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					AN INVESTIGATION OF FACTORS LIMITING THE BIOSPARGEMEDIATED IN-SITU BIOREMEDIATION OF HYDROCARBON CONTAMINATED GROUNDWATER

A Master’s Thesis Presented to the Faculty California Polytechnic State University San Luis Obispo

In partial fulfillment of the requirements for the degree of Master of Science in Civil and Environmental Engineering

By Jason G. Waudby August 2003

COPYRIGHT OF MASTER’S THESIS

I reserve the reproduction rights of this thesis for a period of three years from the date of submission. I waive all reproduction rights after that time span has expired.

Jason G. Waudby

Date

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MASTER’S THESIS APPROVAL

TITLE:

AN INVESTIGATION OF FACTORS LIMITING THE BIOSPARGE-MEDIATED IN-SITU BIOREMEDIATION OF HYDROCARBON CONTAMINATED GROUNDWATER JASON G. WAUDBY AUGUST 2003

AUTHOR: DATE SUBMITTED:

THESIS COMMITTEE MEMBERS:

Dr. Yarrow Nelson

Date

Dr. Sam Vigil

Date

Dr. Thomas Ruehr

Date

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ABSTRACT

AN INVESTIGATION OF FACTORS LIMITING THE BIOSPARGEMEDIATED IN-SITU BIOREMEDIATION OF HYDROCARBON CONTAMINATED GROUNDWATER

JASON G. WAUDBY

A horizontal biosparge system was installed as part of the Guadalupe Restoration Project (GRP), near Guadalupe, California to stimulate the bioremediation of petroleum hydrocarbon contaminated groundwater. Minimal reductions of the total petroleum hydrocarbon (TPH) concentrations in the groundwater plume have been observed over the three years of biosparge operation. An assessment of possible biological/chemical factors limiting the biodegradation of contaminants at the biosparge site was performed. A series of experiments were conducted to determine if biodegradation is being limited by: a lack of hydrocarbon degrading microorganisms, inorganic nutrient (N, P, K, S) concentrations, insufficient dissolved oxygen supply, or the biodegradability of the petroleum constituents in the groundwater. These issues were addressed using laboratory experiments with diluent-contaminated groundwater from the TB8 site and measuring biodegradation using a combination of respirometry and TPH analyses. Respirometry experiments showed considerable carbon dioxide (CO2) production and concomitant TPH loss in nonamended groundwater samples from the biosparge site, indicating hydrocarbon degrading microorganisms are present in the groundwater and biodegradation was occurring. No increase in total CO2 production was observed in samples with added nutrients, inoculum, or nutrients and inoculum, over the 28-day experiment. TPH biodegradation was observed in all sample sets. Nutrient addition (N, P, K, and S) increased TPH degradation rates only slightly for both inoculated and noninoculated groundwater, but this increase was not statistically significant. Nutrient and inoculum addition were not necessary for contaminant biodegradation, at least in the short term. Consistent stirring of experimental samples also increased CO2 evolution. Decreasing dissolved oxygen (DO) concentration decreased CO2 production, but considerable CO2 production was observed even in samples with DO concentrations as low as 0.5 mg/L. Zero order rate constants of 19.8 µg/L-day (R2 = 0.87) and 24.0 µg/Lday (R2 = 0.88) were observed for the overall non-amended biodegradation of hydrocarbons in long-term experiments (124 and 103 days, respectively) to determine the ultimate biodegradability of the TPH constituents in the groundwater. In this long-term experiment, TPH degradation rates decreased significantly after initial observed biodegradation. Inorganic nutrient analyses for N and P were non-detectable after approximately 117 days of incubation. However, no increase in TPH biodegradation was observed (14 days after nutrient addition) during a supplementary investigation of nutrient limitations at the end of the long-term experiments. Thus, while short-term

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biodegradation rates were adequate and not limited by nutrients and microorganisms in the site groundwater, the problem of long-term biodegradability should be addressed in future research.

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ACKNOWLEDGEMENTS

I want to thank my family and friends for their patience, support and encouragement through this and all of my academic adventures.

I want to thank Dr. Yarrow Nelson for his guidance, patience, and approachable demeanor; Miss Wendy Martin for her continuous help and motivational support; Mr. Bunkim Chokshi for sharing his respirometer expertise; and Mr. Doug Allen for his assistance with the experimental logistics.

I want to give a special thanks to Unocal for their support and funding of our research. Their funding has enabled me to conduct this interesting experiment.

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TABLE OF CONTENTS

List of Tables .......................................................................................................................x List of Figures ................................................................................................................... xii 1 INTRODUCTION .........................................................................................................1

2 BACKGROUND ...........................................................................................................3 2.1 Historical Background of the Guadalupe Restoration Project Site ........................3 2.2 Historical Background of the TB8 Biosparge System at the GRP .........................4 2.3 Characterization of Diluent Contamination at the TB8 Biosparge Site .................5 2.4 Biosparge as an In Situ Bioremediation Technology .............................................5 2.4.1 Factors Limiting the Effectiveness Biosparging .............................................6 2.5 Aerobic Hydrocarbon Biodegradation....................................................................7 2.5.1 Nutrient Requirements for Effective Biodegradation of Hydrocarbons..........9 2.5.2 Addition of Inoculum to Enhance the Biodegradation of Hydrocarbons......11 2.6 Quantification of CO2 Production as a Measurement of Biodegradation.............12 3 MATERIALS AND METHODS.................................................................................13 3.1 Respirometry Experiments ...................................................................................13 3.1.1 Respirometer Methods...................................................................................13 3.1.2 Experimental Chamber Bottles......................................................................15 3.1.3 Magnetic Stirrers ...........................................................................................15 3.1.4 Temperature Controlled Water Bath .............................................................16 3.2 Respirometer Experiment for Nutrients and Inoculum ........................................18 3.2.1 Sample Groundwater Collection ...................................................................22

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3.2.2 Nutrient Preparation and Addition ................................................................22 3.2.3 Initial TPH and Nutrient Sampling and Analyses .........................................23 3.2.4 Final TPH Sampling Methods .......................................................................25 3.3 Respirometer Experiments for DO Effect ............................................................25 3.3.1 Sample Groundwater Collection ...................................................................26 3.3.2 Respirometer Experiment 2 (EXP2)..............................................................27 3.3.3 Respirometer Experiments 3 and 4 (EXP3 and EXP4) .................................27 3.4 Long-term Biodegradability Experiment..............................................................29 3.4.1 Dissolved Oxygen Delivery System..............................................................31 3.4.2 TPH Sampling and Analysis .........................................................................33 3.4.3 Investigation of the Effects of Stirring and Nutrient Addition on the Long-term TPH Biodegradation in TB8 Groundwater .................................35 4 RESULTS AND DISCUSSION ..................................................................................37 4.1 Nutrients and Inoculation Effects on Short-term TPH Respiration (EXP1).........37 4.1.1 Characterization of Groundwater Samples....................................................37 4.1.2 Respirometry Results for EXP1 ....................................................................40 4.1.3 TPH Results for EXP1...................................................................................45 4.1.4 Quantitative Relationship Between TPH Degraded and CO2 Evolved .........48 4.1.5 Nutrient and Inoculum Effects on Carbon Chain Distribution......................50 4.2 Effect of DO Concentration on Respiration Rates of TPH-Contaminated Groundwater .........................................................................................................58 4.3 Implications of Stirring on Respiration ................................................................61 4.4 Long-term Biodegradability Experiment..............................................................63

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4.4.1 Evaporation Effects on TPH Concentrations During Long-term Biodegradability Experiment.........................................................................67 4.4.2 Results for the Investigation of the Effects of Stirring and Nutrient Addition on the Long-term TPH Biodegradation in TB8 Groundwater .......72 4.4.3 Kinetics of the Long-term Biodegradation of TPH in TB8 Groundwater.....75 5 CONCLUSIONS..........................................................................................................79 6 RECOMMENDATIONS.............................................................................................82 REFERENCES ..................................................................................................................84

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LIST OF TABLES

Table 3.1

Description of Respirometry Bottle Contents for Experiment 1 (EXP1)......................................................................................................19 Sample Descriptions and Initial TPH Concentrations for EXP1 .............38 TPH Carbon Chain Distribution of Initial Groundwater Samples ...........38 Inorganic Nutrient Analysis of TB8 Groundwater...................................40 Total CO2 Production (EXP1 + EXP1A + EXP1B).................................41 Final TPH Sample Descriptions and Results ...........................................45 Nutrients and Inoculum Effects on Respiration and TPH Biodegradation in TB8 Groundwater Samples ........................................48 Data Showing the Relationship Between CO2 Production and TPH Degradation ..............................................................................................49 Final TPH Carbon Chain Distribution in EXP1 (After 28 Days).............51 Oxygen Percentages in Respirometer Refresh Air, Average DO Concentrations, and Corresponding 48-Hour Average CO2 Production Values ....................................................................................59 Measured DO Concentrations and Percent of DO Saturation for the Long-term Experimental Sample Bottles (1), (2), (2A) ...........................63 TPH Carbon Chain Distribution and Total TPH for Bottles (1), (2), and (2A) from the Long-term Biodegradability study. Values were not adjusted for water loss due to evaporation .........................................65 Evaporation Measurements Over Two 14-Day Periods and the Resulting Calculated Evaporation Rates ..................................................67 TPH Carbon Chain Distribution and Total TPH Concentrations for Bottles (1), (2,) and (2A) of the Long- term Biodegradability Experiment Adjusted for Water Loses Due to Evaporation.....................70

