Isolation and Investigation of the Exopolysaccharide from Thauera Sp

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Doctoral Dissertations Graduate School

12-2002

Isolation and Investigation of the Exopolysaccharide from Thauera sp. MZ1T

Michael S. Allen

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Recommended Citation Allen, Michael S., "Isolation and Investigation of the Exopolysaccharide from Thauera sp. MZ1T. " PhD diss., University of Tennessee, 2002. https://trace.tennessee.edu/utk_graddiss/2089

This Dissertation is brought to you for free and open access by the Graduate School at TRACE: Tennessee Research and Creative Exchange. It has been accepted for inclusion in Doctoral Dissertations by an authorized administrator of TRACE: Tennessee Research and Creative Exchange. For more information, please contact [email protected]. To the Graduate Council:

I am submitting herewith a dissertation written by Michael S. Allen entitled "Isolation and Investigation of the Exopolysaccharide from Thauera sp. MZ1T." I have examined the final electronic copy of this dissertation for form and content and recommend that it be accepted in partial fulfillment of the equirr ements for the degree of Doctor of Philosophy, with a major in Microbiology.

Gary S. Sayler, Major Professor

We have read this dissertation and recommend its acceptance:

David R. Raman, Arthur J. Meyers, Steven W. Wilhelm, Jeffrey M. Becker

Accepted for the Council:

Carolyn R. Hodges

Vice Provost and Dean of the Graduate School

(Original signatures are on file with official studentecor r ds.) To the Graduate Council:

I am submitting herewith a dissertation written by Michael S. Allen entitled “Isolation and Investigation of the Exopolysaccharide from Thauera sp. MZ1T.” I have examined the final electronic copy of this dissertation for form and content and recommend that it be accepted in partial fulfillment of the requirements for the degree of Doctor of Philosophy, with a major in Microbiology.

Gary S. Sayler Major Professor

We have read this dissertation And recommend its acceptance:

David R. Raman

Arthur J. Meyers

Steven W. Wilhelm

Jeffrey M. Becker

Accepted for the Council:

Anne Mayhew Vice Provost and Dean of Graduate Studies

(Original signatures are on file with official student records.)

Isolation and Characterization of the Exopolysaccharide

Produced by Thauera strain MZ1T and Its Role in

Flocculation

A Dissertation

Presented for the

Doctor of Philosophy Degree

The University of Tennessee, Knoxville

Michael S. Allen

December 2002

ACKNOWLEDGEMENTS

I would first like to thank my major professor and mentor, Gary Sayler, for his guidance and support throughout this research. I would also like to thank the other members of my committee, Art Meyers, Steve Wilhelm, Raj Raman, Gary Stacey, and

Jeff Becker, for their patience and support. I would particularly like to thank Art Meyers for his support and assistance above and beyond the normal call of duty.

I am very grateful to David Baker for his assistance and insight, and to his students Karen Welch and Benjamin Prebyl. I am also grateful to Curtis Lajoie, Alice

Layton, and Anthony Hay for their assistance early on in this project. I would also like to thank David Nivens for his time, thoughts, and assistance on a variety of aspects of this research, and to all the members of the CEB, each of who have assisted me at some point along the way.

I would also like to thank my friends James Rice, Scott Moser, Chris Elkins,

Cathy Scott, Dan Williams, and Victoria Garrett for their continued patience and unwavering moral support. Finally, I would like to thank Amy Tomaszewski to whom I owe a great deal beyond what is contained in this manuscript.

ABSTRACT

Thauera sp. strain MZ1T is a floc-forming bacterium isolated from the

wastewater treatment plant of Eastman Chemical Company. Its overabundance in that

system in the form of zoogloeal clusters was positively correlated to episodes of poor

dewatering of activated sludge (Lajoie 2000). The specific cause of this problem was

thought to be due to the production of large quantities of hydrophilic exopolysaccharide

(EPS) by MZ1T, which entraps water in the form of a hydrated gel, and results in a

sludge that is resistant to mechanical dewatering.

A method for the reproducible extraction of EPS from pure cultures of MZ1T was

developed. Subsequent investigation of the physical properties of the purified EPS found

that the polymer was highly soluble in water but insoluble in non-aqueous solvents. The

polymer was also found to be thermally stable. Investigations into the interaction of the

EPS with metal cations revealed that the EPS showed a capacity for binding uranium ions in both aqueous and non-aqueous solutions. Additionally, the EPS was found to interact with calcium chloride, resulting in the precipitation of the EPS from solution.

The glycosyl composition of the EPS was determined by gas chromatography– mass spectroscopy (GC–MS) of both the alditol acetate and per-O-trimethylsilyl methyl

glycoside (TMS) derivatives. By these methods, the EPS was found to include:

rhamnose, N-acetylfucosamine, galacturonic acid, N-acetylglucosamine, and trace amounts of glucose.

iii Spectroscopic analyses of MZ1T EPS by one- and two-dimensional nuclear

magnetic resonance (NMR) and Fourier-transform infrared (FT–IR) spectroscopy were

used to support the chemical analyses and to identify the probable linkage of

monosaccharides within the polymer and their respective D- or L- configurations.

Spectroscopic analyses also revealed the presence of an aglycon substituent present on the EPS polymer.

While all the monosaccharides detected in the chemical analyses could similarly be identified in NMR spectra, no through-space interactions were detected between glucose and any of the other monosaccharides. These results, along with data indicating the presence of glucose in gel permeation column fractions in the absence of other monosaccharides, suggest that glucose may be present in the form of a second polysaccharide in the EPS preparations.

Several mutants of MZ1T incapable of, or reduced in, their capacity for floc formation in liquid media were isolated following chemical mutagenesis. Investigations of EPS extracted from these mutants revealed that all of the monosaccharides previously detected in the wild type EPS could also be identified in the EPS of the mutants, indicating that loss of floc forming capacity was not a result of alteration in the glycosyl composition of the EPS. Spectroscopic analysis by FT–IR of the EPS extracted from true floc– mutants did, however, reveal conserved alterations in the spectra of the mutant EPS

relative to that from the wild type. These data suggest that alteration of the linkage or the substitution of the EPS is responsible for the loss of floc-forming capacity in the mutants.

Additionally, it was found that floc– and floc-reduced isolates, unlike the wild

type, were competent to receive broad host range plasmids by conjugal transfer.

iv Colonies of these mutants also exhibited altered colony texture and differential responses to stains and dyes than did the wild type. Taken together, these data suggest a protective role of the EPS of MZ1T, and that mutations resulting in alterations of the EPS broadly affect the cell surface organization, intercellular interactions, and floc-forming capacity.

TABLE OF CONTENTS

I. Introduction……………………………………………………………1

II. Literature Review……………………………………………………...5

a. Activated sludge wastewater treatment systems…………………..5 b. Flocculation-associated problems in the activated sludge system...6 c. Sludge dewatering…………………………………………………8 d. Theory of biological floc formation……………………………….8 e. Mechanisms of biological flocculation ………………………….10 f. EPS biosynthesis…………………………………………………12 g. Roles of EPS in nature…………………………………………...15 h. Factors influencing EPS production……………………………..16 i. Zoogloea…………………………………………………………17 j. The genus Thauera………………………………………………21 k. Goals of this research…………………………………………….27

III. Materials and Methods…………………………………….…………32

a. Bacterial plasmids and strains……………………………………32 b. Culture conditions and storage…………………………………...32 c. Media and chemicals……………………………………………..34 d. Molecular Biology Techniques…………………………………..36 e. DNA sequencing…………………………………………………37 f. BOX polymerase chain reaction (PCR) …………………………37 g. Ribosomal intergenic sequence analysis (RISA).………………..37 h. Determination of doubling time………………………………….38 i. Carbon-source utilization………………………………………...38 j. Carbohydrate concentration determination………………………39 k. Protein concentration determination……………………………..41 l. Isolation and purification of exopolysaccharide…………………41 m. Size Determination of EPS polymer……………………………..43 n. Physical properties of MZ1T EPS……………………………….43 o. Metal binding by EPS……………………………………………44 p. Alditol acetates…………………………………………………...45 q. Per-O-trimethylsilyl methyl glycosides (TMS) …………………47 r. Gas chromatography–mass spectrometry (GC–MS) ……………51 s. Nuclear Magnetic Resonance (NMR) Spectroscopy…………….51

vi t. Fourier-transform infrared spectroscopy (FTIR).………………..52 u. Conjugation ……………………………………………………...52 v. Electroporation.…………………………………………………..53 w. Mutagenesis.……………………………………………………..54 x. Floc+/– screening assays………………………………………….54 y. MZ1T-39A transposon mutagenesis and mapping………………57

IV. Results………………………………………………………………..60

a. Isolation and identification of MZ1T…………………………….60 b. Microscopic examination of MZ1T.…………………………..…60 c. Growth rates and doubling times of MZ1T and selected floc– mutants….………………………………………………….60 d. Carbon utilization in MZ1T and T. aromatica…………………...64 e. Floc formation …………………………………………………...65 f. EPS production by MZ1T………………………………………..65 g. Physical properties of EPS……………………………………….65 h. EPS and metal binding…………………………………………...72 i. Glycosyl composition of EPS……………………………………72 j. Structural characterization of MZ1T EPS………………….…….89 k. Mutagenesis of MZ1T and isolation of floc- mutants……………99 l. Transposome mutagenesis……………………………………….99 m. EPS production in floc- mutants………………………………..101 n. Glycosyl composition of EPS from floc- mutants………………101 o. Spectroscopic analysis of EPS from floc- mutants……………..101 p. Phenotypic properties of floc- mutants…………………………117 q. Conjugation in floc- mutants……………………………………118 r. Transposon mutagenesis of MZ1T-39A………………………..121

V. Discussion…………………………………………………………..124

a. MZ1T as a member of the genus Thauera……………………...124 b. Wild Type MZ1T EPS………………………………………….125 c. EPS extraction from and composition in mutants………………130 d. Additional phenotypic alterations in floc- mutants……………..131 e. Transposon mutagenesis…………………………………..……132

Conclusions…………………………………………………………………133

Works Cited.………..……….….…...…. .………..………………..…….…136

Vita…………………………………………………………………………..152

vii LIST OF FIGURES

1. Figure I-1. Photomicrograph of MZ1T in activated sludge…………………...3 2. Figure II-1 Distance matrix tree of MZ1T and β-Proteobacteria…………….22 3. Figure II-2 Anaerobic degradation of aromatic hydrocarbons……………….25 4. Figure II-3 Benzoyl-CoA pathway. ………………………………………….26 5. Figure III-1 Carbohydrate assay standard curve………………………………40 6. Figure III-2 Protein assay standard curve……………………………………..42 7. Figure III-3 Gas chromatogram of AA standards……………………………..48 8. Figure III-4 Gas chromatogram of TMS standards. ………………………….50 9. Figure III-5 NTG-treated MZ1T kill curve. ………………………………….55 10. Figure III-6 Map of pTNModO-Km. …………………………………………58 11. Figure IV-1 Photo- and electron micrographs of MZ1T from flocculating culture. …………………………………………………………………………..61 12. Figure IV-2 Complex stain of MZ1T floc…………………………………….62 13. Figure IV-3 MZ1T wild type and mutant growth curves……………………..63 14. Figure IV-4 Photograph of MZ1T floc in liquid culture………………………66 15. Figure IV-5 Carbohydrate concentration vs. optical density. ………………...67 16. Figure IV-6 GPC chromatograph of crude EPS. ……………………………..68 17. Figure IV-7 Photographs of crude and purified EPS. ………………………...71 18. Figure IV-8 Metal binding by MZ1T EPS. …………………………………...73 19. Figure IV-9 Gas chromatograph of EPS alditol acetate derivatives. …………75 20. Figure IV-10 Mass spectrum of rhamnose AA derivative...……………………76 21. Figure IV-11 Mass spectrum of N-acetylfucosamine AA derivative…………..77 22. Figure IV-12 Mass spectrum of inositol AA derivative. ………………………78 23. Figure IV-13 Mass spectrum of glucose AA derivative. ………………………79 24. Figure IV-14 Mass spectrum of N-acetylglucosamine AA derivative…………80 25. Figure IV-15 Gas chromatogram of EPS TMS derivatives………………….…82 26. Figure IV-16 Mass spectrum of rhamnose TMS derivative……………………83 27. Figure IV-17 Mass spectrum of N-acetylfucosamine TMS derivative…………84 28. Figure IV-18 Mass spectrum of glucose TMS derivative………………………85 29. Figure IV-19 Mass spectrum of inositol TMS derivative………………………86 30. Figure IV-20 Mass spectrum of N-acetylglucosamine TMS derivative………..87 31. Figure IV-21 Mass spectrum of galacturonic acid TMS derivative……………88 32. Figure IV-22 FT-IR spectrum of MZ1T EPS…………………………………..90 33. Figure IV-23 1D 1H-NMR spectrum of MZ1T EPS……………………………92 34. Figure IV-24 2D 1H-13C HSQC spectrum of MZ1T EPS. ……………………..93 35. Figure IV-25 Anomeric region of the HSQC spectrum from MZ1T EPS……...94 36. Figure IV-26 2D 1H-1H gCOSY spectrum of MZ1T EPS……………………...95 37. Figure IV-27 Anomeric and ring region of the gCOSY spectrum of MZ1T EPS.………………………………………………………………………………96 38. Figure IV-28 2D 1H-1H TOCSY spectrum of MZ1T EPS. ……………………97 39. Figure IV-29 GC of the AA derivatives from MZ1T-20A EPS. ……………..102 40. Figure IV-30 GC of the AA derivatives from MZ1T-26B EPS………………103 41. Figure IV-31 GC of the AA derivatives from MZ1T-27B EPS………………104

viii 42. Figure IV-32 GC of the AA derivatives from MZ1T-30B1 EPS……………..105 43. Figure IV-33 GC of the AA derivatives from MZ1T-35B1 EPS……………..106 44. Figure IV-34 GC of the AA derivatives from MZ1T-37B EPS….…………...107 45. Figure IV-35 GC of the AA derivatives from MZ1T-39A EPS……………....108 46. Figure IV-36 GC of the TMS derivatives from MZ1T-20A EPS……………..109 47. Figure IV-37 GC of the TMS derivatives from MZ1T-26B EPS……………..110 48. Figure IV-38 GC of the TMS derivatives from MZ1T-27B EPS……………..111 49. Figure IV-39 GC of the TMS derivatives from MZ1T-30B1 EPS……………112 50. Figure IV-40 GC of the TMS derivatives from MZ1T-35B1 EPS……………113 51. Figure IV-41 GC of the TMS derivatives from MZ1T-37B EPS……………..114 52. Figure IV-42 GC of the TMS derivatives from MZ1T-39A EPS……………..115 53. Figure IV-43 GC of the TMS derivatives from MZ1T-44b1 EPS…………….116 54. Figure IV-44 Comparison of FT-IR spectra of MZ1T wild-type and mutant EPS……….…………………….…………………………………..119 55. Figure IV-45 Alcian blue stain of MZ1T wild type and mutant colonies…….120 56. Figure IV-46 Relative abundance of monosaccharides in GPC column fractions..…………………………………………….………………………….129

ix LIST OF TABLES

1. Table II-1. Origins and applications of bacterial EPS…………….……30 2. Table III-1. Bacterial strains and plasmids used in this study…………..33 3. Table IV-1. Compounds and tests applied to floc+ and floc- strains……57 4. Table IV-2. Average EPS yield from floc+ and floc- strains.……………69 5. Table IV-3. Chemical shifts of NMR resonances.……….……….……100

x CHAPTER I

INTRODUCTION

Viscous bulking represents a major operational problem in wastewater treatment systems. These episodes result in poor compaction and settling of biomass in the clarifying basin, increased effluent biological oxygen demand (BOD), and poor dewaterability of the waste biosludge (Jenkins 1993). The high water content of the sludge results in increased costs associated with dewatering, incineration, or landfill disposal. Treatment requires the additional costs of adding synthetic polymers and clay to promote compaction and decrease the water content of the sludge.

Episodes of viscous bulking have been associated with excessive exopolymer production by resident microorganisms (Jenkins 1993). One group of organisms historically associated with slime production in activated sludge systems includes strains of the species Zoogloea ramigera (Butterfield 1935, Crabtree 1966, Friedman 1968a,

Jenkins 1993, Lu 2001, Rossello-Mora 1995, Unz 1967).

Identification of Z. ramigera in wastewater treatment systems has most often been made based on the phenotypic trait of “zoogloeal cluster” formation, in which a number of bacteria are encapsulated in a gelatinous, mucopolysaccharide matrix (Butterfield

1935, Dugan 1987, Friedman 1968b, Unz 1967). This has resulted in the isolation of a variety of strains that were misidentified as Z. ramigera. 16S rDNA and phylogenetic analyses of some of these strains later revealed them to be distinct species within the α and β subclasses of Proteobacteria (Shin 1993). Fluorescent in situ hybridization (FISH) and dot-blot analyses using 16S probes specific to these organisms indicated that only the

1 type strain, Z. ramigera ATCC 19544T (strain 106), was found to commonly occur in the activated sludge systems investigated (Rossello-Mora 1995).

The mechanisms responsible for the formation of zoogloeal clusters are not fully understood. The majority of research into this process and the organisms involved has focused primarily on the production of exopolysaccharide (EPS) (Easson Jr. 1987b,

Farrah 1976, Friedman 1969, Unz 1967). Glycosyl composition of the EPS from the few strains for which data are available has revealed that the compositions are unique among strains. However, insufficient data exist to determine if the compositions of the EPS from the various strains possess any common characteristics.

Recently, a zoogloeal cluster-forming organism was isolated from an industrial wastewater treatment system by micromanipulation (Lajoie 2000). This strain, originally thought to be an isolate of Z. ramigera, was designated MZ1T and classified as a member of the genus Thauera by 16S rDNA sequence analysis. Its abundance (as zoogloeal clusters) in the wastewater treatment facility of Eastman Chemical Company was positively correlated to episodes of poor dewaterability of the activated sludge (Figure I-

1) (Lajoie 2000). Unlike the zoogloeal clusters described for Z. ramigera ATCC 19544T;

however, clusters produced by MZ1T are globular, amorphous, and have not been

observed to form extensive, finger-like projections.

It has also been observed that MZ1T forms flocs in pure culture. It is important to

note that flocculation is not necessarily synonymous with floc formation, though the

terms are often used interchangeably (Dugan 1987). The former refers to the aggregation

of separate cells or particles into larger masses, and the latter is generally associated with

the growth of cells within a loosely associated gelatinous matrix. Both conditions are,

Figure I-1. Photomicrograph of MZ1T in activated sludge. MZ1T (green) can be seen in the zoogloeal cluster phenotype. MZ1T was stained by fluorescent in situ hybridization using a fluoroscein-labeled 16S rDNA gene probe (Lajoie et al. 2000). Other bacteria present in the sample were counterstained red using a rhodomine-labeled universal 16S rDNA probe. Photo by I. Gregory.

3 however, associated with extracellular polymeric substances (ECP) (Dugan 1987,

Friedman 1968, Geesey 1983). For the purposes of this work, formation of flocs by

MZ1T is considered “floc formation”, though some combination of floc formation and

flocculation is likely to be involved. Furthermore, floc formation and zoogloeal cluster

formation are operationally defined here to be aspects of the same process, differing

primarily in scale, where zoogloeal cluster formation is considered a microscopic

observation, and floc formation is defined as its macroscopic manifestation.

