in U.S. Patent No. 4,333,263, previously incorporated herein by reference. The term "smaller macroalgae" refers to algae that are smaller than approximately 20 centimeters in height or length. Examples of such algae include Gracilaria (a red algae), Sargassum, and Dictyota (brown algae). Benthic microalgae or a colony dominated by such algae are useful. In certain usage, however, a colony in which a significant percentage or even the majority of the algae are smaller macroalgae are used, particularly where long harvest times are desirable for operational reasons or a coarse diatom-supporting mesh work is desired because of pollution in the form of a high percentage of larger organic particulates.

Still further light-sensitive microorganisms for use in this aspect of the invention include Halobacterium, Oscillatoria or Aphanocapsa species (see Uma and Subramanian (1990) Proc. Natl. Svmp. Cyanobactt. Nitrog. Fix., IARI, New Delhi. Pages 437-444). Exemplary Halobacterium species include Halobacterium cutirubrum Halobacterium denitrificansflalobacterium distributum, Halobacterium halobium, Halobacterium lacusprofundi, Halobacterium mediterranei, Halobacterium noricense, Halobacterium pharaonis, Halobacterium saccharovorum, Halobacterium salinarium, Halobacterium sodomense, Halobacterium trapanicum, Halobacterium vallismortis and Halobacterium volcanii. Exemplary Oscillatoria species include Oscillatoria simplicissim. Exemplary Aphanocapsa species include Aphanocapsa delicatissima, Aphanocapsa elachista, Aphanocapsa endophytica, Aphanocapsa grevillei, Aphanocapsa pulchra, and Aphanocapsa rivularis.

Representative, non-limiting conditions for use of an algal surface biofilm in conjunction with the invention are described in U.S. Patent No. 5,851,398, as referenced above.

5. EXAMPLES

The present invention is illustrated by the following non-limiting examples. It is to be understood that the particular examples, materials, amounts, and procedures are to be interpreted broadly in accordance with the scope and spirit of the invention as set forth herein.

In the following examples, calcium ion and calcite mineral binding by microbial EPS is investigated by isothermal titration calorimetry (ITC). ITC is a thermodynamic technique that quantifies the heat absorbed or released during chemical reactions. The evolved heat flux is proportional to the amount of binding and can be used to calculate thermodynamic parameters, such as enthalpies of binding. Building on the work by Dimova et al. (Dimova et al. (2003). Langmuir 19(15): 6097- 6103), ITC was utilized to quantify calcite binding by EPS produced by natural microbial isolates. Two EPS-producing microorganizsms were collected from stone surfaces and phylogentically profiles. Additionally, a known calcareous-mineral precipitating microorganism, Proteus mirabilis, was investigated. EPS was produced in culture and chemically characterized. Calcite binding by the collected polymers was measured using ITC, and the data were compared to binding by humic acid. A Langmuir adsorption isotherm (Dimova et al. (2003). Langmuir 19(15): 6097- 6103) thermodynamic model of binding of surficial cations describes the EPS-mineral reaction. These procedures or other quantitative methods could also be expanded to algal polymers.

Example 1: Isothermal Titration Calorimetry

Calcite crystals (CaCO₃) are prepared by slow crystallization (Kitano et al. (1962) L Geophvs. Res. 67(12): 4873; Dimova et al. (2003). Langmuir 19(15): 6097- 6103). A super saturated solution of calcium carbonate is prepared by bubbling CO₂ gas through a suspension of 5 g CaCO₃ in 4 L of nanopure water for 60 min with constant stirring at room temperature. Undissolved CaCO₃ is removed by vacuum filtration through #4 (20-25 μm) Whatman filter paper (Middlesex, UK). Bubbling for another 30 min dissolved any remaining particles in the filtered solution. Crystals with well-defined rhombohedral morphology and surface area precipitated after the solution is left in an unsealed container and allowed to equilibrate for 48 h at room temperature (Rudloff et al. (20021. Macromol. Chem. Phvs. 203(4): 627-635). Crystal surface area is quantified by BET analysis, which measured gaseous pressure drop as nitrogen/helium/krypton mixtures are sorbed to the crystal surface. The surface area measurements are consistent with optical microscopy measurements of a large sample set. A perfect crystal without meso- or atomic-scale topographical irregularities is assumed in the optical microscopy calculations (Shiraki et al. (2000) Aquatic Geochem. 6(1): 87-108; Duckworth, et al (2003) Geochim. Cosmochim. Acta 67: 1787-1801).

Solutions of pure ionic calcium are prepared by filtration (0.1 μm, VC grade; Millipore, Billerica, MA) of the calcite suspensions (Dimova et al. (2003). Langmuir 19(15): 6097- 6103). The counter ion (carbonate) is likely still present in solution as controlled by atmospheric equilibrium. The absence of large crystals is confirmed by optical microscopy. Although a 0.1 μm pore size is used for filtration, meso-scale calcite aggregations may have still been present in solution (Dimova et al. (2003). Langmuir 19(15): 6097- 6103). Solution calcium concentration is measured using flame atomic absorption spectroscopy.

Interactions of EPS with aqueous cations (Ca²⁺ _(aq)) and particulars calcite (CaCO_{3 (s}j) interactions are investigated using a VP-ITC calorimeter (MicroCal, Northampton, MA). Solutions are made with nanopure water, previously degassed for 5 min under vacuum. Experiments are performed at a working volume of 1.428 mL, 30°C, and a stir rate of 280 rpm to obviate settling of the crystals. Titrations of the 1% EPS into deionized water, CaCO₃^ suspension, or Ca²⁺ _(aq) solutions are performed in 10 μL aliquots injected over 20 sec with 300 sec between injections. An equilibration time of 300 sec is necessary to return to baseline, presumably because of sluggish EPS binding kinetics. Analysis of the data is performed using MicroCal Origin 7.0.

Microorganisms and Polymer Characterization

EPS producing isolates are chosen from libraries of biofilm-forming microorganisms collected from two mineral surfaces based on their viability in alkaline conditions (10 < pH < 12) and their ability to produce EPS. The culture conditions are selected to favor the Bacillus genus because several members have been demonstrated to be alkaliphilic (Boyer et al. (1973) Internatl. J. System. Bacteriol. 50: 697-703) Internatl. J. System. Bacteriol. 50: 697-703; Kudo and Horikoshi 1983; Horikoshi, K. (1998). Extremophiles: Microbial Life in Extreme Environments. Horikoshi and Grant. New York, Wiley-Liss: 155-180). The ability of these organisms to grow in high pH environments may be a consequence of the protective strategies of Gram positive bacteria, such as the presence of neutralizing membrane-bound sodium pumps (Rrulwich et al. (2001) Biochim. Biophvs. Acta-Bioenerget. 1505(1): 158-168).

The mineral substrate is predominantly calcite. The Cyprus sample is 95% biomicritic calcium carbonate as determined by thin-section polarized light microscopy (Pers. comm.: M. Breuker, 2005, National Park Service.). The Mexico sample is 98% calcite (McNamara et al. (in press) J. Microbiol. Meth. ). An isolate that produced significant quantities of EPS in culture is designated G3 and chosen for future experimentation from a pool of over twenty alkaliphilic microorganisms collected from the sampled historic site in Cyprus. MEX244.1 is selected from a pool of over 200 epilithic biofilm bacteria collected from the Maya site of Ek' Balam in Mexico (McNamara et al. (in press) Microb. Ecol.). Amplified 16S rDNA gene sequences are aligned with those of other alkaliphilic Bacilli and microorganisms to identify the isolates. Figure 2 represents the phylogenetic relationships based on partial 16S rDNA sequence (1412 base pairs) of two isolates MEX 244.1 and G3. Tree is constructed in PAUP by neighbor-joining method using Jukes-Cantor corrections. Bootstrap values based on 1000 replicates each (for distance and parsimony) are shown for branches with >50% support. The sequences used for tree construction are submitted to GenBank for G3 (accession #AY987935) and MEX244.1 (accession #AY987936).

Although the two isolated alkaliphilic microorganisms are phylogenetically very similar, differing in two base positions; they produced EPS with different chemistries. The assembled sequences G3 and MEX244.1 most closely resembled those of B. cereus and B. thuringiensis. The relationships of these isolates to other cultured alkaliphilic Bacilli are shown in Figure 2. The EPS of the several isolates also had differing binding capacities for calcite.

EPS produced in culture is harvested from the isolates. The monosaccharide residues and linkages of the EPS samples are analyzed. The EPS samples are large macromolecules with several monosaccharide types (Table 2), and complex branching structures. EPS from G3 is dominantly a polymannose, while EPS from MEX244.1 and P. mirabilis contained larger amounts of other monosaccharides. The detection of glucosamine in the G3 EPS suggests that the polymer is a part of a glycoprotein. The EPS samples from the isolates had molecular weights of at least 167 kD; this value is used in calculations of molarity. GC-MS chromatograms of the polymers from G3 and MEX244.1 had only a single peak, which suggested the presence of a single purified polysaccharide and the absence of contaminating macromolecules. In contrast, the EPS from P, mirabilis appeared to have two components. The bacteria may also have produced other, smaller oligosaccharides. If this is the case, they are not collected by the purification procedure or they are present in insignificant quantities in comparison to the large EPS polymer.