Table 4.1 Table 4.2 Table 4.3 Table 4.4 Table 4.5 Table 4.6

Table 4.7

Table 4.8 Table 4.9

Table 4.10

Table 4.11

Table 4.12

Table 4.13

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Table 4.14

TPH Carbon Chain Distribution and Total TPH Concentrations for Bottles 1 and 2A Comparing the Biodegradation in TB8 Groundwater with and without Added Nutrients (1) and with and without Stirring (2A). All Values are Adjusted for Water Loses Due to Evaporation......73

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LIST OF FIGURES

Figure 2.1 Figure 3.1

Location of the Guadalupe Restoration Project .....................................3 Experimental respirometer set-up used to measure CO2 evolution in sample bottles containing TB8 groundwater ...................................14 VWR 200 Mini Stirrer used in the respirometry experiments.............16 Temperature controlled water bath apparatus showing .......................17 An 85.7-L Light Blocking Tedlar® bag with On/Off valve used to dispense refresh gas to the respirometer during experiments to investigate DO effect on TPH biodegradation.....................................26 Diagram showing the system used to deliver air/N2 mixtures of various O2 concentrations to the Micro Oxymax Refresh Air Input to control DO concentrations in respirometry samples........................28 Experimental incubator set-up for the long-term biodegradability experiment............................................................................................31 Diagram of the DO delivery system used to stimulate the biodegradation of TPH contamination in TB8 groundwater sample bottles during the long-term biodegradability experiment...................32 A digital image of the DO delivery system used to aerate TB8 groundwater samples during the long-term biodegradability experiment............................................................................................33 Long-term experimental Bottle (2A) with an active magnetic stir bar and bubbling air stones. The lines on the bottle represent 1 L increments (TPH sample volume) and served as a basis for evaporation quantification ...................................................................36 Initial TPH carbon chain distributions for Nonamended and Inoculated samples VOC Collection and Water Supply Systems .......39 Total CO2 production (EXP1+EXP1A+EXP1B) versus time for TB8 groundwater with and without nutrient, inoculum, and nutrient/inoculum amendments ...................................42

Figure 3.2 Figure 3.3 Figure 3.4

Figure 3.5

Figure 3.6

Figure 3.7

Figure 3.8

Figure 3.9

Figure 4.1

Figure 4.2

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Figure 4.3

The average total CO2 production for nonamended groundwater, and groundwater with added nutrients, inoculum and nutrient/inoculum.................................................................................44 A comparison of initial and final average total TPH concentrations in Non-amended TB8 groundwater and TB8 groundwater with added Nutrients, Inoculum, or Nutrients/Inoculum .............................46 Effect of nutrient amendment on biodegradation of TPH fractions in inoculated TB8 groundwater. Actual TPH concentrations of each fraction remaining are presented for duplicate samples..........53 Effect of nutrient addition on biodegradation of TPH fractions in noninoculated TB8 groundwater .........................................................54 Effect of inoculation on the biodegradation of TPH fractions in groundwater samples with no nutrients added. A comparison of the percent of initial TPH remaining in nonamended TB8 groundwater samples and TB8 groundwater with added inoculum.....56 Effect of inoculum addition on biodegradation of TPH fractions for groundwater with nutrients added. A comparison of the percent of initial TPH remaining in TB8 groundwater samples with added Nutrients and TB8 groundwater samples with added Nutrients and Inoculum........................................................................57 Effect of DO concentration on CO2 production rates during TPH biodegradation for TB8 groundwater. Rates are averaged over the 48-hour runs. Two-dimensional error bars indicate standard deviation of both the CO2 production rates and the measured DO concentrations ......................................................................................60 Cumulative CO2 production versus time with a reduction of CO2 produced by the Series 3 triplicate sample after the magnetic stirrer failed at approximately 24-hours .........................................................61 Cumulative CO2 production vs. time showing a reduction of CO2 production in duplicate samples containing inactive magnetic stirrers 24-hours ...................................................................................62 TPH concentrations vs. time during the long-term biodegradation of TB8 groundwater without nutrient or inoculum addition. Concentrations were not adjusted to account for the evaporation of water from the sample bottles..........................................................66

Figure 4.4

Figure 4.5

Figure 4.6

Figure 4.7

Figure 4.8

Figure 4.9

Figure 4.10

Figure 4.11

Figure 4.12

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Figure 4.13

TPH concentrations vs. time adjusted for water loss due to evaporation during the long-term biodegradation of TB8 groundwater without nutrient or inoculum addition ...........................71 TPH concentrations versus time for all TPH sample events (including those for the investigation of the effects of stirring and nutrient addition on TPH biodegradation. All values are adjusted for evaporation .....................................................................................74 The Natural Log of TPH Concentration vs. Time for experimental bottles (1) and (2) over the first 6.6 days of the Long-term Biodegradability experiment. Linear trendlines for bottles (1) and (2A) showing first order rate constants (day-1) and respective R2 values. ..................................................................................................77 TPH vs. Time for all nonamended (no nutrients or stirring) TB8 groundwater TPH sampling events during the Long-term Biodegradability experiment. Linear trendlines for bottles (1) and (2A) showing zero order rate constants (µg/L-d) and respective R2 values. ..................................................................................................78

Figure 4.14

Figure 4.15

Figure 4.16

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CHAPTER 1 INTRODUCTION

Biosparging is an in-situ remediation technology used to biodegrade organic constituents dissolved in groundwater and adsorbed to soil within the saturated zone and capillary fringe at contaminated sites. In biosparging, aerobic biodegradation of contaminants by indigenous microorganisms is stimulated by the injection of air (or oxygen) into the saturated zone. Biosparging is predominantly used for the remediation of mid-weight petroleum hydrocarbon (e.g., diesel fuel) contamination in the subsurface (USEPA 1994).

As part of the Guadalupe Restoration Project (GRP), near Guadalupe California, a pilot scale horizontal biosparge system was installed to test the effectiveness of biospargeenhanced biodegradation of petroleum hydrocarbons. This site is heavily contaminated by a C10 to C30 hydrocarbon mixture previously used at the oil field as a diluent for the viscous crude oil being extracted. The biosparge operation at the Tank Battery 8 (TB8) site was the special focus of this thesis. After three years of biosparge operation, the concentrations of total petroleum hydrocarbons (TPH) in the groundwater have remained relatively unaffected by the biosparging. Possible physical limitations of the biosparge system at TB8 were investigated in a companion study (Coffey, 2003). Biological factors were addressed in this thesis using a series of laboratory experiments with groundwater from the TB8 site.

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The laboratory studies were performed to address the following questions: 1. 2. 3. 4. Are hydrocarbon-degrading organisms present in TB8 groundwater? Is TPH degradation in TB8 groundwater nutrient limited? How does CO2 production correlate with TPH degradation? Are dissolved oxygen (DO) concentrations present in TB8 groundwater sufficient for TPH biodegradation? 5. 6. Does TPH biodegradation stop at a threshold TPH concentration? What are the kinetics of TPH biodegradation under experimental conditions?

These questions were addressed by performing a series of laboratory experiments using respirometry and TPH analyses of groundwater from the TB8 biosparge site. In

experiment 1 (EXP1), short-term respirometry was used to examine nutrient and inoculum requirements for biodegradation at the TB8 site by measuring CO2 production and TPH degradation. In respirometry experiments 2 – 4 the production of CO2 was measured and compared for samples containing varied concentrations of DO. A longterm laboratory biosparge experiment was also conducted to investigate the kinetics and long-term biodegradability of the petroleum hydrocarbons present in the TB8 groundwater.

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CHAPTER 2 BACKGROUND

2.1 Historical Background of the Guadalupe Restoration Project Site The GRP is located on California’s Central Coast, north west of the city of Guadalupe and along the southern edge of San Luis Obispo County (Figure 2.1). The site was

previously the home of the Guadalupe Oil Field (GOF), operated by Unocal Oil Company for nearly 50 years.

Figure 2.1: Location of the Guadalupe Restoration Project.

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To facilitate the extraction and transportation of the highly viscous crude oil at the GOF, a diesel/kerosene hydrocarbon mixture, referred to as "diluent", was used as a thinning agent. A system of storage tanks and pipelines was used to distribute the diluent

throughout the oil field. Over the years, this extensive storage and transport system aged, and considerable quantities of diluent were released into the surrounding environment. In 1990, the discovery of an oil sheen on the beach prompted Unocal to reduce oil production at the GOF and permanently discontinue the use of diluent. By 1994, Unocal had become aware of the extensive contamination, and all production at the site was discontinued. Shortly thereafter, the remediation efforts began and the first steps were taken towards what is now referred to as the GRP.

2.2 Historical Background of the TB8 Biosparge System at the GRP In April 1999, a horizontal biosparge system was installed at the TB8 site to stimulate the biodegradation of diluent contaminated groundwater. The TB8 site was chosen due to the accessibility of the dissolved phase diluent plume within the relatively shallow groundwater aquifer (8 – 20 feet below ground surface) (Coffey, 2003). Design

parameters for the TB8 biosparge system were based upon results from previous pilot tests conducted within the GRP. The biosparge system delivers 500 to 600 scfm of air to the saturated zone via three horizontal sparge wells. Measurements of groundwater DO in monitoring wells down gradient from the air injection wells range from 0.2 to 7.4 mg/L (Coffey, 2003).

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2.3 Characterization of Diluent Contamination at the TB8 Biosparge Site The diluent contaminant present at the TB8 site has a hydrocarbon range from C6 to > C30, with the majority (70 %) of the hydrocarbons falling within the C10 – C25 (diesel) range. Analysis of TPH data from the TB8 site suggests long-term weathering has substantially degraded the dissolved phase diluent compounds (LFR, 1996). Recent TPH concentrations measured in groundwater up gradient of the TB8 biodsparge site ranged from approximately 2 – 13 mg/L (C10 – C40) (Coffey, 2003).