The goal of this research was to understand the role of MZ1T in viscous bulking

episodes and poor dewaterability of waste activated sludge. To achieve that end, the

following research objectives were proposed: 1) to isolate and identify the nature of the extracellular polymer produced by MZ1T, 2) to determine the glycosyl composition of the isolated exopolysaccharide, 3) to determine what EPS characteristics, if any, are conserved among the closely related zoogloeal cluster-forming bacteria, 4) and to determine what attributes of the EPS are necessary for floc formation in MZ1T.

4 CHAPTER II

LITERATURE REVIEW

A. Activated sludge wastewater treatment systems

The most common method for the secondary treatment of wastewater is the activated sludge process (Jenkins 1993). This process generally consists of two main components: 1) the aeration basin and 2) the clarifying or settling basin. First, wastewater enters the aeration basin, which serves as a biological reactor for the removal of dissolved and suspended pollutants by resident micro- and macroorganisms that make up the activated sludge. Air is introduced into this basin to maintain aerobic conditions for degradation and assist in mixing. Mixed effluent from the aeration basin is then transported to the clarifying basin. The role of this second basin is the separation of treated wastewater from activated sludge solids by sedimentation. The treated wastewater may then be directly discharged to a watershed or municipal water system, disinfected, further processed for the removal of specific nutrients or contaminants, or reused. Waste activated sludge that has accumulated by sedimentation on the bottom of the clarifying basin is continuously removed. One fraction is further processed by mechanical dewatering followed by incineration or landfill disposal, while the remainder is returned to the aeration basin to maintain the process.

The process of flocculation facilitates the separation of activated sludge from treated wastewater. Flocculation is the aggregation of colloidal and dissolved materials into a mass that settles from solution or suspension (Dugan 1987, Pavoni 1972).

Successful removal of the flocs from the treated wastewater produces a clear effluent

5 with low biological oxygen demand (BOD). The overall efficiency of the activated sludge system is therefore dependent on flocculation (Jenkins 1993, Jorand 1995, Pavoni

1972).

B. Flocculation-associated problems in activated sludge systems

Several problems associated with flocculation may arise in the activated sludge treatment process. Investigation of these problems tends to focus on the roles of bacteria, since these organisms make up the bulk of the activated sludge flocs (Unz 1987). One problem of activated sludge processes is that of dispersed growth in which the organisms fail to aggregate or form small “pinpoint flocs” that settle poorly (Watanabe 1999).

These conditions typically result in the generation of a turbid effluent (Jenkins 1993).

A second type of problem may result when the sludge blanket rises or portions of the activated sludge solids float. These problems may be caused by excessive production of gases (e.g. N2) in the clarifying basins and the subsequent trapping of bubbles in the sludge blanket, or by the production of foams (Jenkins 1993). These conditions typically result in the formation of scum or foams on the surface of the clarifying basin.

A third problem associated with activated sludge systems is bulking. This condition can be subdivided into two types, filamentous or viscous bulking. The former results from the overabundance of filamentous organisms such as Sphaerotilus natans in the activated sludge. The filaments of these organisms extend outside of the sludge floc into the bulk fluid resulting in poor settling and compaction of the activated sludge and a high sludge volume index (SVI) (Jenkins 1993). The sheath surrounding the filaments of

S. natans is composed of EPS, and transposon insertions that result in non-filamentous

6 phenotypes have been mapped to a glycosyl transferase gene (Suzuki 2002).

Complementation of this gene restores the filamentous phenotype.

Viscous, or non-filamentous bulking, is the result of overproduction of extracellular slime by resident microorganisms in the activated sludge system. This condition results in reduced settling in the clarifying basin and difficulties in mechanical dewatering of the waste sludge (Jenkins 1993). No direct evidence has been reported linking Zoogloea ramigera directly with episodes of viscous bulking, though the organism is known to produce large quantities of EPS and is often identified in activated sludge systems (Lu 2001, Rossello-Mora 1995, Unz 2002). Recent evidence has, however, positively correlated the abundance of the zoogloeal-cluster forming bacterium

Thauera sp. MZ1T with episodes of poor dewaterability of activated sludge at an industrial wastewater treatment facility (Lajoie 2000).

Treatment for settling problems in activated sludge systems generally involves the addition of chemicals and polymers. In the case of filamentous bulking, chlorination or addition of hydrogen peroxide has been used (Jenkins 1993). This technique is possible because the large surface area to volume ratio of the filamentous organisms makes them highly susceptible to chemical oxidation/disinfection treatment (Jenkins 1993).

Conversely, non-filamentous bulking or turbid growth has been treated by the addition of synthetic cationic polymers or minerals (Jenkins 1993). These compounds help bridge small and diffuse flocs, thereby increasing density and improving the settling characteristics of the sludge blanket in the clarifying basin. Biodegradable microbial polymers that can serve as bioflocculants are currently under development (Dremlin

1999).

C. Sludge dewatering

Following removal of the activated sludge from the clarifying basin, the sludge is

typically mechanically dewatered to aid in its final disposal in either a landfill (by

reducing weight and volume) or by incineration (by increasing burning efficiency).

Effective dewatering of the sludge is, therefore, dependent upon the interactions of water

within the sludge.

Water in sludge can be free or bound within the sludge matrix (Colin 1995,

Vesilind 1994). The extent to which water is bound and unable to be removed by

mechanical dewatering is a function of the composition of the sludge (Vesilind 1994).

Poxon and Darby suggested that it is the biochemical characteristics of the extracellular

polyanion (ECPA) component of the sludge rather than the total ECPA concentration that influences sludge dewaterability (Poxon 1997). Recent work showing increased sludge dewaterability at pH 2.5 supports this conclusion (Chen 2001).

D. Theory of biological floc formation

Activated sludge flocs consist of both biological (e.g. bacteria, fungi) and non- biological (i.e. inorganic compounds) components (Forster 1976, Jenkins 1993, Unz

1987). Their aggregation may be based on ionic or hydrophobic interactions, or by physical entanglement within extracellular polymeric material (Liao 2001, Poxon 1997,

Sobeck 2002, Unz 1987).

The actions of extracellular polymeric substances (abbreviated here as “ECP”

rather than “EPS” for clarity) are the primary mechanisms by which cells establish and

8 maintain associations between microorganisms and between microbes and surfaces

(Geesey 1983). These polymers exist as a highly hydrated (98% water) gel encapsulating

one or multiple species of microorganisms. Investigations of activated sludge samples

taken from two municipal treatment plants found that cell mass constituted only 10-15% of the total organic matter, the remainder being composed primarily of ECP (Frolund

1996). The composition of the ECP responsible for aggregation may include proteins, nucleic acids, carbohydrates, lipids, or various combinations of these (Unz 1987).

The composition of the ECP from a given species, however, should not be thought of as fixed. For example, the Gram negative, β-Proteobacterium Thiobacillus ferrooxidans has been shown to alter the composition of its ECP to conform to the nature of the substrate to which it is attached. When grown on hydrophobic, elemental sulfur, the ECP is composed of fatty acids with little or no detectable carbohydrates (Sand

1999). Conversely, ECP of pyrite-grown cells contain carbohydrates including uronic acids, which interact with iron(III) during pyrite degradation (Gehrke 1998, Sand 1999).

Considerable emphasis has been placed on the role of ions and their interaction with microbial extracellular polymers during biological flocculation, or bioflocculation

(Dugan 1987, Forster 1976, Pavoni 1972, Sobeck 2002, Unz 1987). Three main theories have arisen to explain these interactions. The first of these is the Double Layer theory

(Adamson 1990). This theory describes flocculation as a colloidal development, in which a charged particle is coated with a layer of counter ions. A diffuse, second layer then develops, which decreases the electric potential of the internal charge, and promotes aggregation by short-range attractive forces. Increasing ionic strength compresses the diffuse layer and better facilitates flocculation.

9 A second theory involving cation-induced bioflocculation is the Divalent Cation

Bridging (DCB) theory (McKinney 1952). This theory suggests that divalent cations in

solution form bridges between negatively charged polymers (e.g. acidic sugar or amino

acid residues) present on and around microorganisms. Support for this theory comes

from experiments showing that monovalent sodium ions can displace divalent cations and

result in deterioration of the floc structure (Higgins 1997). Additionally, the divalent

cations Ca2+ and Mg2+ have been shown to improve settling and dewatering of activated sludge, and their presence has been shown to increase populations of bacteria adhering to surfaces (Simoni 2000, Sobeck 2002).

A third theory of cation-induced bioflocculation is the Alginate theory (Bruus

1992). This theory involves the specific interaction of the polyuronic acid polysaccharide alginate specifically with Ca2+ ions in the formation of alginate gels. Alginate-producing

strains as well as uronic acids have been identified in activated sludge systems (Frolund

1996, Horan 1986, Pike 1971).

E. Mechanisms of biological flocculation

Cellular aggregation of the Gram negative, α-Proteobacterium, Azospirillum brasilense in pure culture has been positively correlated to the presence of outer membrane proteins (OMP) (Burdman 1999). Sonicated extracts of these cells showed decreased aggregative capacity after treatment with proteinase K.

Mutants of this strain that were non-fluorescent in the presence of the carbohydrate-binding dye calcofluor were unable to form flocs (Michiels 1990). Data

also indicated that the EPS content was positively correlated to the extent of aggregation,

10 which suggested a possible interaction of EPS and OMP (Burdman 1998). These data are

consistent with the hypothesis that aggregation in this species may involve the interaction of EPS with cell surface lectins, examples of the latter having been isolated from this species (Castellanos 1998).

Nucleic acids have also been implicated in the flocculation of some bacterial species. The Gram negative, photosynthetic α-Proteobacterium Rhodovulum sp. was shown to accumulate large quantities of nucleic acids extracellularly (Watanabe 1998).

Flocs of these cells in pure culture were disrupted by treatment with RNase and DNase,

but not by treatment with amylase, pectinase, cellulase, protease, or trypsin, suggesting

the involvement of nucleic acids in flocculation (Watanabe 1998).

Exopolysaccharide (EPS) is another ECP component known to influence

intercellular aggregation. Pseudomonas aeruginosa, for example, produces alginate—an

extracellular polysaccharide composed of mannuronic and guluronic acids (Evans 1973).

Genetic mutation affecting the O-acetylation of alginate has been shown to prevent

formation of natural biofilms by this bacterium (Nivens 2001). Production of EPS has

also been shown to positively correlate to flocculation in Zoogloea MP6 (Unz 1976b).

Mutants of two other bacterial strains previously considered members of the species Z. ramigera that were deficient in floc formation were found to have altered EPS levels and compositions (Easson Jr. 1987a, Easson Jr. 1987c). Additionally, isolation of large quantities of EPS from activated sludge suggests that these polymers play an active role in flocculation within these systems (Forster 1976, Frolund 1996, Goodwin 1989, Horan

1986, King 1990).

11 F. EPS biosynthesis

Biosynthesis of EPS in Gram-negative bacteria involves the sequential addition of

sugars onto a lipid carrier in the periplasm. The oligosaccharide of the sugar-lipid

complex, usually comprising a repeating unit, is then polymerized and translocated across

the outer membrane to the exterior of the cell. This general process appears to be

conserved among Gram-negative bacteria for EPS, cell wall, and O-antigen biosynthesis

(Gacesa 1998, Glucksmann 1993, Ielpi 1993, Sutherland 1972, White 1995).

The first components necessary in the production of EPS or O-antigen are

monosaccharides. In most cases, glucose serves as the starting material from which all

other hexose sugars are derived. For example, in colanic acid biosynthesis by

Escherichia coli K-12, guanosine diphosphate (GDP)-D-glucose is first converted to

GDP-D-mannose, followed by a three-step conversion to yield GDP-L-fucose

(Adrianopoulos 1998, Stevenson 1996). When not present in the growth medium,

glucose is presumably produced via gluconeogenesis (White 1995).

In this process, monosaccharides are phosphorylated and the resultant

phosphosugar is transferred directly to a nucleotide triphosphate (NTP) to form a sugar

nucleotide (NDP-X) (Mengin-Lecreulx 1993, Sutherland 1972). In the case of the

production of the EPS xanthan by the bacterium Xanthomonas campestris, a phosphate

from ATP is transferred to glucose (Glc) to form glucose-6-phosphate (Glc-6-P) by the

enzyme glucokinase. The enzyme phosphoglucomutase converts Glc-6-P to glucose-1-

phosphate (Glc-1-P), which is then added to uridine triphosphate (UTP) by uridine

diphosphate (UDP)-glucose pyorphosphorylase to form UDP-glucose (Harding 1993,

Koplin 1992). Some Glc-6-P is also converted to fructose-6-phosphate, which is then

12 isomerized to mannose-6-phosphate (Man-6-P) by phosphomannose isomerase. Man-6-P

is converted to mannose-1-phosphate (Man-1-P), which is then added to GTP to form

guanosine diphosphate (GDP)-mannose (Harding 1993, Koplin 1992).

Following the production of sugar nucleotides, the first monosaccharide in the

repeating unit is transferred to bactoprenol (i.e. undecaprenyl phosphate), a C55 isoprenoid phosphate that serves as a lipid carrier for the polymerization and translocation of oligosaccharides during the production of peptidoglycan, lipopolysaccharide, and EPS (Sutherland 1972, White 1995). The first step of this process in the production of xanthan is the addition of glucose in the form of UDP- glucose to bactoprenol (lipid-P), producing lipid-P-P-glucose and releasing uridine monophosphate (UMP) (Ielpi 1993). In this manner, with each step requiring a different enzyme, a second glucose, followed by a mannose, a glucuronic acid, and a final mannose residue are sequentially added to form a pentasaccharide repeating unit anchored to the cytoplasmic side of the inner membrane (Ielpi 1993, Katzen 1998).

Similar biosynthetic pathways have been described for sphingan EPS (e.g. gellan, welan, rhamsan, etc) from Sphingomonas species, the EPS of Rhizobium lugumuminosarum, and

succinoglycan from Rhizobium meliloti (Glucksmann 1993, Pollock 1998).

During the producion of xanthan, the mannose residues of the repeating unit are

O-acetylated prior to polymerization. GumF acetylates the inner mannose residue while

the outer residue is acetylated by GumG (Katzen 1998). The repeating unit is then

further modified by the addition of pyruvate to the outer mannose by the protein GumL.

The modification of an EPS by the addition of acetyl, pyruvyl, or succinyl moieties

esterified to one or more individual monosaccharides in the polymer appears to be a

13 common phenomenon in bacteria (Skjåk-Braek 1986, Stankowski 1993, Sutherland 1985,

Troyano 1996). These modifications can affect the functionality of the polymer, as is

seen in the lack of proper biofilm formation in P. aeruginosa mutants that do not O-

acetylate their alginate (Nivens 2001), or in defective nodule invasion by R. meliloti

mutants that do not succinylate their EPS (Leigh 1987).

Following the production of the lipid-linked repeating units, translocation of the

repeating units to the periplasm and subsequent polymerization of the polysaccharide

must ensue to yield the final EPS. Strains of X. campestris with mutations in gumB, gumC, or gumE genes were shown to accumulate the pentasaccharide repeating units, but to be incapable of xanthan production (Katzen 1998). Based on hydrophobicity plots of deduced amino acid sequence and homology to known proteins, it was postulated that these gene products are membrane-bound proteins involved in the polymerization and translocation of xanthan. Some of these mutants were found to be lethal in the wild type, and could only be isolated as conditional mutants. By contrast, algK mutants of P.

aeruginosa could not produce alginate but were found to secrete uronic acids, suggesting

this gene may play a role in EPS polymerization (Sumita 1998). The mechanism of final

transfer of the growing polysaccharide across the outer membrane is not well understood.

The biosynthesis of bacterial alginate by P. aeruginosa is slightly different from

that described above. Its EPS, alginate, is not composed of a repeating unit. Instead,

alginate is produced as a polymer of polymannuronate, which is then epimerized in the

C5 position to periodically introduce guluronic acid residues into the polymer (Evans

1973, Franklin 1994).

14 The genes involved in the biosynthesis of a given EPS are typically clustered together on the chromosome or on megaplasmids (Chitnis 1993, Glucksmann 1993,

Harding 1987, Long 1988, Peñaloza-Vázquez 1997). Surprisingly, given their large number, these biosynthetic genes are often transcribed as one or only a few operons

(Chitnis 1993, Glucksmann 1993, Katzen 1996). Accessory genes whose protein products are necessary for translocation of the EPS polymer and/or are used in the production of other extracellular polysaccharides (e.g. O-antigen), however, may be located within other, distant operons (Glucksmann 1993, Ye 1994).

G. Roles of EPS in nature

The roles of exopolysaccharides in nature are as diverse as the organisms that produce them. One important role is in surface attachment and the formation of biofilms.

Biofilm formation by Pseudomonas aeruginosa is facilitated by the production of alginate (Costerton 1994). This EPS provides a microniche during bacterial surface colonization, as well as produces a diffusion barrier for solutes and nutrients. Laboratory experiments with co-cultures have illustrated that the formation of a biofilm can influence competition among multiple species and facilitate coexistence, even when growth rates are very different (Banks 1991, Stewart 1997). Individual species grown in dual species biofilms have also been shown to become co-embedded in EPS, thereby becoming partially protected from a single, specific glycanase (Skillman 1999).

The barrier presented by the EPS also provides some protection from antimicrobial compounds (Stewart 1994). Further evidence for a protective role of EPS comes from studies with Acinetobacter sp., which showed that the EPS-protected cells

15 from freezing, heat, drying, pH changes, biocides, detergents, and heavy metals (Pirog

1997). The latter was not found to be the case for Sphingomonas paucimobilis R40, however, where production of the EPS gellan resulted in decreased tolerance to sublethal concentrations of copper (Cu2+) relative to a nonproducing strain (Richau 1997). EPS may not be a barrier to bacteriophage, however, since endodepolymerases that degrade host-specific EPS have been isolated from virus particles (Nelson 1988).

EPS plays an integral part in plant-associated bacteria. For example,

Agrobacterium tumafaciens strains deficient in production of succinoglycan were unable to form normal crown gall tumors (Cangelosi 1987). In symbiotic organisms, mutations in EPS synthesis adversely affect proper establishment of symbioses (Finan 1985, Leigh

1985). Plant pathogenicity of bacteria has also been shown to be affected by loss of EPS biosynthesis (Chou 1997, Hayward 1991).

H. Factors influencing EPS production

EPS production in a number of bacterial strains has been shown to be affected by carbon-to-nitrogen (C:N) ratios within the growth medium (Casas 2000, Degeest 1999,

Huang 1994, Parsons 1971, Roseiro 1993, Unz 1976b, Williams 1980). For example, xanthan production was found to be optimal under nitrogen-limiting conditions (Roseiro

1993). Similarly, polysaccharide to protein ratios increased in E. coli biofilms with increased C:N ratio (Huang 1994). Other factors reported to influence EPS production in bacteria include: the carbon source, growth rate, pH, ionic strength of the medium, trace element concentration, surface attachment, and temperature (Barbaro 2001, Mengistu

1994, Petry 2000, Roseiro 1993, Uhlinger 1983, Vandevivere 1983).

16 Growth phase has also been shown to influence EPS biosynthesis. EPS

production was also found to increase “markedly” following mid-exponential growth

phase in Pseudomonas NCIB 11264 (Williams 1980). For P. atlantica, the largest

accumulation of EPS was found to occur in stationary phase (Uhlinger 1983). Similar

results were reported for Zoogloea ramigera (Unz 1976b).

Some bacteria have been reported to possess the capacity to alter their EPS during

their growth phase. Increased length of fermentation time was found to result in

increased degrees of acetylation and pyruvylation as well as increased polymer size of

xanthan from batch cultures of Xanthomonas campestris (Casas 2000). P. atlantica was found to alter the relative abundance and concentration of uronic acids in its EPS during its growth cycle (Uhlinger 1983). Unlike these minor alterations, a completely new EPS polymer was detected under conditions of phosphorus starvation in Rhizobium meliloti, in addition to the better-studied succinoglycan (Rüberg 1999, Zhan 1989).

