Online Monitoring of Aerobic Denitrification of Pseudomonas

ONLINE MONITORING OF AEROBIC DENITRIFICATION OF PSEUDOMONAS

AERUGINOSA BY NAD(P)H FLUORESCENCE

A Thesis

Presented to

The Graduate Faculty of The University of Akron

In Partial Fulfillment

of the Requirements for the Degree

Master of Science

Qing Xia

December, 2005 ONLINE MONITORING OF AEROBIC DENITRIFICATION OF PSEUDOMONAS

AERUGINOSA BY NAD(P)H FLUORESCENCE

Qing Xia

Thesis

Approved: Accepted:

______Advisor Department Chair Lu-Kwang Ju Lu-Kwang Ju

______Committee Member Dean of the College Bi-min Zhang Newby George K. Haritos

______Committee Member Dean of the Graduate School Ping Wang George R. Newkome

______Date

ABSTRACT

In cystic fibrosis airway infection, Pseudomonas aeruginosa forms microaerobic biofilm and undergoes significant physiological changes. It is important to understand the bacterium’s metabolism at microaerobic conditions. Continuous cultures of P. aeruginosa (ATCC 9027) maintained at different dissolved oxygen concentrations (DO) and three different dilution rates (D) were studied for the effects of DO and D on various culture properties, especially on aerobic respiration and denitrification. The DO was varied from 0 mg/L (completely anoxic condition) to 2.2 mg/L, and measured with optical sensors that could accurately determine very low DO based on oxygen-quenched luminescence. The studied dilution rates were 0.026 h-1, 0.06 h-1 and 0.13 h-1. The strain was found to perform aerobic denitrification; while the specific nitrate and nitrite reduction rates decreased with increasing DO, denitrification persisted even at relatively high DO levels (1-2.2 mg/L) at different D. In the presence of nitrate, the Monod constant for DO (i.e., the critical DO at which the specific oxygen uptake rate (OUR) is half of the maximum rate) was practically zero (< 0.001 mg/L) for this P. aeruginosa strain. Aerobic denitrification appeared to function as an electron-accepting mechanism supplementary or competitive to aerobic respiration. The shift of culture’s respiratory mechanism was also clearly detected with a fluorometer targeting at intracellular

NAD(P)H, i.e., the reduced coenzymes nicotinamide adenine dinucleotides (phosphate).

Comparatively, the NAD(P)H fluorescence was highest at the anoxic,

iii

denitrifying condition (NFUDN), lowest at fully aerobic conditions (NFUOX), and intermediate fluorescence (NFU) at conditions where both denitrification and aerobic respiration occurred. Representing a quantitative measure of the culture’s “fractional approach” to the fully denitrifying state, the normalized fractions (NFU -

NFUOX)/(NFUDN - NFUOX) were correlated with the calculated fractions of electrons accepted by denitrification. The denitrification-accepted fractions of electrons increased with the NFU fractions: the increases were gradual at larger DO levels (DO ≥ 0.1 mg/L), but much sharper at lower DO at three different dilution rates. The fluorescence fraction changed more rapidly than the electron fraction at very low DO levels (< 0.001 mg/L).

The results demonstrated that online NAD(P)H fluorescence was a feasible technique for effective monitoring and quantitative description of the microaerobic state of microorganisms.

ACKNOWLEDGEMENTS

I would like to express my deep appreciation to my academic advisor, Dr. Lu-

Kwang Ju, for his encouragement, precious suggestions, sufficient patience to guide, and financial support during my master’s study. I am also thankful to Dr. Bi-min Zhang

Newby and Dr. Ping Wang for serving as my committee members as well as for their suggestions on this work.

I also appreciated Mr. Fan Chen’s help for the experiment setup and development.

I am also grateful for Mr. Nathan, K. Klettlinger and Mr. Nicholas, J. Hammilton’s helps in all analytical experiments.

I would like to thank my group members, Ms. Lin Huang and Ms. Shuyan Qiu, for their friendship and assistance.

I am eternally grateful to my beloved husband Fan Chen for his love, understanding, and whole-hearted support during entire graduate study at the University of Akron. I also thank my parents and sisters for their continuous support from China during my study.

TABLE OF CONTENTS

Page

LIST OF FIGURES ...... ix

CHAPTER

I INTRODUCTION...... 1

1.1 Scope of Research...... 4

1.2 Research Objectives...... 5

1.3 Structure of Thesis ...... 5

II LITERATURE SURVEY...... 7

2.1 Biofilm Formation ...... 7

2.2 Two Luminescent Techniques ...... 10

2.2.1 Fluorescence ...... 10

2.2.2 NAD(P)H Fluorescence...... 12

2.2.3 Influence of Various Factors on Fluorescence in Solutions ...... 15

2.2.4 Luminescence Technique for DO Measurement ...... 18

2.2.4.1 The Principles of the Fiber-optic Oxygen Meter ...... 18

2.3 Biosurfactant Production...... 20

2.3.1 Structures and Properties of Biosurfactants...... 20

2.3.2 Synthesis of Biosurfactants...... 21

2.3.2.1 Microorganisms for Rhamnolipid Production ...... 21

2.3.2.2 Choice of Carbon Substrates and Limiting Nutrients...... 22

2.3.2.3 Temperature and pH Effects ...... 23

2.3.2.4 Foaming in Biosurfactant Fermentation ...... 23

2.3.3 Rhamnolipid Types and Rhamnolipid Production...... 24

2.3.3.1 Rhamnolipid Formation...... 25

2.4 Respiration ...... 27

2.4.1 Aerobic Respiration ...... 27

2.4.2 Anaerobic Respiration ...... 29

2.4.2.1 Denitrification...... 29

2.4.3 Microaerobic Denitrification ...... 32

III MATERIALS AND METHODS ...... 34

3.1 Microorganism and Medium...... 34

3.2 Experimental Setup and Continuous Culture...... 35

3.3 Analytical Methods...... 37

3.3.1 Cell Concentrations and Cell Dry Weight Analysis ...... 39

3.3.2 Ammonium and Nitrate Analysis ...... 39

3.3.3 Nitrite Analysis...... 40

3.3.4 Glucose Analysis ...... 40

3.3.5 Rhamnolipids Analysis ...... 41

3.3.6 DO and OUR Measurement...... 41

3.3.7 Culture Fluorescence Measurement...... 42

3.3.8 Calculations...... 43 vii

IV AEROBIC, MICROAEROBIC, ANAEROBIC DENITRIFICATION OF PSEUDOMONAS AERUGINOSA...... 46

4.1 Introduction...... 46

4.2 Cell Metabolism and Respiration Mechanism at D = 0.026h-1, D = 0.06 h-1 and D = 0.13 h-1 ...... 48

4.2.1 Materials and Methods...... 48

4.2.2 Cell Properties in Continuous Cultures Maintained at Different DO and D……………………………………………………………………..49

4.2.3 Respiration Mechanism and Energy Generation at Different DO and D . 66

4.2.4 Culture Fluorescence ...... 70

4.2.5 Examine Transition of Electron-Accepting Mechanism Using NAD(P)H Fluorescence...... 74

4.2.6 Combined two Fractions at D = 0.06 h-1...... 78

V CONCLUSIONS ...... 81

5.1 Conclusions...... 81

5.2 Recommendations for Future Work...... 85

BIBLIOGRAPHY...... 86

viii

LIST OF FIGURES

Figure Page

2.1 Five-stage models for biofilm development ...... 9

2.2 The cyclic nature of coenzymes NAD(P) and NAD(P)H...... 14

2.3 Fluorescence of P. aeruginosa under different electron accepting conditions...... 16

2.4 Oxygen quenching of the fluorescence of tris(2, 2’-bipyridiyl) ruthenium (II) dichloride ...... 19

2.5 Four types of rhamnolipids found in Pseudomonas species ...... 26

2.6 Biosynthesis of rhamnolipids...... 28

3.1 Cell concentration profiles at the different antifoam concentrations...... 36

3.2 Experimental setup...... 38

3.3 Electron acceptance and ATP formation in respiratory chain of P. aeruginosa...... 45

4.1 Steady-state concentrations measured in continuous cultures maintained at different DO (D = 0.026 h-1)...... 50

4.2 Steady-state concentrations measured in continuous cultures maintained at different DO (D = 0.06 h-1)...... 51

4.3 Steady-state concentrations measured in continuous cultures maintained at different DO (D = 0.13 h-1)...... 52

4.4 Steady-state concentrations measured in continuous cultures maintained at different air circulation rates. (D = 0.06 h-1)...... 53

4.5 Specific rates of oxygen uptake, nitrate reduction and nitrite reduction determined for continuous cultures maintained at different DO (D = 0.026 h-1)...... 56

4.6 Specific rates of oxygen uptake, nitrate reduction and nitrite reduction determined for continuous cultures maintained at different DO (D = 0.06 h-1)...... 57

4.7 Specific rates of oxygen uptake, nitrate reduction and nitrite reduction determined for continuous cultures maintained at different DO (D = 0.13 h-1)...... 58

4.8 Specific rates of nitrate reduction (NAR), nitrite reduction (NIR) and oxygen uptake (OUR) at different air circulation rates (D = 0.06 h-1) ...... 59

4.9 Cell yield and ATP generation rates determined for continuous cultures maintains at different DO (D = 0.026 h-1)...... 61

4.10 Cell yield and ATP generation rates determined for continuous cultures maintains at different DO (D = 0.06 h-1)...... 62

4.11 Cell yield and ATP generation rates determined for continuous cultures maintains at different DO (D = 0.13 h-1)...... 63

4.12 Calculated culture properties, including specific rhamnolipid production rate, ATP generation rate, cell yield (YX/S), ATP yield (YATP/S), and ATP yield per -1 cell yield (YATP/X), at different air circulation rates (D = 0.06 h )...... 64

4.13 Specific ATP generation rates determined for continuous cultures maintained at different DO and D...... 68

4.14 An example profile of NAD(P)H fluorescence responding to brief perturbation from the continuous culture’s steady state to aerobic and anoxic conditions (The continuous cultures shown here was maintained at DO of 0.1 mg/L and D of 0.026 h-1) ...... 71

4.15 Total and specific culture fluorescence intensities observed at fully denitrifying (anoxic) and aerobic conditions, plotted against the corresponding cell concentrations or DO to show the effects of background fluorescence from fluorophores other than NAD(P)H (D = 0.026 h-1)...... 73

4.16 Correlation between two different indicators to the culture’s “extent” of denitrification: the fraction of electrons accepted by denitrification versus the NFU fraction (at D = 0.026 h-1, D = 0.06 h-1 and D = 0.13 h-1 ...... 75

4.17 Correlation between denitrification-accepted fraction of electrons and NAD(P)H fluorescence fraction for continuous cultures at D = 0.06 h-1 under practically zero-DO conditions...... 76

4.18 Correlation between denitrification-accepted fraction of electrons and NAD(P)H fluorescence fraction for continuous cultures at D = 0.06 h-1 under various DO and aeration conditions. (Results were from two sets of experiments, with the presumed connection drawn as the dashed line, using feed media with two different glucose concentrations.) ...... 80

CHAPTER I

INTRODUCTION

Oxygen availability is an important factor in the regulation of bacterial behavior and distribution in the environment, and for bacterial cultivation. Accordingly, the uneven distributions of water flow, nutrients, and microbial populations generate a continuous and dynamic spectrum of aerobic, microaerobic, and anaerobic/anoxic conditions in the frequently heterogeneous, complex environments. The ecological fate of different organic compounds varies differently with the changing dissolved oxygen

(DO) level (1). For example, oxygenated compounds are largely biodegradable in all conditions (at different rates) (2), while highly chlorinated hydrocarbons are more susceptible to sequential degradation: a reductive dechlorination under anoxic conditions

(e.g., by sulfate-reducing bacteria or methanogens) followed by the aerobic mineralization (3). Knowing how microbial metabolism for different organic materials changes with the varying DO level is essential to the modeling of their ecological fate for risk assessment and management as well as to the development of advanced bioremediation technology.

Field implementation of petroleum bioremediation should therefore address (1) the dual roles of O2 in biodegradation, i.e., for respiration and as a direct reactant (4), and the interactions of microbial activities under different (aerobic, microaerobic, and anaerobic) environments. For example, in microaerobic conditions, the respiratory need

may be satisfied mainly, if not completely, by alternative electron acceptors such as nitrate so that the scanty O2 available is reserved for driving hydrocarbon transformation.

Once oxygenated, the metabolites are readily degradable even under microaerobic or anaerobic conditions. Knowledge on microaerobic metabolism is therefore essential to understanding the ecological fate of petroleum hydrocarbons and to developing more economical and effective bioremediation technology. The microaerobic condition is, however, ill defined. From the simple point of view of microbial respiration, the aerobic condition corresponds to that when the organism(s) uses O2 as the terminal electron acceptor (aerobic respiration); the anaerobic/anoxic condition corresponds to that when the organism(s) performs fermentation (without external terminal electron acceptors) or uses chemicals other than O2 as terminal electron acceptors (anaerobic respiration) (1).

Accordingly, the microaerobic condition may be defined as the transition condition when the organism(s) performs simultaneous aerobic and anaerobic respiration/fermentation.

Being ubiquitous and having versatile metabolic capability, Pseudomonas aeruginosa was studied in this work. The Gram-negative bacterium P. aeruginosa is an opportunistic pathogen which frequently causes pneumonia (5, 6), nosocomial bloodstream infections (7), and serious lung infection in the cystic fibrosis patients (8, 9).

In cystic fibrosis airway infection, the formation of biofilm by P. aeruginosa confers an inherent antibiotic resistance (10, 11) and the bacterium undergoes the phenotype change from nonmucoid (with flagella or type IV pili) to mucoid (alginate capsule) (12, 13). In addition to the loss of motility, the phenotype transition causes significant changes in the expression of virulence factors. Large amounts of alginate and exopolysaccharides are produced and other virulence factors such as exotoxin A, exoenzyme, elastase, alkaline 2

protease, phospholipase and rhamnolipids are released (14). The alginate and exopolysaccharides help bacteria adhere to the host cells, form the microcolony matrix, evade the host immune system response (8), and protect against phagocytosis, antibodies and antibiotic treatment (13, 15, 16). The dissolved oxygen level (DO) in the environment plays an important role in the phenotype change. It has been reported that the bacteria grow fast without mucoid under aerobic conditions, but turn mucoid and form biofilm matrix under microaerobic or anaerobic conditions in the presence of nitrate

(13, 17). In another study of biofilm formation of P. aeruginosa, the anaerobic biofilm was found 3-fold thicker and 1.8-fold higher in cell viability than the aerobic biofilm

(18). Monitoring the bacterium’s physiological responses at different DO is therefore very important to the understanding of its metabolism and respiratory mechanisms.

Online monitoring of the fluorescence from NAD(P)H, i.e., the reduced coenzymes nicotinamide adenine dinucleotides (phosphate), has been used on the cell culture for many years. Based on the correlation of fluorescence profile with other process parameters, on-line NAD(P)H fluorescence technology will demonstrate to be an ideal tool for monitoring sensitive changes of cellular physiology and providing insight to the shift of e--accepting mechanisms of P. aeruginosa under the microaerobic conditions.

On the other hand, to mineralize aliphatic hydrocarbons, P. aeruginosa strains degrading aromatic and polyaromatic hydrocarbons have also been isolated (1, 3). Many strains of the bacterium produce effective biosurfactants (rhamnolipids) when growing on hydrophobic and hydrophilic substrates (19). Biosurfactants are substances widely used in various industrial processes such as pharmaceutic, cosmetic, petroleum, and food production because of their low toxicity, excellent moisturizing properties, and skin 3

compatibility. Rhamnolipids are among the most effective surfactants known today.

Being extremely effective in emulsifying and solubilizing hydrocarbons, rhamnolipids are recognized for their potential in enhanced oil recovery and in mobilizing non-aqueous phase liquid (NAPL) contaminants in soils and aquifers (4). Rhamnolipids have also been planned as pesticides because of their antibacterial, antifungal, mycoplasmacidal, and antiviral activities, and as the source of the fine chemical rhamnose (20, 21, 22). For example, the use of rhamnolipids biosurfactants for the treatment of leaves of Nicotiana glutinosa infected with tobacco mosaic virus led to a 90% reduction in the number of lesions (6). It has been reported that 1% of rhamnolipids decreased the potato virus X content of systematically infected N. tabacum L. Sansun by 46% and secondarily infected leaves by 43% (6).

The project also examined the production of rhamnolipids by P. aeruginosa under the aerobic, microaerobic, anaerobic denitrifying conditions.

1.1 Scope of Research

This study mainly investigated the rhamnolipids production and the cell metabolism at the different DO levels and three dilution rates based on the glucose biodegradation by P. aeruginosa ATCC 9027. The effects of DO and D on various culture properties were observed in the continuous cultures respectively, especially on aerobic respiration and denitrification. The relationship of the denitrification-to- respiration ratios (respiration mechanism) and NFU fractions was also primarily detected by online NAD(P)H fluorescence techniques at aerobic, microaerobic and anaerobic denitrifiying conditions.