Table 2. Glycosyl composition analysis

G3 EPS MEX 244.1 EPS P.mirabilis EPS

Glycosyl residue Mass Mole %* Mass Mole %* Mass Mole

(μg) (μg) (μg) Vo¹

Arabinose (Ara) 3.1 5.0 7.4 9.4 15.6 16.3

Rhamnose (Rha) Trace n.d. 2.8 3.2 3.4 3.3

Fucose (Fuc) n.d n.d n.d n.d n.d n.d

Xylose (XyI) 2.1 3.4 4.4 5.6 4.9 5.1

Glucuronic acid (GIcA) n.d n.d n.d n.d n.d n.d

Galacturonic acid (GaIA) Trace n.d 7.6 7.5 10.3 8.3

Mannose (Man) 51.8 70.3 49.2 52.2 35.5 30.6

Galactose (Gal) 9.0 12.1 15.9 16.9 26.5 23.0

Glucose (GIc) 3.5 4.7 4.9 5.2 15.4 13.4

N-acetyl glucosamine 4.0 4.5 n.d n.d n.d n.d

(GIcNAc)

Total carbohydrate 73.5 92.2 111.6

¹ Values are expressed as mole percent of total carbohydrate, n.d. - not detected

Example 2: Exopolymer Production and Characterization

EPS is produced by growing the isolates in a 15 L batch fermenter in nutrient broth with constant stirring and aeration for 96 h. Cells are removed from the culture by tangential filtration through a 0.22 [μm membrane filter (Durapore, Pellicon-2, Millipore). The EPS is concentrated 100 x using a 5 kD membrane filter (PLCCC, Pellicon-2, Millipore). Contaminating macromolecules, including DNA, RNA, and proteins, are removed by the method of Goncalves et al. ((2003) BioTechnol. App. Biochem. 37: 283-287). Contaminating salts are removed by centrifugal filtration (10 kD Macrosep filter; Pall, East Hills, NY) and repeated rinsing of EPS retentate with nanopure water. This protocol resulted in purified polysaccharides.

Glycosyl composition and linkage analysis is analyzed using gas chromatography/mass- spectrometry (GC-MS) (York et al. (1985). Colowick and Kaplan. New York, Academic Press. 118: 3-40) of partially methylated alditol acetates (Ciucanu, et al. (1984) Carb. Res. 131(2): 209-217). The EPS molecular weight is determined by size exclusion chromatography. A 1 mg sample of a 10 mg mL^" EPS solution is injected onto a Superose 12 column at a flow rate of 0.40 mL min^"1 in 50 mM ammonium formate at pH 4.8. Dextran standards of 10, 40, 67, and 167 kD are run in tandem with the sample.

Example 3: Isothermal Titration Calorimetrv with Harvested Exopolvsaccharides

A suspension of calcite crystals is grown in a supersaturated calcium solution for ITC analysis. The precipitated calcite is predominantly regular (1014 ) rhombohedral crystals. The surface area of the crystals is 0.39 m g as measured by BET analysis, which is equivalent to 32 x 10^" m L^" in the calcite suspension. A similar value is obtained from optical microscopy measurements, indicating that most of the surface area is in the form of large crystals. No evidence of vaterite precipitation is observed by optical microscopy. Filtration of these calcite suspensions resulted in solutions of aqueous calcium and carbonate equilibrium species. Crystal removal from solutions is confirmed by optical microscopy of multiple samples. This preparation avoided any pH or ionic strength differences between the solutions with and without calcite crystals, which otherwise could have affected the ITC measurements. Solutions with and without calcite crystals had similar concentrations of total calcium (0.97 ± 0.03 mM), indicating that the calcium content of the suspension is dominated by the aqueous species rather than by large calcite crystals.

ITC measurements are performed for EPS from two natural isolates (G3 and MEX244.1) as well as from Proteus mirabilis Hauser. Experiments are conducted in nanopure water, in a solution containing aqueous calcium cations (Ca²⁺ _(aq)), and in a solution containing aqueous calcium cations and precipitated calcite (Ca²⁺ _(aq)). The titrant contained 1% (w/w) EPS solutions. Heat fluxes accompanying the titrations are shown in Figures 3 to 5 (2.4). The several types of EPS had different heat-flux responses during addition to nanopure water. Figure 3 represents the heat flux titration for G3 EPS ( ), MEX244.1 EPS (Δ), P. mirabilis EPS (o), and humic acid (I). In these experiments, biomolecule is progressively added to nanopure water, so the heat flux arises from the hydration process.

EPS from MEX244.1 and P. mirabilis had very little heat-flux when titrated into water. EPS from G3 had a slightly exothermic character that stabilized near the baseline, indicating that this biomolecule is very hydrophilic and that energy is released with hydration. For comparison, titration with humic acid (Alfa Aesar, Ward Hill, MA) is also carried out. The heat flux of humic acid is initially endothermic, which is probably the result of its more hydrophobic nature that requires more energy to successfully disperse in the aqueous milieu. The differences between the observed curves are within the short-term noise range of the ITC (2 nJsec^"1).

The differences among the titration profiles may also have resulted from, in part, pH differences of the injectant solutions. However, due to the complexity of the acid-base chemistry and conformations of EPS, pH is not adjusted nor are buffers employed. Moreover, solution additives, which otherwise could have affected the EPS adsorption to calcite, are also avoided.

Example 4: Titration of EPS / Calcium Cations

In the following experiment, the EPS molecules are titrated into solutions containing aqueous calcium cations. EPS produced by MEX244.1 is initially slightly endothermic that quickly stabilized at the baseline (Figure 4). This behavior is interpreted as a minimal interaction of the EPS with the aqueous cations, which is caused by a combination of restructuring of the EPS in solution and a breaking of water bonds with EPS and dissolved calcium. Importantly, however, the absolute magnitude of the heat flux is small, which indicated that this polymer weakly associated with calcium ions.

In comparison to MEX244.1, EPS from P. mirabilis resulted in a mildly more endothermic event, indicating more interaction with the cations. Figures 4 represents heat flux titration for G3 EPS ( ), MEX244.1 EPS (Δ), P. mirabilis EPS (O), and humic acid (O). In these experiments, biomolecule is progressively added to nanopure water solutions containing 1 mM Ca²⁺ _(aq), so the heat flux arises from a combination of the H₂O- and Ca²⁺ _(aq) - binding by the various biomolecules. Induced conformational changes may also contribute to the heat flux.

The EPS from P. mirabilis stabilizes growing mineral crystals by binding cations (Clapham et al. (1990) J. Crvst. Growth 104: 475-484; Dumanski, et al. (1994) Meet, Immun. 62: 2998- 3003), which is consistent with observations of its ability to associate with calcium ions. The initial endothermic nature of this interaction appears counterintuitive when considering normal ligand-receptor energetics, which are often exothermic. The a priori assumption is that the Coulomb interactions between the positively charged Ca²⁺ _(aq) and negatively charged, electron-donating oxygen species on the EPS (such as hydroxyls and ethers) would be the driving force for these interactions (Chapter 8 and (Perry IV et al. (submitted). L Phys. Chem. - A)). The endothermic character of this reaction instead indicated a more important role for the liberation of water from the hydration shells of this hydrophilic EPS {i.e., exoergic but endothermic; cf. Figure 3) (Dimova et al. (2003). Langmuir 19(15): 6097- 6103).

EPS produced by G3 had a stronger endothermic character than the EPS of P. mirabilis, suggesting a much stronger interaction with the cations that stabilized as an exothermic event (Figure 4). The stabilization of the trend at negative values may be due to continuing rearrangements of the EPS- Ca²⁺ _(aq) folding and cross-linked structures, even though calcium binding had reached equilibrium.

Example 5: Titration of EPS / Calcite Crystals

In the next set of experiments, the EPS molecules are titrated into solutions containing suspended calcite crystals. The presence of calcite resulted in an endothermic shift relative to the heat fluxes in the absence of calcite. Direct observation of the heat flux arising from EPS-calcite interactions is complicated by the substantially greater heat flux resulting from hydration and binding of suspended calcite (Dimova et al. (2003). Langmuir 19(15): 6097- 6103). The heat flux arising from the EPS-calcite interaction can, however, be obtained by taking the difference of the heat flux into calcite suspensions from the heat flux in filtered solutions. The resulting heat flux is shown in Figure 5.

Figure 5 shows that the heat flux associated with surface binding by EPS from G3 became increasingly endothermic but then abruptly switched to increasingly exothermic during the reaction. The data profiles show an increasingly endothermic behavior for 0 < [EPSτ] < 4 - 6 μM followed by an increasingly exothermic behavior for [EPSτ] > 6 μM. These data suggest that, for [EPSτ] < 4 - 6 μM, EPS from G3 preferentially binds to aqueous cations first due to the greater effect of hydration shell disruption when binding aqueous cations compared to surficial cations. For [EPSτ] <4 -6 μM, the concentration of the reactive sites on the EPS approximates the concentration of aqueous cations. Once EPS has bound the aqueous calcium (e.g., 4 -6 μM), it begins to bind to the calcite surface. The inflection point in Figure 2 shows the crossover during the titrations. This behavior is similarly observed for the humic acid-calcite interaction, although the magnitude of the heat-flux resulting from the reaction diminishes. The ability of the EPS to scavenge cations appears to be a relatively fast process that binds the aqueous calcium faster than it is replaced by accelerated dissolution of the calcite crystal through lowering of aqueous saturation (Perry IV, et al. (20041 Env. Sci. Technol. 38: 3040-3046). The descending portion of the data for [EPSτ] > 4 μM is attributed to the binding of calcite reactive surface sites by EPS and eventual surface equilibrium. Li comparison to EPS from GS (D/B), EPS from P. mirabilis (O/Φ) and MEX244.1 (Δ/A) had weak interactions (i.e., small heat flux) with the calcite surface.