I. The genus Zoogloea

The species of the genus Zoogloea are of particular relevance due to their: 1) ability to form flocs and/or zoogloeal clusters, 2) production of exopolysaccharides, 3) phenotypic similarity and phylogenetic relatedness to Thauera, and 4) historic association with wastewater treatment systems. For these reasons, an overview of the Zoogloea is presented here.

The naming of the bacterium Z. ramigera and the first references to zoogloeal

formations were made by H. Itzigsohn in 1868 according to Butterfield (Butterfield

1935). Later isolation of a phenotypically similar organism from activated sludge and its

17 subsequent study in pure culture led to the conclusion that Z. ramigera was the primary

organism involved in the production of sludge flocs (Butterfield 1935). Flocs of the

organism were characterized as possessing a fingered structure as determined by

microscopic examination.

Isolation of another strain, Z. ramigera ATCC 19544 (a.k.a. strain 106) by Unz

and Dondero that produces fingered zoogloeal clusters, was analyzed for the production

of EPS (Farrah 1976, Unz 1967). Glycosyl composition of the extracted EPS produced in

pure culture was found to contain predominately aminosugars and be similar to extracts

from activated sludge. The aminosugars were tentatively identified as glucosamine and

possibly a methyl-pentosamine by paper chromatography. Uronic acids were also

detected by colorimetric assay. Additionally, a second strain, Z. ramigera MP6, was

analyzed and yielded similar results. Tezuka reported similar results from EPS extracts

of an unspecified strain of Z. ramigera, in which the second aminosugar was

hypothesized to be fucosamine (Tezuka 1973).

Early investigation into the growth dynamics of a pure culture of an unspecified

strain of Z. ramigera found a normal increase in turbidity with time followed by the rapid onset of flocculation. This led to an increase in floc size, and the settling of large flocs was marked by a decrease in optical density (Finstein 1967). In a separate experiment,

Strand et al. showed that the addition of cell-free supernatant from wild-type Z. ramigera

ATCC 19544T culture could induce flocculation in a spontaneous floc- mutant (Strand

1988). Later studies on the formation of zoogloeal clusters by Z. ramigera ATCC

19544T by cinephotomicrography found that fingered zoogloeae emerged from the

activated sludge flocs as a result of unidirectional movement and multiplication (Unz

18 1976a). The authors further noted that fingered zoogloeae were not present in the freshly

cultured, activated sludge flocs prior to the experiment. The fingered development was

hypothesized to be an aerotactic response. Nothing is known of the genetics of floc

formation or EPS biosynthesis for this strain, and no reports of genetic manipulation of

any kind have been published.

Friedman and Dugan isolated another strain of Z. ramigera from water samples

(Friedman 1968b). This strain, Z. ramigera 115 (ATCC 25935) produces a

mucopolysaccharide matrix that has been described as being “globular packets of cells”

with fingerlike projections (Friedman 1968b). The isolated EPS from this strain was also

found to be susceptible to cellulase degradation. Poly-β-hydroxybutyrate (PHB) granules

were detected within the cells, and increases in EPS production were later linked to PHB

degradation (Friedman 1968b, Parsons 1971). The thiolase involved in PHB synthesis

has been cloned and studied in detail (Palmer 1991, Williams 1992).

Further investigation of the EPS produced by Z. ramigera ATCC 25935 found it

to behave as a polyelectrolyte with strong affinity to metal cations, and to be a good

stabilizer and emulsifier in oil-in-water systems (Norberg 1982, Stauffer 1980). Glycosyl

composition of the EPS from this strain was found to be composed of D-glucose, D- galactose, and pyruvic acid (Ikeda 1982). Later work indicated the presence of acetate and succinate substituents on the EPS (Troyano 1996). Structural data for the EPS have also been reported (Franzen 1984, Troyano 1996).

Successful transfer of incP plasmids into Z. ramigera ATCC 25935 by conjugation was reported using nitrosoguanadine-treated mutants with altered colony morphology (Easson Jr. 1987b). Although these mutants were found to be capsule and

19 flocculation deficient, they produced a chemically similar EPS that was released directly

into the media and was not tightly associated with the cells (Easson Jr. 1987b). A

plasmid containing DNA sequence that complemented the mutants and restored wild type

phenotype was found to be similar to the pyruvyl transferase gene of Xanthomonas

campestris by DNA-DNA hybridization (Easson Jr. 1987a). This plasmid was later sequenced revealing two open reading frames that showed no homology to any previously identified proteins (Sam-Pin 2000). Later investigation of the EPS from mutant and wild type cultures indicated that solutions of EPS purified from the wild type possess different viscosities than that of mutant strains, though the basic composition remained the same (Kim 1994). These data suggest some alteration in the structure of and/or substitutions on the EPS polymer are responsible for the change in phenotype.

Another strain, Z. ramigera ATCC 19623 (strain I-16-M) was isolated by

Crabtree et al. (Crabtree 1966). This strain forms flocs in pure cultures, but has not been described as producing zoogloeal clusters (Friedman 1968a, Friedman 1968b). This strain, like Z. ramigera strain 115, has also been shown to accumulate PHB granules

(Crabtree 1966, Roinestad 1970).

EPS from Z. ramigera ATCC 19623 is cellulose-like, but specific details on its composition and structure are unknown (Easson Jr. 1987c). Transposon insertion mutants deficient in calcofluor binding, a dye that binds β-1,4 linked glucose molecules, were shown to be floc-, and some mutants to be completely deficient in detectable EPS

(Easson Jr. 1987c). Spontaneous EPS- mutants of this strain were also shown to

commonly occur (Easson Jr. 1987a).

20 Investigations into the environmental distribution of fingered zoogloea-forming

strains identified isolates in wastewater, polluted pond and stream waters, duck feces,

garbage disposal wastes, and fish aquarium waters (Williams 1983). Enrichment

techniques were developed based on the aerobic degradation of aromatic hydrocarbons via the meta-cleavage pathway (Unz 1972). However, the isolation of multiple strains of

Z. ramigera with different EPS compositions and phenotypes necessitated the need for a more detailed investigation of the phylogenetics of the various strains of Z. ramigera

(Crabtree 1966, Friedman 1968a, Kwon 1994, Shin 1993, Tezuka 1973, Unz 1967).

Z. ramigera strains I-16-M, 106, and 115 share the property of floc formation and all were isolated from wastewaters. The strains, however, are biochemically and phenotypically very different (Hiraishi 1992). Strains 115 and the type strain, 106, were found by 16s rDNA sequence analysis to group within the β-Proteobacteria with

Rhodocyclus purpureus as the closest relative (Lajoie 2000, Shin 1993). This assignment places these strains within the same branch as the genus Thauera (Lajoie 2000, Macy

1993). By contrast, strain I-16-M was found to be a member of the α-Proteobacteria, closely related to Agrobacterium species (Shin 1993).

J. The genus Thauera

The genus Thauera was first described in 1993 by Macy et al. and named in honor of the German microbiologist R. Thauer (Macy 1993). The isolated type strain,

Thauera selenatis (ATCC 55363), was found to have the novel capacity to utilize

selenate as an electron acceptor, without interference from nitrate (Macy 1993). The

Figure II-1. Distance matrix tree of MZ1T and β-Proteobacteria. Tree is based on 16S rDNA gene sequence homology. The rDNA sequence of MZ1T was found to have high levels of similarity with other Thauera strains (96-97%). (Source: Lajoie et al. 2000)

22 selenate reductase from this organism has been purified and characterized (Schroder

1997).

Phylogenetic investigation of the isolate by comparison of 16S rDNA sequences

indicated that T. selenatis was most closely related to members of the beta subclass of

Proteobacteria (similarity 86.8%) (Macy 1993)(Figure II-1). The rod-shaped cells were found to be 1.4 µm × 0.56 µm and in some instances to contain poly-β-hydroxybutyrate inclusions. Phenotypic characterization revealed the strain to be Gram negative, rod- shaped, and motile possessing a single polar flagellum, with a G+C content of 66%

(Macy 1993). The strain was found to grow aerobically or anaerobically when selenate or nitrate was present as a terminal electron acceptor. In the process, selenate was reduced to selenite, and nitrate was reduced to N2O.

A number of Thauera species were subsequently isolated by enrichment on

aromatic hydrocarbons under anaerobic conditions. These newly described members of

the genus Thauera have been isolated anoxically on such diverse compounds as 4- hydroxyphenylacetate, cyclic oxygenated monoterpenes, landfill leachate, resin acids, and BTEX compounds (benzene, toluene, ethyl benzene, and xylenes) in the presence of nitrate (Anders 1995, Evans 1991, Foss 1998, Mohn 1999, Scholten 1999, Song 1998).

Strain T1 was isolated anaerobically using toluene as a sole carbon source and nitrate as the terminal electron acceptor (Evans 1991). This strain was later named T. aromatica along with a proposal to modify the Thauera generic description to include the

peritrichous and degenerately peritrichous flagellation as found in the new strain (Anders

1995, Song 1998). T. aromatica was subsequently found to utilize a wide variety of

aromatic hydrocarbons under denitrifying conditions (Anders 1995).

23 Research involving members of the genus Thauera continued to focus on the

anaerobic degradation of aromatic hydrocarbons using nitrate and/or selenate as the

terminal electron acceptor. Degradation of a variety of aromatic carbon compounds

proceeds to the formation of the central intermediate benzoyl-CoA, which is the product

of benzoic acid and coenzyme A (Figure II-2) (Bonting 1996, Gallus 1998, Heider 1997,

Schneider 1997). The reductive cleavage of benzoyl-CoA by benzoyl-CoA reductase is

the first step in a shared aromatic degradative pathway conserved among Thauera species

(Boll 1998, Boll 1997, Breese 1998, Coschigano 1997, Coschigano 1994, Coschigano

1998, Heider 1997, Heider 1998, Laempe 1999). This pathway is shown in Figure II-3.

Several of the genes in the benzoyl-CoA pathway serve a dual-purpose as they are also involved in the aerobic degradative pathways. In these cases, transcription of the majority of these genes is up regulated in the presence of their respective substrates, regardless of the presence or absence of molecular oxygen (Heider 1998).

Of the enzymes specific for anaerobic degradation of aromatic hydrocarbons, no enzyme activity was detected under aerobic conditions (Heider 1998). Benzoyl-CoA reductase production was also found to be strongly down regulated under oxic conditions.

Small quantities of the inactive enzyme, however, were found in aerobically grown cultures (Heider 1998). Furthermore, benzoyl-CoA reductase has been shown to be rapidly inactivated in the presence of oxygen (Heider 1998).

There are no published reports that describe production of mono- or polysaccharides in Thauera species. Similarly, there are no publications specifically dealing with cellular aggregation or flocculation in these bacteria. Two reports, however, have noted the aggregation of different Thauera species in pure culture during late log or

CH 3 CH2OH COOH Aniline CH2 p-Cresol Benzyl Alcohol NH2 Phenyl- acetate OH

CHO COOH COOH CHO CO SCoA

CH2 Phenol OH OH Cl 4-Amino- NH2 benzoate COOH COOH CO SCoA COOH

4-hydroxybenzoate Benzoate CO

OH NH2 CO SCoA Phenylglyoxylate CO SCoA

HOOC Benzoyl-CoA

CH3 COOH

Toluene Benzylsuccinate

Figure II-2. Anaerobic degradation of aromatic hydrocarbons. Anaerobic degradation of a variety of aromatic hydrocarbons by Thauera sp. proceed through the common intermediate, benzoyl-CoA (Heider, 1997).

CoAS l-CoA lohex-1-

C c O 6-Oxocy ene-1carbony NADH + H adation of aromatic hydrocarbons in l-CoA ly e NAD CoAS HOOC

C pim

HO l-CoA O clohex- CoAS droxy

cy HO C 3-Hy

O droxy

6-Hy 1-ene-1carbony O 2 2 H Diagram of the pathway for degr

CoAS l-CoA 2 a-1,5-diene- -CoA C l O clohexy 3 Acety + 1 CO Cy 1-carbony +2Pi 2ADP P T 2H +2A species. Adapted from Breese, 1998. CoAS C l-CoA

O Figure II-3. Benzoyl-CoA pathway. Thauera

Benzoy

26 stationary phase (Foss 1998, Song 1998). For example, T. aromatica strain K 172T was found to display a diverse morphology and to exhibit clumped growth in liquid culture.

This clumped growth phenotype, however, was not found in T. aromatica strain T1 (Song

1998). The mechanism of this aggregation, the extent to which the phenotype is distributed among species within the genus, or its role in the microbial ecology of these organisms have not been described.

K. Goals of this Research

The most common secondary wastewater treatment process in the world is the activated sludge process (Jenkins 1993). Problems associated with this process, therefore, have global importance. In spite of this, the current understanding of the mechanisms resulting in viscous bulking and poor dewaterability are not well delineated, and even less is known about the causative organisms. The importance of this research is further strengthened by the narrow and limited focus of all previous work involving this relatively new group of bacteria.

Although Zoogloea species have been associated with slime production in activated sludge, and viscous bulking has even been called “zoogloeal bulking”

(Eikelboom 1981), no direct correlation has been reported between the presence of a specific strain of Zoogloea and episodes of viscous bulking or poor dewaterability (Unz

2002). The positive correlation of MZ1T with these conditions in the wastewater treatment plant of Eastman Chemical Company represented the first in-depth investigation and rigorous identification of a bacterium closely associated with these problems. Still, nothing was known of the role of MZ1T or any other organism in these

27 activated sludge system upsets, or the exact mechanisms behind them. Viscous bulking and poor dewaterability of activated sludge have been described as being associated with the presence of large quantities of extracellular slime, however the term “slime” in these cases is often not further clarified, and could be composed of very different extracellular compounds (e.g. protein, EPS) (Jenkins 1993).

Though not discussed elsewhere, carbohydrates, along with fatty acids, nucleic acids and proteins, make up the major classes of compounds comprising all living organisms. The study of carbohydrates as polysaccharides, however, has been limited by a lack of techniques for their amplification and analysis as compared with those for other biopolymers such as polypeptides or polynucleotides (Venkataraman 1999). This discrepancy is further exacerbated by the sheer complexity of polysaccharides relative to proteins or nucleic acids. For example, a simple disaccharide composed of only glucose can be linked from any carbon of the first monomer to any carbon of the second via a glycosidic bond resulting in 36 possible connections. This number is doubled when considering that linkages may be either α or β, and raised by an additional factor of 4 if one considers the possible D- or L- configurations of the individual monosaccharides.

The resulting disaccharide composed of the same hexose monomer can therefore have 72 possible configurations. When including the number of possible monosaccharides, the different organic and inorganic substituents, and their possible sites of substitution, several million possible structures could be generated from an oligomer containing fewer than ten monosaccharides.

Even with these complications, bacterial polysaccharides have been shown to be of great commercial importance (Sutherland 1986, Sutherland 1998). Applications for

28 these polymers are found in the food, manufacturing, medical, and chemical industries

(examples of which are listed in Table II-1). Given the relatively few bacterial polysaccharides that have been described relative to the number of bacterial species, continued investigation in this field will undoubtedly result in a wide variety of new polymers and applications.

Based on these factors, this work was designed to investigate the possible mechanisms by which MZ1T might result in viscous bulking and poor dewaterability of activated sludge wastes at Eastman Chemical Company. The following research hypotheses were proposed:

1. MZ1T produces an extractable exopolysaccharide.

2. The EPS of MZ1T possesses the same chemical functional groups as that

produced by Z. ramigera ATCC 19544.

3. EPS is involved in the aggregation of MZ1T cells into clusters and flocs.

To test these hypotheses, a method for the reproducible production and extraction of extracellular material from MZ1T cultures was developed. The extracted material was then subjected to rigorous chemical and spectroscopic analyses to determine its composition and partial structure. These data were then compared with the available literature describing the EPS of zoogloeal cluster and/or floc-forming strains in an effort to delineate key components necessary for this phenotype. Finally, mutants deficient in floc-formation were generated from MZ1T and their EPS extracts were analyzed in an effort to elucidate what components of the EPS, if any, are involved in cellular aggregation MZ1T.

Table II-1: Origins and applications of bacterial exopolysaccharides. (Source: Sutherland, 1998)

Polymer Source Uses/Properties

Xanthan Xanthomonas campestris Suspending agent in foods, pesticides, etc Viscosity control in oil-drilling ‘muds’ Emulsion stabilization in foods, thixotropic paints Foam stabilization in beer, fire fighting fluids Viscosity control in jet printing applications Oligosaccharide preparation

Bacterial Acetobacter xylinum, Wound dressings cellulose Agrobacterium sp. Acoustic membranes

Gellan Sphingomonas Gelling agent in foods, biotechnology paucimobilis Oligosaccharide preparation

Curdlan Agrobacterium sp. Gelling agent in foods, biotechnology Rhizobium sp. Blockage of permeable zones during oil recovery

Dextran Acetobacter sp. Source for dextran-derivatives (e.g. Sephadex) Leuconostoc mesenteroides

Alginate Pseudomonas aeruginosa Gel formation in presence of Ca2+, potential use in food, Azotobacter vinelandii biotechnology, and textile printing applications

Pullulan Pullularia pullulans Film formation and food coatings Oligosaccharide preparation

Hyaluronic acid Streptococcus equi, Eye and joint surgery St. zooepidemicus Hydrating agent in cosmetics and pharmaceuticals

30 Results of this work clearly revealed a fundamentally new and novel

polysaccharide polymer from a relatively unknown class of microorganisms. The EPS

was shown to possess characteristics that may make it potentially useful in industrial

applications (e.g. metal binding). It has also been shown to contain certain common

chemical components with Z. ramigera ATCC 19544 which can be used to draw

conclusions on the mechanisms of cluster formation. EPS extracts from floc– mutants were similarly analyzed and found to possess conserved alterations in their absorbance spectra, though they maintain the same glycosyl compositions. Finally, the floc–mutant

colonies responded differently to a variety of dyes and mutants were found to readily

accept plasmids by conjugal transfer, suggesting the mutants possessed an alteration in

the organization of their cell surfaces. These discoveries have resulted in the creation of

a platform from which future genetic and physiological studies of Thauera MZ1T and

related strains may proceed.

31 CHAPTER III

MATERIALS AND METHODS

A. Bacterial plasmids and strains

Thauera sp. strain MZ1T was originally isolated by Lajoie et al. by

micromanipulation of zoogloeal clusters from activated sludge samples taken from the

wastewater treatment plant of Eastman Chemical Company in Kingsport, Tennessee

(Lajoie 2000). Because MZ1T was not known to have any readily distinguishable

phenotypic characteristics, its 16S rDNA gene was amplified by PCR, cloned into the

TOPO pCR2.1 TA cloning vector (Invitrogen), and sequenced to verify the use of the

proper strain. Following verification, a culture of this strain was preserved for all

subsequent experiments.

Spontaneous rifampicin-resistant mutants of MZ1T were isolated from Stoke’s

agar plates containing 100-µg/mL rifampicin. Colonies isolated from the selective plates

were grown in 10 mL of Stoke’s broth containing 100-µg/mL rifampicin. Colonies were

verified to be MZ1T by 16S rDNA sequence. A stock was prepared of a single strain and all subsequent experiments utilized this spontaneous rifampicin-resistant mutant unless otherwise specified. All bacterial strains and plasmids used in this study are listed in

Table III-1.