2.4 Biosparge as an In Situ Bioremediation Technology Biosparging is an in-situ remediation technology used to stimulate the aerobic biodegradation of organic contaminants by indigenous microorganisms. In aerobic

biodegradation, oxygen is the terminal electron acceptor during the metabolism of organic compounds (Suthersan, 1997). Biosparging provides oxygen to aerobic

microorganisms (and facultative aerobes) in the subsurface by the injection of air (or oxygen) into the saturated zone. The subsurface introduction of air increases the

concentration of dissolved oxygen in groundwater, and promotes the growth of aerobic microorganisms capable of degrading hydrocarbons dissolved in the groundwater and/or adsorbed to soil within the saturated zone and capillary fringe. Biosparging is most often used at sites contaminated by mid-weight petroleum hydrocarbons (e.g., diesel fuel). Lighter hydrocarbons volatilize too rapidly to allow for effective biodegradation, and heavier hydrocarbons generally take longer to biodegrade (USEPA, 1994).

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2.4.1 Factors Limiting the Effectiveness Biosparging Two primary factors may limit the effectiveness of biosparing: 1. the permeability of the soil at the contaminated site and 2. the biodegradability of the contaminants (USEPA, 1994).

The rate of oxygen transfer to the groundwater depends upon the soil permeability. Finegrained soils restrict the flow of air through the smaller pore spaces, decreasing the efficiency of oxygen dispersion. Biosparging is likely to be effective in soils with an intrinsic permeability greater than 10-9 cm2 (USEPA, 1994). During biosparging, soil permeability may be reduced in aquifers containing dissolved ferrous iron (Fe+2) at concentrations greater than 10 mg/L. The dissolved iron is oxidized to Fe+3 in the presence of oxygen, which can precipitate out of solution as iron oxide (rust) and restrict oxygen and groundwater flow by clogging soil pores (USEPA, 1994).

Most petroleum hydrocarbons can be biodegraded (USEPA, 1994).

However, the

chemical structure of the petroleum hydrocarbons to be treated determines the rate of biodegradation. Aliphatic and monoaromatic (nine carbon atoms or less) hydrocarbons are more readily degraded than the larger aliphatic and polyaromatic hydrocarbons (PAHs) (USEPA, 1994). The weathering or “aging” of hydrocarbons can reduce their availability to microorganisms and extend their life in the environment. The presence of C17 and C20 compounds in diesel contaminated soil after 600 days of aerobic incubation was attributed to the “sequestering” of the compounds due to possible aging of the hydrocarbon (Siddiqui and Adams, 2001).

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2.5 Aerobic Hydrocarbon Biodegradation Microorganisms are capable of biodegrading petroleum hydrocarbons under aerobic and anaerobic conditions (Suthersan, 1997). The rate of the biodegradation of organic

constituents is considered more rapid under aerobic conditions (Wilson et al., 2002; Landmeyer et al., 2001; Yerushalmi et al, 2001; Vasudevan and Rajaram, 2001; Dragun, 1998). Therefore, aerobic (or facultative) microbes are considered to be most important (for the bioremediation of petroleum products) (USEPA, 1995). The general process of aerobic biodegradation can be described by the following equation:

Bacteria + Organics + O2 + Nutrients

CO2 + H2O + Biomass + Byproducts

(2.1)

Aerobic biodegradation of hydrocarbons, such as aliphatics, aromatics and PAHs, involves the incorporation of oxygen by microbial cells to produce oxygenase enzymes (monooxygenases and dioxygenases) to break the hydrocarbon bonds (Suthersan, 1997). Therefore, consistent aeration is beneficial in stimulating the complete biodegradation of petroleum products.

Stoichiometric analysis can be used to estimate the theoretical amount of oxygen required aerobically to degrade a given quantity of hydrocarbon, as described in Equation 2.2. Generally, the degradation of 1 mg of a medium length hydrocarbon requires 3 to 4 mg of oxygen (Dragun, 1998).

CxHy +[x + (y/4)] O2

x CO2 + (y/2) H2O

(2.2)

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For example: If dodecane (C12H26) is used to represent TPH present at the TB8 site, the aerobic biodegradation can be written as:

C12H26 + 18.5 O2

12 CO2 + 13 H2O

(2.3)

Molecular Weight Dodecane = 170 mg/mmol Molecular Weight O2 = 32 mg/mmol

Mass of Dodecane Consumed: 170 mg/mmol x 1 mmol = 170 mg Dodecane Mass of Oxygen Consumed: 32 mg/mmol x 18.5 mmol = 592 mg O2

Mass of Carbon Dioxide Produced: 44mg/mmol x 12 mmol = 528 mg CO2

Thus, approximately 3.48 mg of oxygen is required aerobically to biodegrade 1 mg of ‘TPH’ (dodecane) to CO2 and H2O. When the same calculations are performed for the biodegradation of for octadecane (C18H38) as the representative TPH compound, the ratio of oxygen to hydrocarbon consumed is 3.46 mg O2 to 1 mg ‘TPH’. Equations 2.2 and 2.3 account for respiration only, and do not take into consideration the quantity of reactants consumed in microbial cell synthesis.

Aerobic biodegradation has been effective in reducing concentrations of various types of hydrocarbons including those in gasoline, diesel fuel and crude oil. The gasoline

oxygenate methyl tert-butyl ether (MTBE), although once considered recalcitrant in the

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natural environment, has been shown to biodegrade under oxic conditions (Landmeyer et al., 2001; Wilson et al., 2002). Relatively small concentrations of DO can significantly stimulate hydrocarbon biodegradation. For instance, a study of in-situ biodegradation of MTBE in groundwater reported an 83 % reduction of MTBE concentrations when groundwater DO levels were increased from < 0.2 to 2 mg/L (Landmeyer et al., 2001). Under aerobic conditions, benzene, a highly toxic, highly water-soluble monoaromatic hydrocarbon, degrades quite easily (Suthersan, 2002). However, significant

mineralization of benzene occurred under microaerophilic (DO < 2 mg/L) conditions, including the recovery of 50.8 ± 1.4 % of
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C labeled CO2 (from benzene

biodegradation), with the injection of DO at an initial concentration of 1.0 mg/L (Yerushalmi et al., 2001).

2.5.1 Nutrient Requirements for Effective Biodegradation of Hydrocarbons Inorganic nutrients [nitrogen (N), phosphorus (P) and sulfur (S)] are needed to support microbial cell growth and for the production of metabolic enzymes required for biodegradation (Suthersan, 1997; USEPA, 1995). If these nutrients are not available in sufficient quantities in the contaminated aquifer, then nutrients may need to be added to enhance the biodegradation processes. In the absence inorganic nutrients, biodegradation continues in the subsurface due to the natural recycling of elements. However, nutrient limited biodegradation proceeds at a slower rate (Suthersan, 1997).

The most critical inorganic nutrients to the bioremediation of hydrocarbons are nitrogen and phosphorus (Onwurah, 1999). Both N and P are critical in cell growth and account

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for 14 and 3 % (N and P respectively) of the dry weight of a typical microbial cell (Suthersan, 1997; Liebeg and Cutright, 1999). N and P are also the nutrients most often in limited supply in contaminated soil and groundwater. The recommended carbon: nitrogen: phosphorus mole ratios for enhanced biodegradation are in the range of 100: 10: 1 to 100: 1: 0.5 (USEPA, 1995; Liebeg and Cutright, 1999).

Nitrogen and phosphorus are the two inorganic nutrients most commonly added in the bioremediation of hydrocarbon-contaminated sites. Increased biodegradation of

hydrocarbons, following nutrient biostimulation has been well documented. The addition of commercial fertilizer (20: 20: 20: N, P, K, respectively) has substantially enhanced the mineralization of hexadecane and naphthalene (Whyte et al., 2001). The presence of N and P in the experimental media was essential for hydrocarbon degradation in experiments on the biodegradation of diesel and heating oil (Marin et al., 1995). In a highly nutrient-limited marine environment, oil biodegradation rates were increased 3 – 5 times with the introduction of a garden fertilizer formulation containing N: P of 23: 2 (Atlas, 1995). The addition of nutrients to diesel contaminated soils increased bacterial populations to 10 times the population of soils without N and P addition, while increasing respiration (Siddiqui and Adams, 2001).

Although N is generally considered the most limiting inorganic nutrient at hydrocarbon contaminated sites, the nutrient present in the smallest quantity will be the limiting factor of microbial growth (Suthersan, 1997). A study of enhanced bioremediation of PAH contaminated soil reported larger increases in oxygen consumption in contaminated soil

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biostimulated by nutrient solutions containing P as the dominant macronutrient (Liebeg and Cutright, 1999). Phosphorus limitations in the subsurface may develop due to its adsorption to soil because oil binding reduces the transport of P, rendering it unavailable for microbial metabolism.