1.2 Research Objectives

The main goal of research was to investigate the effects of different DO and D for cell metabolism and respiration mechanism of P. aeruginosa and the rhamnolipid production in the glucose-limited continuous culture under the aerobic, microaerobic, and anaerobic/anoxic conditions.

The specific objectives of this project were to:

1. Observe the transition of microbial electron-accepting respiratory mechanism under different DO with NAD(P)H fluorescence technique;

2. Explore effects of dissolved oxygen levels on the production of secondary metabolites (rhamnolipids) by P. aeruginosa.

3. Study the metabolism of P. aeruinosa using glucose as a substrate under microaerobic-denitrifying conditions in continuous-culture systems.

This research also utilized the quenching-based luminescence technique for measuring very low DO in the broth and determined the correlation of denitrification and respiration monitored by online NAD(P)H fluorescence techniques.

1.3 Structure of Thesis

An extensive literature survey of this research was described in the Chapter II, including the structure and nature of the rhamnolipids and biofilm formation. The concepts of the fluorescence technology, luminescence techniques for DO measurement and the synthesis of rhamnolipids etc were also enclosed in the Chapter II. All the materials and methods of whole study as well as the experiment setup were introduced in the Chapter III. In the Chapter IV, the cell properties, cell metabolism and microbial electron-accepting respiratory mechanism at different dilution rates and different DO

conditions were covered first. Second, the effects of different DO and D for the production of rhamnolipids were also discussed. Subsequently, the correlations of denitrification-to-respiration ratios with NAD(P)H fluorescence fractions were represented at anaerobic, microaerobic and aerobic conditions. Moreover, those two fractions obtained from none-zero-DO and zero-DO systems combined with DO and

NFU fractions at the same studied D (0.06 h-1). Finally, a cleaner relationship between those fractions was established to significantly improve the applicability of online

NAD(P)H fluorescence in monitoring and quantitatively describing the microraerobic state of the microorganisms. In Chapter V, conclusions were summarized based on the experimental results of this research and suggestions for further studies of this research were given.

CHAPTER II

LITERATURE SURVEY

The contents of this chapter concentrate on the background information related to this research. The aims of this work are to study the mechanism of respiration and the production of biosurfactant, rhamnolipid, by Pseudomonas aeruginosa ATCC 9027 under the aerobic, microaerobic, anaerobic conditions. The biofilm formation and the association phenotype changes of Pseudomonas aeruginosa are also important to this research and those backgrounds will be briefly introduced in the first section. Two luminescent techniques are used to substantiate the feasibility of the research.

Fundamental concepts of NAD(P)H fluorescence and luminescence technique for DO measurement are covered first. In the subsequent sections, the specific pathway of aerobic and anaerobic hydrocarbon metabolism and the structure, properties of biosurfactants as well as biosynthesis of rhamnolipids are reviewed. The respiratory chain will be involved in the last section.

2.1 Biofilm Formation

The microbial biofilm has received much recent attention, in part owing to scientific communities’ acknowledgements that biofilms are ubiquitous in natural, industrial, and clinical environments. Interest in clinically related research on biofilms has been particularly great. This is not surprising—the difficulty of eradicating biofilm bacteria with antibiotic treatment is a prime concern of medicine. Depending on the

organism and type of antimicrobial and experimental system, biofilm bacteria can be up to a thousand times more resistant to antimicrobial stress than free-swimming bacteria of the same species. Understanding the nature of the increased antimicrobial resistance is a central goal of much clinical and basic research on medically relevant biofilms today (23,

24, 25, 26). Generally, in the chronic infections, bacteria adapt to their environment in a way that makes them both more persistent and less invasive.

Biofilm formation can occur by at least three mechanisms. One is by the redistribution of attached cells by surface motility (27, 28). Results from O’Toole &

Kolter (29) on studies of Pseudomonas aeruginosa mutants suggest that type IV pili- mediated twitching motility plays a role in surface aggregation for this organism. A second mechanism is from the binary division of attached cells (30). As cells divide, new cells spread outward and upward from the attachment surface to form cell clusters, in a similar manner to colony formation on agar plates. A third mechanism of aggregation is the recruitment of cells from the bulk fluid to the developing biofilm (31). The relative contribution of each of these mechanisms will depend on the organisms involved, the nature of the surface being colonized, and the physical and chemical conditions of the environment.

The individual adherent cells that initiate biofilm formation on a surface are surrounded by only small amounts of exopolymeric material, and many are capable of independent movement (29) by means of pilus-mediated twitching or gliding (Figure 2.1

(34), stage 1). These adherent cells are not yet “committed” to the differentiation process leading to biofilm formation, and many may actually leave the surface to resume the planktonic lifestyle. During this stage of reversible adhesion, (32) the bacteria exhibit

Figure 2.1 Five-stage models for biofilm development (34)

several species-specific behaviors, which include rolling, creeping, aggregate formation, and “windrow” formation (28), before they begin to exude exopolysaccharide and adhere irreversibly (Figure 2.1, stage 2). Davies & Geesey (33) showed that in P. aeruginosa the cluster of genes responsible for alginate production is unregulated within 15 min of the cell’s initial contact with the colonized surface and that this genetic event initiates the process of biofilm formation. As biofilms mature they develop the basic microcolony/ water channel architecture that is now well recognized in natural and in vitro biofilms

(Figure 2.1, stage 3), and many cells alter their physiological processes (e.g., grow anaerobically) in response to conditions in their particular niches. Individual microcolonies may detach from the surface or may give rise to planktonic revertants that swim or float away from these matrix-enclosed structures, leaving hollow remnants of microcolonies or empty spaces that become parts of the water channels (Figure 2.1, stages 4 and 5). Additionally, whole microcolonies may naturally break away from the biofilm (detach without any obvious perturbation to the system), although the mechanisms behind this phenomenon are yet unclear. These processes are not necessarily synchronized throughout the whole biofilm but are often localized so that at any one time a small area on the surface may contain biofilm at each developmental stage.

2.2 Two Luminescent Techniques

2.2.1 Fluorescence

Luminescence techniques are commonly used in biotechnology research, environmental testing, industrial applications, and clinical research (35, 36, 37).

Luminescence is the emission of light frequency ν from a state which is 'excited' by some form of energy, corresponding to E2 -E1=hν. It arises from molecular relaxation from 10

electronically excited states of higher energy E2 to ground state E1, which occurs after a substance absorbs light, heat, X-rays, chemical energy, and so on. The term broadly includes the commonly-used categories of fluorescence and phosphorescence (35).

Fluorescence is an emission that results from the return to the ground electronic state from the lowest energy excited singlet state. The emission rate results in a fluorescence lifetime near 10-8 s. On the other hand, phosphorescence is an emission that results from the transition from the triplet excited state to the lowest singlet ground state. Such a transition is not allowed, and its emission rate is slower than that of the former. Typical phosphorescence lifetimes range from 10-3 s to several seconds. So fluorescence is said to occur where emission ceased almost immediately after withdrawal of the exciting source and where there is no thermal cause, whereas in phosphorescence the emission decays for some time after removal of excitation (36).

Fluorescence is the phenomenon in which absorption of light of a given wavelength by a fluorescent molecule is followed by the emission of light at longer wavelengths. The distribution of wavelength-dependent intensity that causes fluorescence is known as the fluorescence excitation spectrum, and the distribution of wavelength- dependent intensity of emitted energy is known as the fluorescence emission spectrum.

Fluorescence detection has three major advantages over other light-based investigation methods: high sensitivity, high speed, and safety (35). The point of safety refers to the fact that samples are not affected or destroyed in the process, and no hazardous byproducts are generated.

2.2.2 NAD(P)H Fluorescence

NADH is a reduced form of nicotinamide adenine dinucleotide (NAD) liberated as a coenzyme in cells, and NAD+ is the oxidizing form of it. NADH is a fluorophore of high fluorescence intensity, with absorption and fluorescence emission maxima at 340 and 450 nm (37), respectively, whereas NAD+ has no fluorescence. The fluorophore of

NADH is somewhat quenched by collision with the adenine nucleus, but when it is linked to a protein, the fluorescence intensity increase about fourfold because of the transformation in its chemical structure accompanying combination with protein (1, 38).

Cells survive by utilizing available resources or “substrates”, like nutrients, nitrates, and oxygen, to generate forms of energy and cellular building blocks useful to them. All living cells contain NAD(P) coenzymes, which serves as the major intermediate electron carriers in cellular metabolism. The cyclic nature of NAD(P) for heterotrophs is summarized in Figure 2.2 (39). Accompanying substrate catabolism, the oxidized forms of coenzymes, NAD(P)+, are reduced to NAD(P)H. NADPH primarily serves as the reducing power in biosynthetic reactions (anabolism). NADH, on the other hand, is oxidized back to NAD+ through the following reactions: In aerobic respiration

(reaction 3 in Figure 2.2), NADH is directly involved in oxidative phosphorylation for

ATP generation. Under anoxic conditions, certain microbial species can use oxidants

3+ 2- 4- 0 such as nitrate/nitrite (also, Fe , CO2/CO3 , SO , S , and some organic compounds) as the terminal electron acceptors. The NAD(P) cycle for nitrate/nitrite respiration thus is

- very similar to that for aerobic respiration, with NOX replacing the role of oxygen

(reaction 2 in Figure 2.2). In anaerobic fermentation, no externally supplied electron acceptor is required. The regeneration of NAD(P)+ from NAD(P)H is coupled with the

reduction of an organic compound that is formed during catabolism. For example, the anaerobic uptake of acetate in a biological nutrient removal (BNR) process may involve the reduction of NAD+ to NADH with the conversion of glycogen to acetyl-CoA via the

EMP (Embden-Meyerhof-Parnas) pathway and the oxidation of NADH with the synthesis of poly(α-hydroxybutyrate) (PHB) from acetyl-CoA (reaction 1 in Figure 2.2)

(75).

While the reduced coenzymes, NAD(P)H, are fluorescent, the oxidized counterparts, NAD(P)+, are not. The fluorescence of NAD(P)H thus depends on the kinetic balance of its generation (by catabolism) and consumption (by respiration and anabolism) and is extremely sensitive to the change of cellular electron-accepting mechanism (29).

NAD(P)H fluoresces emitted at 460 nm when irradiated with 340-nm light (87).

Monitoring of the fluorescence of intracellular NAD(P)H therefore is an effective way to obtain information on biological activity.

The fluorescence technique has been applied to many microbial fermentations and animal cell cultures to generate information on cellular metabolism and to provide better process control in large-scale industrial operations (36, 37, 38, 39, 40, 44, 65).

With an-online fluorometer (BioGuide system, BioChem Technology, King of

Prussia, PA), our group (40, 41, 42) and others (43, 44, 45) have demonstrated this in biological wastewater treatment processes, resting cultures in minimal media, and various fermentations.

Metabolic Intermediates

& NAD(P)H CO 2 Electron Acceptor Catabolites Fluorescent

Anabolism &

Reaction 1, 2, or 3 Catabolism

+ NAD(P) Anabolites , Energy

Substrate Non-fluorescent & Reduced Product

1 2 3 Anaerobic Anoxic Oxic Organic NO X O2 Intermediates

PHA N 2 Alcohols, Acids H2O H O 2

Figure 2.2 The cyclic nature of coenzymes NAD(P) and NAD(P)H

The signals obtained by the fluorometer are composite and complex, partly intrinsic to optical instruments, partly reflecting the optical “dirtiness” and variability of biological broth, and partly caused by the relatively broad excitation/emission bandwidths employed by the fluorometer (46, 47, 48, 49).

Quantitative correlation of the fluorescence, e.g. with the biomass concentration, is therefore difficult and system dependent.

The complexity, however, will not compromise its usage in this study for detecting the shift of electron-accepting mechanisms.

As shown in Figure 2.3, clear and instant fluorescence changes accompany such shifts (40).

The fluorescence was highest under anaerobic (non-denitrifying) condition, lowest under aerobic condition, and intermediate under anaerobic denitrifying condition.

2.2.3 Influence of Various Factors on Fluorescence in Solutions

Solvents: generally solvents affect the fluorescence characteristics of substances greatly.

As aspects of interaction between molecules of solute and solvent, there is the intermolecular transfer of an electron in a fluorophore due to polarity of the solvent, electrostatic action between dipole moments, dispersions, hydrogen bonds and so on.

When the rates of internal conversion and intersystem crossing are affected by these factors, the fluorescence emission spectrum and its quantum yield are variable (36).

Temperature: It is very an important factor to affect the fluorescent signal of fluoro-meter. The higher the temperature in the solution rises, the lower fluorescence

Figure 2.3 Fluorescence of P. aeruginosa under different electron accepting conditions.

intensity becomes by reason of the increasing probability of intermolecular collision and the loss of potential energy due to the radiationless deactivation. As a practical problem accompanying a rise of temperature in the solution, the wavelength of the fluorescence emission spectrum shifts slightly, so it is preferable to measure the fluorescence intensity at lower temperature (36, 37). pH: Generally, the variation of pH in solution has an influence on the fluorescence excitation and emission spectra at the time of transition of the proton.

Quenching: When a fluorophore coexists with some other substances, the fluorescence intensity falls remarkably because the fluorescence quantum yield decrease on account of radiationless deactivation by their mutual collision and other factors. Such phenomena are called quenching. When fluorophore and quencher are expressed as F and Q, respectively, the quenching processes can be written as follows:

F + hνa → F* (Photoabsorption)

F* → F + hνf (Emission of Fluorescence)

F* + Q → F + Q (Quenching by foreign molecule)

F* + F → F + F (Quenching by the same molecule)

Principal examples of the quenching effect for fluorophores are described below.

Transition elements, particularly the colored metal ions such as Cr3+, Fe3+, Ni2+, and Cu2+ and anions like Br-, I- and NO3-, are all quenchers of fluorescence. Generally, the fluorescence intensity of a fluorophore in the gas or liquid state decreases with an increase in its concentration after the fluorophore reaches above a certain level. This phenomenon is termed concentration quenching. Moreover, in solvents with a group as

such C=C or C=O, the quenching effect is smaller than in solvents with C-N, C-Cl, or C-

Br groups (35, 36, 37).

2.2.4 Luminescence Technique for DO Measurement

Oxygen quenches the luminescence of many fluorescent and phosphorescent materials (50, 51), with a hyperbolic dependency (Stern-Volmer relationship) as shown in

Figure 2.4 for tris(2, 2’-bipyridiyl) ruthenium (II) dichloride (52).

The higher sensitivity at low DO is more ideal for microaerobic measurements.

Detection of oxygen at ppb levels has been reported, based on the quench- phosphorescence of trypaflavine (53) and palladium-porphyrine complex (52, 54).

2.2.4.1 The Principles of the Fiber-optic Oxygen Meter (PreSens Precision Sensing

GmbH, Regensburg, German)

Fiber-optic oxygen meter with micro-sensors PSt3 and MEF14 is applied to measure DO at different levels in this study. The principle of the sensor operation is based on the quenching of luminescence caused by collision between molecular oxygen and luminescent dye molecules in the excited states (55).

The collision between the luminophore in its excited state and the quencher

(oxygen) results in radiationless deactivation and is called collisional or dynamic quenching. After collision, energy transfer takes place from the excited indicator molecular to oxygen which consequently is transferred from its ground state (triplet state) to its excited singlet states.

As a result, the indicator molecular does not emit luminescence and the measurable luminescence signal decreases.

Figure 2.4 Oxygen quenching of the fluorescence of tris(2, 2’-bipyridiyl) ruthenium (II) dichloride

2.3 Biosurfactant Production

2.3.1 Structures and Properties of Biosurfactants

Biosurfactants display a range of structures but have the common ability to cause emulsification of oil-water mixture. Accordingly, biosurfactants must be able to dissolve, at least partially, in both water and water-immiscible liquid, thereby affecting a decreased surface tension enabling mixing and microsolubilization (i.e., emulsification). Many microbes appear to produce a complex mixture of biosurfactants, particularly when living on water-immiscible substrates. Among microbes, a majority of biosurfactants are found to be produced by bacteria (56). Generally, biosurfactants are microbial metabolites with typical amphiphilic structure of a surfactant, where the hydrophobic moiety is either a long chain fatty acid, or an α-alkyl-β-hydroxy fatty acid whereas the hydrophilic moiety can be a carbohydrate, an amino acid, a cyclic peptide, phosphate, carboxylic acid, alcohol, etc (57). Physical and chemical properties, surface tension reduction, and stability of the emulsion formed are very important in search for a potential biosurfactant.

These properties are used in evaluating biosurfactants and in screening potential microorganism for biosurfactant production.