B. Culture conditions and storage

All MZ1T strains were grown at 30°C unless otherwise noted, in either Stoke’s medium or Thauera Defined Medium (TDM). All E. coli strains were grown at 37°C 32 Table III-1: Bacterial strains and plasmids used in this study. Strains Relevant Characteristics Source

MZ1T Original Floc+ isolate Lajoie, 2000 MZ1TR Floc+, Rifampicin resistant (rifr) MZ1T This work MZ1T-20A Floc– NTG mutant of MZ1TR This work MZ1T-26B Floc– NTG mutant of MZ1TR This work MZ1T-27A1 Floc– NTG mutant of MZ1TR This work MZ1T-30B1 Floc– NTG mutant of MZ1TR This work MZ1T-35B1 Floc– NTG mutant of MZ1TR This work MZ1T-37B Floc– NTG mutant of MZ1TR This work MZ1T-39A Floc-reduced NTG mutant of MZ1TR This work MZ1T-44B1 Floc-reduced NTG mutant of MZ1TR This work MZ1T-118 Floc– Transposon mutant of MZ1T-39A This work MZ1T-142 Floc– Transposon mutant of MZ1T-39A This work MZ1T-153 Floc– Transposon mutant of MZ1T-39A This work MZ1T-171 Floc– Transposon mutant of MZ1T-39A This work MZ1T-167 Floc– Transposon mutant of MZ1T-39A This work MZ1T-295 Floc– Transposon mutant of MZ1T-39A This work MZ1T-296 Floc– Transposon mutant of MZ1T-39A This work MZ1T-309 Floc– Transposon mutant of MZ1T-39A This work MZ1T-377 Floc– Transposon mutant of MZ1T-39A This work MZ1T-∆39 Floc– Transposon mutant of MZ1T-39A This work MZ1T-λ18 Floc– Transposon mutant of MZ1T-39A This work MZ1T-λ35 Floc– Transposon mutant of MZ1T-39A This work MZ1T-θ18 Floc– Transposon mutant of MZ1T-39A This work MZ1T-θ40 Floc– Transposon mutant of MZ1T-39A This work Escherichia coli JM109 Promega

E. coli HB101 HsdS20(rB-mB-), recA13 Lacks, 1977

E. coli DH5α HsdR17(rB-mB+), recA1 Invitrogen E. coli INVF’ Strain used with TA cloning vector pCR2.1 Invitrogen Thauera aromatica Type strain for genus Thauera ATCC 700265

Plamids Relavent Genotype/Characteristics Source pCR2.1 TA cloning PCR vector Invitrogen pBSHR1 Broad-host-range vector for Gram – bacteria, tcr Hay, 2000 pRK2013 RK2-transfer gene-containing helper plasmid, tcr Figurski, 1979 pRK290 Broad-host-range vector for Gram – bacteria, tcr Ditta, 1980 pRK415 Broad-host-range vector for Gram – bacteria, tcr Keen, 1988 pTNMod-OKm Transposon delivery vector with integral origin of Dennis and Zylstra, Replication, kmr 1998 p142P Plasmid isolated following PstI digest of MZ1T-142, kmr This work p153P Plasmid isolated following PstI digest of MZ1T-153, kmr This work p171P Plasmid isolated following PstI digest of MZ1T-171, kmr This work

33 except in mating experiments with MZ1T, which were cultured at 30°C. Liquid cultures

were grown shaking in 250 mL Erlenmeyer flasks at 150 r.p.m.

Freezer stocks were prepared by adding 0.5 mL samples from an overnight

culture (48 hr. culture for MZ1T) with appropriate antibiotics to sterilized tubes

containing 0.5 mL sterile 50% (v/v) glycerol. Tubes were maintained frozen at –80°C

until ready for use.

C. Media and chemicals

E. coli strains were routinely cultivated in autoclave sterilized Luria-Bertani (LB)

broth (10.0 g tryptone, 5.0 g yeast extract, 10.0 g NaCl per liter, pH 7.5). Filter-sterilized

(0.2 µm) antibiotics were aseptically added to LB broth as necessary. LB agar plates

were made by the addition of 17 gL-1 of agar prior to autoclave sterilization.

Stokes’s broth was used for the routine cultivation of MZ1T strains. One liter of

Stoke’s medium contained: 5 g polypeptone, 0.2 g MgSO4⋅7H2O, 0.15 g Fe(NH4)(SO4),

0.1g sodium citrate, 0.05 g CaCl2, 0.05 MnSO4, 0.01 FeCl3⋅6H2O. The pH of the medium

was then adjusted to 7.2 by the addition of a 5M NaOH solution. For agar plates, 15 gL-1

agar was added to Stoke’s medium. Media solutions were sterilized by autoclaving at

121°C and 15 p.s.i. for 20 minutes, solutions were allowed to cool below 50°C and the

following filter-sterilized vitamin solutions were added to yield final concentrations as indicated: cyanocobalamin, 0.5 mg/L; thiamine hydrochloride, 0.4 mg/L; and biotin, 0.4 mg/L. All vitamin stock solutions were stored in the dark at 4°C.

Thauera defined medium (TDM) was used for the cultivation of MZ1T and mutant strains for all EPS isolation experiments. TDM is a modified version of the 34 medium used by Rabus and Widdel (Rabus 1995). TDM basal medium is composed of:

0.3 g NH4Cl, 0.5 g KH2PO4, 0.5 g MgSO4 ⋅ 7H2O, 0.1 g CaCl2 ⋅ 2H2O, and a carbon source in 1 L of deionized water. Unless otherwise noted, the carbon source for all experiments was 5 g of sodium succinate. This solution was autoclaved at 121°C for 20 min. at 15 p.s.i. and then allowed to cool to below 50°C before the following filter- sterilized stock solutions were aseptically added: 1 mL Trace Element Solution, 1 mL

Tungsten solution, 1 mL Vitamin solution, 1 mL Thiamine solution, 1 ml

Cyanocobalamin solution, and 30 mL of sodium bicarbonate solution.

Trace Element Solution was composed of: 2.1 g FeSO4 ⋅ 7H2O, 5.2 g Na2EDTA,

30 mg H3BO4, 100 mg MnCl2 ⋅ 4H2O, 190 mg CoCl2 ⋅ 6H2O, 24mg NiCl2 ⋅ 6H2O, 25 mg

CuSO4 ⋅ 5H2O, 144 mg ZnSO4 ⋅ 7H2O, 36 mg Na2MoO4 ⋅ 2H2O and 1 L deionized water.

The pH was adjusted to 6.0-6.5 and the solution was sterilized in an autoclave as

described.

The Tungsten solution contained 200 mg NaOH and 6mg Na2WO4 ⋅ 2H2O in 1 L of deionized water. The solution was sterilized in the autoclave as described.

The Vitamin Solution contained 200 mL 20 mM sodium phosphate buffer

(pH 7.1), 8 mg p-aminobenzoic acid, 2 mg D-biotin, 20 mg nicotinic acid, 10 mg calcium

D-pantothenate, and 30 mg pyridoxin hydrochloride. This solution was filter-sterilized

through a 0.2 µm filter and stored in the dark at 4°C.

The Thiamine solution was made by adding 20 mg thiamine hydrochloride to 200

mL of sodium phosphate buffer, pH 3.4. This solution was filter-sterilized through a 0.2

µm filter and stored in the dark at 4°C.

35 The Cyanocobalamin (B12) solution contained 5 mg cyanocobalamin in 100 mL of deionized water. This solution was filter-sterilized through a 0.2 µm filter and stored in the dark at 4°C.

The Sodium Bicarbonate Solution was contained 84 g of sodium bicarbonate 1 L of deionized water. This solution was filter-sterilized through a 0.2 µm filter and stored at room temperature in a sealed container.

Sugar standards for analytical experiments were purchased from Sigma (St. Louis,

MO) except for N-acetyl-D-fucosamine (2-acetamido-2,6-dideoxy-D-galactose), which was a gift from D. Horton. All chemicals and reagents used for glycosyl composition determination were of the highest analytical grade and purchased from Sigma unless otherwise specified. Standard media components were purchased from Fisher Scientific

(Pittsburgh, PA) or Sigma. All vitamins used in media preparations were purchased from

Sigma. Nitrogen and ultra-high purity nitrogen gas were purchased from National

Welders Supply (Charlotte, NC).

D. Molecular Biology Techniques

All molecular biology techniques were performed as outlined in Sambrook et al. unless otherwise noted (Sambrook 1989). All enzymatic reactions and routine purification of nucleic acid involving kits were performed as indicated in the manufacturer instructions. All DNA modifying enzymes were obtained from Promega

(Madison, WI) or Fisher. All PCR reactions were conducted using Ready-To-Go PCR beads (Amersham Biosciences, Piscataway, NJ), unless otherwise specified, on a MJ

Research PTC 200 Peltier thermal cycler (Waltham, MA). 36

E. DNA sequencing

All DNA sequencing was performed by the Molecular Biology Resource Facility at the University of Tennessee, Knoxville, using an ABI Prism 310 Genetic Analyzer (PE

Applied Biosystems, Foster City, CA) with BigDye Terminator Cycle Sequencing v2.0

(PE Applied Biosystems).

F. BOX polymerase chain reaction (PCR)

Because of the lack of readily distinguishable phenotypic characteristics of MZ1T and the risk of cross contamination with rifampicin resistant bacteria, BOX PCR was sometimes used to verify the presence of MZ1T or mutant strains (Louws 1994). When separated on an agarose gel, the pattern of the multiple bands produced in the PCR reaction could be used to differentiate between MZ1T and E. coli or contaminating bacteria. The single primer used for box PCR had the following sequence:

5’-CTA CGG CAA GGC GAC GCT GCT GAC G-3’

The conditions for the PCR reaction were 94°C for 4 min, followed by 35 cycles of

92°C for 45 s, 50°C for 1 min, and 65°C for 6 min.

G. Ribosomal intergenic sequence analysis (RISA)

For the same reasons as described previously for BOX PCR, ribosomal intergenic sequence analysis (RISA) was sometimes used for the routine identification of MZ1T and mutant strains (Gürtler 1996). The PCR conditions for RISA were as follows: 94°C for

37 1 min, followed by 25 cycles of 94°C for 15 s, 56°C for 30 s, and 72°C for 1 min. Primer

sequences used in the reaction were:

1492 forward, 5'-AAGTCGTAACAAGGTA-3' and

23S reverse, 5'-GGGTTBCCCCATTCR-3'.

Following amplification, double restriction digests of the PCR products were

performed. Digest reactions included: 16 µL PCR reaction product, 1.0 µL HhaI, 1.0 µL

RsaI, and 2.0 µL Buffer C (Promega). On some occasions, the enzymes AlaI and MspI with Buffer B (Promega) were substituted or used in an additional reaction. All restriction enzyme concentrations were 10 U/µL, and reactions were allowed to incubate at 37°C from 4 hours to overnight.

H. Determination of doubling time

Growth curves and doubling times were determined based on the increase in optical density at 600 nm (OD600) over time of liquid cultures grown in Stoke’s broth.

OD600 measurements were made using a Beckman Coulter DU-640B spectrophotometer

(Fullerton, CA). Specific growth rate was calculated from the slope of the natural log of

OD600 measurements corresponding to the linear portion of the curve during exponential growth phase, typically between 10 and 16 hours. All experiments were conducted in triplicate. The average slope was used for the mathematical calculation of generation time and the standard deviation for the replicates is reported.

I. Carbon-source utilization

38 Various compounds were tested to determine which among them could be used by

MZ1T as sole carbon sources. Batch experiments were conducted in 100 mL cultures in

250 mL Erlenmeyer flasks in which various carbon compounds were added to the TDM

basal medium at concentrations of 5 g/L. All experiments were conducted under aerobic

conditions unless otherwise indicated. Anaerobic conditions were generated using

Oxyrase (Mansfield, OH) following the manufacturers instructions.

Biolog GN2 microplates (Biolog, Hayward, CA) were also used to screen carbon utilization patterns for MZ1T and Thauera aromatica. Overnight cultures grown in

Stoke’s medium were pelleted by centrifugation, washed two times, and resuspended in carbon source-free TDM to an OD600 of 0.3. 150 µL aliquots of this cell suspension were

transferred to each of the 96 wells of the GN2 microplates. Experiments were conducted

aerobically and in triplicate for both MZ1T and T. aromatica.

J. Carbohydrate concentration determination

Routine determinations of carbohydrate presence or concentration were made

using a modified phenol/sulfuric acid assay (Ames 2000) based on the method of (Dubois

1956). Briefly, 1 mL of sample solution was mixed with 2 mL of 4% phenol (w/v) and

then 5 mL of concentrated sulfuric acid were added. The solutions were mixed by vortex

and allowed to stand at room temperature. After 20 minutes, absorbance measurements

were taken at 488 nm with a DU-640B spectrophotometer (Beckman Coulter). A

standard curve using glucose is shown in Figure III-1.

1.4 Slope = 0.010 1.2 r ² = 0.974

1.0

0.8 ance b

sor 0.6 b A 0.4

0.2

0.0 0 20406080100120 Glucose

Figure III-1. Carbohydrate assay standard curve. Standard curve for the relationship between carbohydrates (as µg glucose) and absorbance using the phenol-sulfuric acid assay. Error bars represent the standard deviations of triplicate samples. The slope of the regression line (solid) is 0.010 with R2 = 0.974. Dashed lines indicate 95% confidence intervals.

40 K. Protein concentration determination

Protein concentrations were determined using the Bio-Rad Protein Assay (Bio-

Rad) as outlined by the manufacturer. Briefly 100-µL samples of MZ1T culture grown in

TDM were added to 5 mL 1:4 assay reagent:deionized water. Samples were mixed with

a vortex for 1 minute and allowed to stand undisturbed at room temperature for 30

minutes. Following incubation, samples were analyzed for absorbance at 595 nm in a

DU-640B spectrophotometer. Standards were prepared using 0, 0.25, 0.5, 0.75, and 1.0

mg/mL bovine serum albumin (BSA) in TDM. All experiments were conducted in

triplicate. A standard curve is shown in Figure III-2.

L. Isolation and purification of exopolysaccharide

1-L cultures of MZ1T grown in TDM and shaken at 30°C for seven days were centrifuged for 15 min. at 8000 × g to pellet the cells. The supernatant fraction was decanted through a nylon mesh cloth to remove any cellular debris, and then concentrated to one-tenth its original volume (100 mL) using a Pellicon 2 Mini Holder filtration

apparatus equipped with a 100-kDa MWCO membrane (Millipore). The retentate, which

contained the putative polysaccharide, was then diluted back to 1 L with deionized water

and the process repeated for a total of three times. The resulting desalted polysaccharide

solution was then frozen at -80°C and lyophilized to yield the crude preparation. For

most analyses, crude, lyophilized polymer was dissolved in 10 mL of 0.25 M ammonium

formate buffer and further purified by liquid chromatography (LC) using a 1 m × 2.5 cm

column packed with fine mesh Bio-Gel P30 resin (Bio-Rad). Flow rate was

0.6 Slope = 0.0499 r ² = 0.971 0.5

0.4 ance

b 0.3 sor b

A 0.2

0.1

0.0 024681012 BSA Figure III-2. Protein assay standard curve. Standard curve for the relationship between bovine serum albumin BSA (in µg) and absorbance using the Bio-Rad Protein Assay. The error bars represent standard deviations of triplicate samples. The slope of the regression line (solid) 0.0499 with R2 = 0.971. Dashed lines indicate 95% confidence intervals.

42 approximately 4 mL/hr. using 0.25 M ammonium formate buffer. Fractions were

monitored with an inline UV absorbance detector at 254 nm and putative polysaccharide-

containing fractions were combined, frozen at -80°C, and lyophilized to remove the ammonium formate buffer. For trimethylsilyl methyl glycoside (TMS) analysis, polymer purified by filtration was lyophilized and further purified by liquid chromatography (LC) using a 60 × 5 cm column in series with a 60 × 3 cm column packed with BioGel P-100

(Bio-Rad Labs). The void volume of the column was estimated to be ~300 mL.

Approximately 30 mg of the lyophilized polymer was dissolved in 7 mL of deionized water and loaded onto the column, which was then gravity-eluted with deionized water at an average flow rate of 0.3 mL/min. Total solids recovery was approximately 70% (21 mg) of the original sample mass.

M. Size Determination of EPS polymer

Size determination of the polysaccharide by gel-permeation chromatography was performed by V-Labs, Inc. (Covington, LA) using 5 TSK-Gel PWXL columns arranged in series with 9:1 0.15 M NaCl:acetonitrile, and a flow rate of 6 mL/min. Sodium polystyrene sulfonate standards were used as a reference.

N. Physical properties of MZ1T EPS

Solubility of EPS was determined by the addition 1.5 mL of various solvents to 1 mg EPS, vortexing for 30 sec. and allowing to stand overnight. Thermal stability was determined using a capillary tube packed with EPS on a melting point apparatus.

43 Carbon-Hydrogen-Nitrogen content for the purified EPS was determined by

Eastman Chemical Company using a Carlo-Erba Model 1106 Elemental Analyzer

following the manufacturers instructions and analyzed with the accompanying Eager 100

software.

O. Metal binding by EPS

Uranyl nitrate

A stock solution of uranyl nitrate was prepared containing 100 mg UO2(NO3) in

50 mL deionized water. One mL of uranyl nitrate was added to triplicate 1 mL solutions

containing 10 mg crude EPS in deionized water. Triplicate controls were prepared by the

addition of 1 mL uranyl acetate to 1 mL of deionized water. Both experiment and control

samples were vortexed for 30 sec. and allowed to sit undisturbed for 5 min. Each

solution was transferred to a pre-washed Millipore (Waltham, MA) Ultrafree centrifugal

filter column containing a Biomax 30–50kDa MWCO membrane and centrifuged at 5000

× g for 1 hr. 1.5 mL of from each tube was added to 10 mL ReadySafe scintillation

cocktail (Beckman) and disintegrations were counted in a LS 6000 Series Scintillation

System (Beckman).

To test the ability of MZ1T EPS to remove uranyl nitrate from non-aqueous solutions, a stock solution was prepared by adding 100 mg UO2(NO3) to 50 mL

methanol. Two mL of this stock solution was then added in triplicate to empty

Eppendorf tubes (Eppendorf, Westbury, NY) (negative controls), tubes containing 8 mg

crude EPS, or tubes containing 8 mg dextran sulfate. Samples were placed in a 30°C

shaking incubator (160 r.p.m.) for two hours. EPS was pelleted from solution by

44 microcentrifugation for 2 min. at 10,000 r.p.m. and 1.5 mL of supernatant was analyzed in a scintillation counter over ten minutes as previously described.

Calcium chloride

Aqueous solutions containing 0.0, 0.01, 0.05, or 0.1 M CaCl2 were added to test tubes containing 0.5 mg crude EPS. Tubes were mixed by vortex for 30 seconds and allowed to stand undisturbed overnight at room temperature. Depth of the precipitate was then measured the following day.

P. Alditol acetates

Alditol acetate samples were prepared as described previously (York 1985).