2.5.2 Addition of Inoculum to Enhance the Biodegradation of Hydrocarbons The addition of inoculum to contaminated soil and water to enhance the biodegradation of hydrocarbons “bioaugmentation” has been debated. The inoculation of a contaminated medium with microorganisms capable of biodegrading petroleum contaminants is generally considered unnecessary (Norris et al., 1994; Suthersan, 1997). The presence of hydrocarbons as a food source naturally selects for those indigenous microbes capable of metabolizing the contaminants (Rahman et al., 2002). Microorganisms indigenous to soils with no history of hydrocarbon contamination have adapted to, and grown on diesel hydrocarbons within 6 days of hydrocarbon addition (Siddiqui and Adams, 2001). One situation where inoculation may prove beneficial is when a medium is contaminated by less biodegradable organic compounds such as PAHs (Norris et al., 1994; Dragun, 1998). The application of a bacterial consortium grown on oily sludge with the addition of a nutrient mixture substantially reduced concentrations of alkanes, aromatics and asphaltene fractions of TPH in in-situ test plots of soil contaminated with oily sludge (Mishra et al., 2000). The enrichment of dominant indigenous hydrocarbon degraders at a site by cultivating the microbes in the laboratory and reintroducing the enriched cultures to contaminated soil increased the aerobic biodegradation of long chain alkanes and naphthalene in laboratory experiments (Li et al., 2000).

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2.6 Quantification of CO2 Production as a Measurement of Biodegradation Carbon dioxide is a by-product of respiration during the biodegradation of organic compounds. The quantification of CO2 production has been used to measure the

biodegradation of petroleum hydrocarbons in numerous laboratory studies (Miles and Doucette, 2001; Whyte et al., 2000; Siddiqui amd Adams, 2001; Namkoong et al., 2001; Hollender et al., 2003). A strong correlation (P < 0.01) existed between CO2 production rates and the biodegradation of diesel fuel (Namkoong et al., 2001). Respiration

experiments measuring the evolution of CO2 have been used to identify nutrient limitations in soil samples contaminated with BTEX and PAHs (Hollender et al., 2003). The measurement of CO2 evolution is generally considered a reliable technique to evaluate biodegradation in laboratory experiments (Miles and Doucette, 2001). However, CO2 evolution may not be completely reliable for measurement of biodegradation in soils containing unknown sinks and sources of CO2 (biomass and soil organic matter) (Sharabi and Bartha, 1993).

In experiments employing the analysis of hydrocarbons degraded, the theoretical amount of CO2 to be evolved from the hydrocarbon biodegradation can be estimated using Equations 2.2 and 2.3 of section 2.5. If dodecane is used to represent the TPH in the system, Equation 2.3 holds true and 12 moles of CO2 are expected to be produced from the biodegradation of 1 mole of ‘TPH’. This ratio converts to 3.11 mg CO2 produced from the biodegradation of 1 mg of ‘TPH’.

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CHAPTER 3 MATERIALS AND METHODS

Groundwater samples used in the experiments presented here were collected from groundwater monitoring wells directly up gradient from the TB8 biosparge system. The groundwater samples were used in experiments designed to simulate conditions at the TB8 site, while allowing for the incorporation of experimental treatment conditions 3.1. Respirometry Experiments 3.1.1. Respirometer Methods The respirometry experiments were conducted using a Micro Oxymax respirometer manufactured by Columbus Instruments, Inc. (Columbus, Ohio). The Micro Oxymax system is a closed-circuit respirometer used to detect low levels of carbon dioxide evolution. The system includes the following: sample pump, single beam, non-dispersive infrared CO2/CH4 Sensor with a CO2 sensor range of 0.0 % to 1.0 %, expansion interface, condensing air drier, in-line sample drier column containing anhydrous magnesium perchlorate reagent [Mg(ClO4)2] and Mirco Oxymax Version 6.09b computer software. A Drierite®, gas drying unit (Hammond Laboratory) containing anhydrous CaSO4 was added to ensure the air used to refresh the head space in the sample bottles was moisture free. A “Columbus Micro Systems” IBM compatible computer was used as the dedicated controller of the Micro Oxymax System, and data recorder. The respirometer system was capable of collecting CO2 production data for up to ten samples for each experiment. A picture of the complete experimental set-up used for the respirometry experiments in this study is shown in Figure 3.1.

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Figure 3.1: Experimental respirometer set-up used to measure CO2 evolution in sample bottles containing TB8 groundwater

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The sample pump for the Micro Oxymax respirometer system was sent to Columbus Instruments, Inc. for service prior to the respirometer experiments in this study. Before proceeding with the respirometer experiments, a two-point CO2 sensor calibration was performed using grade-five nitrogen gas (0 % CO2) and 0.500 % CO2 gas with nitrogen balance (AIRCO Rare and Specialty Gases Dept. and Air Liquide, respectively). Directly preceding each experiment, the sensor volume and the headspace in the test chambers was measured and entered into the computer and a complete “System Diagnostics” was performed per the prescribed “Experiment Checklist” (Columbus Instruments International Corporation, 1994). CO2 production was determined at 2 or 3 hour intervals and raw data was recorded as both µg/mL and µg/mL-hr. The sample headspace was purged and refilled with refresh air after every other sample was taken (every 4 – 6 hours for 2 and 3 hour sample intervals).

3.1.2. Experimental Chamber Bottles 2–L PYREX® bottles (Corning No. 1395) were used as experimental sample vessels. These large bottles were used to accommodate for volume requirements for the final TPH analyses of experimental samples (1 L). Based on the TPH sample requirements,

headspace refresh time and the expected CO2 production rates, a sample volume of 1500 mL was selected for all respirometer experiments.

3.1.3. Magnetic Stirrers Experimental samples were stirred using nine VWR 200 Mini (magnetic) Stirrers (VWR CAT No. 58940-158) (Figure 3.2). The space below the Micro Oxymax condenser was

37

very limited. The stir plates had to be small enough to fit into the area provided, yet powerful enough to mix large groundwater samples effectively. The 200 Mini Stirrer was chosen because of its ability to stir large quantities of liquid (up to 2 L) and its compact size.

Figure 3.2: VWR 200 Mini Stirrer used in the respirometry experiments

3.1.4. Temperature Controlled Water Bath A temperature controlled water bath was used to simulate the approximate average groundwater temperatures at the TB8 biosparge site (19 o C). A Sterilite® plastic storage container (27 x 27x 8 cu inches) was used to construct a water bath large enough to contain nine of the 2-L experimental bottles (Figure 3.3). Nine, 5.5-inch diameter holes

38

were cut into the storage container lid allowing the sample bottles to be inserted into the water bath. The holes allowed access to the sample bottles, while the lid provided added insulation of the sample bottles from fluctuating room temperatures. Rings cut from a two-inch foam pad were used to insulate the necks of the sample bottles and sealing the gap between the water bath lid and the sample bottles (Figure 3.3). The foam rings proved effective in stabilizing the sample bottles over their respective stir plates. Halfinch, rigid foam board was used to mediate any heat transfer from the stir plates to the bottom of the water bath.

39

Figure 3.3: Temperature controlled water bath apparatus.

A water bath recirculator/temperature controller (VWR Scientific Products Model No. 1162) was used to control the temperature of the water bath during the respirometer experiments. The controller had both a cooling element and a heating element to provide precise temperature control. The water bath recirculator was located on the floor,

approximately two feet below the water level in the bath, to allow for passive gravitational flow from the water bath to the controller unit. To ensure even temperature control within the water bath, the incoming water flow was split into two flows (using 0.5 inch silicon tubing) as it was pumped into the water bath. Two pieces of half-inch silicon tubing were used to direct the water flows into the water bath through two holes cut into

40

the lid of the container (Figure 3.3). A drain was inserted into the side panel of the water bath to allow gravitational flow back into the temperature controller. The drain was inserted high on the side of the container so the water level in the water bath exceeded the groundwater level in the sample bottles. During the respirometer experiments, the

temperature was monitored in three zones of the water bath (two by Micro Oxymax temperature probes and the third using a mercury thermometer) to confirm no shortcircuiting of the temperature-controlled water developed between the water inputs and the drain.

3.2. Respirometer Experiment for Nutrients and Inoculum Respirometer Experiment 1 (EXP1) was initiated to evaluate the biosparge-mediated biodegradability of the contaminated groundwater at TB8 on January 22, 2003. The purpose of this experiment was to determine the respiration rate of non-amended TB8 groundwater and to assess whether nutrient (N, P, K, S) addition or inoculum addition would increase the respiration rate. This respirometry experiment was performed using nine (9) sample bottles with contents as described in Table 3.1. The test samples

included four sets of duplicates containing non-amended TB8 groundwater, groundwater with added nutrients, inoculated TB8 groundwater, and TB8 groundwater with both added nutrients and inoculum. Carbon dioxide production was measured on three-hour intervals over an experimental term of 28 days.

Table 3.1: Description of Respirometry Bottle Contents for Experiment 1 (EXP1) Bottle Number Bottle Description and Contents 1 Control - D.I. Water

41

2 3 4 5 6 7 8 9

Nonamended TB8 groundwater Nonamended TB8 groundwater TB8 groundwater plus added nutrients (6.5: 1.7: 1.7: 1.7 mg/L, N: P: K: S) TB8 groundwater plus added nutrients (6.5: 1.7: 1.7: 1.7 mg/L, N: P: K: S) TB8 groundwater plus added inoculum (10 % inoculum by volume) TB8 groundwater plus added inoculum (10 % inoculum by volume) TB8 groundwater plus added nutrients and inoculum (as above) TB8 groundwater plus added nutrients and inoculum (as above)

All experimental bottles initially contained a total of 1500 mL of groundwater (D.I. water for control). Experimental sample bottles not containing inoculum received 1500 mL of

TB8 groundwater. Sample bottles containing inoculated TB8 groundwater received 1500 mL of a mixture of TB8 groundwater and 10 % inoculum water.