Synthetic surfactants are usually classified according to the nature of their polar group. However, biosurfactants are commonly differentiated on the basis of their biochemical nature and the microbial species producing. Major classed of biosurfactants include 1) glucolipids, 2) phospholipids and fatty acids, 3) lipopeptide/lipoprotein, 4) polymeric surfactants, and 5) particulate surfactants. Some microbes produce cell wall bound biosurfactant, to this group of microorganisms belong Candida Lipolytica and C. tropicalis, which produce cell wall bound lipopolysaccharides when growing on n-

alkanes (58, 59, 60) and Rhodococcus erythropolis and several Mycobacterium and

Arthrobacter species, which synthesize nonionic trehalose corynomycolates (61, 62, 63,

64). On the other hand, Pseudomonas and Torulopsis species are well known to excrete extracellular glycolipids, especially rhamnolipids (65, 66, 67, 68, 69, 70) and sophorolipids (71, 72, 73), respectively. Glycolipids are carbohydrates in combination with long-chain aliphatic acids or hydroxy aliphatic acids. The most commonly isolated and characterized biosurfactants are glycolipids, lipopeptides and lipopolysaccharides

(74). Most intensively studied glycolipid surfactants are the sophorolipids and rhamnolipids (70). Certain species of Pseudomonad are known to produce large amounts of the glycolipids, rhamnolipids, which contain one or two molecules of rhamnose sugar- ring linked to one or two molecules of β-hydroxydecanoic acid (75). The presence of rhamnolipids is commonly detected by the reduction of the interfacial/surface tension or by the thin-layer chromatography techniques.

2.3.2 Synthesis of Biosurfactants

2.3.2.1 Microorganisms for Rhamnolipid Production

The microbial species that is inherently capable of producing rhamnolipids is

Pseudomonas aeruginosa. Various P. aeruginosa strains, specifically, DSM 2874,

ATCC 9027, ATCC 10145, UG2, etc., have been reported to be excellent producers of rhamnolipids (76, 77, 78, 79). Rhamnolipids can also be produced by P. aeruginosa or P. putida by genetically cloning the essential genes from P. aeruginosa for the synthesis of rhamnolipid-producing enzymes, but not by Escherichia coli (80). P. aeruginosa isolated from the soil contaminated with polycyclic aromatic hydrocarbons (PAHs) (81). The sludges contaminated by hydrocarbons adjacent to petrochemical industries or crude oil

spills (82) were found to furnish rhamnolipid production. The production of rhamnolipids can be performed in batch, continuous, or semi-continuous processes.

Rhamnolipid production by the growing, free resting, and immobilized cells was studied by Wagner et al. (74). The highest production was obtained with the free resting cells.

2.3.2.2 Choice of Carbon Substrates and Limiting Nutrients

Various carbon sources can be used for rhamnolipids synthesis, for example, n- paraffin (60), glycerol (65), proteose peptone-glucose salts (83), ethanol (84), stearic acid

(85), corn oil (71), etc. Pseudomonas species can also utilize hydrocarbons for growth in aerobic condition (22, 86). However, in the absence of oxygen, the microorganism can not digest the saturated hydrocarbons. This was observed when using hexadecane as the carbon source for P. aeruginosa ATCC 10145 (87). It is known that the initial oxidation step of saturated aliphatic hydrocarbons involves molecular oxygen as a reactant and monooxygenase as the enzyme. One of the atoms of the oxygen molecule is incorporated into the oxidized hydrocarbon as a hydroxyl group, with the second atom of oxygen being reduced to water. Once there is no oxygen available under anaerobic condition, the microorganism cannot accomplish this metabolic pathway.

Different cultivation conditions and medium compositions influence the productivity and crude product composition of rhamnolipids (85, 74). The influence of various nitrogen and carbon sources on the rhamnolipid production from glycerol was reported by Arino et al. (81). It was found that the nitrogen concentrations around 3 – 5 g/L with the mix substrate of glycerol and ethyl dodecanoate yielded the highest rhamnolipid production. The best rhamnolipid production was reported mostly under aerobic nitrogen-limiting conditions, in the presence of excess carbon and phosphorus

concentrations (88, 89, 74). Magnesium limitation also causes rhamnolipid production with the highest yield obtained when the carbon-to-magnesium ratio in the fresh feed to continuous culture is 364 or higher (91). Guerra-Santos et al. (90) reported that the change of potassium, sodium, or calcium concentrations has no effect on the rhamnolipid production. The optimal sodium chloride concentration was 100 mM when P. aeruginosa DSM 2874 was grown on glucose medium (85). Linhardt et al. (71) suggested the optimal iron concentrations of 50-100 µg/L for the production of rhamnolipids by P. aeruginosa UI29791 in the corn oil medium. However, rhamnolipid production from glycerol was reported to be inhibited by organic acid salts such as sodium succinate, sodium citrate, and sodium acetate (66).

2.3.2.3 Temperature and pH Effects

P. aeruginosa can produce rhamnolipids in the temperature range of 23 – 43°C, with the best production reported in the range of 27 – 37°C (85, 90). The cultivation pH influences the degree of foam formation. Above a pH of 6.5, foam formation was much higher than that at lower pH values. The optimum pH for rhamnolipid production is dependent on the strains, for example, the best between 6.2 – 6.4 suggested by Guerra-

Santos et al. (90), while minimal pH effect between 6 and 7 was suggested by Arino et al.

(81).

2.3.2.4 Foaming in Biosurfactant Fermentation

A serious problem in biosurfactant production under aerobic condition is the extensive formation of foams. With the conventional submerged aeration, very stable foams are generated in the presence of rhamnolipids at concentrations greater than 0.1

g/L (87). This causes serious problems, including the high expenditures for foam control and also the decreased productivity.

Due to rapid foaming in large volume and high foam stability, mechanical foam breaker fails to control the problem. On the other hand, chemical antifoam agents may affect downstream processing, especially the performance of filtration unit (92). Their possible influence on cell metabolism and the pollution of the reactor effluent have to be considered as well. This serious foaming problem could be eliminated if there is no or less need of aeration for the culture’s respiration. P. aeruginosa is a facultative aerobe that can use either oxygen or nitrate as the terminal electron acceptor. Nitrate addition may therefore be used to replace the aeration in the process for meeting cells’ respiration demand. The research on the rhamnolipid production under aerobic, microaerobic and anaerobic condition is presented in Chapter IV and Chapter V

2.3.3 Rhamnolipid Types and Rhamnolipid Production

Rhamnolipids were first reported to be produced from the fermentation of glycerol by P. aeruginosa in 1949 (93). Hauser and Karnovsky further investigated the biosynthesis of rhamnolipids and its production in 1954 (94, 66)as shown in Figure 2.5.

Rhamnolipids, the brownish solid material, also fall into this case. They consist of one or two molecules of rhamnose and one or two molecules of β-hydroxydecanoic acid.

Besides the abundant 3-hydroxydecanoic acid moiety shown in Figure 2.5, 3- hydroxyoctanoic, 3-hydroxydodecanoic and 3-hydroxydodecenoic acid moieties were also recently observed in rhamnolipid structures (81).

2.3.3.1 Rhamnolipid Formation

The synthesis of rhamnolipids in vivo using glycerol and sodium acetate as the substrates was investigated by Hauser and Kanovsky (94, 95, 96).

It was found that, in the system using glycerol and sodium acetate as the mixed substrate, the carbons from glycerol can furnish all carbons in both sugar and acid parts of rhamnolipids, while the carbon from acetate can supply only the carbon of the β- hydroxydecanoic acid part.

The rhamnose moiety seemed to be derived by the condensation of two 3-carbon units formed from glycerol and the carbon of fatty acid seemed to be synthesized by fatty acid biosynthesis pathway from 2-carbon units. The de novo rhamnolipid synthesis by the extracted enzymes from P. aeruginosa has been investigated by Burger et al. (65, 97).

These enzymes, called rhamnosyl transferases, catalyze the reactions between the precursors of the sugar and the acid parts in order to form rhamnolipids. The sugar part, which is the rhamnosyl donor, is thymidine diphosphate L-rhamnose (TDP-L-rhamnose) and the acid part is β-hydroxydecanoyl-β-hydroxydecanoate.

Ochsner et al. (98, 99) have further studied the DNA genes involved in rhamnolipid biosynthesis and confirmed the mechanism demonstrated by Burger et al.

The rhlR and rhlI genes were identified as the regulatory genes which are necessary for the transcriptional activation of the rhlA and rhlB genes that encode the rhamnosyltransferase enzyme. The rhlA gene promoter was found to be expressed low in the exponential growth phase, but it increases 20-fold during the stationary phase. The rhlB protein is the catalytic protein of the rhamnosyltransferase. The binding of

O CH CH O 2 C CH CH2 COOH HO O CH (C H ) 3 2 6 (C H2)6

OH OH CH3 CH3 Rhamnolipid 1

O CH CH2 COOH HO O CH 3 (C H2)6 Rhamnolipid 2 OH OH CH3

O CH CH O 2 C CH CH2 COOH HO O CH (C H ) 3 2 6 (C H2)6

CH CH OH O 3 3 HO O CH3 Rhamnolipid 3 OH OH

O CH CH2 COOH HO O CH 3 (C H2)6

CH OH O 3 HO O CH 3 Rhamnolipid 4

OH OH

Figure 2.5 Four types of rhamnolipids found in Pseudomonas species. 26

activated rhlR protein to target sites upstream of the rhlA promotor enhances transcription of the rhlA and rhlB operon and subsequently form the rhamnosyltransferase (rhlAB), which is the key enzyme capable of rhamnolipid biosynthesis. The biosynthesis of rhamnolipids is shown in Figure 2.6.

2.4 Respiration

2.4.1 Aerobic Respiration

Oxygen plays an important role in nature: The aerobic respiration (combustion of glucose) is the main energy source of living organisms (Figure 2.2).In the course of this process, molecular oxygen (oxidation state 0) is reduced to the level of the oxide

(oxidation state -2). In the photosynthesis, this reaction is inversed thanks to solar energy, thus recycling molecular oxygen. The principal source of vital energy is the combustion of glucose during the respiration process, according to:

C6H12O6 + 6O2 → 6CO2 + 6H 2O

This reaction is exothermic and takes place in living organisms by taking a complicated reaction path that allows storing the liberated energy in the form of molecules (ATP).

This biological combustion requires molecular oxygen and produces carbon dioxide and water, the most important substrate of this combustion being glucose (1). In contrast to its technological analogue, biological combustion occurs in 3 well-defined steps: 1) the glycolysis, which transforms glucose (a sugar containing 6 carbon atoms) into 2 pyruvate molecules (3 carbon atoms), 2) the citric acid cycle, which transforms the 2 pyruvate molecules into 6 carbon dioxide molecules, 3) the respiration chain, in which molecular oxygen is reduced to water (1).

O HO O O O HO CH C O CH COOH CH3 CH 2 CH 2 P O P O Thymidine O + (CH 26) (CH 26) HO HO - - CH CH O O 3 3 Thymidine-diphospho-rhamnose (TDP-rhamnose) beta-Hydroxydecanoyl-beta-hydorxydecanoate

Rhamnosyltransferase 1

TDP

rhamnosyl- O transferease

O CH CH2 C O CH CH2 COOH rhl rhl rhl rhl HO O A B R I CH (CH ) (CH ) Genes 3 26 26 CH CH autoinducer 3 3 synthetase HO HO

Rhamnolipid 1 L-rhamnoosyl- beta-hydroxydecanoyl- beta-hydroxydecanoate TDP-rhamnose

Rhamnosyltransferase 2

TDP

O CH CH2 C O CH CH2 COOH HO O CH 3 (CH 26) (CH 26) CH CH 3 3 HO O

HO O Rhamnolipid 3 CH3 L-rhamnosyl-L-rhamnosyl beta-hydroxydecanoyl- beta-hydroxydecanoate HO HO

Figure 2.6 Biosynthesis of Rhamnolipids. 28

2.4.2 Anaerobic Respiration

- - An anaerobic process is where microorganisms utilize oxides of N (NO3 , NO2 ,

3+ 4- 4+ N2O), Fe (Fe ), S (SO ), or Mn (Mn ) as alternative electron acceptors when O2 is absent (soil). In aerobic conditons oxidation proceeds , ultimately with the formation of carbon dioxide and water. The oxidation of anaerobic respiration is usually incomplete and various organic compounds are produced including methane, and ethylene and acetic acid. All these oxidation/reduction reactions are driven by the soil microorganism. As the soil becomes more anaerobic the demand for alternative electron acceptors increases,

- 4+ 3+ 2- and reduction occurs in the order: O2>NO3 >Mn >Fe >SO4 >CO2 (100).

2.4.2.1 Denitrification

- - An anaerobic process is where microorganisms utilize oxides of N (NO3 , NO2 ,

- 4+ 3+ 2- and reduction occurs in the order: O2>NO3 >Mn >Fe >SO4 >CO2 (100).

- - The terms refers to the reduction of NO3 or NO2 to N2 and oxides of nitrogen by

- microbial activity (biological denitrification), and to the chemical reduction of NO2 and other unstable nitrogen compounds (chemical denitrification). Biological denitrification is often called dissimilatory denitrification to distinguish it from assimilatory

denitrification in which microorganism assimilate and reduced nitrate as a first step in protein synthesis. The process of denitrification is a stepwise reduction:

- - NO3 ----> NO2 ---> X ---> N2O---> N2 (2.4.1) where the intermidiate X might be nitric oxide, NO, though this is not certain. Many bacterial genera possess enzymes for each step in the reduction of nitrate to nitrogen gas.

Some take the reduction only to nitrite, and others may be unable to reduced nitrous oxide. Many genera with the capability to denitrify are found in soils, but it is difficult to know which are the most active. Species of Pseudomonas and Alcaligenes have commonly been isolated and bacteria of both genera are known to denitrify (101).

Most denitrifying bacteria are facultatives anaerobes; they use organic substances as a source of energy and are therefore hetertrophs. In aerobic conditions they use O2 as the electron acceptor but can adapt to the absence of oxygen, or to low concentration of oxygen, aby using oxygenated forms of nitrogen, for example nitrate, as alternative electron acceptors, which are thereby reduced (100, 101).

The reduction of nitrate to nitrogen gas required a series of enzymes. The enzyme that reduces nitrate to nitrite is called nitrate reductase, and nitrite is reduced in turn by nitrite reductase. In the course of reduction the gases nitric oxide, NO, and nitrous oxide,

N2O, are evolved and diffuse into the atmosphere. Under the most reducing conditions the main product is nitrogen gas. Nitrous oxide is implicated in global warming, and soil is its main source.

Nitrous oxide reductase has been found to be inhibited by acetylene. Use has been made of this observation to measure denitrification. By blocking the final step in the reduction, total denitrification can be measured as N2O production, which is possible with sensitive gas chromatograohs, whereas measurement of comparatively small amounts of

N2 is almost imposible because of its high background concentration in the atmosphere

(101, 102).

Denitrification requires the presence of nitrate, metabolized carbon compounds and the almost complete absence of oxygen at the site if reduction. Nitrate is formed by nitrification or might be added in fertilizer. Soil organic matter, plant roots and organic manures provide metabolizable carbon compounds. The concentration of oxygen is reduced to a sufficiently low level when the soil air is displayed by water, as after heavy rainfall or irrigation, or flooding. The soil does not need to be devoid of oxygen because denitrification will occur at microsites that are anerobic, for example within water- saturated aggregates or where an energy-rich substrate cause oxygen depletion, even though the soil as a whole contains oxygen. The rate of denitrification increases with temperature and is highest at a soil pH between about 6 and 8. Under more acid conditons not only is it slower but the ration of N2O : N2 is higher, either because of chemical reduction of nitrite to N2O or inhibition of N2O reductase. It has also been found that nitrate inhibits N2O reductase, which results in a higher ration of N2O : N2 in its presence

(103).

2.4.3 Microaerobic Denitrification

Low-oxygen or even anaerobic conditions are found below the oxygenated surface waters of lakes and oceans and, except for the deep sea, aquatic sediments are anoxic below a thin oxidized surface layer. These enviroments, in which oxygen access is restricted or absent, are inhabited by oxygen-sensitive bacteria well adapted to these conditions.

In recent years, interest has been directed toward studying P. aeruginosa under low dissolved oxygen, or microaerobic, conditions (in our group). Unfortunately, the methods used to control microaerobic conditions in laboratory experiments and to monitor low concentrations of residual dissolved oxygen are not standardized, and the extent of oxygen limitation varies considerably from study to study. For example, initial dissolved oxygen (DO) concentrations of 2 and a poised constant level of less than 1 mg/l were both considered to be microaerobic in separate studies (26, 28) while in another study, microaerobic conditions were created by adding sufficient levels of biodegradable material to an aerobic (DO = 8.9 mg/L) solution so that the oxygen demand exceeded the oxygen available and DO was depleted over time (2). In a fourth study, cultures were incubated in sealed serum bottles under 98% N2:2% O2 headspace (47). Chawala has also conducted a preliminary experiment to confirm the existence of microaerobic denitrification by P. aeruginosa (104). The results have shown under the microaerobic conditions studied, the denitrification rate decreased with decreasing DO for hydrocarbon. With decreasing DO, this step limited the rate of hydrocarbon oxidation, which in turn slowed the electron donation to drive the electron-accepting denitrification.

On the other hand, the oxygen repression/inhibition of denitrifying enzymes is expected

to occur at high DO (105). The denitrification rate for oxygenated, anaerobically degradable substrates (such as glucose or fatty acid) probably decreases always with increasing DO because of the prevailing oxygen repression/inhibition effect. In this study, the luminescence techniques will be used to measure the low DO involved as well as to monitor the transition of microbial electron-accepting mechanism based on glucose substrate.