Individual 1-mg samples of EPS were placed in Pyrex screw cap test tubes and 10 µl of a

2-µg/µl inositol solution was added to each tube as an internal standard. 0.5 mL of 2M- trifluoroacetic acid (TFA) was added to each sample-containing tube. The tubes were sealed with Teflon lined screw caps and the samples were heated at 121°C for 3 hours in a heating block to hydrolyze the polysaccharides. Samples were removed from the heating block and allowed to cool before being dried under a stream of ultra high purity

(UHP) nitrogen gas to remove the acid. Residual acid was removed by the addition and subsequent drying of 250 µL isopropanol and then repeated a second time. The dried, hydrolyzed samples of free monosaccharides were dissolved in freshly prepared 0.25 mL of 1 M ammonium hydroxide solution (Fluka, Ronkonkoma, NY) containing 10-mg/mL sodium borodeuteride (Sigma). The samples were capped and incubated at room temperature for 1 hr. Following reduction, residual sodium borodeuteride as reacted with 45 glacial acetic acid to form borate. Glacial acetic acid was added dropwise until bubbling ceased. Borate was removed by the addition of 0.5 mL of methanol:glacial acetic acid

(9:1) followed by drying under UHP N2. This process was repeated for a total of three

times. Following the third time, solvent was switched to methanol (100%) and dried to

completeness for a total of three additional times. Following reduction of the

monosaccharides, the samples were acetylated as follows: 250 µl of pyridine was added to each tube followed by the addition of 250 µl of acetic anhydride. Samples were capped and heated at 50°C for 10 min. After acetylation, samples were removed from the heating block and allowed to cool. 1 mL of toluene was added to each tube and the samples were dried under UHP N2. 1 mL of 0.2 M sodium carbonate was added and each

tube was swirled to neutralize any residual acid. One mL of dichloromethane (Sigma)

was added to each tube and the samples were agitated to extract the alditol acetates. The

top, aqueous layer was removed and discarded, and the process was repeated two more

times by the addition of 1 mL HPLC-grade water (Fisher). On the last wash, the samples

were allowed to stand ten minutes and then the non-aqueous bottom phase was removed

by pipette and transferred to a new screw-cap test tube. Samples were dried under UHP

N2 and then dissolved in 100 µl acetone. Freshly dissolved samples were immediately

transferred to GC vials, capped with Teflon-lined caps, and held until being analyzed by

gas chromatography/mass spectroscopy.

Positive controls for alditol acetate samples were prepared by adding 10 µl of 10

µg/µl solutions of sugar standards to test tubes containing 20 µg inositol. Negative controls were prepared which contained only inositol. Controls were treated identically

46 and at the same time as experimental samples. A gas chromatogram of the alditol acetate

derivatives generated from the combined monosaccharide standards is shown in Figure

III-3.

Q. Per-O-trimethylsilyl methyl glycosides (TMS)

1 mg EPS samples were transferred to Pyrex screw cap test tubes containing 10 µl of a 2 µg/µl inositol solution. These samples were frozen at –80°C for a minimum of 30 minutes and then dried under vacuum on a Freezone 6 lyophilizer (Labconco, Kansas

City, MO) for at least 3 hours to remove residual water. 500 µL of 1 M hydrogen chloride in anhydrous methanol (methanolic HCl) was added to each tube, which was then flushed with UHP N2 and sealed with a Teflon-lined screw cap. Tubes containing

samples were transferred to a heating block and held at 80°C for 16 hours for hydrolysis

of the polysaccharides.

Methanolic HCl was prepared using the Instant Methanolic Kit (Alltech, State

College, PA). Briefly, a 250 mL Pyrex two-neck flask that had been pre-baked at 300°C to remove residual water was allowed to cool and then one neck was outfitted with a stopper containing a glass pipette connected to a UHP N2 tank. With nitrogen gas

flowing, the flask was transferred to an ice bath on a stir plate and a Teflon-coated stir bar was added. 5 mL of anhydrous methanol was added and allowed to cool for a few minutes. 700 µL of acetyl chloride was added drop wise to the continuously stirring, N2- flushed flask. Following the reaction, the reagent was transferred to N2-flushed, screw

. s d r a GlcNAc d n a t s

e d i r a h c c itol; Glc, glucose; and Glc a s o Inos n o m

f o

s e v i t a v i r e d

e t a t e c -acetylfucosamine; Inos, inos a

Rha l N

o , t c i d A l a N

c e u h t F

f ; o e

s o m a e. n r m g a o t h a r

, m a o h r h R

c glucosamin :

l e s r a a

G s

n . -acety o 3 i - N t I a I i I

v e e r r u b g b i A F

GlcNAc,

48 cap test tubes. Tubes of reagent that were not used immediately were stored at –20°C in

a sealed bag containing Dry-rite for up to three months.

Following hydrolysis, samples were allowed to cool and then dried under a flow

of UHP N2 gas placed in a blow-down apparatus held in a water bath that was maintained

at 40-45°C. Residual acid was removed by the addition of 250 µL methanol followed by

drying under UHP N2 two times. Aminosugars were N-acetylated by the addition of 200

µL methanol, 20 µL pyridine, and 20 µL of acetic anhydride (Sigma). Samples were then incubated at room temperature for six hours. After N-acetylation, samples were dried under UHP N2. A 200 µL aliquot of Tri-Sil TMS reagent (Pierce, Rockford, IL) was then

added to each tube, which was then placed in a heating block at 80°C for 20 minutes.

Samples were removed, allowed to cool, and flushed with UHP N2 until just dry. Dried samples were dissolved in 100 µl hexane, transferred to GC vials containing glass inserts, immediately capped with Teflon-lined aluminum caps, and held at room temperature until analyzed by gas chromatography/mass spectroscopy. Positive and negative controls were prepared as described except for the following: 1) for positive controls, 10 µL of a

10-µg/µL solution of each sugar standard was added plus 10 µl of a 2-µg/µL solution of inositol 2) and for negative controls, only the inositol was added. A gas chromatogram of the TMS derivatives generated from the combined monosaccharide standards is shown in

Figure III-4.

49 r peaks Glc, ;

. mine e majo a lfucos -acety N amine. Only th -acetylglucos N derivatives of monosaccharide standards GalA, galacturonic acid; FucNAc, rnal standard; and GlcNAc, rhamnose; s are: Rha, Gas chromatogram of the TMS n Figure III-4. Abbreviatio glucose; Inos, Inositol inte are labeled for clarity.

50 R. Gas chromatography–mass spectrometry (GC–MS)

Derivatives were analyzed on a HP 6890 Series gas chromatograph (Hewlett-

Packard, Palo Alto, CA) outfitted with a DB-5MS capillary column (J&W Scientific,

Folsom, CA; 30m × 0.25 mm i.d.; 0.25 µm film thickness) and an HP 5973 mass

selective detector in the scan mode. Helium was used as the carrier gas at flow rate of 1

mL/min. For both alditol acetate and TMS methyl glycoside derivatives, the following

temperature profile was used: Initial temperature, 150°C for 2 min, then increasing at

2°C/min to a final temperature of 240°C where it was held for 3 min. A post run

temperature of 300°C was maintained for 3 min. before injection of subsequent samples.

S. Nuclear Magnetic Resonance (NMR) Spectroscopy

Unless otherwise mentioned, 1 mg of EPS sample was dissolved in 1 mL of D2O, to which 2 µL of deuterated trifluoroacetic acid (TFA-d) was added. Samples were

lyophilized and dissolved in 1 mL D2O and 2 µL of TFA-d two times, and then dissolved in 650 µL of D2O for analysis. All experiments were run at 25°C, unless otherwise

noted, using a Varian Inova 600 MHz (Palo Alto, CA) spectrometer. All spectra were

acquired and processed using standard Varian software (VNMR 6.1B). 1H and 13C chemical shifts were referenced to acetone (δH 2.225, δC 31.08). Total correlated

spectroscopy (TOCSY) spectra were acquired at mixing times of 20, 50, 80, and 150 ms.

Heteronuclear spin quantum-coupled (HSQC) and gradiant correlated (gCOSY) spectra

were generated with 269 ms. acquiring times. NOE spectra (NOESY) were generated

with mixing times of 90 ms.

T. Fourier-transform infrared spectroscopy (FTIR)

Samples containing 1.5 mg of purified EPS were dissolved in 100 µl water and applied to Janos ECRAN IR screen cells. Cells were dried under vacuum desiccation for

72 hours prior to analysis. Spectra were generated on a Bio-Rad FTS-60A Fourier- transform infrared spectrophotometer (Hercules, CA) using single-beam transmittance with 16 scans/s resolution.

U. Conjugation

Bacterial matings were carried out essentially as described by Coschigano et al.

(Coschigano 1994). Briefly, 10–mL cultures of donor, recipient, and helper strains when appropriate, were grown to an approximate optical density at 600 nm (OD600) of 0.5 in

Stoke’s broth with 100-µg/ml rifampicin for MZ1T or LB with suitable antibiotics for E. coli strains. Cells were centrifuged as described to form pellets, the supernatant fraction was decanted, and the cell pellets were suspended in an equal volume of phosphate buffered saline (PBS, pH 7.0). This washing was repeated twice and on the third time, donor, recipient, and helper strains were combined into a single tube and centrifuged together. The supernatant fraction was decanted, and the pellet was suspended in 1 mL

PBS. Aliquots containing 100 µL of cell suspension were applied to autoclave sterilized

25 mm, 0.2 µm filter disks on non-selective Stoke’s agar plates and incubated at 30°C for

48 hrs. Following incubation, filter disks were aseptically removed and transferred to sterile Eppendorf tubes. 1 mL of PBS was added to each tube and vortexed to suspend

52 cells. Aliquots containing 100 µL of cell suspension were then spread plated onto

Stoke’s agar plates containing 100-µg/mL rifampicin and a second appropriate antibiotic.

These plates were then incubated for 48 hrs. at 30°C and screened for the appearance of colonies.

V. Electroporation

Electrocompetent cells of MZ1T were prepared using the method outlined for

Psuedomonas putida CYM318 as outlined in the ECM 600 Electrocell Manipulator operator’s manual (BTX, San Diego) and stored at –80°C until ready for use. For the experiment, electrocompetent cells were thawed on ice and 1 µL of EZ::TN kanr

Transposome solution (Epicentre, Madison, WI) was added to the cells and gently mixed with the pipette tip. As a control, 1 µL of deionized water was added to a second reaction mixture. The solutions were then transferred to ice-cold, sterile, electroporation cuvettes with 2 mm gaps (Molecular BioProducts) and electroporated using an ECM 600

Electrocell Manipulator (BTX). Following electroporation, 1 mL of ice-cold Stoke’s broth was immediately added, and the cuvettes were incubated on ice for a minimum of

10 min. Cultures were then transferred to culture tubes, one additional mL of Stoke’s broth was added, and tubes were allowed to incubate shaking at 30°C and 150 r.p.m. for

12 hours. Following incubation, cultures were plated onto Stoke’s agar plates containing

50-µg/mL kanamycin and incubated at 30°C for 48 hrs.

53 Electrocompetent E. coli strains were prepared as described in the E. coli DH5α

protocol in the ECM 600 Electrocell Manipulator operator’s manual. Conditions for

electroporation were 2.5 kV using the R5 setting as outlined in the protocol.

W. Mutagenesis

Chemical mutagenesis on rifampicin resistant MZ1T was carried out using N- methyl-N’-nitro-N-nitrosoguanidine (NTG) from Sigma Chemical Co. in a manner similar to that outlined by Coschigano and Young (Coschigano 1997). Briefly, cells from frozen stock were grown in Stoke’s broth for 48 hours to mid-log phase. 10 mL of MZ1T culture was centrifuged at 5000 × g to pellet the cells. Cells were resuspended in 9.9 mL of 100mM sodium citrate buffer (pH 5.5) and 0.1 mL of NTG stock solution in the same buffer was added to yield a final NTG concentration of 100 µg/mL. Tubes were incubated without shaking for 30 minutes (Figure III-5). The cells were pelleted by centrifugation and washed three times with 100 mM potassium phosphate (pH 7.0).

Treated cells were serially diluted and plated on Stoke’s agar plates. Resultant colonies were picked and visually screened for the loss of floc-forming capacity in 10 mL culture tubes containing 2 mL Stoke’s broth.

X. Floc+/– screening assays

Staining solutions were prepared and 5 mL of staining solution was applied to

plates containing both floc+ and floc- strains of MZ1T. The solution was allowed to

remain on the plate for 5 min. and then gently rinsed with deionized water for 1 min.

Stains used and their concentrations are shown in Table IV-1. 54 Stoke’s agar plates for the experiments were generated by either spot inoculation into

rows of the same strains using 15 µL of culture broth, or by using 100 µL of a 50:50

mixture of floc+ and one floc- strain spread onto the plates. The plates were then

incubated for 48 hr. prior to testing. When using a mixture of floc+ and floc- strains,

replica plates were first generated to allow screening of individual colonies for floc

formation in broth tubes.

Y. MZ1T-39A transposon mutagenesis and insertion mapping

Transposon mutants of floc-deficient mutant MZ1T-39A were generated by

conjugal transfer of pTNMod-OKm (Figure III-6) as described. Kanamycin-resistant

transconjugants were then screened for complete loss of floc formation in Stoke’s broth.

Genomic DNA preparations from mutants containing the transposon TNMod-

OKm were digested using 100 ng genomic DNA, 5 µL of the appropriate buffer, 10U of enzyme, and brought to a total volume of 20 µL with HPLC-grade water (Fisher) (Dennis

1998). The digests were incubated at 37°C for 2 hrs.

Following digestion, the samples were heated to destroy the restriction enzymes as outlined by their respective manufacturers (typically 65°C for 15 min). Digested genomic DNA fragments were then self-ligated. For self-ligation reactions, the following were added to tubes containing 20 µL DNA digest solution: 2.7 µL T4 DNA ligase, 10

µL ligase buffer, and 67.3 µL HPLC-grade water. Ligation reactions were then incubated at 16°C overnight. DNA samples from ligation reactions were precipitated by the

Figure III-5. NTG-treated MZ1T kill curve. Semi-log plot of MZ1T cell number when exposed to 100 µg/mL NTG. Samples were taken at 15 minute intervals, washed two times with 100mM potassium phosphate (pH 7.0), serially diluted in PBS, and plated on Stoke’s agar plates. Cell number values after 75 minutes were too few to count (<20 colonies/ plate without dilution).

Table IV-1. Compounds and tests used to differentiate floc+ and floc- colonies. Compound/Test Concentration/Source Result Alcian Blue 1% in 95% ethanol Stain could be washed from floc+ colonies Sudan Black 5% in 95% ethanol Stain could be washed from floc- colonies Safranin Gram Stain Kit-BBL No difference Evans Blue 1% in 95% ethanol No difference Gram’s Iodine Gram Stain Kit-BBL No difference Copper sulfate 20% in H20 No difference Cobalt chloride 10% in H20 No difference India Ink Undiluted No difference Nigrisin Undiluted No difference Carbol Fuschin 1% in 95% ethanol No difference UV light N/A No difference Crystal Violet Gram Stain Kit-BBL No difference Calcofluor White As supplied Floc- colonies dissolved, floc+ unaffected Potassium 0.1M Floc- colonies dissolved, floc+ unaffected hydroxide

IR I MCS I pMB1 OriR

Tn5 t np

pTnModO-Km

5028 bp

kanR

MCS II RP4 OriT IR II

Figure III-6. Schematic map of the pTnModO-Km transposon delivery vector. Abbreviations are as follows: IR I, inside inverted repeat; IR II, outside inverted repeat; MCS I and II, multi-cloning sites; RP4 OriT, RP4 origin of transfer; pMB1 OriR, narrow- host range origin of replication from plasmid pMB1; kanR, aminoglycoside phosphotransferase; Tn5 tnp, transposase gene from transposon TN5.

58 addition of 10 µL 3.0M potassium acetate solution (pH 7.0) and 250 µL absolute ethanol, followed by overnight incubation at –20°C. Precipitated DNA in microcentrifuge tubes

(Eppendorf) was centrifuged at 14,000 × g for 10 minutes. The supernatant was decanted and the pellet was washed in 80% ethanol. The suspension was centrifuged as before, the supernatant fraction was decanted, the pellet was air dried for five minutes, and dissolved in 20 µl HPLC-grade water.

For electroporation into E. coli JM109, 17.5 µL of electrocompetent cells were added to each 20-µL DNA solution and mixed gently with the pipette tip. The cell-DNA solutions were aseptically transferred to 2 mm gap electroporation cells (Molecular

BioProducts) and electroporated at 2.5 kV as previously described.

Representative samples of the resulting colonies were picked and subcultured in

LB broth with antibiotics. Small-scale plasmid preparations were performed on 2-mL aliquots of the cell cultures using the Wizard Mini-Prep Kit (Promega). Directions for the plasmid prep were followed as outlined for cell lines containing the endA genotype, with the exception that the guanidine hydrochloride/isopropanol column wash was omitted.

Sequencing of the purified plasmids was performed using the following primers:

pTNMod-OKmLeft: 5’–TT TCC TGG TAC CGT CGA CAT–3’

pTNMod-OKmRight: 3’–GAG ACA CAA CGT GGC TTT CC–3’

DNA Primers were purchased from Sigma-Genosys (The Woodlands, TX) and had

calculated Tm values of 60.4°C and 60.7°C, respectively.

59 CHAPTER IV

RESULTS

A. Isolation and identification of MZ1T

Thauera MZ1T was originally obtained from stock cultures provided by Lajoie et

al. Because MZ1T was not known to have any readily distinguishable phenotypic characteristics, its 16S rDNA gene was amplified by PCR, cloned into the pCR2.1 TOPO

TA cloning vector (Invitrogen) and sequenced to verify possession of the proper strain.

B. Microscopic examination of MZ1T

Photomicrographs of pure cultures of MZ1T revealed the presence of large quantities of extracellular material in post-flocculating cultures. Cells are visible both within and outside of the floc mass. The extracellular material is visible as long tendrils

(>100 µm) extending from the floc mass (Figure IV-1A). Electron micrographs indicated that the extracellular material was polymeric in nature and loosely associated with the cell surface (Figure IV-1B). Differential staining using alcian blue and carbol fuschin indicated that the extracellular material was an acidic polysaccharide (Figure IV-2).

C. Growth rates and doubling times of MZ1T and selected floc– mutants.

The doubling time of MZ1T was determined to be approximately 3.67 hr at 30°C.

Growth rates and doubling times for the wild type and selected floc– mutants are shown

in Figure IV-3. Error values for growth rates represent the standard deviations among

triplicate samples. Statistical analysis of the of mutants versus the wild type using a two-

Y X

Figure IV-1. Photo- and electron micrograph of MZ1T from flocculation culture. A) Photomicrograph of post-flocculating culture of MZ1T showing x) polymeric tendrils from a mass of cells and y) the location of cells both inside and outside of the floc mass (1000X). B) A scanning electron micrograph of a single cell of MZ1T from a post-flocculating culture showing association of polymeric material to the cell surface (23,000X)

Figure IV-2. Complex staining of a post-flocculating pure culture of Thauera sp. MZ1T. Binding of alcian blue to the interstitial material indicates its composition to be an acidic polysaccharide. Cells are counterstained red.

62 8

7.5 WT 7 35B1 6.5 39A 37B1 6 1000X LN OD 20A 5.5

5 0 5 10 15 20 Time (hrs.) A

1.4 1.2 1 WT 0.8 35B1 39A 0.6 37B1 0.4 20A 0.2 Optical Density (600 nm) 0 0 5 10 15 20 25 30 B Time (hrs.)

C Strain Growth Rate Doubling Time (hrs) MZ1T WT 0.189 ± 0.002 3.67 35B1 0.152* ± 0.001 4.55* 37B1 0.182 ± 0.014 3.82 39A 0.185 ± 0.013 3.75 20A 0.183* ± 0.001 3.79*

Figure IV-3. MZ1T wild-type and mutant growth curves. A) Growth curves of MZ1T wild type and mutant strains. Error bars represent standard deviation among triplicate samples. B) Specific growth rate was calculated as the slope of the log-transformed optical density measurements taken during exponential phase (~6–16 hours). Error bars indicate standard deviations among replicates. C) Table showing calculated growth rate and doubling times for wild type (WT) and mutant strains. *Denotes mean value was found to be significantly different from that of the wild type at the 95% confidence level using a two-tailed Student’s t-test. 63 tailed Student’s t-test indicated that the mean growth rates of mutants 20A (p = 0.0086) and 35B1 (p = 5.51 × 10-6) were significantly different than that of the wild type at the

95% confidence level.