Groundwater sample bottles were prepared by first shaking a five gallon groundwater container vigorously for two minutes prior to each sample bottle being filled. The groundwater was then dispensed into a clean 2-L beaker to approximately 1800 mL. The groundwater was then poured into a 1000-mL graduated cylinder at measured 1000 and 500 mL aliquots. Prior to measuring the 500 mL sample portions, the groundwater remaining in the 2000 mL beaker was swirled to reincorporate any material, potentially separating from the solution. Any groundwater remaining in the 2-L beaker after the

individual sample had been dispensed was returned to the five-gallon container to be mixed again. Once each sample bottle had received the prescribed quantity of groundwater (1500 mL), a clean Teflon-coated magnetic stir bar was added to each bottle

42

and the samples were re-capped and placed into the temperature controlled water bath to acclimate.

Diluent-contaminated Guadalupe groundwater used in a previous phytoremediation experiment was added as inoculum at 10 % of the total liquid volume (bottles 6 – 9). The phytoremediation experiment involved recirculating diluent contaminated groundwater (approximately 3.6 mg/L TPH initially) through microcosms filled with clean sand from the C8 phytoremediation site of the GRP. The experiment ran for over 105 days and

considerable TPH biodegradation occurred in the test microcosms containing sand only. The recirculated microcosm water was expected to contain well-acclimated hydrocarbondegrading microbes.

A total of 800 mL of inoculum water was required for the EXP1 respirometer experiment (600 mL for respirometry and 200 mL for TPH analyses of initial samples). More than 300 mL of inoculum water was removed from the recirculating reservoirs of each of the three microcosms containing sand only. This provided a total of 900 – 1000 mL of inoculum. The inoculum water was thoroughly mixed in a 2-L beaker to homogenize the sample. A 1000 mL graduated cylinder was used to dispense 800 mL of the mixed inoculum water to a 3.5-gallon glass carboy bottle containing 7.2-L of well-mixed TB8 groundwater. The inoculated TB8 groundwater was thoroughly mixed and 1500 mL portions were dispensed to experimental sample bottles 6 – 9 in 1000 and 500 mL aliquots. The remaining inoculated groundwater was reserved for TPH analysis.

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Ambient air (20.9 % O2) was used to refresh the headspace in the sample bottles for this respirometry experiment. The sample chambers were contained in a temperature

controlled water bath as described at an average temperature of 18.8 ± 0.4 oC. All sample bottles were stirred with magnetic stirrers.

Several operational problems were encountered with the respirometer during this first experiment. This resulted in discontinuous monitoring of CO2 production but allowed intermittent measurement of respiration. On January 25, 2003, the computer controlling the experiment was found shut off. The experiment was immediately restarted (EXP1A) using the same water samples. No respiration measurements were taken between January 24 at 09:00 and January 25 at 06:30 (21.5 hours). On February 3, 2003, an investigation of sporadic CO2 and O2 levels revealed a clogged drier line due to excessive moisture content. Respiration measurements were discontinued until the problem was remedied. The experiment was restarted the same day using the same water samples (EXP1B). No respiration measurements were taken on February 3 between at 11:00 and 16:00 (5 hours). On February 24, 2003, the computer controlling the respirometer was found shut off again. No CO2 samples had been recorded since February 21 at 08:30. The samples were maintained on stirrers with temperature control and adequate oxygen for the entire experimental period (1/22/03 – 2/24/03) – only the measurement of CO2 production, was interrupted (28 days of CO2 data were collected).

44

3.2.1. Sample Groundwater Collection BJF Services collected the groundwater used in this experiment from monitoring well S8, directly up gradient from the TB8 biosparge site, on January 20, 2003. The groundwater sample was delivered on the same day to the California Polytechnic State University Environmental Engineering Laboratory. The sample was stored in its sealed five-gallon plastic container at 4 o C for two days until it was used for experimentation.

3.2.2. Nutrient Preparation and Addition Nutrients were added to test bottles 4, 5, 8, and 9 to yield nutrient concentrations of 6.46: 1.67: 1.67: 1.67 mg/L (ppm), N: P: K: S, respectively. Nutrients were added in the form of Miracle Grow® (Miracid with trace elements and 30-10-10 % N-P-K respectively), plus ammonium sulfate (24 % S, 21 % N).

Calculations for Miracle Grow® addition were based on a desired 5 mg/L (5 ppm) of nitrogen. The calculations resulted in a required Miracle Grow® quantity of 25 mg to be added to the nutrient amended sample bottles. The quantity of ammonium sulfate required to supply sulfur at the equivalent level of the phosphorus and potassium contributed by the Miracle Grow® (1.67 mg/l) was calculated to be 10.417 mg. A measurement of 1.0147 g of ammonium sulfate was added to 100 mL of D.I. water to produce a 10.417-mg/L ammonium sulfate stock solution. A 1 mL aliquot of the

solution was then pipetted to the nutrient amended sample bottles.

45

3.2.3. Initial TPH and Nutrient Sampling and Analyses ZymaX Envirotechnology, Inc. performed TPH and inorganic nutrient analyses of theTB8 groundwater samples used in EXP1. The samples were analyzed for TPH using a combination of gas chromatography with mass spectrophotometry (State of California method similar to EPA Method 8015), after extraction in methylene chloride (EPA Method 3510). The TPH was quantified against diluent standards, over an analytical range of C8 – C40 with a Practical Quantitation Limit (PQL) of 50 µg/L. Inorganic nutrients analyses were performed to quantify the initial content of macronutrients in the TB8 groundwater. The initial samples were analyzed for ammonia-N (EPA Method 350.3); nitrate-N (EPA Method 300.0); nitrite-N (EPA Method 300.0); ortho-phosphate (EPA Method 300.0); and sulfate (EPA Method 300.0).

Surrogate recovery values were reported for each TPH sample analysis. A surrogate is a compound having similar physical and chemical properties as does the sample target analytes, but is not normally found in the natural environmental samples. The surrogate is applied to each sample at a known concentration and is recovered with the analytes (in this case TPH) during the analysis. The percent surrogate recovery is used as a

qualitative indicator of how well the extraction and analysis method is working, and surrogate recovery values between 50 and 120 % are generally considered acceptable (by ZymaX envirotechnology, Inc.). However, some target analyte and surrogate compound recoveries have shown strong correlation, and the use of such correlations to reduce bias in analytical samples has been suggested (Butler et al., 1995). To date, the use of surrogate recovery values is not recommended for use as an overall correction factor for

46

analyte recovery. However, according to the Assistant Laboratory Director of ZymaX envirotechnology, Inc., the surrogate value for a particular sample can be considered to be representative of the extent of target analyte (in this case TPH) recovery (Michael S. Ng, July 30, 2003).

Four (4) initial water samples were taken on January 22, 2003 (2 – non-amended TB8 groundwater and 2 – TB8 groundwater with 10 % inoculum water) for TPH and inorganic nutrient analyses. The 1 L amber glass sample bottles were filled between experimental sample preparations to ensure representative samples had been collected. The 1 L bottles were filled slowly, minimizing aeration, until a bead of water extended beyond the mouth of the sample bottle. The samples were cleared of any noticeable bubbles and tightly sealed. The samples were stored at 4 oC, for less than three (3) hours, until they were delivered to ZymaX Envirotechnology, Inc. for analyses.

3.2.4. Final TPH Sampling Methods Prior to final TPH sampling, the temperature control for the water bath was decreased to the lowest possible setting to reduce any further biological degradation. A Sension 6 digital dissolved oxygen meter (Hach Company p/n 51850-23), equipped with a Sension DO/Temperature electrode (Hach Company p/n 51970-03), was used to measure the dissolved oxygen and temperature in the experimental chambers before final sampling. The concentration of dissolved oxygen in the samples was measured for each bottle at a temperature of approximately 14 oC. A sample volume of 1 L was collected from each experimental bottle using the same sampling protocol described. All samples were stored

47

at 4 oC for 17 hours, until they were delivered to ZymaX envirotechnology, Inc. for final TPH analyses. No final nutrient analysis was performed.

3.3. Respirometer Experiments for DO Effect A series of three, 48-hour respirometer experiments (EXP2, EXP3, EXP4) were performed to investigate the effects of DO concentration on the rate of CO2 production for TB8 groundwater. DO concentrations of experimental samples were manipulated by using N2/air mixtures of varying composition for refresh air in the respirometer. The N2/air gas mixtures were prepared using two 85.7 L Tedlar® bags (light blocking with on/off valve), filled with gas mixtures of various oxygen concentrations (Figure 3.4). Since the samples were stirred continuously, equilibrium was maintained resulting in controlled DO concentrations. At the end of each run, the DO concentration in the sample waters was measured, and the CO2 production data were compiled. The average 48-hour CO2 production rates for all four respirometer experiments (EXP1-EXP4) were compared (including the non-amended runs described in Section 3.2).

48

Figure 3.4: An 85.7-L light blocking Tedlar® bag with on/off valve used to dispense refresh gas to the respirometer during experiments to investigate DO effect on TPH biodegradation.

3.3.1. Sample Groundwater Collection for DO Experiments The groundwater used in the DO experiments (EXP2 – EXP4) was part of a second round of samples delivered by BJF Services. The samples were again collected from

monitoring well S8, directly up gradient from the TB8 biosparge system, on March 19, 2003. Two resealable plastic sample containers (5-gallon and 6.5-gallon) were delivered on the same day to the California Polytechnic State University Environmental Engineering laboratory. The sample containers were stored in a laboratory refrigerator at approximately 5 oC until they were used for experimentation. All experiments using groundwater from these samples (EXP2 – EXP4 and the Long Term Biodegradability experiment) were initiated within the EPA recommended 7-day hold time limit for TPH sample storage.