CHAPTER III

MATERIALS AND METHODS

3.1 Microorganism and Medium

The microorganism used in this study was P. aeruginosa (ATCC 9027). The stock culture was maintained at 4 °C after lyophilization in 10% skim milk. For activation the culture was maintained on the PTYG agar plates (Peptone 5g/L, Tryptone

. . 5g/L, Yeast Exatract 10 g/L, Agar 15 g/L, MgSO4 7H2O 0.6 g/L, CaCl2 2H2O 0.07 g/L) and subcultured regularly. For inoculum’s preparation, a loop of cells from an agar slant was transferred into a test tube containing 10 mL of 3% TSB solution and incubated at

25°C for 24 hours. The cells with the medium were added to a 500-mL Erlenmeyer flask with 100 mL of a basal medium supplemented with 2 mL of 400 g/L glucose and pH adjusted to 7.0. The basal medium consisted of: glucose, 20 g/L; NH4Cl, 3 g/L; K2HPO4,

0.7 g/L; MgSO4•7H2O, 1 g/L; and FeSO4, 0.45 mg/L. The magnetic stirrer was used at

300 rpm and the broth was incubated at room temperature (24 ± 2 °C) for one day. Then the culture was used as the inoculum to the batch culture systems. The medium for batch culture had the following composition: glucose, 10 g/L; NaNO3, 10 g/L; NH4Cl, 6 g/L;

KH2PO4, 0.7 g/L; NaCl, 0.5 g/L; MgSO4•7H2O, 0.18 g/L; CaCl2, 0.01 g/L; MnCl2•4H2O,

0.01 g/L; FeSO4, 0.01 g/L; and antifoam (Trans-280, Trans-Chemco, Inc., Bristol, WI),

0.5 g/L. After growing at room temperature (24 ± 2 °C) in a magnetically stirred (2000-

mL) fermentor for seven days, the culture was ready to used for the continuous culture fermentation. The glucose concentrations of the fresh feeding media are decreased in gradient following by the increased dilution rates in order to keep the gluocsed-limited cluture avoid the effect of the diution rates. The glucose concentrations of different continuous systems are 10 g/L at D = 0.026h-1, 8 g/L at D = 0.06 h-1, 6 g/L at D = 0.13h-1.

A relatively high concentration of NH4Cl was included so that NaNO3 was consumed only for denitrification, not for N-source of assimilation (105). The antifoam was found necessary because of the extreme foaming associated with the rhamnolipids produced by P. aeruginosa. The results of antifoam study were shown in Figure 3.1.

The experiment was carried out in two days and 100 mL medium and the recipes were: glucose, 20 g/L; NH4Cl, 3 g/L; K2HPO4, 0.7 g/L; MgSO4•7H2O, 1 g/L; and FeSO4, 0.45 mg/L. The antifoam concentrations, 0 g/L, 0.01 g/L, 0.05 g/L, 0.1 g/L, 0.2 g/L and 0.5 g/L, were used in the experiment respectively.

The cell concentration profiles were very similar at the different antifoam concentrations. It concluded that the antifoam had no detectable effects on cell metabolism, at the concentration employed.

3.2 Experimental Setup and Continuous Culture

The experimental setup was shown in Figure 3.2. The continuous culture was conducted in a 2-L glass fermentor with the working volume of 0.7 L of medium. The fermentation pH was maintained at 6.5 ± 0.1 by automatic addition of either NaOH or a mixed solution of HNO3 and NaNO3. The pH control was achieved by using a set of pH probe and controller (Ingold Mettler Toledo, LaGrange, IL).

6 0 g/L 5 0.01 g/L 0.05 g/L 4 0.1 g/L 3 0.2 g/L 0.5 g/L 2

1 C ell Concentration (g/L) 0 0 1020304050 Time (h)

Figure 3.1 Cell concentration profiles at the different antifoam concentrations

Temperature was maintained at 35.0 ± 0.2 °C. The flow rate of influent air stream, to the headspace of the fermentor, was kept at a constant 60 mL/min. To maintain the continuous culture at different DO levels, an adjustable air pump was used to circulate the air from the fermentor headspace through (in the given order) a cooling condenser, the circulation pump, a sterile 0.22-µm filter, and an air stone placed at the bottom of the fermentor, through which the air was introduced back into the broth as fine bubbles. The filter was used to ensure sterility, and the condenser to reduce moisture so that the filter would not be wet and clogged by the water condensed otherwise in the circulation line. There was also a separate exhaust gas stream leaving the fermentor, whose oxygen concentration was measured using an oxygen analyzer (FC-1B Oxygen

Analyzer, Sable Systems, Henderso, NV) for determination of the culture’s oxygen uptake rate (OUR) from oxygen balance (as described later in more details). The OUR results would be less accurate if the airflow rate was high and the determination depended on very small differences between two large concentrations.

3.3 Analytical Methods

Glucose consumption was studied with Pseudomonas aeruginosa ATCC 9027 continuous cultures maintained in aerobic, microaerobic, and anaerobic/denitrifying condition (at different DO levels). To control cell concentration, the glucose concentration in the fresh medium is carefully calculated to be the limiting nutrient. The three dilution rates used in this study were kept at 0.026, 0.60, and 0.13 h-1. Each dilution run begins with highest DO (fully aerobic), followed by lower DO level until zero DO

(the anaerobic condition) is reached. Five to six DO levels will be chosen to

Fresh Air Refrigerated Water Bath/Circulator Air Circulation Pump

PH Controller

PC O Analyzer 2

Acid/Base DO Meter PC NAD(P)H Fluorometer Effluent Fresh Medium Temperature Stir Plate Controller Air Stone

Luminescence Based DO Sensor

Liquid Flow Gas Flow Signal Cable

Figure 3.2 Experimental setup

measure the cell properties and culture fluorescence. After setting each new air circulation rate (as described above), the culture was maintained for at least 4-5 days

(corresponding to the time for replacing the broth volume thrice) and then the cell concentration was followed daily for at least 3 more days to ensure the reach of a constant steady-state cell concentration. Culture fluorescence was measured and periodical samples of about 10 mL were taken from the fermentor. The samples were centrifuged to separate cells from the supernatants, at 10,000 rpm for 10 minutes. Cell and nitrite concentrations were analyzed immediately. The remaining supernatants were frozen at -18°C for other analyses later.

3.3.1 Cell Concentrations and Cell Dry Weight Analysis

For cell concentrations, the optical densities (OD) of broth samples, after a known-fold dilution to the right linear range (0.1-0.6), were measured at 460 nm and converted to cell dry-weight concentrations according to a pre-established calibration curve. The cell dry-weight concentrations were determined by washing the cell pellets

(collected by centrifugation) once with deionized water, and then drying the washed cells to constant weight in an aluminum weighing-dish (Fisher Scientific) at 110 °C for at least

3 h. The calibration curve was established by drying the washed cells at different OD values.

3.3.2 Ammonium and Nitrate Analysis

- The ammonium-N and NOx -N (including both nitrate-N and nitrite-N) analyses were made using an ammonia electrode (M-44325, Markson Science (106, 41). The ammonium concentrations could be measured accurately in a wide range (1-1000 mg/L

+ - - of NH4 -N) while the NOx concentrations only in 1-20 mg/L of NOx -N. The sample was 39

therefore diluted to the proper range prior to the analysis and poured into a U-tube with a small magnetic stirring bar for mixing. The ammonia electrode was inserted through one end of the U-tube and the reagent was added from the other end. 0.3 mL of an ionic- strength adjusting solution (ISA, Fisher Scientific Co) and 1.5 mL of 10 N NaOH were added. The steady mV reading was recorded and converted to the ammonium concentration using a pre-established calibration (Appendix A-1). For nitrate analysis, 0.3 mL of titanous chloride solution (20%, Fisher Scientific Co.) was added to reduce nitrate

(and nitrite) to ammonia. The next steady reading (mV) was taken. It corresponded to the combined ammonium, nitrate concentrations present in the medium.

3.3.3 Nitrite Analysis

The nitrite analysis followed the standard method described by Gerhardt (111).

5 mL of the sample was placed in a test tube. 0.1 mL of sulfanilamide 1% APHA (5173-

5, LabChem Inc.) was added and the sample allowed to stand in the dark for 2-8 minutes.

0.1 mL of N-1 (Naphthyl)-ethylenediamine dihydrochloride, 0.1% APHA (7132-6,

LabChem Inc.) solution was then added, mixed immediately, and allowed to stand for at least 10 more minutes. Then the absorbance was measured by the spectrophotometer

(UV/VIS Spectrophotometer, Perkin Elmer Lambda 3B) at 543 nm.

3.3.4 Glucose Analysis

The glucose concentrations were determined by using the enzymatic glucose assay kit obtained from Sigma Diagnostics (Procedure No. 510). The glucose oxidase method is based on the simultaneous use of the enzymes, glucose oxidase and peroxidase, in two concurrent reactions. A chromogenic oxygen acceptor, o-Dianisidine, is able to form a more intense brown color when oxidized by peroxidase. The original glucose

concentration is proportional to the intensity of the color formed. This intensity can be measured with UV-Vis spectrophotometer at wavelength between 425-475 nm.

3.3.5 Rhamnolipids Analysis

The analysis of rhamnolipids used in this research work was similar to that used by Wu (87), Osman (84) and Chayabutra (104). To quantify the rhamnolipids, the sample was adjusted to pH 2.0 with 1 N HCl and extracted with ethyl acetate of double volume at room temperature. The organic phase was then dried at 40°C. The residue was re- dissolved in 0.05 M sodium bicarbonate solution. This aqueous phase was quantified for the concentrations of rhamnolipids as rhamnose concentration, following the standard anthrone method (107). 3.3 mL of the anthrone solution (2 g/L of anthrone in concentrate sulfuric acid) was prepared and mixed with 1.7 mL sample solution at 5°C.

The mixture was heated at 95°C for 16 minutes, before the absorbance at 625 nm was measured by a UV/VIS Spectrophotometer (Perkin-Elmer, Lambda 3B).

3.3.6 DO and OUR Measurement

DO was measured by optical micro-sensors PSt3 and MEF14 (PreSens Precision

Sensing GmbH, Regensburg, German), according to the quenching of luminescence caused by collision between oxygen and luminescent dye molecules in the excited state

(51). The oxygen-sensitive dye was immobilized in silicone matrix (125 µm in thickness) and attached onto a flexible transparent polyester foil. A small piece (5 mm × 5 mm) of the autoclavable sensing matrix was glued inside the glass wall of fermentor with silicone glue. The oxygen concentration of the broth in contact with the sensing matrix was then monitored from outside through the fermentor wall using an optical fiber, and the data

logged and analyzed with computer software. PSt3 had a larger measurement range (0-

45 mg/L, i.e., 0-500% air-saturation) but lower accuracy (0.01 mg/L). MEF14 had a smaller range (0-1.8 mg/L, i.e., 0-20% air-saturation) but higher accuracy (0.001 mg/L).

In this study, PSt3 was used at DO above 0.6 mg/L for its more stable signals while

MEF14 was used for more accurate measurements at lower DO.

Oxygen uptake rate (OUR) was calculated from the material balance on oxygen, using the measured oxygen concentrations and gas flow rates of the influent and effluent gas streams. The gas-phase oxygen concentrations were measured with an oxygen analyzer (FC-1B Oxygen Analyzer, Sable Systems, Henderson, NV) having an accuracy of 0.0001%. The gas flow rates were measured with a flow meter (Valved Acrylic

Flowmeters, Cole-Parmer, Vernon Hills, IL) that was controlled by a gas flow controller

(Mass Flow Meter/Controller Electronics, Sable Systems, Henderson, NV) having an accuracy of 0.2%.

3.3.7 Culture Fluorescence Measurement

Culture fluorescence was monitored by an online fluorometer (the BioGuide

System, BioChem Technology, Inc., King of Prussia, PA). The fluorometer was designed for monitoring the fluorescence of intracellular NAD(P)H, with excitation wavelength of 340 ± 20 nm and emission wavelength of 400 – 480 nm. After the culture reached the steady state under a specific DO, the fluorescence intensity was recorded

(denoted here as NFU, i.e., in Normalized Fluorescence Unit). Both the influent air valve and circulation air pump were then turned off. Cell respiration quickly depleted the DO in the medium and turned the culture to fully anoxic denitrifying condition, causing a sharp increase in fluorescence to a level at the denitrifying condition (NFUDN). Next, a

high airflow rate was introduced to create the fully aerobic condition and the corresponding fluorescence (NFUOX) was also recorded.

3.3.8 Calculations

As described above, the following properties were measured/determined

-1 0 experimentally: dilution rate (D, h ); nitrate-N concentration in the fresh feed (CNA , mg/L); nitrate-N and nitrite-N concentrations in the effluent broth (CNA and CNI, mg/L);

A nitrate-N concentration in the acid added for pH control (C NA , mg/L), average rate of the acid addition (QA, L/h), and the broth volume (V, L).

Accordingly, nitrate and nitrite reduction rates (NAR and NIR, in mmol/L-h) of the continuous culture could be calculated from the material balances:

0 A A NAR = [(CNA − CNA ) * D + CNA * (Q /V )] /14

0 A A NIR = [(CNA − CNA − CNI )* D + CNA *(Q /V )]/14 where 14 is the molecular weight of N.

Oxygen uptake rate (OUR, in mmol O2/L-h) was determined from oxygen balance:

OUR = (G /V )*(Cin − Cout ) where G is the gas flow rate (L/h), and Cin and Cout are the gas-phase oxygen concentrations (mmol/L) in the influent and effluent gas streams.

Cell yield from glucose (Yx/s) was calculated as:

X Y = x / s S 0 − S where X (g/L) is the steady-state cell concentration, and S0 and S (g/L) are the glucose concentrations in the fresh feed and the broth, respectively. 43

For bioenergetics of P. aeruginosa, the known pathways for electron acceptance and ATP generation (from respiratory chain) are shown in Figure 3.3 (108).

Accordingly, the ATP formation rate in respiratory chain (FRATP, in mmol/L-h) and the fraction of electrons accepted by anaerobic respiration could be calculated using the NAR (to nitrite), NIR, and OUR determined above:

FRATP = NAR + 3* NIR + 6*OUR

Rateof e− thru denitrification Fraction = Rateof e− thru aerobicrespiration + Rateof e− thru denitrification

2* NAR + 3* NIR = 4*OUR + 2* NAR + 3* NIR

The specific rates, per unit cell concentration, of the above volumetric rates (per unit culture volume) were calculated by dividing the volumetric rates by the corresponding cell concentration (X).

The denitrification-accepted e- fraction, (2*NAR + 3*NIR)/(2*NAR + 3*NIR+

4*OUR), is a quantitative indicator for the “extent” of denitrification. In this study, it was correlated with the culture fluorescence fraction, (NFU - NFUOX)/(NFUDN - NFUOX), another quantitative measure of the culture’s “fractional approach” to the completely denitrifying state.

Figure 3.3 Electron acceptance and ATP formation in respiratory chain of P. aeruginosa

CHAPTER IV

AEROBIC, MICROAEROBIC, ANAEROBIC DENITRIFICATION OF PSEUDOMONAS AERUGINOSA

4.1 Introduction

Monitoring the bacterium’s physiological responses at different DO and D is very important to understand its metabolism and respiratory mechanisms. Cell respiration is the process by which the chemical energy of food molecules is released and partially captured in the form of ATP. Carbohydrates, fats, and proteins can all be used as fuels in cell respiration, but glucose is most commonly used as an example to examine the reactions and pathways involved. In this work, the cell metabolism and respiration mechanism are investigated in the glucose-limited continuous culture.

Respiration has some types, for example aerobic respiration, microaerobic respiration, anaerobic respiration, and photorespiration (109). Most of cells continue to respire when deprived of oxygen. Some researchers have reported the cell activity in the aerobic respiration, which is higher than that observed in the anaerobic condition (110,

111). Some studies showed that the use of nitrate as terminal electron acceptor is not totally inhibited by the very high dissolved oxygen concentration (DO) that present in the medium (112). The enzymes of nitrate respiration are only formed when specific conditions for denitrification are present (105). Anaerobic conditions alone are sufficient to partially derepress all of the enzymes involved in the nitrate respiration, but the presence of nitrate or nitrite intensifies the response. The synthesis of denitrifying enzymes is repressed in the presence of oxygen. Oxygen blocks the synthesis of

respiratory nitrate reductase in E. coli at the level of transcription and later step of enzyme formation (113). Although the derepression of denitrifying enzymes is thought to require strict anaerobiosis, the activity of some of the performed enzymes in the pathway may persist under aerobic conditions. It has been known that oxygen inhibits the reduction of nitrate and the formation of nitrogen gas by denitrifying bacteria (114,

110, 111, 113). Addition of oxygen to cultured cell suspensions or reaction mixtures containing extracts of denitrifiers results in inhibition of the pathway at some level and thus a rapid loss of the capacity for reduction of nitrate and other nitrogen oxides (113).

However, denitrification proceeded during aerobic conditions (at very high DO > 1.0 mg/L) in our experiments (It will be discussed later.).