D. Carbon utilization in MZ1T and T. aromatica

MZ1T was found to grow aerobically in carbon source-free TDM when amended with the following compounds: sodium succinate, sodium fumarate, α-ketoglutarate, oxaloacetate, sodium benzoate, and tributyric acid. In addition, MZ1T could utilize sodium benzoate anaerobically when nitrate was present as an electron acceptor. MZ1T could not utilize the following substrates in TDM basal media: dextrose, fructose, sucrose, lactose, or mannitol. Interestingly, MZ1T could not grow on agar plates made from TDM (15 g/L agar), though it could grow on either Stoke’s agar plates or TDM broth. Similarly, no growth was detected when the agar was substituted with noble agar or electrophoresis grade agarose.

Carbon-utilization experiments were also conducted on both MZ1T and T. aromatica using GN2 Microplates (Biolog). Utilization patterns were analyzed visually and ranked as high, medium, or low depending upon the extent of color change of the redox-indicating dye. MZ1T was found to utilize the following substrates to a high degree (bright, uniformly red color): cis-aconitic acid, D-gluconic acid, α-ketoglutamic acid, gulonic acid, D-alanine, L-alanine, L-asparagine, L-aspartic acid, and α- D-glucose.

MZ1T was found to utilize the following substrates to a medium degree (uniform pink color): Tween 40, Tween 80, methyl pyruvate, citric acid, liaconic acid, D,L-lactic acid,

64 propionic acid, succinic acid, alaninamide, L-alanyl-glycine, L-glutamic acid, glycyl- L-

glutamic acid, L-histidine, L-leucine, L-proline, L-serine, D,L-carnitine, γ-amino butyric

acid, inosine, putrescine, and glycerol. MZ1T could also utilize the following substrates

to a low degree (barely visible color change to light pink): mono-methylsuccinate, β-

hydroxybutyric acid, acetic acid, formic acid, β-hydroxyphenylacetic acid, α-ketobutyric

acid, α-ketovaleric acid, bromosuccinic acid, L-ornithine, L-phenylalanine, L-threonine,

urocanic acid, uridine, thymidine, phenyl ethylamine, 2-amino ethanol, and 2,3-

butanediol.

Carbon utilization patterns for T. aromatica differed for only a few compounds.

T. aromatica was found to be capable of utilizing all carbon sources previously noted for

MZ1T. Additionally, unlike MZ1T, T. aromatica was found to be able to utilize D- fructose and L-arabinose at high and low levels, respectively.

E. Floc formation

Floc formation by MZ1T was always observed for cultures grown in either TDM

or Stoke’s media (Figure IV-4). The floc formation occurred after cultures reached an

OD600 of approximately 1.0. OD600 was found to decrease following the onset of floc

formation, as shown in Figure IV-5.

Figure IV-4. Photograph of MZ1T in liquid culture. The tube on the left shows the entire cellular contents aggregated into a single floc mass. The tube on the right was vigorously shaken indicating that mechanical disruption of the floc does not result in resuspension of the cells or increased turbidity of the solution equivalent to that achieved prior to floc formation.

1.4

1.2

0.8

0.6

Absorbance 0.4

0.2

0 0 20 40 60 80 100 120 Time (min)

Figure IV-5. Carbohydrate concentration vs. optical density. A) Changes in total carbohydrate concentration (), and total protein (), vs. OD600 of MZ1T culture grown in TDM with succinate over time. The decrease in OD600 at ~36 hr. coincides with the onset of floc formation.

Figure IV-6. Gel permeation chromatogram of crude EPS preparation. The large peak at 50.67 min. was estimated to be approximately 260 kDa in size. The smaller peak at 77.60 min. is approximately 1200 Da.

Table IV-2. Average yields of EPS from floc+ and floc- strains. MZ1T EPS was recovered from 1-L cultures of MZ1T wild type (WT) and mutant strains. Yield values are averages of at least three replicates, accept where noted. Error values represent standard deviations. **Indicates that the mean value is significantly different from the mean of the wild type at a 95% confidence interval. ***Indicates that the mean is significantly different from that of the wild type at a confidence interval of 90%. *Represents values from single experiments.

Strain EPS Yield (mg/L) MZ1T WT 93.6 ± 40.8 20A 85.3 ± 46.9 26B1 51.8 ± 19.0 27A 118.5 ± 31.6 30B1 119.0* 35B1 41.1*** ± 12.3 37B1 14.1** ± 3.5 39A 90.0 ± 18.3 44B1 76.1*

69 F. EPS production by MZ1T

The average yield of extractable EPS is shown in Table IV-2. Total carbohydrate

content as measured by the phenol-sulfuric acid assay versus the increase in the OD600 of

TDM-grown cultures of MZ1T is shown in Figure IV-5.

G. Physical properties of EPS

The crude EPS of MZ1T was found to have a fluffy, off-white, cotton-like appearance. This material was submitted to V-Labs (Covington, LA) for fractionation and size determination. The chromatogram (Figure IV-6) revealed the presence of two carbohydrate-containing peaks. The first of these peaks, eluting at approximately 50 minutes, was detected using a refractive index detector as well as by UV absorbance. Its size was estimated to be 260,000 Da by comparison to the elution times of sodium polystyrene sulfonate standards of known size. A second peak eluting at approximately

76 minutes was also detected by refractive index. This peak, however, had no UV absorbance. The size of the material in the second peak was estimated to be approximately 1200 Da.

The presence of a second smaller peak led to the inclusion of a membrane filtration step using a 100 KDa MWCO membrane in the EPS isolation procedure in order to focus on the 260 KDa polymer of interest. The resultant material possessed a slightly lighter appearance, as shown in Figure IV-7A. Further purification by gel

permeation chromatography resulted in an almost pure white product (Figure IV-7B).

Figure IV-7. Photographs of crude and purified EPS. A) crude and B) column purified preparations of MZ1T EPS.

71 The thermal stability of the EPS was also measured. EPS was found to remain visibly unchanged until reaching a temperature of 200°C, where it began to turn brown in color. The material was completely charred upon reaching 205°C.

The EPS was found to readily dissolve in deionized water at concentrations greater than 10 mg/mL. Viscosity was observed to increase at concentrations greater than

20 mg/mL. Increase in the pH above 7.0 reduced the rate and extent of solubility of the

EPS in water. Samples of EPS could not be dissolved in organic solvents including: dimethyl sulfoxide (DMSO), acetone, toluene, ethanol, methanol, hexane, or ethyl acetate.

Elemental analysis was performed on both crude and column purified EPS samples. The crude EPS was found to be composed of 34.77% carbon, 6.29% hydrogen, and 7.10% nitrogen. Purified EPS contained 35.27% carbon, 6.84% hydrogen, and

5.03% nitrogen.

H. EPS and metal binding

Crude preparations of EPS were found to bind some metals. Binding experiments with uranyl nitrate in deionized water indicated that 36.5% ± 3.2% of uranyl nitrate was removed from solution in the presence of EPS with a contact time of 10 minutes (Figure

IV-8A). Uranyl nitrate could similarly be removed from solution in 100% methanol by interaction with EPS as shown in Figure IV-8B. Approximately 75.6% ± 0.7% of the uranyl nitrate was removed from the methanol solution after two hours of shaking in the presence of EPS. This value is nearly twice the removal of uranyl nitrate by an

Calculated % U Total Uranium U Remaining Estimated U A Bound to (Control) After Treatment Bound to EPS Polymer

EPS (aqueous) 490.5 ± 31.5 311.4 ± 16.1 179.1 36.5% ± 3.2%

EPS (CH3OH) 22110 ± 891.8 5400.5 ± 159.2 16709.5 75.6% ± 0.7% Dextran sulfate 12961.7 ± 210.6 9148.3 41.4% ± 1.0% (CH3OH)

Precipitate depth 4 0.01 0.05 0.1 Calcium chloride Concentration (M) B

Figure IV-8. Metal binding by MZ1T EPS. Uranyl nitrate was dissolved in deionized water or methanol both with and without EPS present. For experiments in aqueous solutions, a solution of EPS was mixed with a solution containing uranyl nitrate and allowed to mix for 10 minutes as described. The EPS was then removed by centrifugation through a 40 KDa MWCO membrane and the uranium concentration of the eluent was determined by scintillation counting. Values are disintegrations per minute. For experiments in methanol, polymer (EPS or dextran sulfate) was removed by centrifugation as described and the uranium concentration of the supernatant was determined by scintillation counting. Values for methanol experiments represent disintegration counts over ten minutes. Standard deviations of triplicate experiments are given. B) Precipitate depth of EPS from aqueous solution as a function of final CaCl2 concentration. No precipitate was detected in the absence of CaCl2.

73 equivalent mass of the negatively charged polysaccharide polymer dextran sulfate (41.4%

± 1.0%).

EPS remained in aqueous solution indefinitely. EPS in solution could be

precipitated by the addition of CaCl2 at concentrations greater than 0.01 M. Increasing

the concentrations of CaCl2 to 0.05 M and 0.1 M, respectively, resulted in a decreasing of

the volume of precipitate with increasing CaCl2 concentration, as shown in Figure IV-8C.

I. Glycosyl composition of EPS

The glycosyl composition of MZ1T EPS was determined by both the alditol acetate and

trimethylsilyl methyl glycoside methods.

Alditol acetate (AA) derivatives from MZ1T EPS

The gas chromatogram of the alditol acetate derivatives generated from MZ1T

EPS using a hydrolysis time of 2 hours revealed the presence of three major

monosaccharide peaks with mass spectra indicative of monosaccharides, as well as one

major peak from the inositol internal standard. Extension of the hydrolysis time to three

hours subsequently revealed the presence of a fourth putative monosaccharide peak at

approximately 32 minutes, as shown in Figure IV-9. Further extension of the hydrolysis

time to four hours did not reveal additional peaks but instead resulted in the decreased

relative abundance of other peaks, suggesting the beginning of degradation of these monosaccharides.

The mass spectrum of peak I in Figure IV-9 is shown in Figure IV-10A. Its early elution relative to the other putative monosaccharides, as well as the presence of ions

- N e; s V. GlcNAc tate derivatives from ; Glc, gluco d r a d n IV. Glc a t s

l a III. Inos n r e t follows: Rha, rhamnose; FucNAc, n i

l II. FucNAc o t i s ogram of the alditol ace s are as o n n i

, glucosamine. s l o n I

; e acety n i N- Gas Chromat m a I. Rha s o c u f l y t e c Figure IV-9. MZ1T EPS. Abbreviatio a and GlcNAc,

3.00E+04 129 171 2.50E+04

2.00E+04

1.50E+04 115 1.00E+04 157 87 188 231 5.00E+03 Relative Abundance 0.00E+00 m/z A

2.50E+05 129 171 2.00E+05

1.50E+05

1.00E+05 115 157 188 231 5.00E+04 87 Relative Abundance

0.00E+00

B m/z

Figure IV-10. Mass spectrum of rhamnose AA derivative. A) Mass spectrum of the putative alditol acetate derivative of rhamnose (Retention time 15.808 min.). Individual m/z values for selected ions are labeled for comparison. B) Mass spectrum of the alditol acetate derivative of a rhamnose standard (Retention time 15.872)

2.50E+04 85

2.00E+04 98 145 1.50E+04

1.00E+04 61 182 242 Relative Abundance 5.00E+03 302

0.00E+00 m/z A

100000 85

80000 98 60000 145

40000 61 182 Relative Abundance 20000 242 302

0 B m/z

Figure IV-11. Mass spectrum of the N-acetylfucosamine AA derivative. A) Mass spectrum of the putative N-acetylfucosamine alditol acetate derivative from MZ1T EPS (elution time = 22.489 min). M/z values for selected ions are included for clarity. B) Mass spectrum of the alditol acetate derivative of a N-acetylfucosamine standard.

3.50E+04 168 3.00E+04

2.50E+04 210 2.00E+04 115 126 1.50E+04

1.00E+04 Relative Abundance 5.00E+03 270 0.00E+00 A m/z

6.00E+04 168 5.00E+04

4.00E+04 210 3.00E+04 115 126

2.00E+04 Relative Abundance 1.00E+04 270 0.00E+00 B m/z

Figure IV-12. Mass spectrum of the inositol AA derivative. A) Mass spectrum of the alditol acetate derivative of the inositol internal standard when added to a preparation of wild type EPS (Retention time 24.415 min.). B) Mass spectrum of the alditol acetate derivative of the inositol internal standard when analyzed with other purified monosaccharide standards (Retention time 24.451 min.).

3.50E+03 115 3.00E+03

2.50E+03 145 2.00E+03 187 103 1.50E+03 218 289 1.00E+03 260 Relative Abundance 5.00E+02 362 0.00E+00 A m/z

2.50E+05 115 2.00E+05

1.50E+05 145 187 103 1.00E+05 218 289

Relative Abundance 5.00E+04 260 362 0.00E+00 B m/z

Figure IV-13. Mass spectrum of the glucose AA derivative. A) Mass spectrum of the putative alditol acetate derivative of glucose (Retention time 25.730 min.). Individual m/z values for selected ions are labeled for comparison. B) Mass spectrum of the alditol acetate derivative of a glucose standard (Retention time 25.949 min.).

1.60E+04 85 1.40E+04 1.20E+04 1.00E+04 145 8.00E+03 103 6.00E+03 318 4.00E+03 Relative Abundance 61 259 2.00E+03 0.00E+00 m/z

2.50E+05 85

2.00E+05 145 1.50E+05 103 1.00E+05 318

Relative Abundance 61 259 5.00E+04

0.00E+00 m/z

Figure IV-14. Mass spectrum of the N-acetylglucosamine AA derivative. A) Mass spectrum of the putative alditol acetate derivative of N-acetylglucosamine (Retention time 31.754 min.). Individual m/z values for selected ions are labeled for comparison. B) Mass spectrum of the alditol acetate derivative of a N-acetylglucosamine standard (Retention time 31.944).

80 with an m/z of 171, suggested that the monosaccharide was a deoxy hexose sugar. The

elution time and mass spectrum of peak I was found to match that of a rhamnose

standard, as shown in Figure IV-10B.

The mass spectrum of AA peak II from Figure IV-9 is shown in Figure IV-11A.

The presence of an ion with m/z of 145 indicated the presence of an N-acetyl group on the monosaccharide. Furthermore, its early elution relative to the other monosaccharides suggested that it too was a deoxy hexose sugar. By comparison to a synthetic standard (a gift from D. Horton to D. Baker), it was determined that the GC elution time and mass spectrum of peak II matched that of N-acetyl fucosamine (2-acetamido-2,6-dideoxy- galactopyranose) as shown in Figure IV-11B.

The mass spectrum and elution profile on the gas chromatogram of peak III indicated that this peak was the alditol acetate derivative of the internal standard inositol.

The mass spectra for peak III and for pure inositol alone are shown in Figure IV-12. The mass spectrum of peak IV from Figure IV-9 is shown in Figure IV-13. As indicated, its mass spectrum as well as its GC elution time matches that of the monosaccharide glucose.

The mass spectrum of the hydrolysis-resistant peak V from Figure IV-9 is shown in Figure IV-14. The presence of a predominant ion with m/z of 145 suggested the unknown was an N-acetylated hexose sugar. The presence of the N-acetyl can also explain the resistance of this monosaccharide to hydrolysis. The elution time and mass spectrum of peak V was found to match that of N-acetylglucosamine.

81 . mine, Inos, inositol internal a fucos l acety - N ycoside derivatives of hydrolyzed MZ1T EPS onic acid;, FucNAc, e. ucosamin l trimethylsilyl methyl gl O- nose; GalA, galactur acetylg N s are as follows: Rha, rham n Abbreviatio Figure IV-15. Gas chromatograph of per- standard; Glc, glucose; and GlcNAc,

5.00E+05 204 4.50E+05 4.00E+05 3.50E+05 3.00E+05 2.50E+05 73 2.00E+05 133 147 217 Abundance 1.50E+05 1.00E+05 59 189 5.00E+04 0.00E+00 A m/z

3.00E+06 204 2.50E+06

2.00E+06

1.50E+06

1.00E+06 73

Abundance 133 147 217 5.00E+05 189 59 0.00E+00

B m/z

Figure IV-16. Mass spectrum of the rhamnose TMS derivative. A) Mass spectrum of the putative rhamnose TMS derivative from MZ1T EPS. B) Mass spectrum of the TMS derivative of a rhamnose standard.

9.00E+04 173 8.00E+04

7.00E+04

6.00E+04

5.00E+04 73

4.00E+04 131 247 3.00E+04 Abundance 147 2.00E+04 204 1.00E+04 59 0.00E+00 m/z A

250000

200000 173

150000

100000 73 131 247 Abundance 147 50000 204 59 0

B m/z

Figure IV-17. Mass spectrum of the N-acetylfucosamine TMS derivative. A) Mass spectrum of the putative N-acetylfucosamine TMS derivative from MZ1T EPS. B) Mass spectrum of the TMS derivative of synthetic N-acetylfucosamine standard.

4.50E+04 4.00E+04 204 3.50E+04 3.00E+04 2.50E+04 2.00E+04 73 1.50E+04 133 217 Abundance 147 1.00E+04 5.00E+03 377 0.00E+00 m/z A

2.00E+06 204 1.80E+06 1.60E+06 1.40E+06 1.20E+06 1.00E+06 73 8.00E+05 217 133 Abundance 6.00E+05 147 4.00E+05 2.00E+05 377 0.00E+00 m/z B

Figure IV-18. Mass spectrum of the glucose TMS derivative. A) Mass spectrum of the putative glucose TMS derivative from MZ1T EPS. B) Mass spectrum of the TMS derivative of synthetic glucose standard.

4.50E+03 305 4.00E+03 217 3.50E+03 73 3.00E+03 2.50E+03 147 318 2.00E+03

Abundance 1.50E+03 1.00E+03 432 5.00E+02 507 0.00E+00 A m/z

9.00E+05 305 8.00E+05 217 7.00E+05 6.00E+05 5.00E+05 73 318 4.00E+05 147

Abundance 3.00E+05 2.00E+05 432 1.00E+05 507 0.00E+00 B m/z

Figure IV-19. Mass spectrum of the inositol TMS derivative. A) Mass spectra of the TMS derivative of the inositol internal standard in an EPS sample and B) from a positive control. Retention times were 25.202 and 25.281 min. respectively.

120000 173 100000

80000

60000 73

Abundance 40000 131 147 204 20000 115 226 259

0 m/z

1000000 173 800000

600000 73 400000

Abundance 131 200000 147 204 226 259 115 0 m/z

Figure IV-20. Mass spectrum of the N-acetylglucosamine TMS derivative. A) Mass spectrum of the putative N-acetylglucosamine TMS derivative from MZ1T EPS. B) Mass spectrum of the TMS derivative of synthetic N-acetylglucosamine standard.

1.20E+05 217 1.00E+05

8.00E+04

6.00E+04 73 4.00E+04 Abundance 133 2.00E+04 159 234 277

0.00E+00 A m/z

7.00E+05 217 6.00E+05

5.00E+05

4.00E+05 3.00E+05 73

Abundance 2.00E+05 133 1.00E+05 159 234 277

0.00E+00 m/z B

Figure IV-21. Mass spectrum of the galacturonic acid TMS derivative. A) Mass spectrum of the putative galacturonic acid TMS derivative from MZ1T EPS (elution time 12.341 min). B) Mass spectrum of the TMS derivative of a galacturonic acid standard (elution time 12.392).