3.3.2. Respirometer Experiment 2 (EXP2) Experiment 2 (EXP2) was initiated on March 20, 2003 and consisted of three 1.5 L samples of non-amended TB8 groundwater and one D.I. water control. The sample bottles were placed into a temperature controlled water bath at an average temperature of 18.6 ± 0.1 oC. All sample bottles were stirred with magnetic stirrers. The Tedlar® bags

49

used for the refresh air supply contained a pre-mixed gas with 8.02 % O2 with N2 balance (Scott Specialty Gases, Inc.). Carbon dioxide production was measured on three-hour intervals over an experimental term of 48 hours.

3.3.3. Respirometer Experiments 3 and 4 (EXP3 and EXP4) Sample DO concentrations for EXP3 and EXP4 were manipulated by using air/N2 gas mixtures of varying O2 concentrations for the refresh gas in the respirometer. Concentrations of 1.0 % and 0.25 % O2 in the refresh air were selected for EXP3 and EXP4, respectively. Calculations were performed to determine the quantities of air/N2 to be mixed to yield 1.0 % and 0.25 % O2 in the 85.7 L Tedlar® bags. The gas mixtures were prepared by filling the two Tedlar® bags with measured air/N2 flows from a Dasibi® gas calibrator. The filled Tedlar® bags were then connected in parallel to the respirometer refresh line via a plastic T fitting. Imperial Eastman ¼-inch Poly-Flo tubing (low O2 permeability polyethylene) was used for all fittings connecting the refresh bags to the respirometer refresh air input vent (Figure 3.5). ________________________________________________________________________

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Tedlar R Bags

1/4 “ PE Tubing T Fitting Refresh Air Input Micro Oxymax Sample Pump

Drierite Gas Drying Unit

Figure 3.5: Diagram showing the system used to deliver air/N2 mixtures of various O2 concentrations to the Micro Oxymax Refresh Air Input to control DO concentrations in respirometry samples.

Experiment 3 (EXP3) was initiated on March 23, 2003 and consisted of three 1.5 L samples of non-amended TB8 groundwater and one D.I. water control. The sample bottles were placed into a temperature-controlled water bath at an average temperature of 18.8 ± 0.3 oC. All sample bottles were stirred with magnetic

stirrers. The Tedlar® bags used for the refresh air supply contained 1.0 % O2 in an air/N2 mixture. Nitrogen gas was bubbled through the experimental samples prior to beginning the respirometer experiment to reduce the initial DO concentrations near to the concentrations expected by the % O2 in the refresh air. Carbon dioxide

51

production was measured on two-hour intervals over an experimental term of 48 hours.

Experiment 4 (EXP4) consisted of five 1.5 L samples of nonamended TB8 groundwater and one D.I. water control. The two additional groundwater samples were used to assess stirring on CO2 production and contained inactive stir bars. The investigation of stirring was prompted by the observation of a drop off in CO2 production in EXP2 when a magnetic stirrer failed. The sample bottles were placed into a temperature controlled water bath at an average temperature of 18.8 ± 0.1 o C. The Tedlar® bags used for the refresh air supply contained 0.25 % O2 gas. Again, N2 was bubbled through the samples to reduce the initial DO concentrations in the sample waters. Carbon dioxide production was measured on three-hour intervals over an experimental term of 48 hours. After 48 hours, the inactive stir bars were activated to identify any changes in the CO2 production due to stirring. The experiment was operated for an additional 117 hours.

3.4. Long-term Biodegradability Experiment An experiment was conducted to investigate the kinetics of the long-term biosparge mediated biodegradation of the contaminated groundwater at the TB8 when ample DO concentrations are maintained. An additional objective of the experiment was to

determine if biodegradation stops or slows down after initial biodegradation (all experiments described previously examined relatively short term degradation), and to determine at what concentration biodegradation slows down. To answer these questions,

52

experimental chambers capable of containing large volumes of groundwater were required to accommodate for regular TPH sampling and analyses.

The long-term biodegradability experiment was initiated on March 19, 2003. Two fivegallon glass carboy bottles initially containing 12 L of non-amended diluentcontaminated TB8 groundwater were subjected to specific environmental conditions for several months. To mimic groundwater temperature conditions at the TB8 biosparge site, the two bottles were incubated in an Equatherm Incubator (Curtin Matheson Scientific Inc.) at approximately 19 oC (Figure 3.6). No nutrient or microbial amendments were added during the initial stages of this experiment.

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Figure 3.6 Experimental incubator set-up for the Long-term biodegradability experiment

3.4.1. Dissolved Oxygen Delivery System The bottles were left open to the atmosphere, and air was bubbled through the water using aquarium air pumps (Elite 800 Air Pump, ART. # A-800, minimum output of 1500 cc/min at 2.5 psi). For each bottle, air was pumped through silicone air tubing (Hagen Art # A-1124) to a T fitting. The T fitting split the airflow to two pieces of silicone 54

tubing, each leading to a flow control valve (Lee’s Aquarium and Pet Products, Airline Control Kit, valves and T fittings). Each valve controlled the flow through a length of tubing capped with an air dispersion stone (Figure 3.7 and Figure 3.8). The air stones were used to release the air to the groundwater in the form of small bubbles, thereby increasing the oxygen transfer to the water. Dissolved oxygen was monitored to

determine the concentrations of DO provided to the hydrocarbon-degrading microbes present in TB8 groundwater.

________________________________________________________________________

Air P ump

Valves

Air S tone s

_______________________________________________________________________

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Figure 3.7: Diagram of the dissolved oxygen delivery system used to stimulate the biodegradation of TPH contamination in TB8 groundwater sample bottles during the long-term biodegradability experiment

Figure 3.8: A digital image of the dissolved oxygen delivery system used to aerate TB8 groundwater samples during the long-term biodegradability experiment

3.4.2. TPH Sampling and Analysis Before each TPH sample was taken, the bottles were covered and shaken to create a homogenous solution. Approximately 1200 mL of the groundwater was dispensed into a clean 2000 mL beaker. The groundwater was then poured into a 1-L amber glass sample bottle. The 1-L TPH sample bottles were filled slowly until a bead of water extended beyond the mouth of the bottle. The samples were cleared of any noticeable bubbles and

56

tightly sealed. The samples were stored at 4 oC, for less than 3 hours, until they were delivered to ZymaX envirotechnology, Inc. for analyses (see section 3.2.3. for TPH analysis methods). Any groundwater remaining in the 2000 mL beaker after the individual sample had been dispensed was swirled to reincorporate any material, possibly separated from the solution, and returned to the experimental bottle from which it was taken originally.

Initial TPH analyses were conducted plus two subsequent incremental TPH analyses (at 1.6 d and 6.6 d). Between the third sample (t = 6.6 d) and the fourth sample, an accident occurred during routine dissolved oxygen monitoring. Bottle 2 was broken while mixing the sample and the water was lost. Shortly thereafter, another bottle (Bottle 2A) was established in the incubator using left over water from the same initial groundwater sample. The goal of the additional bottle was to provide replication for the investigation into the rate of degradation at lower TPH concentrations and to determine if a TPH concentration exists where degradation no longer occurs at appreciable rates. An initial TPH sample of Bottle 2A was taken on April 9, 2003, and a second sample was taken 28 days later with the fourth Bottle 1 sample. The last scheduled TPH samples were performed on July 7th (Sample 5 for Bottle 1, Sample 3 for Bottle 2A).

Due to concerns of water evaporation effecting the TPH concentrations, an investigation of the water loss due to evaporation was begun on July 7, 2003. Lines were drawn on the experimental bottles marking the level of water at the time (t = 0). Exactly 14 days later, a measurement of the amount of D.I. water required to return the water level to the

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original marking was performed. On the same day (July 21, 2003), an additional TPH sample of the long-term experiment was taken as the initial TPH sample for an investigation of the effects of nutrients and stirring on the slowing biodegradation. After the sample was taken, a second line was drawn on the experimental bottles to quantify the evaporation rate once again. Exactly 14 days later (August 3, 2003), a measurement of water lost to evaporation over the two-week period was performed using this method. From these measurements, the average daily evaporation rates were calculated for each bottle. The evaporation rates were used to adjust the TPH concentration for losses of water due to evaporation.

3.4.3. Investigation of the Effects of Stirring and Nutrient Addition on the Long-term TPH Biodegradation in TB8 Groundwater

After the long-term experiment had run for 124 days, experimental conditions were varied to investigate the effects of stirring and nutrient addition on the slowing TPH biodegradation rates. After the initial TPH sample was secured on July 21, 2003 using the sampling protocol described in Section 3.2.2, nutrients were added to Bottle 1 at concentrations similar to those in EXP1 (no ammonium sulfate was added due to the discovery of high sulfate levels (130 mg/l) in the TB8 groundwater (see Table 4.3 of section 4.1.1). At the same time, a Teflon coated magnetic stir bar was activated in Bottle 2A to determine if stirring would increase the availability of the TPH and DO for microbial biodegradation. Stirring was not initially possible for the long-term

biodegradability study, due to the large volumes of groundwater used in the experiment. The DO delivery systems were the only source of mixing prior to the stir bar in Bottle 2A

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being activated. On August 3, 2003, final TPH samples were taken to determine if the rate of TPH biodegradation had been increased due to the stirring or nutrient addition.

Figure 3.9: Long-term experimental Bottle (2A) with an active magnetic stir bar and bubbling air stones. The lines on the bottle represent 1 L increments (TPH sample volume) and served as a basis for evaporation quantification.