The investigation of rhamnolipid production, on the other hand, was another main point in this research, especially under microaerobic and anaerobic conditions. Some research workers found that the rhamnolipid production displayed higher values at a very low DO or none-DO under minimal aeration (112). This observation was especially significant for the rhamnolipid-associated pathogenicity of P. aeruginosa, which resides in microaerobic to anoxic biofilms in airway mucus of cystic fibrosis patients (115).

Up to now, luminescence techniques have been used wildly in biotechnology research, environmental testing, industrial applications, and clinical research because of their high sensitivity, high speed, and safety. Online monitoring technique of the fluorescence from NAD(P)H, i.e., the reduced coenzymes nicotinamide adenine dinucleotides (phosphate), was used in our group (40, 41, 42) and others groups (43, 44,

45) that demonstrated that in biological wastewater treatment processes, resting cultures in minimal media, and various fermentations. The background of NAD(P)H fluorescence

and luminescence technique of DO measurement have been described in Chapter II. The optics-based DO meter and NAD(P)H fluorometer were applied to examine the cell metabolism and respiratory mechanism of the bacteria, Pseudomonas aeruginosa, at different DO levels under diverse dilution rates in this research. Two quantitative indicators, the normalized fraction (NFU-NFUOX)/(NFUDN-NFUOX) and the fraction of electrons (e-) accepted by denitrification (2*NAR + 3*NIR)/(4* OUR + 2*NAR +

3*NIR), were correlated to represent the culture’s denitrifying state (NFU was the fluorescence level of a specific continuous culture, and NFUOX and NFUDN were the briefly perturbed fluorescence levels at fully aerobic and anaerobic-denitrifying conditions, respectively.). Compared to DO, the NAD(P)H fluorescence was shown to be a much more sensitive and useful indicator for the microbial activity under the microaerobic conditions.

4.2 Cell Metabolism and Respiration Mechanism at D = 0.026h-1, D = 0.06 h-1 and D

= 0.13 h-1

4.2.1 Materials and Methods

The microorganism, culture media, and experimental methods used were similar to those reported previously (Chapter III). Briefly, the microorganism studied was P. aeruginosa

ATCC 9027. The feed for the continuous culture were different at different dilution rates in order to maintain the glucose-limited systems. The nutrients at D = 0.026h-1 had the following composition: glucose, 10 g/L; NaNO3, 10 g/L; NH4Cl, 6 g/L; KH2PO4, 0.7 g/L;

NaCl, 0.5 g/L; MgSO4•7H2O, 0.18 g/L; CaCl2, 0.01 g/L; MnCl2•4H2O, 0.02 g/L; FeSO4,

0.01 g/L; and antifoam (Trans-280, Trans-Chemco, Inc., Bristol, WI), 0.5 g/L. The culture media at D = 0.06 h-1 and D = 0.13 h-1 were similar to those described above but

glucose concentrations used in the system were 8 g/L and 6 g/L, respectively. For the systems at D = 0.06 h-1 with different air circulation rates, all the composition of the culture were increased 1.2 times to ensure to maintain the cell growth rate (glucose, 12 g/L; NaNO3, 10 g/L; NH4Cl, 7.2 g/L; KH2PO4, 0.84 g/L; NaCl, 0.6 g/L; MgSO4•7H2O,

0.216 g/L; CaCl2, 0.012 g/L; MnCl2•4H2O, 0.024 g/L; FeSO4, 0.012 g/L; and antifoam

(Trans-280, Trans-Chemco, Inc., Bristol, WI), 0.5 g/L).

The samples taken periodically from the continuous cultures were measured for concentrations of cells, glucose, ammonium, nitrate, nitrite and rhamnolipid.

These measured concentrations were used to calculate nitrate reduction rate

(NAR), nitrite reduction rate (NIR) and their specific rates (per unit cell concentration).

Oxygen concentrations in the inlet and outlet streams were measured and used to calculate OUR.

ATP formation and the electrons accepted by denitrification were calculated from

NIR, NAR, OUR.

The methods for analyses and calculations were the same as those reported previously (Chapter III).

4.2.2 Cell Properties in Continuous Cultures Maintained at Different DO and D

The steady-state culture properties measured in continuous cultures maintained at different DO levels and D were summarized in Figure 4.1, Figure 4.2, Figure 4.3 and

Figure 4.4.

The dilution rates used (0.026 h-1, 0.06 h-1 and 0.13 h-1) were very low when

-1 compared to the culture’s maximum specific growth rate (µmax = 0.23 h ), estimated

2.2 0.20 2.0 1.8 1.6 1.4 0.15 -N (g/L) -N (g/L) - -N (g/L) 1.2 - + 2 3 4 1.0 NO NO NH 0.8 0.10 0.6 0.4 A 0.2 0.0 0.05 0.00.20.40.60.81.01.21.4

0.20 4.0 1800 3.8 1600 (NFU) 0.15 3.6 1400 3.4 1200 3.2 0.10 1000 3.0 800 Cell (g/L) Cell Rhamnolipid (g/L) 2.8 600 0.05 2.6 400 2.4 B Culture Fluorescence 0.00 2.2 200 0.0 0.2 0.4 0.6 0.8 1.0 1.2 1.4 DO (mg/L)

Figure 4.1 Steady-state concentrations measured in continuous cultures maintained at different DO (D = 0.026 h-1)

1.8 0.010

1.6 A 0.008 1.4

1.2 0.006

-N (g/L) 1.0 3 NO2-N (g/L)

NO3-N (g/L) 0.004 NH 0.8 0.6 0.002 0.4 0.2 0.000 0.00.51.01.52.02.5 0.040 3.5 1000

0.035 3.0 900

0.030 2.5 800

0.025 2.0 B 700

0.020 1.5 600

0.015 1.0 500 Cell Concentration (g/L)

0.010 0.5 400 Culture Fluorescence (NFU) Rhamnolipid Concentration (g/L) 0.005 0.0 300 0.0 0.5 1.0 1.5 2.0 2.5 DO (mg/L)

Figure 4.2 Steady-state concentrations measured in continuous cultures maintained at different DO (D = 0.06 h-1)

0.010 2.5

0.008 2.0 A

1.5 0.006 -N (g/L) 3

NO3-N (g/L) 0.004 (g/L) NO2-N NH 1.0

0.5 0.002

0.0 0.000 0.0 0.5 1.0 1.5 2.0 0.24 3.0 640

0.22 620 2.5 600 0.20 580 2.0 0.18 560 0.16 1.5 540

0.14 520 1.0 500 0.12 B

Cell Concentration (g/L) 480 0.5 0.10 460 (NFU) Fluorescence Culture Rhamnolipid Concentration (g/L) 0.08 0.0 440 0.0 0.5 1.0 1.5 DO (mg/L)

Figure 4.3 Steady-state concentrations measured in continuous cultures maintained at different DO (D = 0.13 h-1)

1.6 0.5

1.4 (A) 0.4 1.2 0.3 -N (g/L)

1.0 - 2 -N (g/L) -N (g/L) - + 3 4 0.8 0.2 NO NO NH 0.6 0.1 0.4

0.2 0.0 0 100 200 300 400 500 600 2.0 4.5 7 (B) 4.0 6 1.5 3.5 5

3.0 4 1.0 Cell (g/L) 2.5 3 Glucose (g/L)

Rhamnolipid (g/L) 2.0 2 0.5 1.5 1

0.0 1.0 0 0 100 200 300 400 500 600 Air Circulation Rate (mL/min)

Figure 4.4 Steady-state concentrations measured in continuous cultures maintained at different air circulation rates. (D = 0.06 h-1)

from the exponential growth observed in the batch fermentation (The data were not shown here.). At such low dilution rates, the concentrations of the limiting nutrient, glucose, were practically zero in the broths (They were not reported in the Figure 4.1.) except at the zero-DO points (Figure 4.2 and Figure 4.3) and the cell concentrations at zero-DO points were much lower than those at non-zero-DO points (The

µ * S equation, µ = D = max (µ is the cell growth rate, S is the substrate), can be used to K + S explain that phenomena.).

As shown in Figure 4.1A, Figure 4.2A, Figure 4.3A and Figure 4.4A, ammonium

+ -1 concentrations were present in excess (0.6 – 1.6 g/L of NH4 -N at D = 0.026 h , 0.82 g/L

+ -1 + -1 - 1.3 g/L of NH4 -N at D = 0.06 h , 1.1 g/L – 1.5 g/L of NH4 -N at D = 0.13 h , and 0.85

+ -1 – 1.34 g/L of NH4 -N at D = 0.06 h with different air circulation rates) to ensure that nitrogen source from ammonium was enough for cell growth and nitrate was consumed only for respiration, because the assimilatory nitrate reductases are repressed by ammonium (105).

In the systems of three different dilutions (Figure 4.1A, Figure 4.2A, Figure

4.3A), the nitrate concentrations at DO = 0 mg/L were much higher (except that D =

0.026 h-1) than those at non-zero-DO levels. It was because cell denitrification caused nitric acid adding and made the nitrate concentrations of the broths increased. The nitrate concentrations increased with increasing DO (non-zero-DO systems) (0.2 - 1.2 g/L of

-1 -1 NO3-N at D = 0.026 h , 0.5 - 0.8 g/L of NO3-N at D = 0.06 h , and 0.5 – 0.75 g/L of

-1 NO3-N at D = 0.13 h ), indicating a decrease in the nitrate reduction rate (NAR). The decreasing trends of calculated specific NAR were shown in Figure 4.5, Figure 4.6,

Figure 4.7 and Figure 4.8. The decrease with increasing DO was consistent with the well-known repression and inhibition effects of oxygen on nitrate reductases (114). It indicated that increasing aeration or DO levels hold denitrification, in general, and leads to a higher remaining nitrate concentration. Denitrification, however, persisted (a specific NAR of 0.2-0.3 mmol/g-h at D = 0.026 h-1, a specific NAR of 1.3-1.6 mmol/g-h at D = 0.06 h-1, a specific NAR of 3.2-3.6 mmol/g-h at D = 0.13 h-1, a specific NAR of

2.2-2.4 mmol/g-h at D = 0.06 h-1 with different air circulation rates) at relatively high DO levels ( 1-2.2 mg/L). Although rare, similar phenomena, termed aerobic denitrification, have been observed in several microbial species/strains (112, 116), including P. aeruginosa (112, 117). The occurrence of aerobic denitrification in the current study was also supported by the observations associated with pH changes: The culture pH was observed to decrease under fully aerobic conditions, but increase under fully anoxic denitrifying conditions because of the removal of nitric acid. The acid addition rates required for pH control in the continuous cultures were found to increase with decreasing

DO. For the system maintained at D = 0.026 h-1 with DO of 0.6 mg/L, no addition of either base or acid was necessary, indicating that both aerobic respiration and denitrification took place and the two mechanisms reached a balance in their effects on culture pH.

Nitrite concentrations were given in Figure 4.1A, Figure 4.2A, Figure 4.3A and

Figure 4.4A and the profiles remained at the relatively low values (0.09-0.10 g/L of

— -1 — -1 — NO2 N at D = 0.026 h , 0-0.003 g/L of NO2 N at D = 0.06 h , 0-0.002 g/L of NO2 N

-1 — -1 at D = 0.13 h , 0 g/L of NO2 N at D = 0.06 h with different air circulation

3.0

2.5 ]

-1 2.0

1.5 Specific NIR Specific NAR Specific Specific OUR Specific 1.0 [mmol (g*h)

0.5

0.0 0.00.20.40.60.81.01.21.4

DO (mg/L)

Figure 4.5 Specific rates of oxygen uptake, nitrate reduction and nitrite reduction determined for continuous cultures maintained at different DO (D = 0.026 h-1)

4.5 4.0 3.5 3.0 2.5 Specific NIR Specific OUR Specific NAR 2.0 [mmol/g-h] 1.5 1.0 0.5 0.0 0.0 0.2 0.4 0.6 0.8 1.0 1.2 1.4 1.6 1.8 2.0 2.2

DO (mg/L)

Figure 4.6 Specific rates of oxygen uptake, nitrate reduction and nitrite reduction determined for continuous cultures maintained at different DO (D = 0.06 h-1)

5 ] -1 4

Specific NIR Specific 3 Specific OUR Specific Specific NAR Specific

[mmol (g*h) 2

0 0.00.30.60.91.21.51.8

DO (mg/L)

Figure 4.7 Specific rates of oxygen uptake, nitrate reduction and nitrite reduction determined for continuous cultures maintained at different DO (D = 0.13 h-1)

1.8

6.0 1.6

5.4 1.4

4.8 1.2

4.2 1.0

3.6 0.8 Specific NIR [mmol/(g-h)] Specific NAR [mmol /(g-h)] Specific OUR [mmol /(g-h)] [mmol OUR Specific 3.0 0.6

2.4 0.4

1.8 0.2 0 100 200 300 400 500 600 Air Circulation Rate (mL/min)

Figure 4.8 Specific rates of nitrate reduction (NAR), nitrite reduction (NIR) and oxygen uptake (OUR) at different air circulation rates (D = 0.06 h-1)

rates) in three different D systems. The specific nitrite reduction rates calculated were also shown in Figure 4.5, Figure 4.6, Figure 4.7 and Figure 4.8. They were only slightly smaller than those for nitrate reduction at the same D conditions. The zero nitrite concentrations interpreted that all of the nitrite formed from nitrate reduction was immediately reacted, therefore, NAR = NIR (The nitrite formed from nitrate reduction was immediately converted through the subsequent pathway in denitrification (to NO,

N2O and N2), leaving only a relatively constant residual concentration.).

As shown in Figure 4.1B, Figure 4.2B, Figure 4.3B and Figure 4.4B, the steady state cell concentrations increased significantly as DO increased from fully denitrifying conditions to aerobic conditions (Cell concentrations increased from 2.4 to 3.6 g dry cells

/L as DO increased from 0 mg/L to ~ 0.6 mg/L at D = 0.026 h-1, from 2.0 to 3.2 g dry cells /L as DO increased from 0 mg/L to ~ 0.1 mg/L at D = 0.06 h-1, from 0.5 g/L to 2.6 g dry cells /L as DO increased from 0 mg/L to ~ 0.08 mg/L at D = 0.13 h-1, from 1.5 to 4.0 g dry cells/L as the air circulation rate increased from 0 to 250 ml/min at D = 0.06 h-1 with different air circulation rates.), and then remained relatively constant at higher DO levels.

The calculated cell yields (from consumed glucose) reflected a similar trend

(Figure 4.9, Figure 4.10, Figure 4.11 and Figure 4.12A), and can be attributed to the higher energy (ATP) yield of aerobic and microaerobic respiration than denitrification.

(More discussion was given later.) The culture fluorescence (Figure 4.1B) measured by the NAD(P)H fluorometer also exhibited the similar curve at D = 0.026 h-1 and D= 0.06 h-1, however, the trends of the culture fluorescence weren’t consistent with the

0.40 45 0.38 ) 40 0.36 ] X/S 0.34 35 -1 0.32 30 0.30 Cell Yield (Y 0.28 25 [mmol (L*h) 0.26

20 ATP Gerneration Rate 0.24 0.22 15 0.0 0.2 0.4 0.6 0.8 1.0 1.2 1.4 DO (mg/L)

Figure 4.9 Cell yields and ATP generation rates determined for continuous cultures maintained at different DO (D = 0.026 h-1)

80.0 0.40

0.38 ) 70.0 X/S

] 0.36

-1 60.0 0.34 50.0 0.32 Cell Yield (Y ATP Generation Rate ATP Generation 40.0 0.30 [mmol (L*h) 0.28 30.0 0.26 20.0 0.0 0.2 0.4 0.6 0.8 1.0 1.2 1.4 1.6 1.8 2.0 2.2

DO (mg/L)

Figure 4.10 Cell yields and ATP generation rates determined for continuous cultures maintained at different DO (D = 0.06 h-1)

120 0.5 ) 100 0.4 X/S ] -1 80 0.3

60 0.2 Cell Yield (Y [mmol (L*h)

ATP Generation Rate 40 0.1 20 0.0 0.0 0.3 0.6 0.9 1.2 1.5 1.8

DO (mg/L)

Figure 4.11 Cell yields and ATP generation rates determined for continuous cultures maintained at different DO (D = 0.13 h-1)

.38 80 .36

) 70 .34 X/S .32 60 .30 .28 50 [mmol/(L-h)]

Cell Yield (Y (A) .26 ATP Generation Rate 40 .24 .22 30 0 100 200 300 400 500 600

140 500 )

) (B) 120

450 ATP/X

ATP/S 100 400

80 g] 350

[mmol/g] 60 mmol/ 300 [ ATP Yield (Y 40

20 250

0 200 ATP Yield per Cell (Y 0 100 200 300 400 500 600 Air Circulation Rate (mL/mim)

Figure 4.12 Calculated culture properties, including specific rhamnolipid production rate, ATP generation rate, cell yield (YX/S), ATP yield (YATP/S), and ATP yield per cell yield -1 (YATP/X), at different air circulation rates (D = 0.06 h ).

cell concentration trends at D = 0.13 h-1 (Figure 4.3B) although they increased significantly as DO increased from the zero-DO value to none-zero-DO values at the same D conditions. The phenomena clearly reflected the composite and complex nature of the fluorescence signals, partly intrinsic to optical instruments, partly reflecting the optical “dirtiness” and variability of biological broth, and partly caused by the relatively broad excitation/emission bandwidths employed by the fluorometer (46). The observations indicated that the total culture fluorescence measured was primarily from some interfering fluorophores other than NAD(P)H, and the interferences were much stronger under aerobic than anaerobic conditions. It demonstrated that the background culture fluorescence has limitation to reflect the cell properties and the normalized

NADH fluorescence method is a good approach to monitor the cell metabolism.