A calcium-binding mass balance is developed from information about the G3 polymer structure (see Table 3). It showed that, for [EPSτ] < 4 - 6 μM, the concentration of EPS reactive sites approximates the number of aqueous calcium ions. Chemical characterization of the polymer permits estimation of the maximum calcium binding capacity ((EPS • Ca²⁺)_max) per mole of polymer via:

where MW_EP_S is the molecular weight of the EPS, z_<j and 1₅ are the relative percentages and MWe and MW ₅ are the average molecular weights of the six- and five-membered monosaccharides detected, respectively, and q is the coordination of the binding reaction. Octadentate coordination of EPS monosaccharides around a single calcium ion is assumed because this coordination has been observed as a maximum binding capability for other natural polymers (Gregor et α/. (1996). Water Res. 30(6): 1319-1324). The calculated result is that EPS produced by G3 binds 220 moles of calcium per mole polymer. For [EPSτ] = 4 μM, the concentration of octadentate reactive sites on the G3 approximates the concentration of aqueous cations. This calculation supports the earlier statement of the role of aqueous calcite EPS binding in the ascending portion of the heat-flux data (Figure 5).

Example 6: Langmuir Binding Model

A two-parameter Langmuir model can be fit to the data to determine an adsorption constant (K_0Js) and specific enthalpy ( AH_sw-f) for the EPS-calcite interactions. A revised Langmuir type isotherm (Dimova et al. (2003). Langmuir 19(15): 6097- 6103), a commonly used descriptor of surface adsorption, is used to explain the adsorption of biomolecule to the calcite surface. The model assumes that there is a single type of reaction site on both the biomolecule and calcite. This simplification of the system, in which both reactive species are heterogeneous due to complex monosaccharide arrangements and branching structures of EPS and the complex surficial features on calcite, is nevertheless valuable for quantification of the

EPS-calcite interaction and adequately accounts for the empirical results. The Langmuir model is applied to later injections (after the inflection point) because chelation of aqueous cations affects the heat flux in the early injections of the titration. Model-fit lines are shown in Figure 5. The fitted values for the adsorption constants and enthalpy are given in Table 3 for EPS from GS, MEX244.1, and P. mirabilis, and humic acid.

Table 3. Fitted parameters for biomolecule reactions with calcite

Macromolecule Ka₁₁₅(M^'1) AHW (J m^"2) R²

G3 EPS 1.52 X lO⁵ 2.33 0.99

MEX 244.1 EPS 5.25 x lO⁴ 1.12 0.82

P. mirabilis EPS 5.0O x IO⁴ 1.33 0.95

Humic acid* 3.00 x l0⁵ 0.99 0.99

Note that molecular weight is not determined due to the heterogeneity of the sample. Example 7: Controlled Precipitation of Calcite by Bacterial Exopolymers

Calcite mineralization (dissolution and precipitation) impacts environmental, geological, and hydrogeological systems. Calcite is an important reservoir of carbon, and mineralization affects global carbon cycling (Schlesinger, W. H. (1997). Biogeochemistry: An Analysis of Global Change. San Diego, Academic Press.), the chemistry of marine systems (Pilson, M. (1998). An Introduction to the Chemistry of the Sea. Upper Saddle River, NJ, Prentice Hall.), the local pH and alkalinity of terrestrial environments (Stumm, W. et al. (1996). Aquatic Chem. New York, Wiley), hydrologic complexity in reactive transport modeling (Stumm, W. (1992). Chemistry of the Solid- Water Interface. New York, Wiley), the fate and transport of anthropogenic pollutants, especially heavy metals (Elzinga, et al. (2002). Geochim. Cosmochim. Acta 66: 3943- 3954), and deterioration of stone cultural heritage materials (S aiz- Jimenez, C. (1993). Atmos. Envir. 27B: 77-85).

Bacteria, Archaea, cyanobacteria, algae, fungi, and lichens affect calcite mineralization through production of metabolic by-products (Christensen et al. (1990). Physical and Chemical Properties in Biofilms. Biofilms. W. G. Charaklis and K. C. Marshall. New York, Wiley; Perry rV et al. (2005) J. Am. Chem. Soc. 127: 5744-5745). Many biologically derived compounds, affect mineralization processes, including lipids and phospholipids (Suess, E. (1970) Geochim. Cosmochim. Acta 34: 157-168.; Frye, et al. (1993) Chem. Geol. 109: 215-226), proteins (Suess, E. (1970) Geochim. Cosmochim. Acta 34: 157-168.), humics (Hoch et al. (2000) Geochim. Cosmochim. Acta 64: 61-72.), and other carboxylated compounds (Franzen et al. (1999) FEMS Microbiol. Letters 173: 395-402: Wu et al (2002) Langmuir 18: 6813-6820.), though the effect varies widely with chemical identity and aqueous conditions (Compton et al (1993) J. Coll. Inter. Sci. 158: 439-445: Thomas et al. (1993^s) Chem. Geol. 109: 227-237). Exopolysaccharides (EPS), which are a dominant exudates of bacteria growing on biofilms on surfaces, have been observed to bind to cations on mineral surfaces, both inhibiting and promoting dissolution (Thomas et al (1993) Chem. Geol. 109: 227-237; Welch, et al (1994) Geomicrobiol. J. 12: 227- 238; Barker et al (1997) J. F. Banfield and K. H. Nealson. Washington, D.C., Minearlog. Soc. Am. 35: 391-428.; Perry IV, et al (2004) Env. Sci. Technol. 38: 3040-3046). However, how these mechanisms and reactions precisely work is not well articulated (Welch, et al (1994) Geomicrobiol. J. 12: 227- 238; Banfield et al (1999). Proc. Natl. Acad. Sci. (USA) 96(7): 3404-3411 ; Hemming, et al (2001) Water Sci. Technol. 43(6): 9-16).

Many organisms mediate inorganic precipitation and crystallization by selective application of organic compounds to exert detailed control over the structure, orientation, growth kinetics, and nucleation of inorganic crystals (Berman et al. (1988) Nature 331(6156): 546-548; Mann et al (1993) ScL 261(5126): 1286-1292; Belcher et al. (1996) Nature 381(6577): 56- 58). Altered precipitated calcite morphology can be necessary for organism survival and may confer an ecological advantage, these materials are known as biogenic minerals. However, compounds produced by organisms can also affect calcareous mineral morphology simply as the result of chemical interactions between the forming minerals and the biomolecules. Microorganisms often mediate destructive and constructive processes on the same substrate due to the heterogeneity of biofilms, which are communities of microorganisms attached to the stone surface in an anchoring matrix of excreted EPS (Costerton et al. (1999) ScL 284(5418): 1318- 1322). Mature biofilms on mineral surfaces are temporally, spatially, and taxonomically dynamic communities that often contain local zones of concentration and pH variability caused by metabolism of the resident microorganisms. The importance of biofilms and microorganisms in the formation of calcareous materials is well documented. Bacteria also have been observed to precipitate calcareous (Cacchio et al (2004) Geomicrobiol. J. 21: 497-509) and other types of minerals (Banfield et al (2000) ScL 289(4 August): 751-754) by stabilization of nucleation events (Clapham et al. (1990) J. Cryst. Growth 104: 475-484), cation recruitment, or increasing the local alkalinity (Stickler, D. (1996) Biofouling 9(4): 293-305; Stocks-Fischer et al. (1999) Soil Biol. Biochem. 31: 1563-1571; Banfield et al. (2000) ScL 289(4 August): 751-754;

Canaveras et al. (2001) GeomicrobioL J. 18: 223-240; Contos et al. (2001) Geomicrobiol. J. 18: 331-343; Rodriguez-Navarro et al. (2003) APP. Environ. Microbiol. 69: 2182-2193; Cacchio et al. (2004) Geomicrobiol. J. 21: 497-509).

Exopolysaccharides (EPS) produced by environmental stone-colonizing biofilm bacteria can control calcite crystal morphology during precipitation. Additionally, altered crystal morphology is observed in the presence of EPS produced by Proteus mirabilis, a medically important bacterium often associated with pathogenic biomineralization (Dumanski, et al. (1994) Infect. Immun. 62: 2998- 3003). The effect of these bacteria is compared to other naturally occurring biomolecules, namely lipids, proteins, and polysaccharides. The exacting control of the EPS affects the current understanding of the role and specificity of bacteria in environmental mineralization reactions.

Bacterial isolates G3 and MEX244.1 are isolated from the surfaces of cultural heritage limestone surfaces. P. mirabilis (#51286) is obtained from the American Type Culture Collection (Manassas, VA). EPS from the isolates are collected and purifed, according to previously published methods (Perry IV et al. (submitted). Env. Sci. Technol.). Briefly, the EPS is produced in a batch fermenter and collected by tangential filtration. Contaminating molecules, including proteins, nucleic acids, and salts, are removed by sequential ethanol precipitations and salts are removed by repeated centrifugal rinsing with nanopure deionized water (18.3 MΩ cm; Barnstead, Debuque, IA). To bind 100 mg / L calcium cations (typical concentration of tap water) using G3 and MEX244.1 polymers, 0.35 g / L of exopolysaccharide have to be produced by these bacteria. Humic acid (41747), curdlan (C-7821), and starch (S-516) are all used as provided by their manufacturers (Alfa Aesar, Ward Hill, MA; Sigma- Aldrich, St. Louis, MO; Fisher Scientific, Fair Lawn, NJ, respectively). Cells are prepared by growth of a preculture in nutrient broth for 24 hrs., centrifugation of 1 mL of the preculture, and 3X rinsing by resuspension with nanopure water.