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CHAPTER 4 RESULTS AND DISCUSSION

4.1 Nutrients and Inoculation Effects on Short-term TPH Respiration (EXP1) 4.1.1 Characterization of Groundwater Samples The groundwater used for this experiment (collected from monitoring well S8 at TB8) was highly turbid with a brownish orange (rusty) color, and emitted a mild hydrocarbon smell. Analysis of the two initial non-amended TB8 groundwater samples resulted in an average TPH concentration of 7350 ± 71 µg/L (Table 4.1). TPH analysis of the two initial TB8 groundwater samples containing ten percent inoculum water resulted in an average TPH concentration of 6800 ± 141 µg/L. The addition of 10 % inoculum water to the TB8 groundwater resulted in an average reduction in TPH concentration of 7.5 % because of the low TPH concentration in the inoculum. Since the TPH concentration decreased by 7.5 % instead of 10 % the inoculum water likely contained TPH at detectable levels. The TPH compounds present in all samples were limited to the C10 – C32 hydrocarbon range. For both pairs of initial TPH sample results (Inoculum and Nonamended), a decrease in surrogate recovery corresponded to a decreased concentration of TPH (Table 4.1).

Simulated distillation of the TPH compounds contained in the water samples showed similar TPH carbon chain distributions for the initial samples (Table 5.2). The majority of TPH in the TB8 groundwater samples was in the C14 – C24 range for both sample sets (Table 4.2 and Figure 4.1).

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Table 4.1: Sample Descriptions and Initial TPH Concentrations for EXP1 Lab Sample Surrogate Result (ug/L) Number Description Recovery (%) TPH Average Standard Hydrocarbon TPH Deviation 141 71 Range C10-C32 C10-C32 C10-C32 C10-C32 30408-1 (1) 10 % Inoculum 74 6700 6800 30408-2 (2) 10 % Inoculum 77 6900 30408-3 (3) Nonamended 73 7400 7350 30408-4 (4) Nonamended 70 7300 Practical Quantitation Limit (PQL) is 50 µg/l TPH for all samples. Analysis Hydrocarbon Range C8 – C 40 Samples were collected 1/22/2003

Table 4.2: TPH Carbon Chain Distribution of Initial Groundwater Samples Lab Number 30408-1 30408-2 30408-3 30408-4 Description (1) 10 % Inoculum (2) 10 % Inoculum (3) Nonamended (4) Nonamended C Range Hydrocarbon Concentration (µg/L) C10-C12 91 83 104 109 C12-C14 363 355 409 405 C14-C16 1061 1076 1182 1138 C16-C18 1530 1551 1633 1575 C18-C20 1631 1666 1773 1739 C20-C24 1544 1612 1701 1712 C24-C28 392 442 468 484 C28-C32 88 115 130 138 C32-C36 ND ND ND ND C36-C40 ND ND ND ND Total 6700 6900 7400 7300

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2000 1800 Initial Nonamended 1600 TPH Concentration (ug/L) 1400 1200 1000 800 600 400 200 0 C10-C12 C12-C14 C14-C16 C16-C18 C18-C20 C20-C24 C24-C28 C28-C32 Carbon Chain Range Figure 4.1 Initial total petroleum hydrocarbon carbon chain distributions for non-amened and inoculated samples Initial 10 % Inoculum

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Inorganic nutrient analyses of TB8 groundwater samples had total nitrogen levels of approximately 1 mg/L with the majority occurring in the form of nitrate (Table 4.3). The concentration of sulfate in all four samples was 130 mg/L. phosphate analyses were non-detectable (ND) for all four samples. Results for the Ortho

Table 4.3: Inorganic Nutrient Analysis of TB8 Groundwater Constituent Method PQL Nutrient Concentrations (mg/L) (mg/L) (1) 10 % (2) 10 % (3) (4) Inoculum Inoculum Nonamended Nonamended EPA 350.3 EPA Nitrate as N 300.0 EPA 300.0 Nitrite as N EPA Ortho Phosphate 300.0 EPA Sulfate 300.0 Ammonia-N 0.1 0.5 0.5 0.5 1.0 0.2 0.7 ND ND 130 0.2 0.9 ND ND 130 0.2 1 ND ND 130 0.2 1 ND ND 130

4.1.2 Respirometry Results for EXP1 Significant CO2 production was observed for all samples except the D.I. water control throughout the three portions of EXP1 (Figure 4.2). Cumulative CO2 production values for each of the three portions of EXP1 were added to acquire total cumulative CO2 production values for each sample (Table 4.4). The values in Table 4.4 represent the total CO2 collected from each sample over the complete 28-day experimental period. The highest average CO2 production was observed in the non-amended and the samples

63

amended with nutrients only (255.2 ± 3.9 µg/mL and 257.2 ± 6.2 µg/mL, respectively) (Figure 4.3). The samples containing nutrients and inoculum produced an average of 232.2 ± 4.0 µg/mL CO2. The samples containing inoculum only produced the least amount of CO2 of any of the test samples with 218.8 ± 1.9 µg/mL. The total background CO2 produced in the sample control was 1.9 µg/mL.

The addition of inoculum with or without nutrients slightly decreased total CO2 production over the 28-day experiment (Figure 4.2). For the noninoculated groundwater samples, added nutrients did not significantly increase the amount of total CO2 produced (Figure 4.3). For the inoculated samples, only a slight nutrient benefit was observed (Figure 4.3).

Table 4.4: Total CO2 Production for Nutrient/Inoculum Experiment (EXP1) Bottle Control Nonamended Nutrients Inoculum Nutrients/Inoculum Contents and Number (1) (2) (3) (4) (5) (6) (7) (8) (9) Total CO2 Production 1.9 257.9 252.5 252.9 261.6 220.1 217.5 235.0 229.4 (µg/mL) Average CO2 (µg/mL) 1.9 255 257 219 232 Standard Deviation (µg/mL) 3.9 6.2 1.9 4.0

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250 225 200
CO2 Production (µg/mL)

175 150 125 100 75 50 25 0 0 100 200 300 400 Time (hours) 500 600 700

Control: D.I. Water Unamended Unamended Nutrients Nutrients Inoculated Inoculated Nutrients/Inoculated Nutrients/Inoculated

Figure 4.2: Total CO2 production (EXP1+EXP1A+EXP1B) versus time for TB8 groundwater with and without nutrient, inoculum, and nutrient/inoculum amendments

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In the early stages of the experiment, the CO2 production was fastest for the non-amended TB8 groundwater samples (Figure 4.2). After approximately 72 hours, the rates of CO2 production for the nutrient amended samples (with and without inoculum) were approximately equal to those of the nonamended samples. Shortly thereafter, the CO2 production rates for the nutrient amended samples surpassed thoses for the non-amended samples. Over the last 10 days of the 28-day experiment, the rates of CO2 production in the nutrient amended samples was an average of 30.8 % higher (P = 0.0031) than those for samples not receiving nutrients. These increased rates of CO2 production in the nutrient amended samples (later in the experiment) may represent evidence of nutrient limitations for the long-term biodegradtion of TPH in the TB8 groundwater and/or a partial lag phase due to increased growth of microbes in the presence of higher nutrient levels in the groundwater. However, because of the faster respiration of the nonamended samples in the first 48 hours of the experiment, the nutrient amended samples had not significantly surpassed the non-amended samples in total CO2 production by the end of the experiment (Figure 4.2).

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300 255 250 Average Total CO2 Production (ug/mL) 232 219 200 257

150

100

50

2 0 Control Non-amended Nutrients Sample Description Inoculum Nutrients/ Inoculum

Figure 4.3: The average total carbon dioxide production for nonamended groundwater, and groundwater with added nutrients, inoculum and nutrient/inoculum.

67

4.1.3 TPH Results for EXP1 The final TPH analyses (at 28 days) had significant TPH degradation for all samples (Table 4.5 and Figure 4.4). Unfortunately, ZymaX envirotechnology personnel accidentally mixed together a sample from the non-amended set and a sample from the nutrient set. Therefore, only one sample of each of these could be analyzed and statistical analysis for those sample treatments are not possible. Based on the TPH results, the addition of nutrients appears to increase the

degradation rates slightly (Figure 4.4). However, for inoculated groundwater, the effect of nutrient addition was not statistically significant (see error bars in Figure 4.4, P-value = 0.43), and statistics are not available to determine the significance of nutrient addition in the case of non-inoculated groundwater. The variability of the surrogate recovery values was matched by similar differences in TPH concentrations recovered, indicating much of the observed variability of TPH data was due to

Table 4.5: Final TPH Sample Descriptions and Results Lab Number Sample Description 30744-1 30744-2 30744-4 30744-5 30744-6 30744-7 (2) & (4) Mixed (3) Non-amended (5) Nutrients (6) Inoculum (7) Inoculum (8) Nutrients/Inoculum Surrogate Recovery (%) 79 81 76 79 75 71 Result (µg /L) 4200 4400 3700 4300 4000 3100 C10-C32 C10-C32 C10-C32 C10-C32 C10-C32 C10-C32 C10-C32

30744-8 (9) Nutrients/Inoculum 80 4000 *Samples 2 & 4 were inadvertently mixed together by Zymax personnel. Practical Quantitation Limit (PQL) is 50 µg/l TPH for all samples. Samples were collected 2/24/2003

68

8000

7350
7000

7350 6800 6800

Initial

Final

6000

TPH Concentration (ug/L)

5000

4400
4000

4150 3700

3550

3000

2000

1000

0

Non-amended

Nutrients Sample Discription

Inoculum

Nutrients/Inoculum

Figure 4.4: A comparison of initial and final average total TPH concentrations in Non-amended TB8 groundwater and TB8 groundwater with added nutrients, inoculum, or nutrients/inoculum. *Duplicate samples were not available for these cases, consequently no error bars are presented.