The rhamnolipid production, on the other hand, displayed complex phenomena at different DO levels and D. For example, in the systems of D = 0.026 h-1 the rhamnolipid concentration nearly tripled when the culture was shifted from the anoxic condition

(without any aeration) to a very low DO (0.1 mg/L) under minimal aeration, however, in the systems of at D = 0.06 h-1 with different air circulation rates, the rhamnolipid concentration in the system without air circulation (~ 0.9 g/L) was significantly higher than the concentrations in the systems with air circulation (< 0.2 g/L). There were at least two possible reasons for the higher rhamnolipid production. The first was the effect of glucose availability. In the high rhamnolipid-producing system (DO ~ 0, no air circulation), there was remaining glucose that could support the rhamnolipid synthesis, while glucose was practically exhausted in all of the low producing systems. The second possible factor was the extent of aeration. In both studies, rhamnolipid production

appeared to favor the more anaerobic condition. Overall, it was visible that the rhamnolipid production at low DO (anaerobic or microaerobic conditions) levels was higher than that at higher DO (aerobic conditions) levels at the same D conditions. The positive effect of extremely low DO or none-DO on rhamnolipid synthesis remains to be further investigated. The observations of three dilution rates are significant not only for enhancing the productivity of rhamnolipids in industrial production but also for addressing the role of rhamnolipid-associated pathogenicity of P. aeruginosa, as the low

DO conditions may prevail in human lungs (118).

4.2.3 Respiration Mechanism and Energy Generation at Different DO and D

The calculated values of specific OUR observed in continuous cultures maintained at different DO and D were shown in Figure 4.5, Figure 4.6, Figure 4.7 and

Figure 4.8, together with the specific reduction rates of nitrate and nitrite. The specific

OUR increased from 0.38 to 1.58 mmol/g dry cells-h rapidly with increasing air circulation rates (DO close to 0 mg/L) and remained at the relatively constant level at D =

0.06 h-1 ( in Figure 4.8). At the same studied dilution rate(zero-DO conditions), the maximum specific OUR of this P. aeruginosa strain was ~ 2.8 mmol O2/g cells-h (in

Figure 4.6). In the none-zero systems of D = 0.026 h-1 and D = 0.13 h-1, the specific

OUR also approached the maximal level ~ 1.4 mmol O2/g dry cells-h and ~4.2 mmol

O2/g dry cells-h at the lowest DO (0.1mg/L) under non-anaerobic conditions, respectively. From above the observations, the Monod constant for DO (i.e., the critical

DO at which the specific OUR is half of the maximum rate) was therefore practically zero (< 0.001 mg/L) at an approximate air circulation rate of 400 ml/min for this P. aeruginosa strain, (estimated by interpolation with the curve shown in Figure 4.8). As

mentioned earlier, the consumption rates of nitrate and nitrite decreased with increasing

DO but remained at a significant residual rate (~ 1/8 of the maximal rates at D = 0.026 h-

1, ~ 1/4 of the maximal rates at D = 0.06 h-1 and ~1/2 of the maximal rates at D = 0.13 h-1

) even at high DO levels (> 1.0 mg/L).

Denitrification was therefore not completely repressed or inhibited by oxygen, and could function as a competitive or supplementary electron-accepting and energy- generating mechanism to aerobic respiration.

The profiles of cell yields and ATP generation rates were shown in Figure 4.9,

Figure 4.10, Figure 4.11 and Figure 4.12A.

The two profiles were relatively parallel, both increasing with increasing DO at the same D.

Some points of cell yields appeared to be lower than the levels expected from the corresponding ATP generation rate in these trends, for example at DO = 0.1 mg/L in the systems of D = 0.026 h-1 as well as at DO of 20 ml/min and 40 ml/min in the culture of D

= 0.06 h-1 with different air circulation rates.

The additional energy (and material resources) might have been diverted to the synthesis of other metabolites. For example, a significantly higher rhamnolipid concentration was obtained at this condition at D = 0.026 h-1 (Figure 4.1).

Some other metabolites might exist in the broths also. More investigations on the low DO metabolism are warranted. (More details will be given in Chapter IV)

The specific ATP generation rates (SRATP) at different DO levels under different dilution rates were given in Figure 4.13.

)

(

a R

e 20 n

e 0.14 G 10 0.12

P 0.10 T 1 )

A 0.08 -

c 0 0.06 h

i (

f i 0.04 D c 1.5

e 1.0 0.02 p 0.5 S D 0.00 O (m 0.0 g/L) D = 0.026 h-1 D = 0.06 h-1 D = 0.13 h-1

Figure 4.13 Specific ATP generation rates determined for continuous cultures maintained at different DO and D

The obviously increasing trends of specific ATP generation rates were observe as expected from the continuous cultures with increasing D due to µ was equal to D in the continuous operations, suggesting the faster cells grow , the more energy is need to be generated. However, the profiles of specific ATP generation rates stayed at the similar levels compare at the same D conditions (except DO = 0 mg/L), those values were 10 mmol/g-h at D = 0.026 h-1, ~22 mmol/g-h at D = 0.06 h-1 and ~ 42 mmol/g-h at D =0.13

-1 h . The SRATP in the zero-DO systems were slightly smaller than those in the higher DO systems. There were two reasons: one was that cells grew much slower at the zero-DO level, therefore, the needs of ATP energy were lower at the zero-DO level than at the high-DO levels, the other one was that cell metabolic changes might occur in the systems and transfer the additional energy to compose some metabolites (rhamnolipids, etc.).

The steady-state culture properties measured in continuous cultures maintained at

DO < 0.001 mg/L and different air circulation rates were summarized in Figure 4.4 (A and B). The calculated cell yield (YX/S), specific rhamnolipid production rate, ATP generation rate, ATP yield (YATP/S), and ATP yield per cell yield (YATP/X) were given in

Figure 4.12 (A and B). All of the above properties were plotted aligned with the increasing air circulation rates. The profiles of YX/S (cell yield from consumed glucose) corresponded with overall ATP generation rates in Figure 4.12A, seemingly suggesting that the lower cell yields in the systems with low air circulation rates were caused by the lower energy (ATP) generation. However, closer data analyses revealed that YATP/S

(ATP yield from consumed glucose) was relatively constant around 100 mmol/g (~ 50% of the 206 mmol/g predicted from 37 moles of ATP formed per mole of glucose catabolized aerobically, with glucose being converted to pyruvate via Entner-Doudoroff

pathway (1)). The lower YX/S at no or very low air circulation (0 – 20 mL/min) resulted from their larger YATP/X (specific ATP generation required for cell growth), ~ 40% larger than those at air circulation rate ≥ 60 mL/min (Figure 4.12B).

The higher YATP/X at no or very low air circulation suggested that more energy was consumed for synthesis of extracellular products, which was supported by the results of rhamnolipid production as described in the above.

4.2.4 Culture Fluorescence

The Culture fluorescence in continuous systems of this study were recorded by

NAD(P)H fluoremeter.

The total intracellular NAD(P)H concentration in a culture depends on both the cell concentration and the specific NAD(P)H concentration (per unit cell concentration).

The latter depends on the fraction of the reduced coenzymes [NAD(P)H] in the overall coenzyme pool [NAD(P)H + NAD(P)+]”(34).

NAD(P)H fluoresces emitted at 460 nm when irradiated with 340-nm light (87).

Monitoring of the fluorescence of intracellular NAD(P)H therefore is an effective way to obtain information on biological activity.

NAD(P)H fluorescence signals can therefore be used for monitoring the changing cell concentration (43) and/or cellular activity, especially the electron-accepting mechanism which significantly affects the rate of NAD(P)H oxidation (consumption)

(119).

1294 Air On NFUDN 1292

1290 NFU

1288

1286

1284 NFUOX Culture Fluorescence 1282 Air Off 1280 8 9 10 11 12 13 14 Time (min)

Figure 4.14 An example profile of NAD(P)H fluorescence responding to brief perturbation from the continuous culture’s steady state to aerobic and anoxic conditions (The continuous cultures shown here was maintained at DO of 0.1 mg/L and D of 0.026 h-1)

For example, as shown in Figure 4.14 (for the continuous culture at DO = 0.1 mg/L and D = 0.026 h-1), the steady-state fluorescence (NFU) dropped sharply and instantaneously (to NFUOX) when the aeration rate was increased to make the system fully aerobic. Several minutes later, the aeration was completely shut down. The microbial respiration depleted DO and the fluorescence increased (eventually to NFUDN) as the culture entered the fully denitrifying condition. Although clearly identifiable, the fluorescence changes associated with the shift of electron-accepting mechanisms represented very small fractions (< 1%) of the total fluorescence intensities. One of the observations demonstrated that occurrence in Figure 4.15 (for the continuous culture at D

= 0.026 h-1), where the total and specific culture fluorescence intensities (observed at several continuous cultures maintained at different DO) were plotted against the corresponding steady-state cell concentrations or DO. While there appeared to be a general trend of increasing fluorescence with increasing cell concentration, the specific fluorescence (per unit cell concentration) was relatively constant only at non-anaerobic conditions (~ 410-485 NFU-L/g, for DO ≥ 0.1 mg/L) and was significantly lower at the anaerobic condition (~ 170 NFU-L/g) (Figure 4.15). The phenomena clearly reflected the composite and complex nature of the fluorescence signals, partly intrinsic to optical instruments, partly reflecting the optical “dirtiness” and variability of biological broth, and partly caused by the relatively broad excitation/emission bandwidths employed by the fluorometer (46). The observations indicated that the total culture fluorescence measured was primarily from some interfering fluorophores other than NAD(P)H, and the interferences were much stronger under aerobic than anaerobic conditions.

DO (mg/L)

0.0 0.2 0.4 0.6 0.8 1.0 1.2 1.4 OX 2000 500 OX 1800 450 NFU 1600 400 1400 Specific NFU 350

or 1200

1000 300 DN

800 DN 250 NFU 600 200 400

200 150 Specific NFU Specific 2.2 2.4 2.6 2.8 3.0 3.2 3.4 3.6 3.8 4.0 Cell Concentration (g/L)

Figure 4.15 Total and specific culture fluorescence intensities observed at fully denitrifying (anoxic) and aerobic conditions, plotted against the corresponding cell concentrations or DO to show the effects of background fluorescence from fluorophores other than NAD(P)H (D = 0.026 h-1)

4.2.5 Examine Transition of Electron-Accepting Mechanism Using NAD(P)H

Fluorescence

The above complexity did not compromise the use of culture fluorescence in this study for detecting the shift of electron-accepting mechanisms. The normalized NFU fractions, (NFU-NFUOX)/(NFUDN-NFUOX), were utilized to use analyze the cell metabolism under aerobic, microaerobic and anaerobic conditions. As shown in Figure

4.16 and Figure 4.17, the NAD(P)H fluorescence exhibited step changed upon the shift of microbial electron-accepting mechanisms at the different DO and D levels. The fluorescence level was higher under the anaerobic-denitrifying condition (NFUDN) than that under fully aerobic condition (NFUOX). If NADH was oxidized partly by nitrate/nitrite and partly by oxygen, the fluorescence should be between NFUDN and

NFUOX and the normalized fraction (NFU - NFUOX)/(NFUDN - NFUOX) represents a quantitative measure of the “fractional approach” of the culture to the completely denitrifying state.

With the well-known repression/inhibition effects on denitrification, DO was expected to have a significantly negative effect on the above NFU fraction.

As described earlier, simultaneous aerobic respiration and denitrification was observed to occur in the P. aeruginosa strain used in this study, even under relatively high DO.

With the measurements made in this study, another quantitative indicator for the culture’s “extent” of denitrification could be calculated, i.e., the fraction of electrons

1.0

.6 *OUR+2*NAR+3*NIR) .4

0.0 0.0.2.4.6.81.0 (2*NAR+3*NIR)/(4 (NFU-NFUOX)(NFUDN-NFUOX)

D = 0.026 h-1 D = 0.06 h-1 D = 0.13 h-1

Figure 4.16 Correlation between two different indicators to the culture’s “extent” of denitrification: the fraction of electrons accepted by denitrification versus the NFU fraction (at D = 0.026 h-1, D = 0.06 h-1 and D = 0.13 h-1)

N2, 2000 mL/min 1.0

0 mL/min 0.9 20 mL/min Fraction -

60 mL/min 0.8

250 mL/min 0.7

Denitrification-Accepted e Denitrification-Accepted 600 mL/min

0.6 0.7 0.8 0.9 1.0 (NFU-NFU )/(NFU -NFU ) OX DN OX

Figure 4.17 Correlation between denitrification-accepted fraction of electrons and NAD(P)H fluorescence fraction for continuous cultures at D = 0.06 h-1 under practically zero-DO conditions.

accepted by denitrification (out of the total electrons accepted by both aerobic respiration and denitrification) as described earlier in the Calculations section: (2*NAR +

3*NIR)/(4* OUR + 2*NAR + 3*NIR). Both the NFU fraction and the electron acceptance fraction have values between 0 and 1, with 0 corresponding to fully aerobic metabolism and 1 to fully (anaerobic) denitrifying metabolism (The fully aerobic condition was difficult to be reached in our systems because of the air circulation pump limitation. NFUOX were NAD(P)H florescence measured at DO = 1.5 mg/L, DO = 5 mg/L and DO = 6 mg/L corresponding to the continuous cultures which D were equal to

0.026 h-1,0.06 h-1 and 0.13 h-1 respectively. NFU Fraction didn’t start from zero at D =

0.06 h-1 and D = 0.13 h-1). The two fractions are plotted in Figure 4.16 and Figure 4.17 for potential correlation. The data do not fall on the “ideal” diagonal, which would correspond to a directly proportional relationship. Nonetheless, the fraction of electrons accepted by denitrification increased with an increase in the normalized NFU fraction at three different D. The increase at DO ≥ 0.1 mg/L was gradual, followed by a much sharper increase at the lower DO range as D was equal to 0.026 h-1 and D was equal to

0.13 h-1 in the studied continuous cultures. For the systems at the studied D of 0.06 h-1, the same phenomena occurred (as described above) at both of none-zero-DO and zero-

DO with different air circulation rates continuous cultures. With increasing air circulation rates of zero-DO systems, the denitrification-accepted e- fraction dropped from 0.95 to 0.65 and the NFU fraction decreased from 0.86 to 0.64 ( Figure 4.17), indicating the culture’s shifting from the fully anaerobic, denitrifying condition to aerobic conditions. At air circulation rates ≤ 20 ml/min, the change in fluorescence fraction was more sensitive than that in denitrification-accepted e- fraction (from 0.72 to 0.85). At 77

larger air circulation rates, the opposite was true and the values only changed from 0.65 to 0.72. The culture container had a large diameter (14 cm) and a shallow culture depth

(7 cm). Even at no air circulation condition, the surface aeration from the fresh air stream was enough to provide oxygen that accepted ~ 5% of the electrons and lowered the fluorescence fraction by ~ 15%.

With the measurements obtained at D = 0.023 h-1, 0.06 h-1 and 0.15h -1, the clearer relationship was established between two quantitative indicators (the denitrification- accepted e- fraction and the NFU fraction). The gradual increase could be observed in the plots from the aerobic to microaerobic conditions, and then a sharp increase followed from the microaerobic to anaerobic conditions at the same D. The online NAD(P)H fluorescence meter can be used to monitor and quantitatively describe the cell metabolism at different states. It is very feasible to utilize NAD(PH) fluoremeter to detect the “microaerobic” state of microorganisms when DO meter is out of detecting limitation.

4.2.6 Combined Two Fractions at D = 0.06 h-1

In Figure 4.18, the fractions obtained at D = 0.06 h-1 with different air circulation rates of zero-DO systems were combined with those obtained at the same dilution rate with non-zero DO systems. (To have non-zero DO, these systems were conducted using higher air circulation rates and a feed medium with a lower glucose concentration to give lower cell concentrations.) The profile obtained in this study for systems with non-zero

DO was a rather flat curve (less sensitive changes in the e--accepting fraction than in the fluorescence fraction). The most sensitive changes in the denitrification-accepted e- fraction occurred at the fluorescence fractions of ~ 0.6 – 0.72 while the DO levels can not

be detected (Apparatus limitation). The significance of the unique shape of the correlation between the two fractions remains to be further elucidated. Nevertheless, the results demonstrated again that the applicability of the online NAD(P)H fluorescence in monitoring and quantitatively describing the sensitive microaerobic state of microorganisms.