88 Per-O-trimethylsilyl (TMS) methyl glycoside derivatives from MZ1T EPS

The gas chromatogram of TMS derivatives generated from MZ1T EPS is shown in Figure IV-15. Unlike the AA procedure, the ring is not reduced in the TMS methyl glycoside procedure. This results in multiple peaks for each monosaccharide because of mutarotation of the anomeric carbon upon opening and closing of the ring, as well as the formation of both 6-membered pyranose and 5-membered furanose ring structures.

While this results in a more complex gas chromatogram, all of the monosaccharides previously identified by analysis of alditol acetate derivatives could also be identified as their TMS methyl glycoside derivatives by comparison of both their elution profile and of their mass spectra with that generated from known standards (Figures IV-16–20).

In addition to the four monosaccharides previously identified as their alditol acetated derivatives, peaks indicative of an uronic acid were also detected in the TMS analysis.

This monosaccharide would not be detected in the alditol acetate analysis under the conditions used in those experiments. This monosaccharide was identified as galacturonic acid based on the elution profile of the unknown peaks in the gas chromatogram, as well as on comparison of their respective mass spectra with those generated from a pure standard (Figure IV-21).

J. Structural characterization of MZ1T EPS

FT–IR

The Fourier-transform infrared (FT–IR) transmittance spectrum from purified MZ1T EPS is shown in Figure IV-22. Band assignments were made based on Nivens et

of purified MZ1T EPS -1

1060 ittance spectrum from 2000–400 cm

1550

\ 1650 1735 Figure IV-22. FT–IR transm

90 al. and Pouchert (Nivens 2001, Pouchert 1981). Bands consistent with a polysaccharide were detected at 1250 cm-1 (C–O–C stretching), 1100–1000 cm-1 (C–OH stretching), and

1060 cm-1 (C–H stretching of alcohols). IR bands at 1650 cm-1 (Amide I) and 1545 cm-1

(Amide II) are attributed to the amide bond of the N-acetyl groups. Additionally, the IR band at 1735 cm-1 (C=O stretching of esters) suggests the presence of esterified

substituents on the polymer.

One-dimensional 1H-NMR

The one-dimensional (1D) 1H-NMR spectrum generated from MZ1T EPS was consistent with that of a complex polysaccharide (Figure IV-23). Multiple peaks within the anomeric region (4.5-5.2 ppm) indicated the presence of multiple monosaccharides and/or multiple glycosyl linkages within the polymer. Congestion in the anomeric

region, however, prevented exact identification of the monosaccharides present.

The 1D 1H-NMR spectrum also revealed the presence of specific functional

groups within the polysaccharide. Two split peaks at 1.18 and 1.24 ppm, respectively,

indicated the presence of two methyl groups of 6-deoxy hexose sugars and peaks in the

region of 1.943-2.090 ppm indicated the presence of N-acetyl groups (see below).

One-dimensional 13C-NMR

Insufficient signal was generated from a one-dimensional 13C scan to yield an

NMR spectrum. Increasing the concentration of EPS to five mg/mL resulted in increased

sample viscosity but no increase in signal intensity. Increasing temperature of the sample

also had no effect.

H-NMR spectrum of MZ1T EPS. 1 Figure IV-23. One-dimensional

Figure IV-24. Two-dimensional 1H-13C HSQC NMR spectrum of MZ1T EPS. The 1H spectrum is shown on the y-axis and the 13C spectrum is shown on the x- axis.

Figure IV-25. Anomeric region of the 1H-13C HSQC NMR spectrum showing peaks corresponding to five anomeric signatures. The y-axis scale represents the ppm shift for the 1H signals.

Figure IV-26. Two-dimensional 1H-1H gCOSY spectrum of MZ1T EPS.

Figure IV-27. Anomeric and ring region of the gCOSY spectrum of MZ1T EPS. Spectrum shows connectivity among ring protons of the individual monosaccharides comprising the EPS polymer.

Figure IV-28. Two-dimensional 1H-1H TOCSY NMR spectrum of MZ1T EPS.

97 Two-dimensional NMR

The HSQC spectrum of MZ1T EPS reveals several features consistent with the

identification of four principal carbohydrate residues: N-acetylfucosamine, N- acetylglucosamine, rhamnose, and galacturonic acid (Figures IV-24–25). There are six anomeric resonances from 98.66 ppm to 103.5 ppm, with four of higher intensity. There are two peaks at 53.08 and 46.32 ppm that are attributable to C-2 resonances of N- acetylglucosamine and N-acetylfucosamine, respectively (Bartodziejska 1998, Gamian

2000, Gunawardena 1998, Jann 1993, Knirel 1996, Knirel 1998, Perepelov 2000a,

Perepelov 2000b, Shashkov 2000a, Shashkov 2000b, Vinogradov 1992). There are at least three resonances at 20–23 ppm that are attributable to N-acetyl methyl groups. In addition, there are two carbon resonances at 23.30 and 17.95 ppm that are indicative of methyl groups as found on N-acetylfucosamine and rhamnose. There is also a resonance at 71.37 ppm with a corresponding 1H chemical shift of 5.66 ppm, which suggests that

one carbohydrate hydroxyl group is esterified. This evidence is consistent with an IR

absorbance at 1735 cm-1, which also indicates the presence of an ester carbonyl group in

the EPS (Nivens 2001, Pouchert 1981).

Analysis of the gCOSY and TOCSY spectra added further detail to the sugars

present in the polymer (Figures IV-26–28). Provisional assignments were made, and

although the absolute stereochemistries (D or L series) have not been determined, the

assignments are consistent with similar compounds assigned in the literature (Table IV-

3). The anomeric resonances of both of the hexosamines were identified by determining

connectivity to the C-2/H-2 crosspeaks in the HSQC spectrum. Thus, the peak at 98.66

ppm is attached to the N-acetylfucosamine ring and suggests that these resonances are

98 attributable to α-L-N-acetylfucosamine. Similarly, the resonance at 100.23 ppm is

attributed to α-D-N-acetylglucosamine. The two remaining major anomeric resonances

are determined to be β-L-rhamnose (Bartodziejska 1998, Gunawardena 1998, Jann 1993,

Knirel 1998, Shashkov 2000a, Shashkov 2000b) and β-D-galacturonic acid

(Bartodziejska 1998, Gunawardena 1998, Knirel 2000, Stroop 2002). The minor

anomeric resonance at 103.29 ppm is also identified as that of β-D-galacturonic acid based on the similarity of chemical shifts and to the major galacturonic acid component in this sample as well as similar compounds published in the literature. The other minor crosspeak in the anomeric region of the HSQC spectrum is attributed to the presence of

β-D-glucose in the sample (Cerantola 1999, Marshall 2001, Perepelov 2000a, Shashkov

2000b). Provisional assignments of chemical shifts are listed in Table IV-3.

K. Mutagenesis of MZ1T and isolation of floc- mutants

After treatment with NTG, colonies were individually screened for loss of floc-

forming capacity in 10 mL test tubes containing 2 mL Stoke’s broth. Of approximately

500 colonies screened, eight putative floc- mutants were isolated. Their strain

designation and EPS yields are listed in Table IV-2. Since floc-formation is one of the

few distinguishing characteristics of MZ1T, floc- mutants were verified by RISA and/or

BOX PCR. Additionally, 16S rDNA was amplified, cloned, and sequenced from mutant isolates 20A, 26B, and 39A.

Of the eight chemically derived floc- mutants originally isolated, mutants 39A and

44B1 were later found to form flocs to a greatly reduced degree and only after a period of

Table IV-3. Chemical shifts of NMR resonances. Predicted D- or L- configurations are indicated.

δ, 13C δ, 1H ppm Provisional Assignment ppm

5.11 98.66 Anomeric resonance, α-L-N-acetylfucosamine

4.93 100.23 Anomeric resonance, α-D-N-acetylglucosamine

4.65 102.5 Anomeric resonance, β-L-rhamnose

4.61 103.44 Anomeric resonance, D-glucose

4.50 103.26 Anomeric resonance, β-D-galacturonic acid (major)

4.42 103.29 Anomeric resonance, β-D-galacturonic acid (minor)

4.25 46.32 H-2/C-2, α-L-N-acetylfucosamine

4.48 53.08 H-2/C-2, α-D-N-acetylglucosamine

2.10 20.95 N-acetyl methyl resonance

2.04 22.82 N-acetyl methyl resonance

1.95 22.98 N-acetyl methyl resonance

1.22 23.30 Methyl resonance (6-deoxyhexose)

1.20 17.95 Methyl resonance (6-deoxyhexose)

100

3–4 days compared with 2 days for the wild type strain. Flocculation in these two

mutants is limited to only a modest amount of cell-aggregation while the remainder of the

culture broth remains turbid.

L. Transposome mutagenesis

Electroporation of the Transposome transposon deliver vector (Epicentre) into

electrocompetent MZ1T resulted in the isolation of kanamycin-resistant MZ1T colonies.

From 1 µg of the transposon DNA, however, only 47 colonies were isolated. All colonies were found to form flocs in liquid broth.

M. EPS production in floc- mutants

As shown in Table IV-2, all floc- mutants produced some quantity of extractable

EPS. EPS quantities shown in Table IV-2 represent averages of separate EPS extractions from the supernatant fractions of three individual 1-L cultures. Mean yields from mutants 37B1 and 35B1 were found to be significantly different from the mean yield of the wild type at the 95% and 90% confidence levels, respectively.

N. Glycosyl composition of EPS from floc- mutants

The glycosyl composition of EPS isolated from each of the eight putative floc-

mutants was determined using the alditol acetate and TMS methods as previously

101

Ac, - N m GlcNAc tives fro e; and GlcN s a ; Glc, gluco Glc Inos standard the derivatives of their respective FucNAc the alditol acetate deriv follows: Rha, rhamnose; FucNAc, f am o r eviated as r matog ha e R

mine; Inos, inositol internal 9. Gas chro a arides are abb glucosamin l fucos l acety - Figure IV-2 MZ1T-20A EPS. Peaks representing monosacch N acety

102

- N m tives fro e; and GlcNAc s a ; Glc, gluco Inos Glc the derivatives of their respective the alditol acetate deriv follows: Rha, rhamnose; FucNAc, f FucNAc am o r eviated as r amine matog Inos, inositol internal standard glucos ; l Rha mine acety 0. Gas chro a arides are abb - N fucos l Figure IV-3 MZ1T-26B EPS. Peaks representing monosacch GlcNAc acety

103

Ac, - N m tives fro e; and GlcN GlcNAc s a ; Glc, gluco Glc Inos standard the derivatives of their respective the alditol acetate deriv follows: Rha, rhamnose; FucNAc, f FucNAc am o r eviated as r matog e. Rha mine; Inos, inositol internal 1. Gas chro a arides are abb glucosamin l fucos l -acety Figure IV-3 MZ1T-27B EPS. Peaks representing monosacch N acety

104

Ac, - N m GlcNAc tives fro a ; Glc, glucose; and GlcN d Glc Inos the derivatives of their respective FucNAc the alditol acetate deriv follows: Rha, rhamnose; FucNAc, f am o r eviated as r matog e Rha mine; Inos, inositol internal standar 2. Gas chro a arides are abb glucosamin l fucos l acety - Figure IV-3 MZ1T-30B1 EPS. Peaks representing monosacch N acety

105

- N m GlcNAc tives fro e; and s a ; Glc, gluco Glc the derivatives of their respective al standard Inos the alditol acetate deriv follows: Rha, rhamnose; FucNAc, f FucNAc am o r eviated as r amine. matog glucos l Rha mine; Inos, inositol intern 3. Gas chro a arides are abb -acety N fucos l Figure IV-3 MZ1T-35B1 EPS. Peaks representing monosacch acety GlcNAc,

106

- N m tives fro e; and s a GlcNAc ; Glc, gluco Glc Inos al standard the derivatives of their respective the alditol acetate deriv follows: Rha, rhamnose; FucNAc, f FucNAc am o r eviated as r amine. matog Rha glucos l mine; Inos, inositol intern 4. Gas chro a arides are abb -acety N fucos l Figure IV-3 MZ1T-37B EPS. Peaks representing monosacch acety GlcNAc,

107

; d r anda t rnal s te GlcNAc m MZ1T-39A EPS. l in tives fro inosito ccharides are abbreviated as a Inos, ; Glc amine os c Inos tol acetate deriv tylfu FucNAc -ace the aldi glucosamine. f N l es of their respective monosa am o r -acety N matog Rha 5. Gas chro Figure IV-3 Peaks representing the derivativ follows: Rha, rhamnose; FucNAc, Glc, glucose; and GlcNAc,

108

Ac, GlcNAc ak for each andard; and t Inos ly the major pe on , Inositol internal s Glc TMS derivatives from MZ1T-20A nose; GalA, galacturonic acid; FucN r clarity, FucNAc amine. Fo GalA led. e e; Glc, glucose; Inos glucos l Rha aride is lab -acety N fucosamin l -acety GlcNAc, monosacch Figure IV-36. Gas chromatogram of EPS. Abbreviations are: Rha, rham N

109

itol e; Inos, Inos GlcNAc r peak for each Abbreviations are: Inos y, only the majo -acetylfucosamine; Glc, glucos es from MZ1T-26B EPS. N Glc Ac, ucosamine. For clarit l FucNAc -acetylg GalA N led. e Rha aride is lab Rha, rhamnose; GalA, galacturonic acid; FucN internal standard; and GlcNAc, monosacch Figure IV-37. Gas chromatogram of TMS derivativ

110

itol e; Inos, Inos GlcNAc r peak for each Abbreviations are: Inos y, only the majo -acetylfucosamine; Glc, glucos es from MZ1T-27A EPS. N Glc Ac, FucNAc ucosamine. For clarit l -acetylg N GalA led. e Rha aride is lab Figure IV-38. Gas chromatogram of TMS derivativ Rha, rhamnose; GalA, galacturonic acid; FucN internal standard; and GlcNAc, monosacch

111

itol eviations are: GlcNAc e; Inos, Inos r peak for each Inos y, only the majo -acetylfucosamine; Glc, glucos Glc es from MZ1T-30B1 EPS. Abbr N Ac, FucNAc ucosamine. For clarit l GalA -acetylg N eled. ab l Rha s i ide ar

Figure IV-39. Gas chromatogram of TMS derivativ Rha, rhamnose; GalA, galacturonic acid; FucN internal standard; and GlcNAc, monosacch

112

, s ose; Ino c eviations u l GlcNAc Inos , only the major peak arity -acetylfucosamine; Glc, g N Glc cNAc, GalA vatives from MZ1T-35B1 EPS. Abbr -acetylglucosamine. For cl FucNAc N ic acid; Fu n o

r GalA e is labeled. id r Rha amnose; GalA, galactu nosaccha

Figure IV-40. Gas chromatogram of TMS deri are: Rha, rh Inositol internal standard; and GlcNAc, for each mo

113

e ternal l in Inosito GlcNAc eviations are: Rha, r each monosaccharid Inos ucose; Inos, l r peak fo the majo mine; Glc, g ly a n from MZ1T-37B EPS. Abbr Glc fucos l acety - N e. For clarity, o FucNAc glucosamin l GalA -acety uronic acid; FucNAc, N Rha led. e Figure IV-41. Gas chromatogram of TMS derivatives rhamnose; GalA, galact standard; and GlcNAc, is lab

114

itol e; Inos, Inos GlcNAc r peak for each Abbreviations are: Inos y, only the majo -acetylfucosamine; Glc, glucos Glc es from MZ1T-39A EPS. N Ac, FucNAc ucosamine. For clarit l -acetylg GalA N eled. ab l Rha s i aride

Figure IV-42. Gas chromatogram of TMS derivativ Rha, rhamnose; GalA, galacturonic acid; FucN internal standard; and GlcNAc, monosacch

115

- N ride is a

- c N A N c l Inos G

\_ monosacch andard; and GlcNAc, t ternal s Glc l in S derivatives from MZ1T-44B1 EPS. major peak for each FucNAc Inosito GalA, galacturonic acid; FucNAc, ly the GalA ucose; Inos, l rhamnose; r clarity, on Rha s are: Rha, mine; Glc, g n a glucosamine. Fo fucos l l

Figure IV-43. Gas chromatogram of TM Abbreviatio acety acety labeled.

116 described. Both methods revealed the presence of the same monosaccharides previously

detected in EPS extracts from floc+ MZ1T (Figures IV-29–43).

O. Spectroscopic analysis of EPS from floc- mutants

EPS extracts from the eight putative floc- mutants of MZ1T were examined by FT–IR.

Spectra from the individual extracts could be visually separated into two categories based on conserved alterations in the transmittance spectrum. The first group includes EPS samples producing spectra similar to that generated from wild type floc+ EPS extracts.

Strains producing EPS with spectra in the first group include the wild type, mutant 39A,

and mutant 44B1. Interestingly, these are the same two mutants later found to produce

some flocs, albeit at greatly reduced amounts and with a delayed onset relative to the wild

type.

The second group includes the other six mutants. The spectra generated from the

EPS samples isolated from this group all contain conserved alterations in the

transmittance spectrum around 800–400 cm-1 (Figure IV-44). A second conserved

alteration in these spectra relative to the first group is apparent in a reversal of the relative

intensities of amide I (1650 cm-1) and amide II (1545 cm-1)

P. Phenotypic properties of floc- mutants

As stated previously, six of the putative floc- mutants were found to be floc- under

all conditions. Mutants 39A and 44B1 were found to floc late and to a greatly reduced

extent.

Colonies of floc- mutants were found to respond differently to staining on plates.

117 Alcian blue stain applied to Stoke’s agar plates containing colonies of both floc+ and floc-

MZ1T was found to wash easily from floc+ colonies while leaving the floc- colonies

stained blue (Figure IV-45). A similar but opposite staining result was obtained when

using the stain Sudan Black, in which the floc+ colonies retained the stain, and the floc- colonies could be rinsed clear. Results using Sudan Black were not as visually obvious as those obtained from Alcian Blue, but could be better differentiated under UV light.

Application of calcofluor white, a stain known to fluoresce under UV irradiation when bound to β-1,4 linked glucose, showed no fluorescence for either wild type or mutant colonies. However, it was discovered that floc- colonies dissolved on contact

with the stain and were washed from the plate. Floc+ colonies remained tightly adhered

to the plate. It was later found that this dissolution was due to the solvent carrier of the

dye, and the results could be reproduced using 0.1M potassium hydroxide (KOH). The

results of the dye and KOH experiments included not only the six true floc- mutants, but

also the floc-reduced mutants 39A and 44B1.

An additional difference in colony morphology was observed between the wild

type and the eight floc- mutants. While picking colonies of floc+ and floc- mutants it was also observed that the floc- mutants were soft with a stringy texture. This is in stark

contrast to the consistency of wild type colonies, which remain hard and difficult to

physically separate.

Q. Conjugation in floc- mutants

Unlike the wild type, it was found that floc- mutants 20A, 26B, and the floc-

reduced mutant 39A could be successfully mated using E. coli strains harboring broad

118

Figure IV-44. Comparison of FT-IR spectra of MZ1T wild-type and mutant EPS. Floc- mutants 35B1, 26B, and 20A have altered spectra relative to the wild type in the regions of 800–400 cm-1. EPS spectra from the floc-reduced mutants 39A and 44B1 appear similar to the wild type.