Crystallization is performed (Albeck et al. (1993) J. Am. Chem. Soc. 115(25): 11691- 11697). Briefly, solutions of 7.5 mM CaCl₂ and purified bacterial exopolysaccharides (and other model biomolecules) at 0.01% - 0.1% (w/w) are prepared and aliquoted into sample wells containing coverslips. In a sealed container, carbon dioxide (from ammonium carbonate) is allowed to slowly diffuse into the solutions. Coverslips with crystals are removed at different times (< 3 days), lightly rinsed with nanopure water and, after sputter coating with Pt/Pd for 90 sec, are visualized using a scanning electron microscope (LEO). Crystal faces are determined by measuring angles of intersecting faces on replicate samples (n < 10). Projections are created by commercially available software packages of the atomistic arrangements of the determined combinations of expressed faces (CrystalMaker, Oxfordshire, U.K.) and the external crystal forms (Shape, Kingsport, TN).

EPS produced by natural bacterial isolates collected from calcareous limestone surfaces are able to affect the morphology of precipitated calcite crystals. Controlled precipitation reactions in the absence of biomolecule additives resulted in the formation of regular calcite rhombohedra expressing the (10[bar]14) family of faces. Figure 6 represents calcite precipitation: in the absence of EPS (A), and in the presence of 0.01% (w/w) solutions of EPS produced by Proteus mirabilis (B), isolate G3 (C) and MEX244.1 (D). The organisms associated with (C) and (D) are identified as Bacillus spp. Precipitation in the presence of a variety of EPS produced by natural isolates results in formation of alternate crystal morphologies (Figures 6B, 6C, and 6D).

A naturally isolated Bacillus sp. (G3) resulted in expression of a combination of the (10[bar]14) and (10[bar]l) family of faces. Although the reaction conditions are below the high concentrations of calcium which typically result in pitted and defect-ridden rhombohedra (Dickinson et al. (2002) J. Crvst. Growth 244(3-4): 369-378), the presence of pitted surfaces on the (10[bar] 14) faces of the crystals precipitated in the presence of G3 EPS may indicate local areas of supersaturation caused by cation recruitment caused by the EPS. EPS produced by a Bacillus sp. isolate (MEX244.1) resulted in a crystallographically non-specific morphology where the edges appear to be destabilized (Figure ID), which is likely the result of Otswald ripening or a non-specific back dissolution reaction. EPS produced by P. mirabilis resulted in expression of a combination of the (10[bar]14) and the (10[ba]l) family of faces. P. mirabilis has been implicated as a causative agent in medical pathological calcification (Dumanski, et al. (1994) Infect. Immun. 62: 2998- 3003).

The differences in the observed crystal morphologies are likely the result of chemical recognition of the different polymer structures and growing mineral faces. The chemical structure of the EPS produced by the isolated microorganisms is different in terms of the monosaccharide residue and linkages present. The arrangements of the cation-chelating regions on the polymers are controlled by the polymer structure. Modification of the monosaccharide residues and linkages changes the shape and repeat of these regions. Additionally, different monosaccharides will present different electron-donating moieties in the regions for interaction with the cations.

Other molecules with biological origins commonly found in the environment are assayed for their ability to control crystal morphology during precipitation. Common environmentally occurring molecules of biological origin, including curdlan and lichenan, did not control precipitate morphology in a crystallographically specific manner. Figure 7 represents calcite precipitation in the presence of biological molecules, including 0.1% (w/w) solutions of curdlan (A; inset is higher magnification of the corner vertex), lichenan (B), and humic acid (C). The absence of substantially crystallographically altered morphologies in A and B is apparent. However, humic acid causes a dramatically altered crystal morphology. Curdlan, a commonly produced bacterial polysaccharide, and lichenan, a polysaccharide produced by lichens which are often associated with mineral surfaces (Saiz-Jimienez, C. (1999) Geomicrobiol. J. 16: 27-37), both appear to interact non-specifically with the crystal edges (Figures 7 A and 7B). However, humic acid dramatically affected the morphology of the precipitated crystal (Figure 7C). The crystallographic specificity of the EPS produced by P. mirabilis, G3, andMEX244.1 indicates that there may be a functional benefit to the controlled structures of these biomolecules.

Matching of the different primary, secondary, and tertiary structures of the EPS and the molecular structure of calcium and carbonates of the calcite crystal may affect binding to steps on different mineral faces. Several factors determine the magnitude of the EPS-calcite binding energy including: EPS glycosyl identity, branching structure, molecular weight, hydrophobicity, and electron-donating moiety presence (Perry IV, Estroff, et al. (in prep) Biogeochem.; Perry IV et al. (submitted). Env. Sci. Technol.); and calcite calcium spacing, step riser angle, and electrostatic interactions (De Yoreo et al (2004) ScL 306: 1301-1302.). Due to the heterogeneous nature of the EPS, complete structural characterization is difficult but there are significant differences in their glycosyl ratios and linkages (Perry IV et al. (submitted). Env. Sci. Technol.). All of the EPS are dominantly polymannose with the MEX244.1 and P. mirabilis polymers having increased ratios of arabinose, galactose, glucose and galacturonic acid components. The different chemistries of the polymers affect cation-binding and crystallographic recognition with the calcite. Binding to different microscopic steps by EPS is reflected in changes of the macroscopic crystal morphology (De Yoreo et al (2004) Sci. 306: 1301-1302.). These natural EPS molecules do not contain charged moieties, such as carboxylic acids, which is a difference between the chemical structure of these EPS molecules compared to other polymers demonstrated to interact with calcite (Albeck et al. (1996) Conn. Tiss. Res. 35: 365- 370 [419-424].). The charged moieties have previously been reported as the reactive centers of aqueous and surface cation chelation (Davis et al. (2003) Env. Sci. Technol. 37(2): 261-267). While these interactions are important in binding, recent work has shown that other electron-rich moieties, such as hydroxyls and ethers (Perry IV, Estroff, et al. (in prep) Biogeochem.), which are present on the EPS molecules, and the enthalpic effects of specific crystallographic matching (Perry IV et al. (submitted). Env. Sci. Technol.) may also play an important role in EPS interaction with the calcite surface.

The EPS in this study have different chemical compositions and, hence, different spatial arrangements that are controlling their ability to associate with steps on different calcite faces. It should be noted that the composition of EPS in these biofilms is genotypically, phenotypically, and environmentally regulated and the chemical structure varies by microorganism, growth stage, nutrient abundance, and other environmental stimuli (Christensen et al. (1990). Physical ^■ and Chemical Properties in Biofilms. Biofilms. W. G. Charaklis and K. C. Marshall. New York, Wiley). Hence, the crystallographic recognition events presented here should only be expected when using the reaction preparations and procedures. However, it also indicates that the incredible species diversity and abundance (Whitman et al. (1998) Proc. Natl. Acad. Sci. (TJSA) 95: 6578-6583) of environmental microorganisms and, hence, EPS chemical diversity may result in an unexpected diversity of morphology of carbonaceous precipitates induced by microorganisms .

The presence of different morphologies of calcite caused by bacteria in the environment affects the understanding of the role of microorganisms in calcite and mineral transformation processes. The morphology of calcite materials determines its physico-chemical properties, such as solubility and catalytic activity (Stupp et al. (1997) ScL 277(5330): 1242-1248). Crystallographic control of secondary precipitation reactions may affect the solubility and environmental persistence of the precipitates. Additionally, bacteria have been observed to align themselves with particular mineral morphological features such as surface defects (Lύttge et al. (2004) App. Env. Microbiol. 70(3): 1627-1632) and cleavage planes (Edwards et al. (2001) Chem. Geol. 180: 19-32). The specific recognition of bacterial EPS for different mineral faces may allow different bacterial species to attach to different mineralogical features (Edwards et al. (2001) Chem. Geol. 180: 19-32), thereby affecting spatial dispersion to different ecological niches (Henriksen et al. C2003) Am. Mineralog. 88(11-12): 2040-2044).

In addition to several other fabrication techniques (Taubert et al. (2003) J. Phvs. Chem. B 107: 2660- 2666.), several groups have attempted to replicate bio-inspired methods to control crystal morphology (Belcher et al. (1996) Nature 381(6577): 56-58; Orme et al. (20QΪ) Nature 411: 775-779; De Yoreo et al (2004) ScL 306: 1301-1302.). Modern technologies require ^■ innovative control methods for the fabrication of complex crystalline materials with precise localization of particles, nucleation density, size, and morphology, all of which affect material performance (Heuer, et al. (1992) ScL 255(5048): 1098-1105; Bunker et al. (1994) ScL 264(5155): 48-55; Mann et al (1996) Nature 382(6589): 313-318; Stupp et al. (1997) ScL 277(5330): 1242-1248). Specific crystallography of inorganic materials determines their physico-chemical properties, such as solubility, catalytic activity, and optical properties. Functional optimization of the material is possible through the use of additives, which can induce a specific morphology through inhibition and promotion of selected crystal faces. Additives selectively stabilize certain crystal phases through molecular recognition, which is mediated by electrostatic, geometric, and stereochemical interactions. Controlled fabrication of inorganic solids with microscopic regularity could result in materials with optimized mechanical, optical, electric, and catalytic performance. Polysaccharides may provide new routes for the controlled fabrication of advanced inorganic materials.

Extracellular polymers produced by microbes are able to affect calcite precipitation by specific crystallographic recognition. These findings suggest that microbes may use EPS to actively attach to certain crystallographic features. Additionally, the secondary precipitates induced by microbes may have altered chemical properties, such as solubility, than the bulk calcite structure, further complicating reactive transport modeling.