2.5 2.0 1.5 1.0

DO (mg/L) 0.5 0.0 1.0

0.8 Fraction -

0.6

0.4

0.2 Denitrification-Accepted e

0.0 0.2 0.4 0.6 0.8 1.0 (NFU-NFUOX)/(NFUDN-NFUOX)

Figure 4.18 Correlation between denitrification-accepted fraction of electrons and NAD(P)H fluorescence fraction for continuous cultures at D = 0.06 h-1 under various DO and aeration conditions. (Results were from two sets of experiments, with the presumed connection drawn as the dashed line, using feed media with two different glucose concentrations.)

CHAPTER V

CONCLUSIONS

5.1 Conclusions

The cell metabolisms and respiration of the strain of P. aeruginosa ATCC 9027 were studied at aerobic, microaerobic and anaerobic conditions in this project. The phenomena of this strain used in this study were shown that cells perform aerobic denitrification with the glucose-limited continuous culture at different dilution rates

(0.026h-1, 0.06 h-1, 0.13 h-1) and different DO levels. The zero-DO systems with different air circulation rates at D = 0.06 h-1 also were observed after the Monod constant for DO

(i.e., the critical DO at which the specific OUR is half of the maximum rate) was found apparently lower than 0.1 mg/L. On-line NAD(P)H fluorescence technology was demonstrated to be an ideal tool for monitoring sensitive changes of cellular physiology and providing insight to the shift of e--accepting mechanisms of P. aeruginosa under the microaerobic conditions. The following conclusions can be drawn from those studies.

1. The ammonium concentrations were sufficient to ensure that the nitrate was only

devoted for respiration in all the continuous and steady-state systems at three different

D levels. Nitrite formed from nitrate reduction was immediately consumed and

caused that specific NAR values were similar with specific NIR. The decreasing

trends of specific NAR, NIR were shown with increasing DO or air circulation rates

in the Figure 4.5, Figure 4.6, Figure 4.7, and Figure 4.8. Those decrease depended

on the repression and inhibition effects of oxygen reductases, however, denitrification 81

still persisted (specific NAR ≠ 0) even at relatively high DO levels (1.3-2.2 mg/L) in

those continuous systems. Aerobic denitrification appeared to function as a

mechanism of electron acceptance that was supplementary or competitive to aerobic

respiration.

2. For this P. aeruginosa strain, the Monod constant for DO (i.e., the critical DO at

which the specific OUR is half of the maximum rate) was therefore practically zero

(< 0.001 mg/L) at an approximate air circulation rate of 400 ml/min (Zero-DO

systems, Apparatus limitation).

3. The steady-state cell concentrations increased significantly as DO increased from 0

mg/L to 0.1-0.6 mg/L in the three different D systems (~ 0.6 mg/L at D = 0.026 h-1, ~

0.1 mg/L at D = 0.06 h-1 and at D = 0.13 h-1, and ~ 250 ml/min at D = 0.06 h-1 with

different air circulation rates (zero-DO systems)), and then maintain relatively

constant at high DO levels (3.6 g dry cells /L in Figure 4.1, 2.7 g dry cells /L in

Figure 4.2, 2.6 g dry cells /L in Figure 4.3 and 4.0 g dry cells/L in Figure 4.4). Those

reflected the higher energy (ATP) yields generated at the aerobic respiration

conditions than at the denitrification conditions. Cell yields and ATP generation rates

increased with increasing DO in Figure 4.9, 4.10, 4.11 and 4.12 but the cell yields

sometimes appeared to be lower than the levels expected from the corresponding ATP

generation rates such as when DO was 0.1 mg/L at D = 0.026 h-1 and DO were 0.1,

0.5, 1.0 1.6 mg/L at D = 0.06 h-1. As we known, the preeminent role of ATP is to

capture and transfer the free energy within cells. Cell metabolic changes might occur

in the systems and transfer the additional energy to compose some metabolites

(rhamnolipids, etc.).

4. The specific ATP generation rates at different DO levels under different dilution rates

were given in Figure 4.13. As be expected, the obviously increasing trends of

specific ATP generation rates profiles were observed with increasing D, due to µ was

equal to D in the continuous systems and the faster cells grow, and the more energy

will be consumed. The values of specific ATP generation rates maintained the

parallel levels at different DO levels with the same dilution rate. It indicated that ATP

energy consumption rates and generation rate were consistent with the cell

metabolism and growth at the same conditions.

5. The rhamnolipids, one of the most popular biosurfactants, were produced during the

stationary phase of cell growth even at high DO levels. The high concentrations were

shown at very low DOs (under microaerobic conditions), for example at 0.1 mg/L-

DO systems with D = 0.026 h-1 and 0.13 h-1 as well as at 0 mg/L-DO system with D =

0.06 h-1. At zero-DO systems with different air circulation rates (Figure 4.4B), the

rhamnolipid concentration in the system without air circulation (~ 0.9 g/L) was

significantly higher than the concentrations in the systems with air circulation (< 0.2

g/L). There were at least two possible reasons for the higher rhamnolipid production.

The first was the effect of glucose availability. In the high rhamnolipid-producing

system (DO ~ 0, no air circulation), there was remaining glucose that could support

the rhamnolipid synthesis, while glucose was practically exhausted in all of the low

producing systems. The second possible factor was the extent of aeration. Overall, in

both studies, rhamnolipid production appeared to favor the more anaerobic and

microaerobic conditions. The responsible mechanism(s) remains to be further

explored. The observation was significant not only for enhancing the productivity of

rhamnolipids in industrial production but also for addressing the role of rhamnolipid-

associated pathogenicity of P. aeruginosa, which resides in microaerobic to anoxic

biofilms in airway mucus of cystic fibrosis patients (120, 121).

6. The online NAD(P)H fluorescence techniques were used for detecting the shift of

electron-accepting mechanism. Representing a quantitative measure of the culture’s

“fractional approach” to the fully denitrifying state, the normalized fractions (NFU -

NFUOX)/(NFUDN - NFUOX) were correlated with the calculated fraction of electrons

accepted by denitrification (Figure 4.16 and Figure 4.17). Both the fluorescence

fraction and the e- fraction would have values between 0 and 1, with 0 corresponding

to fully aerobic metabolism and 1 to fully anaerobic, denitrifying metabolism. The

fractions of electrons accepted by denitrification, (2*NAR + 3*NIR)/(4*OUR +

2*NAR + 3*NIR), increased gradually with the fluorescence fractions at larger DO

levels, then the fluorescence fractions changed more rapidly than the e- fraction at

very lower DO levels, in other words, the increases were much sharper in lower DO

ranges at three different D levels. The unique shapes of correlations between the two

fractions were reproducible at three different dilution rates. The correlation facilitates

the applicability of online NAD(P)H fluorescence in monitoring and quantitatively

describing the sensitive microaerobic state of microorganisms.

7. The two fractions at D = 0.06 h-1obtained from zero-DO systems with different air

circulation rates and none-zero-DO systems were combined with DO and NFU

fractions in Figure 4.18. The profile with non-zero DO was a rather flat curve (less

sensitive changes in the e--accepting fraction than in the fluorescence fraction). The

most sensitive changes in the denitrification-accepted e- fraction occurred at the

fluorescence fractions of ~ 0.6 – 0.72 while the DO levels can not be detected

(Apparatus limitation). The significance of the unique outline of the correlation

between the two fractions remains to be further elucidated. Nevertheless, the results

demonstrated the applicability of the online NAD(P)H fluorescence in monitoring and

quantitatively describing the sensitive microaerobic state of microorganisms again.

5.2 Recommendations for Future Work

1. Investigate hydrocarbon metabolism and the second metabolites synthesis by P.

aeruginosa at microaerobic and aerobic conditions.

2. Evaluate the effects of different dilution rates and DO for cell metabolism and

respiratory mechanism of P. aeruginosa on hydrocarbon degradation.

3. Set up a model and discuss the nature competition and distribution of electrons

between O2 respiration and denitrification mechanism in the continuous culture

systems based on the glucose substrate.

BIBLIOGRAPHY

(1) Madigan, M. T.; Martinko, J. M.; Parker, J. Brock Biology of Microorganisms. Prentice-Hall, Upper Saddle River, NJ. 1997.

(2) Kincannon, D. F., E. L. Stover, V. Nichols, and D. Medley. Removal mechanisms for toxic priority pollutants. J. Wat. Pollut. Contr. Fed. 1983, 55, 157-163.

(3) Wilson, R. D., and D. M. Mackay. Diffusive oxygen emitters for enhancement of aerobic in situ treatment. Ground Wat. Monitor. Remed. 2002, 22, 88-98.

(4) Boulding, J. R., (Ed.) EPA Environmental Engineering Source Book. Ann Arbor Press, Chelsea, MI 1996.

(5) Dunn, M.; Wunderink, R. G. Ventilator-associated pneumonia caused by Pseudomonas infection. Clinics in Chest Medicine 1995, 16, 95-109.

(6) Crouch Brewer, S.; Wunderink, R. G.; Jones, C. B.; Leeper, K. V., Jr. Ventilator- associated pneumonia due to Pseudomonas aeruginosa. Chest 1996, 109, 1019-1029.

(7) Edmond, M. B.; Wallace, S. E.; McClish, D. K.; Pfaller, M. A.; Jones, R. N.; Wenzel, R. P. Nosocomial bloodstream infections in United States hospitals: a three-year analysis. Clinical Infectious Diseases 1999, 29, 239-244.

(8) Meluleni, G. J.; Grout, M.; Evans, D. J.; Pier, G. B. Mucoid Pseudomonas aeruginosa growing in a biofilm in vitro are killed by opsonic antibodies to the mucoid exopolysaccharide capsule but not by antibodies produced during chronic lung infection in cystic fibrosis patients. Journal of Immunology 1995, 155, 2029-2038.

(9) Ojeniyi, B.; Hoiby, N.; Rosdahl, V. T. Genome fingerprinting as a typing method used on polyagglutinable Pseudomonas aeruginosa isolates from cystic fibrosis patients. Apmis 1991, 99, 492-498.

(10) Anwar, H.; Strap, J. L.; Chen, K.; Costerton, J. W. Dynamic interactions of biofilms of mucoid Pseudomonas aeruginosa with tobramycin and piperacillin. Antimicrobial Agents and Chemotherapy 1992, 36, 1208-1214.

(11) Suci, P. A.; Mittelman, M. W.; Yu, F. P.; Geesey, G. G. Investigation of ciprofloxacin penetration into Pseudomonas aeruginosa biofilms. Antimicrobial Agents and Chemotherapy 1994, 38, 2125-2133.

(12) Henry, R. L.; Mellis, C. M.; Petrovic, L. Mucoid Pseudomonas aeruginosa is a marker of poor survival in cystic fibrosis. Pediatric Pulmonology 1992, 12, 158-161.

(13) Govan, J. R.; Deretic, V. Microbial pathogenesis in cystic fibrosis: mucoid Pseudomonas aeruginosa and Burkholderia cepacia. Microbiological Reviews 1996, 60, 539-574.

(14) Rumbaugh, K. P.; Griswold, J. A.; Iglewski, B. H.; Hamood, A. N. Contribution of quorum sensing to the virulence of Pseudomonas aeruginosa in burn wound infections. Infection and Immunity 1999, 67, 5854-5862.

(15) Greenberg, E. P. Bacterial genomics. Pump up the versatility. Nature 2000, 406, 947-948.

(16) Ishida, H.; Ishida, Y.; Kurosaka, Y.; Otani, T.; Sato, K.; Kobayashi, H. In vitro and in vivo activities of levofloxacin against biofilm-producing Pseudomonas aeruginosa. Antimicrobial Agents and Chemotherapy 1998, 42, 1641-1645.

(17) Xu, K. D.; Stewart, P. S.; Xia, F.; Huang, C. T.; McFeters, G. A. Spatial physiological heterogeneity in Pseudomonas aeruginosa biofilm is determined by oxygen availability. Applied and Environmental Microbiology 1998, 64, 4035-4039.

(18) Yoon, S. S.; Hennigan, R. F.; Hilliard, G. M.; Ochsner, U. A.; Parvatiyar, K.; Kamani, M. C.; Allen, H. L.; DeKievit, T. R.; Gardner, P. R.; Schwab, U.; Rowe, J. J.; Iglewski, B. H.; McDermott, T. R.; Mason, R. P.; Wozniak, D. J.; Hancock, R. E. W.; Parsek, M. R.; Noah, T. L.; Boucher, R. C.; Hassett, D. J. Pseudomonas aeruginosa anaerobic respiration in biofilms: relationships to cystic fibrosis pathogenesis. Developmental Cell 2002, 3, 593-603.

(19) Tiedje, J. M. x. Ecology of denitrification and dissimilatory nitrate reduction to ammonium. Biology of Anaerobic Microorganisms John Wiley & Sons, New York. 1988, 179-243.

(20) Yamane, T. Enzyme technology for the lipids industry: an engineering overview. Journal of American Oil Chemical Society 1987, 64, 1657-1662.

(21) Haferbury, D., Hommel, R., Kleber, H. P., Kluge, S., Schuster, G., Zschiegner, H. J. Antiphotoviral activity of rhamnolipids from Pseudomonas aeruginosa. Acta Biotechnology 1987, 7, 353-356.

(22) Cairns, W. L., Gray, N. C. C. In: Biosurfactants and Biotechnology, Kosaric N. (ed.). Marce Dekker Inc, New York. 1987.

(23) Gilbert P, Maira-Litran T, McBain AJ, Rickard AH, Whyte FW. 2002. The physiology and collective recalcitrance of microbial biofilm communities. Adv. Microb. Physiol. 46:202–56

(24) Kersulyte D, Struelens MJ, Deplano A, Berg DE. 1995. Comparison of arbitrarily primed PCR and macrorestriction (pulsed-field gel electrophoresis) typing of Pseudomonas aeruginosa strains from cystic fibrosis patients. J. Clin. Microbiol. 33:2216–19

(25) Mah TF, O’Toole GA. 2001. Mechanisms of biofilm resistance to antimicrobial agents. Trends Microbiol. 9:34–39

(26) Stewart PS. 2002. Mechanisms of antibiotic resistance in bacterial biofilms. Int. J. Med. Microbiol. 292:107–13

(27) Dalton HM, Goodman AE, Marshall KC. 1996. Diversity in surface colonization behavior in marine bacteria. J. Ind. Microbiol. 17:228–34

(28) Korber DR, Lawrence JR, Lappin-Scott HM, Costerton JW. 1995. Growth of microorganisms on surfaces. In Microbial Biofilms, ed. HM Lappin-Scott, JW Costerton, pp. 15–45. Cambridge, UK: Cambridge Univ. Press

(29) O’Toole GA, Kolter R. 1998. Flagellar and twitching motility are necessary for Pseudomonas aeruginosa biofilm development. Mol. Microbiol. 30:295–304

(30) Heydorn A, NielsenAT, Hentzer M, Sternberg C, Givskov M, et al. 2000. Quantification of biofilm structures by the novel computer program COMSTAT. Microbiology 146:2395–407

(31) Suh SJ, Silo-Suh L, Woods DE, Hasset DJ, West SE, Ohman EE. 1999. Effect of rpoS mutation on the stress response and expression of virulence factors in Pseudomonas aeruginosa. J. Bacteriol. 181:3890–97

(32) Marshall KC, Stout R, Mitchell R. 1971. Mechanisms of the initial events in the sorption of marine bacteria to surfaces. J. Gen. Microbiol. 68:337–48

(33) Davies DG, Geesey GG. 1995. Regulation of the alginate biosynthesis gene algC in Pseudomonas aeruginosa during biofilm development in continuous culture. Appl. Environ. Microbiol. 61:860–67

(34) Gilbert P, Maira-Litran T, McBain AJ, Rickard AH, Whyte FW. 2002. The physiology and collective recalcitrance of microbial biofilm communities. Adv. Microb. Physiol. 46:202–56

(35) J. Wilson, J. F. B. H. Optoelectronics, an introduction. Englewood Cliffs, N.J. : Prentice-Hall 1989, 113-155.

(36) Mason, W. T. Fluorescent and luminescent probes for biological activity : a practical guide to technology for quantitative real-time analysis. London ; San Diego : Academic Press 1993, 12-34, 360-378.

(37) Ichinose, N., Schwedt, G., Schnepel, F. M., and Adachi, K. Fluorometric analysis in biomedical chemistry: trends and techniques including HPLC applications. John Wiley & Sons, Inc. 1991.

(38) Ju, L.-K., and Sundararajan, A. Liquid-film time constant assessment for kLa measurements in respiring fermentation broths. Chem. Eng. Comm. 1995, 131, 161-170.

(39) Ju, L.-K. Monitoring of the biological nutrient removal process by an on-line NAD(P)H fluorometer. Biotechnol. Prog. 1995, 11, 541-551.

(40) Ju, L.-K., Trivedi, H.K. Monitoring of denitrification by Pseudomonas aeruginosa using ononline fluorescence technique. Biotechnol. Techn. 1992, 6, 549-554.

(41) Trivedi, H. K., Ju, L.-K. Study of nitrate metabolism of Escherichia coli using fluorescence. Biotechnol. Prog. 1994, 10, 421-427.