119

39A 26B 20A WT

Figure IV-45: Alcian blue stain of MZ1T wild type and mutant colonies. Columns include four replicates of from left to right: MZ1T-39A, MZ1T-26B, MZ1T-20A, and Floc+ MZ1T

120 host range plasmids in the presence of another E. coli strain carrying the helper plasmid

pRK2013. These results are similar to those reported by Easson et al. using chemically

derived floc- mutants of Z. ramigera 115 (Easson Jr. 1987a). Tri-parental conjugal transfer into MZ1T chemical mutants was successful using the following plasmids: pRK290 (Ditta 1980), pRK415 (Keen 1988), and pBSHR1 (Hay 2000). Conjugal transfer similarly occurred when transferring the cosmid vector pLAFR3. Additionally, bi-parental mating with E. coli carrying the transposon delivery vector pTnMod-OKm was also successful. No colonies were obtained from simultaneous experiments using floc+ MZ1T.

R. Transposon mutagenesis of MZ1T-39A

Because of the difficulties inherent in identifying mutations generated by

chemical means, and because the reduced floc-forming mutant 39A could be mated by

conjugal transfer, it was decided to further mutagenize this strain by mating it with E. coli

carrying the transposon delivery vector pTnMod-OKm (Figure III-6) (Dennis 1998). The

resultant colonies were then screened in culture tubes containing 2 mL Stoke’s broth for

the complete loss of floc formation. Nearly 900 colonies were screened in this manner

yielding an additional fourteen putative floc- mutants listed in Table III-1. Of these, mutants 118, 142, 153, and 171 were randomly chosen for further study.

A characteristic of this transposon is an internal origin of replication. This facet is useful since one can digest the genomic DNA containing the inserted transposon with enzymes specifically chosen to cut outside the region of insertion, followed by self- ligation of the fragments into circular DNA, and finally electroporation of the resulting

121 circular fragments into a strain capable of maintaining the plasmid. Isolation of the

proper fragment is made possible by antibiotic selection on media containing 50-µg/mL kanamycin. The resulting colonies should contain the plasmid with the transposon and its integral antibiotic resistance gene, as well as DNA from the regions flanking the transposon insertion site. The flanking regions can then be sequenced and used to map the site of transposon insertion.

To locate the site of transposon insertion in floc- mutants of MZ1T-39A, genomic

DNA was isolated and digested with a restriction enzyme, self ligated, and transformed

into E. coli JM109. Restriction enzymes used included one or more of the following:

BamHI, PstI, ApaI, Nde1, and EcoRV. Self-ligated plasmids were recovered by

electrotransformation into E. coli from Pst1 digests of the genomic DNA preparations

from MZ1T-39A floc– mutants 142, 153, and 171. No plasmids were recovered from

digests with other enzymes or with strain MZ1T-118. The resulting plasmids were

named according to the following methodology: “p” for plasmid, “171” for the strain

designation, and “P” representing the first letter of the restriction enzyme used.

Sequencing of the isolated plasmids revealed only vector sequence from the site of

priming inside the multi-cloning sites, through the inverted repeats, and into the vector.

No MZ1T-39A genomic DNA was detected.

The sequence data suggest that transposition did not occur. Newly isolated floc– strains of MZ1T-39A, however, were kanamycin resistant and verified to be strains of

MZ1T via RISA, BOX PCR, and 16S rDNA sequence. One possibility is that the plasmid was integrated into the chromosome by homologous recombination. Several lines of evidence support this conclusion. The first is that the plasmid contains a narrow-

122 host range origin of replication limiting the likelihood of successful replication in MZ1T.

Secondly, plasmid DNA could not be isolated from newly-floc– strains of MZ1T 39A, though it was easily purified from the original donor and final recipient E. coli strains.

Next, no transconjugants were detected following mating of MZ1T transformants with E. coli JM109 by biparental mating. Finally, no plasmid was successfully transformed into

E. coli when the genomic DNA preparations from MZ1T-39A floc– strains were digested with enzymes that did not cut the vector sequence. If circular plasmid DNA were present in the DNA preparations and remained uncut through the digestion step, it would be expected that these intact plasmids would result in a high number of transformants after the final electroporation. It can be hypothesized that, in the latter case, the resultant DNA fragments were too large to be efficiently self-ligated and transformed into E. coli.

123 CHAPTER V

DISCUSSION

A. MZ1T as a member of the genus Thauera

MZ1T was found by 16S rDNA homology to belong to the genus Thauera (Lajoie

2000). The carbon-utilization pattern of MZ1T, its isolation from wastewater, and its capacity for anaerobic degradation of sodium benzoate when using nitrate as a terminal electron acceptor as reported here, all support this conclusion.

T. aromatica strain T1 has been reported to have peritrichous flagellation and strain K-172 to be degenerately peritrichous (Evans 1991). T. selenatis has also been described as possessing a single, polar flagellum (Macy 1993). However, no evidence of flagellation was detected in transmission electron photomicrographs of MZ1T.

Carbon utilization patterns of MZ1T were found to be very similar to those of T. aromatica. The only exceptions were found to be in the utilization of sugars as sole carbon sources. T. aromatica could use D-fructose and L-arabinose as carbon sources

whereas MZ1T could not. Both were found to utilize α-D-glucose on GN2 plates, but

dextrose did not support growth of MZ1T in TDM. The reason for this apparent

discrepancy is not known, but the inability of MZ1T to utilize these sugars may have

important ramifications for its EPS production.

Floc formation was found to occur during stationary phase in both media tested,

whether under aerobic or anaerobic conditions. While no reports have described floc

formation in other Thauera to the extent seen in MZ1T, cellular aggregation and

clumping in some Thauera species has been reported (Foss 1998, Song 1998).

124 Species within the genus Thauera are most notable for their ability to degrade aromatic hydrocarbons anaerobically using nitrate as a terminal electron acceptor (Anders

1995, Coschigano 1994, Evans 1991, Scholten 1999, Song 1998). Like MZ1T, a number of these species have also been isolated from wastewater treatment systems. Some reports have noted the aggregation of different Thauera species in pure culture during late log or stationary phase (Foss 1998, Song 1998). MZ1T also aggregates in pure culture following the onset of stationary phase, suggesting that this may be a widespread phenomenon within the genus Thauera.

B. Wild Type MZ1T EPS

Visible and electron photomicrographs indicated that cells of MZ1T from flocculating cultures were surrounded by an extracellular polymeric material. This interstitial material was determined to contain acidic polysaccharide by differential staining with alcian blue. Isolation and investigation of the physical properties of the polysaccharide revealed several interesting characteristics of the EPS. Metal binding as evidenced by removal of uranyl nitrate from both aqueous and non-aqueous solutions suggested that the EPS polymer might have some utility for remediation of metal contaminated liquids. Precipitation of EPS from aqueous solution upon increasing addition of calcium chloride also suggested that the polymer may remain in solution until sufficient quantities of metals are bound, which could facilitate metal contaminant removal from wastewaters. Insolubility of the EPS in non-aqueous liquids might also allow the construction of woven EPS filters for applications in the removal of metal cations from these solutions.

125 Composition analysis of the purified wild type EPS indicates that it is composed

of 35.27% carbon, 6.84% hydrogen, and 5.03% nitrogen by mass, with the remainder

comprising oxygen and other contaminating or possibly bonded elements (e.g.

phosphorus). The molar ratio of carbon to nitrogen was then calculated to be

approximately 8:1. This value is the C:N ratio present in both N-acetylglucosamine and

N-acetylfucosamine. However, this does not account for the carbon present in the two (or

three, if including glucose) non-nitrogen containing monosaccharides detected in the

EPS. This could be explained if two aminosugars were in fact, di-aminosugars. Such

species, however, were not detected by GC-MS of the aldtiol acetate or TMS chemical

derivatives. An alternative case could be made in which the undefined aglycone

substituent on the EPS polymer contained a nitrogenous functional group, though a

review of the literature found no precedent for such a modification in bacterial

exopolysaccharides. The most likely explanation for the excess quantity of nitrogen

detected is incomplete removal by sublimation of the ammonium formate buffer used

during column purification of the EPS.

Chemical and NMR spectroscopic analyses identified rhamnose, galacturonic

acid, N-acetyl-glucosamine, and N-acetyl-fucosamine as components of the extracted

EPS. Provisional assignments of the D- or L- configuration and linkages between monosaccharides have been provided (Table IV-3). Glucose was also identified based on

GC-MS analyses of alditol acetate and TMS methyl glycoside derivatives.

Aminosugars similar to those found in MZ1T EPS have been shown to be components of Z. ramigera ATCC 19544T and strain MP6 (Farrah 1976). Additionally,

Tezuka described a polysaccharide containing glucosamine and possibly fucosamine

126 from an unspecified strain of Z. ramigera (Tezuka 1973). Considering the presence of

aminosugars in the EPS of MZ1T and these cluster-forming strains, but not in the EPS of

the flocculating but non-cluster-forming bacterium Duganella zoogloeoides (i.e. Z. ramigera ATCC 25935 or strain 115), these monosaccharides likely play an important functional role in the formation of zoogloeal clusters.

The presence of N-acetyl-fucosamine is of particular interest. Fucosamine has been hypothesized to exist in Zoogloea-like organisms but never shown conclusively

(Tezuka 1973). This is most likely due to its rarity and in difficulties obtaining purified material for use as a suitable standard. In that study, glucosamine was also identified. It should be noted that in the study by Tezuka, the chemical analyses performed would have removed any N-acetyl groups present, leaving only the aminosugars. Given the poor state of phylogenetic differentiation among Zoogloea-like strains at that time, it is also entirely possible that Tezuka had actually isolated a member of the genus Thauera, possibly even MZ1T.

The presence of galacturonic acid in the EPS of MZ1T is also notable.

Interactions of the carboxylic acid functional groups of uronic acids with divalent metal cations such as Ca2+ are believed to be the basis for gel formation in alginate (Bruus

1992). Uronic acids have also been detected in the EPS of Z. ramigera strains 106

(ATCC 19544T) and MP6 (Farrah 1976).

Pseudomonas aeruginosa produces a bacterial alginate consisting of mannuronic

and guluronic acids (Evans 1973). Though this organism is well known to form biofilms,

there are no published reports on its forming zoogloeal clusters (Costerton 1994). This

suggests that the uronic acid alone is not responsible for zoogloeal cluster formation, but

127 interactions between the uronic acid and one or both of the aminosugar components may be involved.

Glucose was identified in both alditol acetate and TMS methyl glycoside preparations of EPS by GC-MS analysis. TMS analyses of column-purified fractions indicated that glucose was present in samples both with and without the other identified monosaccharides components (Figure IV-46). The latter case suggests that glucose may be a component of a second polysaccharide polymer co-purifying in the EPS preparation.

Conversely, the steady relative ratio of glucose to other identified monosaccharides over a range of fractions could also suggest that glucose may also be a component of the main polysaccharide chain. However, no evidence to support this latter hypothesis in the form of through-space interactions among neighboring monosaccharides was found in the

NMR data.

Both IR and NMR spectroscopy indicated the presence of a possible aglycone

(e.g. non-sugar) substituent esterified to the EPS of MZ1T. Acyl groups in bacterial alginate have been shown to affect biofilm formation in P. aeruginosa, and may therefore influence zoogloeal cluster formation (Nivens 2001).

Duganella zoogloeoides (Z. ramigera strain 115) produces an exopolysaccharide composed of glucose and galactose residues (Friedman 1968b, Friedman 1969, Parsons

1971). Its EPS was subsequently found to contain pyruvyl, acetyl, and succinyl substituents O-esterified to the main polymer chain (Franzen 1984, Ikeda 1982, Troyano

1996). This floc-forming organism has also been described as producing a mucopolysaccharide matrix surrounding its cells, and is a member of the Rhodocyclus branch of the β-Proteobacteria along with Z. ramigera ATCC 19544T and members of the

128 80 % Rha % GalA % FucNAc 60 % Glc % GlcNAc (%)

nce 40 unda

20 ve Ab lati Re

36 37 43 53 60 70-74 98-101 113-120 125-128 141-147 Fraction

0.6

0.5

0.4

g) 0.3 ss (m a 0.2 M

0.1

0.0 B

0 20 40 60 80 100 120 140 160 180 200 Fraction Figure IV-46. Relative abundance of monosaccharides in GPC column fractions. A) Percent relative abundance of individual monosaccharides (as their TMS derivatives) in gel permeation chromatography (GPC) column fractions. B) GPC elution profile showing that the majority of the polymer was eluted from a single peak composed of fractions 36–65.

129 genus Thauera (Friedman 1968a, Hiraishi 1992, Shin 1993). The data from both these

organisms suggest that aglyconic substituents may also play a role in floc formation in

MZ1T.

C. EPS extraction from and composition in mutants

Mutants of MZ1T that were reduced or deficient in flocculation were isolated

following chemical mutagenesis. Surprisingly, all mutants produced some level of

extractable EPS. Furthermore, all five monosaccharides previously identified as their

alditol acetate or TMS derivatives could similarly be found in the EPS preparations from

the mutants.

Of the mutants tested, 20A and 35B1 were found to have significantly different

growth rates than the wild type (Figure IV-3). Mutant 37B1 however, which had a

growth rate similar to that of the wild type, was the only mutant to produce significantly

less EPS (95% confidence level). EPS yield from 37B1 was approximately 15% that of

the wild type and may explain the loss of floc forming capacity in this strain.

Mean EPS yield from mutant 35B1 was also found to be significantly less than

the wild type when using a confidence level of 90%. This latter result is likely linked to

the lower growth rate of the mutant relative to the wild type, and suggests that other non-

EPS related mutations might be involved.

The major difference detected among EPS samples isolated from floc– mutants,

floc-reduced mutants, and the wild type strain were conserved alterations in their respective FT–IR spectra (Figure IV-44).

130 D. Additional phenotypic alterations in floc- mutants

Unlike the wild type, floc– mutants 20A and 26B, and floc-reduced mutant 39A could be successfully mated by bi-and triparental conjugal transfer. This observation is consistent with those results found by Easson for chemically derived floc– mutants of

Zoogloea ramigera 115 (Duganella zoogloeoides) (Easson Jr. 1987a). Additionally,

colonies of these MZ1T mutants were found to respond differently to stains and dyes

compared with the wild type, and to have a much softer colony texture. These results

suggest that the mutations have resulted in substantial alterations of the organization or

composition of the surface of the cells.

The changes in the cell surface are not fully explained by the analyses of the EPS

extracted from these mutants. All monosaccharides detected in the EPS of the wild type

were also identified in EPS from the mutants. In addition, though the FT-IR spectra of

EPS from mutants 20A and 26B were altered relative to the wild type, the spectrum of

EPS from mutant 39A was not. It therefore appears likely that the mechanisms behind

the loss off wild type levels of floc formation among these mutants are different. Future

complementation analysis of the mutants will be necessary to support this hypothesis.

The FT-IR spectral data suggest that EPS from mutants 20A and 26B are structurally

altered in some way, possibly by changes in the extent or type of aglycone substitution.

The phenotype of 39A and the data on its EPS may be explained by alterations in the

attachment of the EPS to the cell surface, or by changes in the genetic regulatory

elements resulting in delayed onset of EPS production. The last point being highly

possible given that plate and mating experiments were conducted after a maximum of 48

131 hours of growth, whereas extraction of EPS from liquid cultures occurred after seven days of incubation.

E. Transposon mutagenesis

Because of the reduced, yet reproducible, manner of flocculation produced by mutant 39A, as well as its capacity for conjugal transfer, it was chosen for further transposon mutagenesis. Transconjugants were screened for a complete floc– phenotype in the hopes that the site of insertion could then be mapped. Fourteen mutants were isolated and of, these, four were chosen for more rigorous investigation. Of these four, genomic DNA was extracted, digested, self-ligated to yield circular DNA, and electroporated into an E. coli host for three of the strains. Sequencing of the isolated plasmids, however, indicated that transposition did not occur. The most likely explanation is that the plasmids inserted into the chromosome by homologous recombination. Further investigation by Southern hybridization or other means is needed to test this hypothesis. Then, mapping of the insertion site and characterization of the disrupted genes affecting floc formation can occur.

132 CHAPTER VI

CONCLUSIONS

The exact role of zoogloeal cluster or floc formation in the ecology of Thauera

MZ1T is not known. Conditions within the activated sludge system, however, would be

expected to select for more strongly aggregating variants. It is possible, though untested,

that zoogloeal cluster formation may produce anaerobic zones within the cluster matrix

and thereby facilitate degradation of aromatic hydrocarbons by the oxygen-labile benzoyl

CoA mediated pathway (Heider 1998). Similar anaerobic zones are known to occur in

the interior of biofilms (Costerton 1994). If true, this hypothesis might also be extended

to Zoogloea species, which have also been shown to be capable of aromatic degradation

and denitrification (Unz 1972).

The composition of the EPS from MZ1T has been shown to have similarities to

that reported for Z. ramigera ATCC 19544T and some closely related strains. These data

have been used to make inferences about the physical and chemical interactions leading

to the formation of zoogloeal clusters. The presence of the negatively charged carboxyl

groups of the galacturonic acid residues explains at least one mechanism of metal binding

and might be involved in divalent cation bridging among polymer strands. The role of

the aminosugars, apparently conserved between the zoogloeal cluster forming bacteria Z.

ramigera 106 and MZ1T, is not completely known. Certainly not all bacteria producing

EPS containing aminosugars form zoogloeal clusters, but aminosugars may present a

partial positive charge for direct interaction with the anionic carboxyl groups previously described. The role of aglycone substituents in biofilm formation and their presence in

133 the EPS of floc-forming organisms also suggests a role for the currently undefined aglyconic moiety in zoogloeal cluster formation and in floc formation of MZ1T.

FT–IR spectral data derived from EPS samples of floc– and floc-reduced mutants

indicate that a modification of the EPS has taken place in floc– mutants. Moreover, this

modification is apparently conserved among all the floc– mutants but absent in the floc-

deficient mutants 39A and 44B1. These data suggest a direct role of EPS in floc

formation. The reduction of floc formation in the floc-reduced mutants might be

explained if the EPS produced by the floc-deficient strains were no longer associated

with the cell surface or not to the same extent as in the wild type. This could explain the

apparent contradiction of reduced floc formation in a mutant with a wild type level of

EPS production and no detectible difference in EPS composition.

Though much is now known of the composition of the EPS produced by MZ1T,

only inferences have been made of the linkage of the individual monosaccharides in the

polymer. Similarly, no identification has been made of the aglycone substituents present

on the polysaccharide. Continued investigations of these questions in both the wild type

and mutant strains are needed for a full understanding of the role of EPS in the

intercellular interaction of MZ1T.

In addition to the findings reported here on MZ1T, its EPS composition, and its

capacity for floc formation, the generation of both floc– and floc-reduced mutants has

produced the necessary precursors for the future identification of the genes involved in

floc formation. This situation is further strengthened by the finding that, unlike the wild

type, these mutants are amenable to genetic manipulation. This has resulted in the

creation of a complete platform for continued genetic study in MZ1T, as well as a

134 roadmap for application to research problems involving similar strains. These tools may now be more broadly applied to aspects of biodegradation, environmental remediation, prediction of wastewater plant upsets, environmental issues in carbon sequestration, and possible commercial polymer production in MZ1T and related bacteria.

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151 VITA

Michael Allen was born in Ft. Worth, Texas in 1971. He graduated with a Bachelor of

Science degree in Biology with special emphasis on Ecology and Environmental Science from the University of Texas at Arlington in 1997. He then pursued his Ph.D. in

Microbiology at the Center for Environmental Biotechnology and the University of

Tennessee.

152