Example 8: Removal of Water-Borne Minerals by an Algal Polymer

In this study, alginic acid was selected as a model environmental polysaccharide to study the effects of a biologically produced polymer on calcite dissolution, due to its well- characterized chemistry, commercial availability, and is present in the environment. It is a straight-chain, hydrophilic, colloidal, polyuronic acid composed of guluronic (G) and mannuronic (M) acid residues configured in poly-G, poly-M, or alternating GM blocks, which is capable of chelating aqueous cations. Approximately 20-50% of polysaccharides produced in a wide sampling of marine and terrestrial bacteria were uronic acids. Alginic acid is a dominant environmental polymer produced by seaweed in marine environments, and by the bacterium Pseudomonas aeruginosa, which is a ubiquitous environmental bacterium. Furthermore, it has been demonstrated that alginic acid specifically interacts with crystallographic features of calcite and can increase the dissolution rate of other minerals.

The electron donating moieties of the alginic acid polymer, such as carboxyls and hydroxyls, chelate aqueous cations such as Ca⁺² (Figure 1 of Davis et al. (2003)). The proportion of mannuronic (M) and guluronic (G) sugar residues and their macromolecular conformation determine the physical properties and the affinity of the polymer for cation binding. The carboxyl functional groups of poly-G alginates have appropriate spacing and geometry for cation binding, and poly-G aglinates have a higher affinity for divalent cation binding than their poly-M counterparts. The buckeled model of poly-G conformation explains the greater binding. An egg-crate conformation of the guluronic regions of two polymer chains forms around calcium ions. Planar poly-M blocks also bind calcium, although the binding is less ordered than poly-G regions because they do not have the correct special and geometric arrangement for chelation sites.

An artificial seawater solution was created following the instructions of INSTANT OCEAN and by use of the same. Sufficient alginic acid was added to create a 1% solution and was immediately filtered using a 5kD tangential filter. Serial repeats on the treated souce solution demonstrated that alginic acid could be used to remove salt from water, thereby reducing conductivity and mineral hardness. Figure 8 shows the steady decrease in conductivity associated with this removal of water-borne minerals by serial passage of the artificial seawater over the alginic acid preparation.

Example 9: Treatment System for Microbial Removal of Water-Borne Minerals

The treatment system uses microorganisms to demineralize water. The core process of the cartridges are shown in Figure 9. Water flows into a chamber containing microorganisms (e.g., algae or bacteria). The chamber may have pre-filter to maintain the purity of the culture(s) contained within the chamber. There also may be preconditioning chemical treatment to control the growth of the microorganisms and/or the binding and precipitation of minerals. These treatments may include, but are not limited to: pH regulation, addition of nutrients and/or chemicals, removal of contaminants. The water provides nutrients and/or trace nutrients for growth of the microorgansisms. The microbes grow and produce exopolymer. Water-borne minerals attach to the produced exopolysaccharides. These polymers may be associated with the surface of the microbial cells, with surfaces within the chamber, or the polymers may be free- floating. Minerals may also attach to the surface of the microbial cells. As the bound mineral content of the polymers increases, polymer bridging across cations and cross-linking causes flocculation and/or precipitation of the polymer-salt/mineral/metal complex.

At the effluent point of the first chamber there is a filter with appropriate flow sizing for the requirements of the system and the ability to retain (hold back) microbials cells, while allowing transmission (flow through) of exopolymers, salts, and polymer-salt complexes. This filter may be an impact or tangential or other type of filter. The pore size of this filter is likely around 0.2μm. The second chamber contains only polymers and salt.

At the effluent of the second chamber is another filter that is or proper pore sizing and performance to retain (hold back) the polymer-salt complexes and allow flow of water. This water contains fewer minerals than the inlet water. In this embodiment, the microbes and the polymer-salt complexes are collected in separate collections streams, e.g. from chamber one and chamber two, respectively. This allows for separate treatment of each waste stream. There may be beneficial use of this microbial waste for of animal feed, human consumption, fuel pellets, and/or ethanol/biofuel source and the polymer-salt complex for food additives, and/or an alternative salt source.

In another example, there is only a single chamber, in which all of the previously describes reactions occur, and the polymer-salt complex is collected simultaneously to the collection of the microorganisms. There may be beneficial use of this microbial/polymer/salt complex as well, in the use of the collected polymer salt complex as animal feed, for human consumption, in food additives, as alternative salt source, as fuel pellets, and/or as anethanol/biofuel source.

In another example, all of these processes are conducted in a series of large in- or above- ground holding tanks without the use of cartridges.

The design of a cartridge for use in the system is describe below. System Cartridge Design

Each cartridge is designed to maximize the aforementioned process. An exemplary design for a system cartridge is shown in Figure 10. The cartridges are available in a variety of sizes for ease of scaling to given systems. They may be offered as small as 1 gallon. The cartridge may contain inlet and outlets to contain internal conditions such as, but not limited to, nutrients, pH, and dissolved oxygen. The cartridges may include flow regulators to control addition of chemicals and/or nutrients to the cartridge, and thus that may be custom tuned for removal of different dissolved solids.

The cartridge may also contain substratum for the microorganisms to attach to and grow on. The exterior of the cartridge may also be transparent to sunlight for the cultivation of photosynthetic microorganisms. The cartridge may also contain materials and/or surface to maximize diffusion and microbial exposure to sunlight. The cartridge may also contain flow- regulation devices to pulse or vary flow conditions to ensure proper mixing. The cartridge may also contain technologies to minimize filter membrane fouling, such as vibration sources. The cartridge may also contain ultraviolet light sources for sterilization of effluents or to control surface growth on membranes or other places where it is not desired. The device may also contain conductivity meter(s) for monitoring water mineral/metal content and cartridge performance. The device may contain other types of detectors for monitoring internal and external conditions; these detectors may include, but are not limited to: temperature, pressure, oxygen, metals, and/or redox meters. The cartridge may also contain standardized wireless transmission devices, such as ZigBee type, for transmission of cartridge parameters. The cartridges may also contain batteries to power the included electronics. The cartridges may also contain devices to insure exclusive use with skids from the same supplier; these technologies may include, but are not limited to: RFID and proprietary connectors. The cartridges are designed for ease of transport, and ease of connection to the skids described below.

The cartridge may contain an outlet for release of the microbial-polymer-mineral complex or multiple outlets for the microorganisms and the polymer-mineral complex. These outlets may drain into collection tanks or be pumped away for additional use. Skid Design

The system is assembled on a single skid consisting of a series of cartridges. An exemplary design for a skid is shown in Figure 11.The skids are modular and can be combined in series and/or in parallel. The skid may be oriented to hold the cartridges in vertical or horizontal arrangements. The specific configuration of the cartridges in the system will be determined by the end performance specifications, including impurities present, inlet and outlet concentrations, and flow rate required.

The system may also include pre-treat and post-treat systems as needed based on the feed water quality. The skid may contain flow-reduction and/or energy capture devices to control the inlet flow. The skid may contain associated devices including, but not limited to, an integrated pre-treatment system, pump, and control system (alternatively, the system could be integrated with OEM RO systems). The skid may contain an electronic control and communication system for monitoring performance, maximizing performance, and user-interfacing. The skid may contain the proper physical and electronic systems for interfacing with OEM systems.

The skid may contain rigid or tubing connections for interfacing with the cartridges, or the additive, or waste-collection systems. The waste-collection system may be incorporated on the skid or an additional modular system.

Example 10: Treatment System as Reverse Osmosis Add-On Component

Reverse osmosis (RO) is a water purification technique invented in the 1970s that has become one of the best water filtration systems known (GE Water, "What Is Reverse Osmosis," GE Water Company Website, http://www.gewater.com/library/tp/833_What_Is.jsp, accessed March 2006). It removes particles from solution by the application of a large pressure gradient across a filter. Mechanical pressure, usually supplied by a pump, forces the water from an area of high solute concentration across a membrane to an area of low concentration.

The membranes used in RO consist of a dense polymer layer of microscopic thickness that allows only water to pass through. The water passes through the polymer membrane by diffusion, and ions and other solutes are left behind. RO is capable of separating bacteria, salts, sugars, proteins, dyes, and other particles that have a molecular weight of greater than 150-250 Daltons (GE Water, "What Is Reverse Osmosis," GE Water Company Website, http://www.gewater.com/library/tp/833_What_Is.jsp, accessed March 2006). For the thermoelectric power generation market specifically, the quality of the water used to produce the steam that will turn a turbine to produce electricity is critical. Impurities can cause problems such as scaling that reduce the amount of electricity that can be produced. This costs the power plants (and their customers) money and increases the amount of fuel that must be consumed to provide the necessary amount of electricity. In the extreme cases, impurities in the process water lead to damage and downtime that prevent a plant from producing electricity at all. RO pretreats water that goes to the boiler, which boils the water (from fossil fuel combustion), that moves the steam turbines. The water is then cooled via cooling towers and can be cycled back to the boiler.

Improvements in RO efficiency reach a point of diminishing return at high concentrations of solute because of process limitations. The add-on component to current reverse osmosis (RO) systems is installed on the effluent reject water stream; it is advantageous for the customer by decontaminating the effluent reject water ordinarily sent to receiving waters or zero level discharge systems. This decontaminated water can then be recycled back into the system, supplementing the original water stream and thus reducing the volume of water purchased from the original municipal water source as well as the volume of contaminated water that is discharged (See Figure 12).

While reverse osmosis (RO) can be a relatively economical water purification technology, a significant capital cost can be disposal charges associated with discarding the effluent reject water discharge, including paying sewage fees. By reducing the volume and contamination level of this stream, the water system provides further value to the customer in reduced sewer charges. In addition, the water system reduces the level of local pollution by decreasing the amount of contaminated water that is discharged into the environment.