(42) Ju, L.-K., Yang, X., Lee, J.F., Armiger, W. B. Monitoring of the biological nutrient removal process by an on-line NAD(P)H fluorometer. Biotechnol. Prog. 1995, 11, 545- 551.

(43) Zabriskie, D. W., Humphrey, A.E. Estimation of fermentation biomass concentration by measuring culture fluorescence. Appl. Environ. Microbiol. 1978, 35, 337-343.

(44) Scheper, T., Gebauer, A., Schugerl, K. Monitoring of NADH-dependent culture fluorescence during the cultivation of Escherichia coli. Chem. Eng. J. 1987, 34, B7-B12.

(45) Winter, E. L., Rao, G., Cadman, T.W. Relationship between culture redox potential and culture fluorescence in Corynebacterium glutamicum. Biotechnol. Techn. 1988, 2, 233-236.

(46) Li, J.-K., Humphrey, A.E. Factors affecting culture fluorescence when monitoring bioreactors. J. Ferment. Bioeng. 1992, 74, 104-111.

(47) Groom, C. A., Luong, J.H.T., Mulchandani, A. On-line culture fluorescence measurement during the batch cultivation of poly-b-hydroxybutyrate producing Alcaligenes eutrophus. J. Biotechnol. 1988, 8, 271-278.

(48) Srinivas, S. P., Mutharasan, R. Inner filter effects and their interference in the interpretation of culture fluorescence. Biotechnol. Bioeng. 1987, 30, 769-774.

(49) Wang, N. S., Simmons, M.B. Characterization of an on-line commercial fluorescence probe: Modeling of the probe signal. Biotechnol. Bioeng. 1991, 38, 1292- 1301.

(50) Guilbault, G. G. Fluorescence: Theory, Instrumentation, and Practice. M. Dekker, New York. 1967.

(51) Surgi, M. R. In: Applied Biosensors, Wise, D.L. (ed.). Butterworth Publishers, Stoneham, MA. 1989.

(52) Wolfbeis, O. S., Weis, L.J., Leiner, M.J.P., Ziegler, W.E. Fiber-optic fluorosensor for oxygen and carbon dioxide. Anal. Chem. 1988., 60, 2028-2030.

(53) Zakharov, I. A., Aleskovskii, V.B. The sensitivity of the phosphorescent determination of oxygen in gases. Zh. Anal. Khim. 1965, 20, 700-703.

(54) Stangelmayer, A. (PreSens Precision Sensing GmbH) personal communication. 2001.

(55) Corp., Z. Product information. North Salt Lake, UT. 1993.

(56) Syldatk, C.; Wagner, F. Production of biosurfactants. Surfactant Science Series 1987, 25, 89-120.

(57) Hommel, R. K.; Ratledge, C. Biosynthetic mechanisms of low-molecular-weight surfactants and their precursor molecules. Surfactant Science Series 1993, 48, 3-63.

(58) Fukui, S.; Tanaka, A. Metabolism of alkanes by yeasts. Advances in Biochemical Engineering 1981, 19, 217-237.

(59) Osumi, M.; Fukuzumi, F.; Teranishi, Y.; Tanaka, A.; Fukui, S. Series of yeast microbodies. II. Development of microbodies in Candida tropicalis during incubation in a n-alkane medium. Archives of Microbiology 1975, 103, 1-11.

(60) Perry, J. J. Metabolism of cyclic alkanes. In: R. M. Altas (ed. ), Petroleum microbiology 1984, Macmillan, New York., 61-97.

(61) Kilburn, J. O.; Takayama, K. Effects of ethambutol on accumulation and secretion of trehalose mycolates and free mycolic acid in Mycobacterium smegmatis. Antimicrobial agents and chemotherapy 1981, 20, 401-404.

(62) Kretschmer, A.; Bock, H.; Wagner, F. Chemical and physical characterization of interfacial-active lipids from Rhodococcus erythropolis grown on n-alkanes. Applied and Environmental Microbiology 1982, 44, 864-870.

(63) Rapp, P.; Bock, H.; Wray, V.; Wagner, F. Formation, isolation, and characterization of trehalose dimycolates from Rhodococcus erythropolis grown on n-alkanes. Journal of General Microbiology 1979, 115, 491-503.

(64) Suzuki, T.; Tanaka, K.; Matsubara, I.; Kinoshita, S. Trehalose lipid and a-branched- b-hydroxy fatty acid formed by bacteria grown on n-alkanes. Agricultural and Biological Chemistry 1969, 33, 1619-1627.

(65) Burger, M. M.; Glaser, L.; Burton, R. M. The enzymic synthesis of a rhamnose- containing glycolipid by extracts of Pseudomonas aeruginosa. Journal of Biological Chemistry 1963, 238, 2595-2602.

(66) Hauser, G.; Karnovsky, M. L. Studies on the production of glycolipide by Pseudomonas aeruginosa. Journal of bacteriology 1954, 68, 645-654.

(67) Hisatsuka, K.; Nakahara, T.; Sano, N.; Yamada, K. Formation of rhamnolipid by Pseudomonas aeruginosa and its function in hydrocarbon fermentation. Agricultural and Biological Chemistry 1971, 35, 684-690.

(68) Wagner, F.; Behrendt, U.; Bock, H.; Kretschmer, A.; Lang, S.; Syldatk, C. Production and chemical characterization of surfactants from Rhodococcus erythropolis and Pseudomonas sp. MUB grown on hydrocarbons. Microb. Enhanced Oil Recovery 1983, 55-60.

(69) Itoh, S.; Honda, H.; Tomita, F.; Suzuki, T. Rhamnolipids produced by Pseudomonas aeruginosa grown on n-paraffin (mixture of C12, C13 and C14 fractions). Journal of Antibiotics 1971, 24, 855-859.

(70) Linhardt, R. J.; Bakhit, R.; Daniels, L.; Mayerl, F.; Pickenhagen, W. Microbially produced rhamnolipid as a source of rhamnose. Biotechnology and Bioengineering 1989, 33, 365-368.

(71) Cooper, D. G.; Paddock, D. A. Production of a biosurfactant from Torulopsis bombicola. Applied and Environmental Microbiology 1984, 47, 173-176.

(72) Goebbert, U.; Lang, S.; Wagner, F. Sophorose lipid formation by resting cells of Torulopsis bombicola. Biotechnology Letters 1984, 6, 225-230.

(73) Tulloch, A. P.; Spencer, J. F. T.; Gorin, P. A. J. Fermentation of long-chain compounds by Torulopsis magnoliae. I. Structures of the hydroxy fatty acids obtained by the fermentation of fatty acids and hydrocarbons. Canadian Journal of Chemistry 1962, 40, 1326-1338.

(74) Wagner, F., Kim, J. S., Lang, S., Li, Z. Y., Marwede, G., Matulovic, U., Ristau, E., Syldatk, C. Production of Surface active anionic glycolipids by resting and immobilized microbial cells. In: Third European Congress on Biotechnology, 1984, 1, 3-9.

(75) Arnulf, T., Steinbuchel, A. Formation of Polyesters Consisting of Medium-Chain- Length 3-Hydroxyalkanoic Acids from Gluconate by Pseudomonas aeruginosa and Other Fluorescent Pseudomonads. Applied and Environmental Microbiology 1990, 56, 3360- 3367.

(76) Syldatk, C., Seigmund, L, Wagner F. Chemical and Physical Characterization of Four Interfacial-Active Rhamnolipids from Pseudomonas spec. DSM 2874 Grown on n- alkane. Seitschrift fuer Naturforchung, C: Biosciences 1985, 40c, 51-60.

(77) Syldatk, C., Matulovic, U., Wagner, F. Biosurfactants - New methods for the production of interfacial-active anionic glycolipids by microorganisms. Biotech-Forum 1984, 1, 58-66.

(78) Zhang, Y., Miller, R. M. Enhanced Octadecane Dispersion and Biodegradation by a Pseudomonas Rhamnolipid surfactant (Biosurfactant). Applied and Environmental Microbiology 1992, 58, 3276-3282.

(79) Dyke, V. M. I., Couture, P., Brauer, M., Lee, H., Trevors, J. T. Pseudomonas aeruginosa UG2 rhamnolipid biosurfactants: structural characterization and their use in removing hydrophobic compounds from soil. Canadian Journal of Microbiology 1993, 39, 1071-1078.

(80) Ochsner, U. A., Reiser, J., Fiechter, A., Witholt, B. Production of Pseudomonas aeruginosa Rhamnolipid Biosurfactants in Heterologous Hosts. Applied and Environmental Microbiology 1995, 61, 3503-3506.

(81) Arino, S., Marchal, R., Vandecasteele, J. P. Identification and production of a rhamnolipidic biosurfactant by a Pseudomonas species. Applied Microbiology and Biotechnology 1996, 45, 162-168.

(82) Pimienta, A. L., Diaz, M. P. M., Carvajal, F. G. S., Grosso, V. J. L. Production of biosurfactants (rhamnolipids) by Pseudomonas aeruginosa isolated from colombian sludges. Ciencia, Tecnologia Futuro 1997, 1, 95-108.

(83) Mulligan, C. N., Gibbs, B. F. Correlation of Nitrogen Metabolism with Biosurfactant Production by Pseudomonas aeruginosa. Applied and Environmental Microbiology 1989, 55, 3016-3019.

(84) Osman, M., Ishigami, Y., Someya, J., Jensen H. B. The bioconversion of ethanol to biosurfactants and dye by a novel coproduction technique. JAOCS 1996, 73, 851-856.

(85) Syldatk, C., Lang, S., Matulovic, U., Wagner, F. Production of Four Interfacial Active Rhamnolipids from n-Alkanes or Glycerol by Resting Cells of Pseudomonas species DSM 2874. Zeitschrift Fur Naturforchung 1985, 40c, 61-67.

(86) Manresa, M. A., Bastida, J., Mercade, M. E., Robert, M., Andes, D. C., Espung, M. J., Guines, J. Kinetic studies on surfactant production by Pseudomonas aeruginosa 44T1. Journal of Industrial Microbiology 1991, 8, 133-136.

(87) Wu, J. Rhamnolipid production by fermentation and its application in enzymatic hydrolysis of cellulose. Ph.D. dissertation, 1997, The University of Akron, Akron, Ohio.

(88) Dalhoff, A., Rehmn, H. J. Studies on regulation of tetradecane oxidation in Pseudomonas aeruginosa II: The effect of glucose on tetradecane oxidation. European Journal of Applied Microbiology 1976, 3, 203-211.

(89) Lemoigne, M. Products of dehydration and of polymerization of -hydroxybutyric acid. Bulletin de la Societe Chimique Biologique 1926, 8, 770-782.

(90) Guerra-Santos, L. H., K?ppeli O., Fiechter, A. Dependence of P. aeruginosa continuous culture biosurfactant production on nutritional and environmental factors. Applied Microbiology and Biotechnology. 1986, 24, 443-448.

(91) Wood, T. M. Properties of cellulolytic enzyme systems. Biochemical Society Transactions 1985, 13, 407-410.

(92) Cabral, J. M. S., Casale, B., Cooney, C. L. Effect of antifoam agents and efficiency of cleaning procedures on the cross-flow filtration of microbial suspension. Biotechnology Letters 1985, 7, 747-752.

(93) Jarvis, F. G., Johnson, M. J. 1949. A glycolipide produced by Pseudomonas aeruginosa. Journal of the American Chemical Society 1949, 71, 4142-4126.

(94) Hauser, G., Karnovsky, M. L. Rhamnose and rhamnolipide biosynthesis by Pseudomonas aeruginosa. Journal of Biological Chemistry 1956., 224, 91-105.

(95) Hauser, G., Karnovsky, M. L. Origin of the carbon atoms of L-rhamnose. Federation Proceedings 1957., 16:, 193-194.

(96) Hauser, G., Karnovsky, M. L. Mode of biosynthesis of a glycolipid. . Federation Proceedings 1953, 12, 215-220.

(97) Burger, M. M., Glaser, L, Burton, R. M. Formation of Rhamnolipids of Pseudomonas aeruginosa. Methods in Enzymology 1966, 8, 441-445.

(98) Ochsner, U. A., Fiechter, A., Reiser, J. Isolation, Characterization, and Expression in Escherichia coli of the Pseudomonas aeruginosa rhlAB Genes Encoding a Rhamnosyltransferase Involved in Rhamnolipid Biosurfactant Synthesis. The Journal of Biological Chemistry 1994, 269, 19787-19795.

(99) Ochsner, U. A., Reiser, J. Autoinducer-mediated regulation of rhamnolipid biosurfactant synthesis in Pseudomonas aeruginosa. Proceedings of the National Academy of Sciences of the United States of America 1995, 92, 6424-6428.

(100) Wild, A. Soils and the Environment: An Introduction. The Press Syndicate of the University of Cambridge, New York 1993, 85-88.

(101) Alexander, M. Introduction to Soil Microbiology, 2nd edition. Wiley, New York 1977.

(102) Paul, R. M., Clark, F.E. Soil Microbiologyand Biochemistry. Academic Press, San Diego 1989.

(103) Wood, M. Soil Biology. Blakie, Glasgow. 1989.

(104) Chayabutra, C., Ju, L.-K. Degradation of n-hexadecane and its metabolites by Pseudomonas aeruginosa under microaerobic and anaerobic denitrifying conditions. Appl. Environ. Microbiol. 2000, 66, 493-498.

(105) Knowles, R. Denitrification. Micribial Rev. 1982, 46, 43-70.

(106) Clesceri, L. S., A. E. Greenberg, and R. R. Trussel. Standard Methods for the Examination of Water and Wastewater. 17th ed., Am. Public Health Assoc., Washington, D.C. 1989.

(107) Ashwell, G. Colorimetric analysis of sugars. In: Methods in Enzymology, Colowick, S. P. and Koplan, N. O. (eds.). Academic Press, New York. 1957, 3, 73-105.

(108) Stouthamer, A. H., J. Van't Riet, and L. F. Oltmann. Respiration with nitrate as acceptor. In Diversity of Bacterial Respiratory Systems. 1980, 2, Knowles, C. J., Ed., CRC Press, Boca Raton, FL.

(109) James, W. O. Cell Respiration. The English University Press Ltd 1971, Chapter 3, 50-65. 94

(110) Pichinoty, F.; d'Ornano, L. Inhibition by oxygen of the biosynthesis of nitrate reductase of denitrifying bacteria. Compt. rend. 1961, 252, 2294-2296.

(111) Kepes, A.; Lemoigne, M. The inhibiting action of oxygen on the utilization of nitrate nitrogen by an aerobic bacillus and the oxidation-reduction potential. Compt. rend. 1948, 226, 1556-1557.

(112) Ka, J. O., J. Urbance, R. W. Ye, T. Y. Ahn, and J. M. Tiedje. Diversity of oxygen and N-oxide regulation of nitrite reductases in denitrifying bacteria. FEMS Microbiol. Lett. 1997, 156, 55-60.

(113) Alexander, K.; Young, I. G. Alternative hydroxylases for the aerobic and anaerobic biosynthesis of ubiquinone in Escherichia coli. Biochemistry 1978, 17, 4750-4755.

(115) Chen, F.; Xia, Q.; Ju, L.-k. Aerobic denitrification of Pseudomonas aeruginosa monitored by online NAD(P)H fluorescence. Applied and Environmental Microbiology 2003, 69, 6715-6722.

(116) Louise, C. B., and J. F. Stuart. Nitric and nitrous oxide reductases are active under aerobic condition in cells of Thiophaera pantotropha. Biochem. J. 1991, 273, 423-427.

(117) Zhao, H. W., D. S. Mavinic, W. K. Oldham, and F. A. Koch. Controlling factors for simultaneous nitrification and denitrification in a two-stage intermittent aeration process treating domestic sewage. Wat. Res. 1999, 33, 961-970.

(118) Hassett, D. J., J. Cuppoletti, B. Trapnell, S. V. Lymar, J. J. Rowe, S. S. Yoon, G. M. Hilliard, K. Parvatiyar, M. C. Kamani, D. J. Wozniak, S.-H. Hwang , T. R. McDermott, and U. A. Ochsner Anaerobic metabolism and quorum sensing by Pseudomonas aeruginosa biofilms in chronically infected cystic fibrosis airways: Rethinking antibiotic treatment strategies and drug targets. Adv. Drug Delivery Rev. 2002, 54, 1425-1443.

(119) Scott, D. A., L. W. Grotyohann, J. Y. Cheung, and R. C. J. Scaduto Ratiometric methodology for NAD(P)H measurement in the perfused rat heart using surface fluorescence. Am. J. Physiol. 1994, 267, H636-644.

(120) Chen, F.; Xia, Q.; Ju, L.-k. Aerobic denitrification of Pseudomonas aeruginosa monitored by online NAD(P)H fluorescence. Applied and Environmental Microbiology 2003, 69, 6715-6722.

(121) Chen, F.; Xia, Q.; Ju, L.-K. Correlate Denitrification-Accepted Fraction of Electrons with NADH Fluorescence of Pseudomonas aeruginosa Performing Simultaneous Denitrification and Respiration at Extremely Low Dissolved Oxygen Conditions. Biotechnology Progress 2004. (in press)