Disposal fees can be quite significant due to the amount of feed water discharged. This reject stream can vary from 40 to 70 percent of the feed water flow for a typical RO process, depending on the salt content of the feed water, pressure, and type of membrane used. Because of these high percentages, the reject stream is almost always a significant volume of water, and the cost of disposal of this reject water is an important part of the feasibility of any RO operation (ABC's of Desalting by O.K. Buros; International Desalination Association).

The reject- water concentrate stream can be beneficially used for irrigation of salt tolerant plants and aquaculture, for dust suppression roadbed stabilization and soil remediation, and for injection into oil well fields. Evaporation ponds allow for collection of solid salts from the concentrate stream and disappearance of the water as water vapor. Often, the salt from these ponds is harvested and transferred directly to a landfill, merely relocating the point-source of saline pollution. These options face significant issues including land use and a lack of economies of scale. Regardless, there is demand for alternative systems because of the unsustainablity of conventional disposal methods.

These systems are often also included with zero level discharge (ZLD) systems. ZLD means that no brackish water concentrate leaves the plant boundaries. Currently, available systems include thermal brine concentrators, spray dryers, high-recovery RO, evaporation ponds, crystallizers, and enhanced evaporation systems. The described device is also suitable for use with these systems.

Equivalents

Those skilled in the art will recognize, or be able to ascertain, using no more than routine experimentation, numerous equivalents to the specific embodiments described specifically herein. Such equivalents are intended to be encompassed in the scope of the following claims.

Claims

CLAIMS What is claimed is:

1. A method of controlling water hardness and scale formation in a water system, comprising: contacting the water system with one or more exopolymer-producing microbes under conditions that allow for the production of the microbial exopolymer, the microbes producing exopolymer and controlling water hardness and scale formation.

2. The method of claim 1, wherein the microbe is an exopolymer-producing bacteria.

3. The method of claim 1 , wherein the microbe is a proteobacteria.

4. The method of claim 3, wherein the proteobacteria is selected from the group consisting of purple bacteria, chemoautotrophic proteobacteria, and chemoheterotrophic proteobacteria.

5. The method of claim 1 , wherein the microbe is a fungus.

6. The method of claim 1, wherein the microbe is a lichen.

7. The method of claim 1 , wherein the microbe is an alga.

8. The method of claim 1, wherein the microbe is a mixture of different types of microorganisms .

9. The method of claim 1, wherein the microbe is an alga selected from the group consisting of Volvox aureus, Volvox carteri, Volvox globactor, Volvox dissipatrix, Volvox tertios, Compsopogon coeruleus, Cladophora crispata, Spirogyra rivularis, Enteromorpha micrococca,

10. The method of claim 1, wherein the microbe is an. Amphora species or a Cocconeis species. Eunotia pectinalis, Melosira varian, and Stigeoclonium tenue.

11. The method of claim 1 , wherein the microbe is a planktonic alga.

12. The method of claim 1, wherein the microbe is an algal surface biofilm-forming alga.

13. The method of claim 12, wherein the algal surface biofilm-forming algae is a Cyanophycota.

14. The method of claim 12, wherein the Cyanophycota is selected from the group consisting of Oscillatoria, Lyngbya, Schizothrix, and Chroococcus Calothrix.

15. The method of claim 12, wherein the algal surface biofilm-forming algae is a green algae.

16. The method of claim 12, wherein the algal surface biofilm-forming algae is a Chlorophycota.

17. The method of claim 16, wherein the Chlorophycota is selected from the group consisting of Ulothrix, Enteromorpha, Spirogyra, Cladophora, Dichotomosiphon, Stigeoclonium, Oedogonium, Mougeotia, and Gloeocystis.

18. The method of claim 12, wherein the algal surface biofilm-forming algae is a Chromophycota.

19. The method of claim 18, wherein the Chromophycota is selected from the group consisting of Melosira, Ctenophora, Asterionella, Eunotia, Amphipleura, Cocconeis, Placoneis, Rhoikoneis, and Bacillaria.

20. The method of claim 12, wherein the algal surface biofilm-forming algae is a Rhodophycota.

21. The method of claim 20, wherein the Rhodophycota is Compsopogon.

22. The method of claim 12, wherein the algal surface biofilm-forming algae is selected from the group consisting of Polysiphonia, Herposiphonia, and Callithamnion.

23. The method of claim 1, wherein the exopolymer-producing microbe is a salt tolerant organism.

24. The method of claim 1, wherein the salt tolerant organism grows in hard or very hard water.

25. The method of claim 24, wherein the very hard water comprises a calcium carbonate concentration of greater than about 180 ppm.

26. The method of claim 1, wherein the exopolymer-producing microbe is a Halobacterium.

27. The method of claim 26, wherein the Halobacterium is selected from the group consisting of Halobacterium cutirubrum, Halobacterium denitrificans, Halobacterium distributum, Halobacterium halobium, Halobacterium lacusprofundi, Halobacterium mediterranei, Halobacterium noricense, Halobacterium pharaonis, Halobacterium saccharovorum, Halobacterium salinarium, Halobacterium sodomense, Halobacterium trapanicum, Halobacterium vallismortis and Halobacterium volcanii.

28. The method of claim 1, the exopolymer-producing microbe is an Oscillatoria.

29. The method of claim 28, wherein the Oscillatoria is Oscillatoria simplicissim.

30. The method of claim 1, the exopolymer-producing microbe is an Aphanocapsa

31. The method of claim 30, wherein the Aphanocapsa is selected from the group consisting of Aphanocapsa elachista, Aphanocapsa delicatissima, Aphanocapsa endophytica, Aphanocapsa grevillei, Aphanocapsa pulchra, and Aphanocapsa ήvularis.

32. A method of controlling water hardness and scale formation in a water system, comprising: contacting the water system with one or more exopolymer-producing bacteria under conditions that allow for the production of the bacterial exopolymer, the bacteria producing exopolymer and controlling scale formation in the water system.

33. The method of claim 32, wherein the exopolymer-producing bacteria is a Proteus species.

34. The method of claim 33, wherein the Proteus species is Proteus mirabilis.

35. The method of claim 32, wherein the Proteus mirabilis is the strain deposited as ATCC #51286.

36. The method of claim 34, wherein the exopolymer-producing bacteria is a Bacillus.

37. The method of claim 36, wherein the Bacillus is selected from the group consisting of Bacillus cerueus and Bacillus thuringiensis.

38. The method of claim 36, wherein the Bacillus is a strain selected from the group consisting ofG3 and MEX244.1.

39. The method of claim 36, wherein the Bacillus is selected from the group consisting of Bacillus gibsonii, Bacillus pseudalcaliphilus, Bacillus pseudofirmus, Bacillus halodurans, Bacillus subtilis, Bacillus benzoevorans, Bacillus simplex, Bacillus horikoshii, Bacillus cereus, and Bacillus thuringiensis.

40. The method of claim 32, wherein the exopolymer-producing bacteria is a Pseudomonas.

41. The method of claim 40, wherein the Pseudomonas is selected from the group consisting of Pseudomonas putida and Pseudomonas aeruginosa.

42. The method of claim 32, wherein the exopolymer-producing bacteria is an Azotobacter.

43. The method of claim 42, wherein the Azotobacter is selected from the group consisting of Azotobacter vinelandii, Azotobacter chroococcum and Azotobacter indicus.

44. The method of claim 1, wherein the exopolymer produced controls water hardness and scale formation by sequestration of ions.

45. The method of claim 44, wherein the ions are earth metals or minerals.

46. The method of claim 45, wherein the ions include earth metal or mineral ions that are normally occurring in earth surface waters selected from the group consisting of sodium, calcium and magnesium.

47. The method of claim 44, wherein the ions include earth metal or mineral ions from anthropogenic sources.

48. The method of claim 44, wherein the ions include heavy metal ions.

49. The method of claim 44, wherein the earth metal or mineral ion is arsenic.

50. The method of claim 1, wherein the exopolymer-producing microbes produce exopolymer and controls water hardness and the formation of scale in the water system by sequestration of cations and the flocculation and/or precipitation of the exopolymer-containing complex.

51. The method of claim 50, wherein the precipitation of the exopolymer-containing complex further results in coprecipitation of associated ions in the water.

52. The method of claim 51, wherein the associated ions in the water include ions selected from the group consisting of chloride, hydroxide, carbonate, bicarbonate, sulfate, and nitrate.

53. The method of claim 51, wherein the associated ions in the water include heavy metal ions selected from the group consisting of chromium, arsenic, and selenium.

54. The method of claim 1, wherein the exopolymer produced controls scale formation by promoting dissolution of existing scale.

55. The method of claim 1, wherein the exopolymer produced controls scale formation by increasing the solubility of newly formed scale.

56. The method of claim 1, wherein the exopolymer produced controls scale formation by inhibiting scale formation on a surface of a water system.

57. The method of claim 1, wherein the exopolymer-producing microbe further produces simple acids that dissolve scale.

58. The method of claim 32, wherein the water system comprises a pipe.

59. The method of claim 32, wherein the water system comprises a heat-transfer system.

60. The method of claim 32, wherein the water system comprises a boiler.

61. The method of claim 32, wherein the water system is a waste-water treatment facility.

62. The method of claim 1, wherein the water system is a power-generation facility.

63. The method of claim 1, wherein the water system is selected from the group consisting of a pulp / paper processing plant, a petrochemical refinery, and a metal refinery.

64. The method of claim 1, wherein the water system is a heat exchanger.

65. The method of claim 64, wherein the heat exchanger is selected from the group consisting of a single pass heat exchanger, a multi-pass heat exchanger, a regenerative heat exchanger, a non- regenerative heat exchanger, a tube heat exchanger, a shell heat exchanger, a plate heat exchanger, a parallel-flow heat exchanger, a cross-flow heat exchanger and a counter-flow heat exchanger.

66. The method of claim 1, wherein the water system comprises one or more chambers in series or parallel, and each of the one or more chambers contains one or more exopolymer-producing microbial populations.

67. The method of claim 66, wherein the microbial populations are retained within the one or more chambers by a high-volume filter.

68. The method of claim 66, wherein the microbial populations are retained within the one or more chambers by a tangential filter.

69. The method of claim 67, wherein one or more of the high-volume filters retain the microbial populations but allow microbial exopolymers and simple acids to pass through the chamber.

70. The method of claim 66, wherein one or more chambers are bounded by high- volume filters that retain the microbial exopolymer.

71. The method of claim 67, further comprising an outlet system for removing cation-saturated microbial exopolymer.

72. The method of claim 1, further comprising supplying the population of one or more exopolymer-producing microbes with one or more nutrients.

73. The method of claim 72, wherein the one or more nutrients include an organic carbon source and a nitrogen source.

74. The method of claim 73, wherein the one or more nutrients is selected from the group consisting of a carbon source, a nitrogen source, a phosphorous source and micronutrients.

75. The method of claim 1, wherein the water system includes an apparatus that is transparent to sunlight to provide energy for growth of photosynthetic organisms contained within the apparatus.

76. A method of controlling water hardness and scale formation in a water system, comprising: contacting the water system with a microbial exopolymer, the exopolymer controlling water hardness and scale formation.

77. The method of claim 76, wherein the microbial exopolymer provided is at least about 50% pure.

78. The method of claim 76, wherein the microbial exopolymer provided is at least about 75% pure.

79. The method of claim 76, wherein the microbial exopolymer provided is at least about 90% pure.

80. An isolated, cation-chelating algal exopolymer having a molecular weight of greater than about 20,000 Daltons.

81. The isolated, cation-chelating algal exopolymer of claim 80, wherein the molecular weight is greater than about 40,000 Daltons.

82. The isolated, cation-chelating algal exopolymer of claim 81, wherein the molecular weight is greater than about 60,000 Daltons.

83. The isolated, cation-chelating algal exopolymer of claim 82, wherein the molecular weight is greater than about 100,000 Daltons.

84. An isolated, cation-chelating bacterial exopolymer having a molecular weight of greater than about 167,000 Daltons.

85. The isolated, cation-chelating bacterial exopolymer of claim 84, wherein the composition is at least about 50% pure.

86. The isolated, cation-chelating bacterial exopolymer of claim 84, wherein the composition is at least about 75% pure.

87. The isolated, cation-chelating bacterial exopolymer of claim 84, wherein the composition is at least about 90% pure.

88. The isolated, cation-chelating bacterial exopolymer of claim 84, wherein the composition is at least about 95% pure.

89. The isolated, cation-chelating bacterial exopolymer of claim 84, wherein the composition is at least about 99% pure.

90. The isolated, cation-chelating bacterial exopolymer of claim 84, wherein the bacterial cation-chelating exopolymer has a mole percent glycosyl composition of at least about 3% xylose, 5% arabinose, 10% galactose, and 30% mannose.

91. The isolated, cation-chelating bacterial exopolymer of claim 84, wherein the exopolymer comprises no carboxylate-containing glycosyl residues and effects cation chelation by conformationally and configurationally positioned arrays of electron pair donating groups.

92. The isolated, bacterial cation-chelating exopolymer of claim 84, wherein the bacterial cation-chelating exopolymer has a calcium-binding capacity of about one cation per 8 glycosyl residues.

93. An isolated, cation-chelating, microbial exopolymer produced by the process comprising: providing one or more exopolymer-producing microbes with light and/or nutrients sufficient to cause the microbes to produce the cation-chelating, microbial exopolymer; and isolating the cation-chelating, microbial exopolymer so produced.

94. The isolated, cation-chelating, microbial exopolymer of claim 93, wherein the microbe is an exopolymer-producing bacterium.

95. The isolated, cation-chelating, microbial exopolymer of claim 93, wherein the microbe is a proteobacterium.

96. The isolated, cation-chelating, microbial exopolymer of claim 93, wherein the microbe is an alga selected from the group consisting of Volvox aureus, Volvox carteri, Volvox globactor, Volvox dissipatrix, Volvox tertios, Compsopogon coeruleus, Cladophora crispata, Spirogyra rivularis, Enteromorpha micrococca, Eunotia pectinalis, Melosira varian, and Stigeoclonium tenue.

91. The isolated, cation-chelating, microbial exopolymer of claim 93, wherein the microbe is from an Amphora species or a Cocconeis species.

98. The isolated, cation-chelating, microbial exopolymer of claim 93, wherein the microbe is an algal surface biofilm-forming alga.

99. The isolated, cation-chelating, microbial exopolymer of claim 98, wherein the algal surface biofilm-forming alga is a Cyanophycota.

100. The isolated, cation-chelating, microbial exopolymer of claim 99, wherein the Cyanophycota is selected from the group consisting of Oscillatoria, Lyngbya, Schizothrix, and Chroococcus Calothrix.

101. The isolated, cation-chelating, microbial exopolymer of claim 98, wherein the algal surface biofilm-forming alga is a Chlorophycota.

102. The isolated, cation-chelating, microbial exopolymer of claim 101, wherein the Chlorophycota is selected from the group consisting of Ulothrix, Enteromorpha, Spirogyra, Cladophora, Dichotomosiphon, Stigeoclonium, Oedogonium, Mougeotia, and Gloeocystis.

103. The isolated, cation-chelating, microbial exopolymer of claim 98, wherein the algal surface biofilm-forming alga is a Chromophycota.

104. The isolated, cation-chelating, microbial exopolymer of claim 103, wherein the Chromophycota is selected from the group consisting of Melosira, Ctenophora, Asterionella, Eunotia, Amphipleura, Cocconeis, Placoneis, Rhoikoneis, and Bacillaria.

105. The isolated, cation-chelating, microbial exopolymer of claim 98, wherein the algal surface biofilm-forming alga is a Rhodophycota.

106. The isolated, cation-chelating, microbial exopolymer of claim 105, wherein the Rhodophycota is Compsopogon.

107. The isolated, cation-chelating, microbial exopolynier of claim 98, wherein the algal surface biofilm-forming alga is selected from the group consisting of Polysiphonia, Herposiphonia, and Callithamnion.

108. The isolated, cation-chelating, microbial exopolymer of claim 93, the exopolymer- producing microbe is a Halobacterium.

109. The isolated, cation-chelating, microbial exopolymer of claim 108, wherein the Halobacterium is selected from the group consisting of Halobacterium cutirubrum, Halobacterium denitrificans, Halobacterium distributum, Halobacterium halobium, Halobacterium lacusprofundi, Halobacterium mediterranei, Halobacterium noricense, Halobacterium pharaonis, Halobacterium saccharovorum, Halobacterium salinarium, Halobacterium sodomense, Halobacterium trapanicum, Halobacterium vallismortis and Halobacterium volcanii.

110. The isolated, cation-chelating, microbial exopolymer of claim 93, wherein the exopolymer- producing microbe is an Oscillatoria.

111. The isolated, cation-chelating, microbial exopolymer of claim 110, wherein the Oscillatoria is Oscillatoria simplicissim.

112. The isolated, cation-chelating, microbial exopolymer of claim 93, wherein the exopolymer- producing microbe is an Aphanocapsa.

113. The isolated, cation-chelating, microbial exopolymer of claim 112, wherein the Aphanocapsa is selected from the group consisting of Aphanocapsa elachista, Aphanocapsa delicatissima, Aphanocapsa endophytica, Aphanocapsa grevillei, Aphanocapsa pulchra, and Aphanocapsa rixmlaris.

114. The isolated, cation-chelating, microbial exopolymer of claim 95, wherein the proteobacterium is selected from the group consisting of purple bacterim, chemoautotrophic proteobacterium, and chemoheterotrophic proteobacterium.

115. The isolated, cation-chelating, microbial exopolymer of claim 93, wherein the microbe is a fungus.

116. The isolated, cation-chelating, microbial exopolymer of claim 93, wherein the microbe is a lichen.

117. An isolated, cation-chelating, bacterial exopolymer produced by the process comprising:

providing one or more exopolymer-producing bacteria with nutrients sufficient to cause the bacteria to produce the cation-chelating, bacterial exopolymer; and

isolating the cation-chelating, bacterial exopolymer so produced.

118. The isolated, cation-chelating, bacterial exopolymer of claim 117, wherein the exopolymer- producing bacteria is Proteus mirabilis.

119. The isolated, cation-chelating, bacterial exopolymer of claim 118, wherein the Proteus mirabilis is the strain deposited as ATCC #51286.

120. The isolated, cation-chelating, bacterial exopolymer of claim 117, wherein the exopolymer- producing bacteria is a Bacillus.

121. The isolated, cation-chelating, bacterial exopolymer of claim 120, wherein the Bacillus is selected from the group consisting of Bacillus cerueus and Bacillus thuringiensis.

122. The isolated, cation-chelating, bacterial exopolymer of claim 120, wherein the Bacillus is a strain selected from the group consisting of G3 and MEX244.1.

123. The isolated, cation-chelating, bacterial exopolymer of claim 120, wherein the Bacillus is selected from the group consisting of Bacillus gibsonii, Bacillus pseudalcaliphilus, Bacillus pseudofirmus, Bacillus halodurans, Bacillus subtilis, Bacillus benzoevorans, Bacillus simplex, Bacillus horikoshii, Bacillus cereus, and Bacillus thuringiensis.

124. The isolated, cation-chelating, bacterial exopolymer of claim 117, wherein the exopolymer- producing bacteria is a Pseudomonas.

125. The isolated, cation-chelating, bacterial exopolymer of claim 124, wherein the Pseudomonas is selected from the group consisting of Pseudomonas putida and Pseudomonas aeruginosa.

126. The isolated, cation-chelating, bacterial exopolymer of claim 117, wherein the exopolymer- producing bacteria is an Azotobacter.

127. The isolated, cation-chelating, bacterial exopolymer of claim 126, wherein the Azotobacter is selected from the group consisting of Azotobacter vinelandii, Azotobacter chroococcum and Azotobacter indicus.

128. The isolated, cation-chelating, bacterial exopolymer of claim 117 having a molecular weight of greater than about 167,000 Daltons.

129. The isolated, cation-chelating, bacterial exopolymer of claim 117 having a purity of at least about 50% pure.

130. The isolated, cation-chelating, bacterial exopolymer of claim 117 having a purity of at least about 75% pure.

131. The isolated, cation-chelating, bacterial exopolymer of claim 117 having a purity of at least about 90% pure.

132. The isolated, cation-chelating, bacterial exopolymer of claim 117 having a purity of at least about 95% pure.

133. The isolated, cation-chelating, bacterial exopolymer of claim 117 having a mole percent glycosyl composition of at least about 3% xylose, 5% arabinose, 10% galactose, and 30% mannose.

134. The isolated, cation-chelating, bacterial exopolymer of claim 117 comprising no carboxylate-containing glycosyl residues and effecting cation chelation by conformationally and configurationally positioned arrays of electron pair donating groups.

135. The isolated, cation-chelating, bacterial exopolymer of claim 117 having a calcium-binding capacity of about one cation per 8 glycosyl residues.

136. A method of producing an isolated, cation-chelating, microbial exopolymer comprising: providing one or more exopolymer-producing microbes with nutrients sufficient to cause the microbes to produce the cation-chelating, microbial exopolymer; and

isolating the cation-chelating, microbial exopolymer so produced.

137. The method of claim 136, wherein the microbe is an exopolymer-producing bacterium.

138. The method of claim 136, wherein the microbe is a proteobacterium.

139. The method of claim 138, wherein the proteobacterium is selected from the group consisting of a purple bacterium, a chemoautotrophic proteobacterium, and a chemoheterotrophic proteobacterium.

140. The method of claim 136, wherein the microbe is a fungus.

141. The method of claim 136, wherein the microbe is a lichen.

142. The method of claim 136, wherein the microbe is an alga.

143. A method of producing an isolated, cation-chelating, bacterial exopolymer comprising:

providing one or more exopolymer-producing bacteria with nutrients sufficient to cause the bacteria to produce the cation-chelating bacterial exopolymer; and

isolating the cation-chelating bacterial exopolymer so produced.

144. The method of claim 143, wherein the exopolymer-producing bacteria is Proteus mirabilis.

145. The method of claim 144, wherein the Proteus mirabilis is the strain deposited as ATCC #51286.

146. The method of claim 143, wherein the exopolymer-producing bacteria is Bacillus.

147. The method of claim 146, wherein the Bacillus is selected from the group consisting of Bacillus cerueus and Bacillus thuringiensis.

148. The method of claim 146, wherein the Bacillus is a strain selected from the group consisting of G3 and MEX244.1.

149. The method of claim 146, wherein the Bacillus is selected from the group consisting of Bacillus gibsonii, Bacillus pseudalcaliphilus, Bacillus pseudofirmus, Bacillus halodurans, Bacillus subtilis, Bacillus benzoevorans, Bacillus simplex, Bacillus horikoshii, Bacillus cereus, and Bacillus thuringiensis.

150. The method of claim 143, wherein the exopolymer-producing bacteria is Pseudomonas.

151. The method of claim 150, wherein the Pseudomonas is selected from the group consisting of Pseudomonas putida and Pseudomonas aeruginosa.

152. The method of claim 143, wherein the exopolymer-producing bacteria is Azotobacter.

153. The method of claim 152, wherein the Azo tobacter is selected from the group consisting of Azotobacter vinelandii, Azotobacter chroococcum and Azotobacter indicus.

154. An apparatus for processing water to control water hardness and scale-formation, comprising:

at least one chamber having at least one input opening for receiving the water from a water source, and at least one output opening for discharging treated water from the chamber;

a source of a cation-chelating, exopolymer-producing microbe disposed within the chamber such that at least a portion of the water passing through the chamber is in fluid communication with the microbe; and

at least one filter for filtering the treated water.

155. The apparatus of claim 154, wherein the microbe is an exopolymer-producing bacterium.

156. The apparatus of claim 154, wherein the microbe is a proteobacterium.

157. The apparatus of claim 156, wherein the proteobacterium is selected from the group consisting of purple bacteria, chemoautotrophic proteobacteria, and chemoheterotrophic proteobacteria.

158. The apparatus of claim 154, wherein the microbe is a fungus.

61

159. The apparatus of claim 154, wherein the microbe is a lichen.

160. The apparatus of claim 154, wherein the microbe is an alga.

161. An apparatus for processing water to control hardness, comprising:

a source of cation-chelating, exopolymer-producing bacteria disposed within the chamber such that at least a portion of the water passing through the chamber is in fluid communication with the bacteria; and

at least one filter for filtering the treated water.

162. The apparatus of claim 161, wherein the chamber retains the bacteria, but not the cation- chelated bacterial exopolymer.

163. The apparatus of claim 161, wherein the chamber retains both the bacteria and the cation- chelated bacterial exopolymer.

164. The apparatus of claim 161, wherein the chamber retains neither the bacteria nor the cation- chelated bacterial exopolymer.

165. The apparatus of claim 161, wherein the filter has a pore size of not more than about 0.2 μm.

166. The apparatus of claim 161, wherein the filter is selected from the group consisting of a 0.2 μm membrane filter, a 5 kD membrane filter, and a combination thereof.

167. The apparatus of claim 161, wherein the source of bacteria provides an exopolymer- producing bacteria.

168. The apparatus of claim 161, wherein the source of bacteria includes a bacterial growth matrix.

169. The apparatus of claim 161, wherein the source of bacteria includes a removable cartridge containing polymer-producing bacteria.

170. The apparatus of claim 161, further comprising at least one additional chamber having a source of bacteria disposed within, for subsequently processing the treated water.

171. The apparatus of claim 161, further including at least one pressure sensor/flow regulator for controlling the water flowing through the chamber.

172. The apparatus of claim 161, wherein the source of bacteria provides one or more exopolymer-producing bacteria that includes a Bacillus.

173. The apparatus of claim 172, wherein the Bacillus is selected from the group consisting of Bacillus cerueus and Bacillus thuringiensis.

\1A. The apparatus of claim 172, wherein the Bacillus is selected from the group consisting of Bacillus gibsonii, Bacillus pseudalcaliphilu, Bacillus pseudofirmus, Bacillus halodurans, Bacillus subtilis, Bacillus benzoevorans, Bacillus simplex, Bacillus horikoshii, Bacillus cereus, and Bacillus thuringiensis.

175. The apparatus of claim 161, wherein the source of bacteria provides one or more exopolymer-producing bacteria selected from the group consisting of Proteus mirάbϊlis (ATCC #51286), Bacillus G3 and Bacillus MEX244.1.

176. The apparatus of claim 161, wherein the source of bacteria provides one or more exopolymer-producing bacteria that includes Pseudomonas.

177. The apparatus of claim 176, wherein the Pseudomonas is selected from the group consisting of Pseudomonas putida and Pseudomonas aeruginosa.

178. The apparatus of claim 161, wherein the source of bacteria provides a population of one or more exopolymer-producing bacteria that includes Λzotobacter.

179. The apparatus of claim 178, wherein the Azotobacter is selected from the group consisting of Azotobacter vinelandii, Azotobacter chroococcum and Azotobacter indicus.

180. The apparatus of claim 161, further comprising a second output opening in the chamber for removal of cation-saturated bacterial exopolymer.

181. The apparatus of claim 180, wherein the chamber is a first chamber and the cation chelating exopolymer-producing bacteria facilitate calcium removal.

182. The apparatus of claim 161, wherein the chamber is a first chamber that facilitates calcium removal.

183. The apparatus of claim 182, further comprising a second chamber that facilitates the formation of less stable forms of scale or other mineral deposits.

184. The apparatus of claim 182, further comprising a second chamber that facilitates the dissolving of scale or other mineral deposits.

185. The apparatus of claim 182, further comprising a second chamber that facilitates the inhibition of scale or mineral deposits formation.

186. The apparatus of claim 182, further comprising a second chamber that facilitates the formation of less stable forms of scale or mineral deposits, a third chamber that facilitates the dissolving of scale or mineral deposits, and a fourth chamber that facilitates the inhibition of scale or mineral deposits formation.

187. The apparatus of claim 182, wherein the apparatus is a part of a reverse osmosis and/or a zero level discharge reverse osmosis system.

188. A method of removing scale-forming minerals in a water system, comprising:

providing exopolymer-producing microorganisms in a chamber having at least one input opening for receiving the water from a water source, at least one output opening for discharging treated water from the chamber, and at least one filter for filtering the treated water; wherein the algae are retained in the chamber; and

passing at least a portion of the water from the water system through the chamber such that the water is in fluid communication with the the exopolymer-producing algae, wherein the exopolymer produced by the algae removes scale-forming minerals by sequestering ions, promoting the dissolution of existing scale, increasing the solubility of newly formed scale, and/or inhibiting scale formation on a surface of the water system.