69

The amount of TPH degraded was calculated for each of the experimental samples used in EXP1 (Table 4.6). The amount of TPH degraded in the single nonamended sample was 2950 µg/L. The TPH degradation for the lone nutrient amended sample was

considerably higher at 3650 µg/L. The least amount of TPH degradation occurred in the inoculum-amended samples at an average value of 2650 ± 354 µg/L. The

nutrient/inoculum samples had an average TPH degradation value of 3250 ± 778 µg/L.

Based on the measured TPH concentrations and the initial nutrient concentrations, the C: N and C: P mole ratios can be compared to those considered adequate for enhanced biodegradation. Considering the average TPH concentration in the groundwater without added inoculum of 7.35 mg/L, and assuming TPH is approximately 80 % carbon, the average concentration of carbon in the non-amended TB8 groundwater samples was calculated to be 5.9 mg/L. Since the total nitrogen measured initially in the non-amended groundwater samples was 1.2 mg N/L, the C: N mole ratio was 100: 17. Ortho phosphate was not detected in the groundwater. Based on the recommended carbon to nutrient mole ratio range of 100: 1: 0.5 to 100: 10: 1 (C: N: P, respectively) necessary to enhance biodegradation (see Section 2.5.1), the biodegradation of TPH in TB8 groundwater is not expected to be limited by nitrogen availability. Phosphorus could be expected to limit the biodegradation potential of TPH in TB8 groundwater. However, if we consider the possibility of the presence of ortho phosphate in concentrations less than the analysis PQL (0.5 mg/L), for example 0.1 mg/L, the C: N: P mole ratio becomes 100: 17: 0.7, which would fall within the range recommended for enhanced biodegradation. Because phosphate has a high affinity for most soils and is known to form precipitates in

70

environments with concentrations of iron, low concentrations of ortho phosphate in TB8 groundwater are expected (Liebeg and Cutright, 1999, USEPA, 1995). The use of a more precise analysis method for the presence of ortho phosphate would be beneficial in understanding the true phosphate limitation on the bioremediation at the TB8 site.

Table 4.6: Nutrients and Inoculum Effects on Respiration and TPH Biodegradation in TB8 Groundwater Samples. TPH Sample Average Standard Final TPH Standard Average Standard Deviation Degraded Degraded Deviation Description Initial TPH Deviation (µg/L) (µg/L) (µg/L) (µg/L) (µg/L) (µg/L) (µg/L)

Non-amended Nutrients Inoculum 1

7350 7350 6800

71

4400 3700

* * 212

2950 3650 2500

2950 3650 2650

* * 354

141

4300

Inoculum 2 6800 4000 2800 Nutrients/ Inoculum 1 6800 3100 3700 141 636 Nutrients/ Inoculum 2 6800 4000 2800 *Duplicate samples were inadvertently mixed together by Zymax personnel.

3250

778

4.1.4 Quantitative relationship between TPH degraded and CO2 evolved: The amount of CO2 produced per unit of TPH degraded was calculated by dividing the total CO2 production for each sample by the amount of TPH degraded in the same sample (Table 4.7). The amount of CO2 produced per unit TPH degraded (averaged over all samples used in EXP1) was 78 ± 9 µg CO2/µg TPH. This value is considerably greater than the CO2/TPH value resulting from stoichiometric calculations using dodecane 71

(C12H26) to represent TPH (3.11 µg CO2/µg TPH) (see Section 2.5). Similar differences between actual and expected values of chemical oxygen demand (COD) occurred in other studies using diluent-contaminated groundwater from the Guadalupe phytoremediation site (Hoffman, 2003). COD is a measure of the oxygen demand for the degradation of chemicals in a media and CO2 is a byproduct of the degradation. At this time it is not clear why the observed CO2 production and COD are so much higher than what is expected from observed TPH degradation.

Table 4.7: Data Showing the Relationship Between CO2 Production and TPH Degradation CO2/TPH Sample Total CO2 TPH Description Produced Degraded (µg/L) (µg/mL) (µg/µg)

Nonamended Nutrients Inoculum 1 Inoculum 2 Nutrients/ Inoculum 1 Nutrients/ Inoculum 2

252.49 261.58 220.14 217.50 235.05 229.38

2950 3650 2500 2800 3700 2800 Average = Std. DEV =

86 72 88 78 64 82 78 9

72

4.1.5 Nutrient and Inoculum Effects on Carbon Chain Distribution The carbon chain distribution of the remaining TPH after 28 days of biodegradation shows the final TPH concentrations within the individual hydrocarbon ranges (Table 4.8). By comparing these distributions to those of the initial samples, the biodegradation of each TPH fraction can be observed. Further, these results can be used to examine the effects of nutrients and/or inoculation on the carbon chain distributions. To examine the effects of nutrients on carbon chain distribution a direct comparison was made between the carbon chains distributions of samples with and without nutrients, both for the inoculated case and the non-inoculated case. This is straightforward because the same initial water sample was used with nutrients and without nutrients (for comparison of non-inoculated samples separately from the comparison of inoculated samples). However,, examination of the effect of inoculum on carbon chain distribution is more difficult because the addition of inoculum changed the initial carbon chain distribution.. Therefore, to make the comparison between initial and final carbon chain distributions for inoculated samples vs. non-inoculated samples, the TPH concentrations in each fraction were normalized and presented as a percentage of the TPH remaining for each fraction.

The effect of nutrient addition on carbon chain distribution was examined for inoculated samples. The average final TPH carbon chain distribution of the samples containing inoculum only and nutrients/inoculum are compared with their initial TPH carbon chain distribution in Figure 4.5. Significant degradation took place with and without nutrient addition, over all carbon chain ranges (Figure 4.5). For all carbon chain ranges, the TPH

73

in the nutrient amended inoculated groundwater was degraded more than with inoculum only. However, none of the carbon ranges had a statistically significant difference between the final TPH values of the experimental treatments (note overlapping error bars in Figure 4.5). A two-sample t-test confirmed no significant biodegradation benefit could be attributed to the addition of nutrients to inoculated TB8 groundwater (P-values ranged from 0.21 to 0.51 over the individual carbon ranges).

Table 4.8: Final Total Petroleum Hydrocarbons Carbon Chain Distribution in EXP1 (After 28 Days) Mixed Nonamended Nutrients Inoculum Nutrients/Inoculum Lab Number 30744-1 Description (2) & (4) C Range C10-C12 C12-C14 C14-C16 C16-C18 C18-C20 C20-C24 C24-C28 C28-C32 C32-C36 C36-C40 Total 22 157 622 1094 1085 902 245 73 ND ND 4200

30744-2 (3) 25 163 657 1153 1131 935 253 83 ND ND 4400

30744-4 (5)

30744-5 30744-6 30744-7 30744-8 (6) (7) (8) (9) 11 114 482 858 798 627 157 53 ND ND 3100 15 148 611 1078 1028 826 218 76 ND ND 4000

Hydrocarbon Concentration (µg/L) 14 28 21 131 167 146 549 639 589 995 1120 1052 970 1090 1027 778 908 845 198 255 234 65 93 86 ND ND ND ND ND ND 3700 4300 4000

The effect of nutrient addition on TPH degradation was examined by comparing the average final carbon chain distribution of the samples containing non-amended groundwater and groundwater with only added nutrients (not inoculated) (Figure 4.6).

74

The hydrocarbon distribution appears very similar to the distribution for the inoculated samples, with considerable degradation apparent over all of the carbon chain ranges. Once again, over all carbon ranges, the TPH degradation was greater for the sample containing nutrients (Figure 4.6). However, because the duplicate samples for both experimental treatments (Nonamended and Nutrients) were mixed together during analysis, there are no statistics to evaluate the significance of this trend. Although the trends are not statistically significant, the consistency of the apparent nutrient benefit on TPH degradation may further signify nutrient limited conditions towards the end of the experiment.

75

1800 1600 1400

Initial Inoculum Final Inoculum Only Final Nutrients and Inoculum

TPH Concentration (µg/L)

1200 1000 800 600 400 200 0 C10-C12 C12-C14 C14-C16 C16-C18 C18-C20 C20-C24

C24-C28

C28-C32

Carbon Chain Range

Figure 4.5: Effect of nutrient amendment on biodegradation of TPH fractions in inoculated TB8 groundwater. Actual TPH concentrations of each fraction remaining are presented for duplicate samples.

76

2000 1800 1600 TPH Concentration (µg/L) 1400 1200 1000 800 600 400 200 0 C10-C12 C12-C14 C14-C16 C16-C18 C18-C20 C20-C24 C24-C28 C28-C32 Carbon Chain Range

Initial Non-amended Final Non-amended Final Nutrients

Figure 4.6: Effect of nutrient addition on biodegradation of TPH fractions in non-inoculated TB8 groundwater.

77

To examine the effect of added inoculum on the carbon chain distribution, the values calculated for the TPH degraded were converted to the percentage of TPH remaining to allow for direct comparisons between samples having different initial TPH concentrations (groundwater with and without 10 % Inoculum). The Percentage of TPH remaining was used to compare the difference in TPH degradation between the non-amended samples and the samples with inoculum (Figure 4.7). The only difference between the

degradation in these samples occurred in the C28 - C32 carbon range, where the inoculation appeared to decrease degradation. However, no statistics were possible for the non-amended sample set for the reasons described. The same method was used to compare the degradation in the nutrient and nutrient/inoculum amended samples (Figure 4.8). No difference occurred in the TPH degradation between the TB8 groundwater with nutrients and inoculum and TB8 groundwater with nutrients only except for slightly less degradation of the C28 – C32 range with inoculum. Again, no statistics were available for the nutrient amended sample set.

At the end of this experiment, the dissolved oxygen levels in the sample waters ranged from 97.3 % to 109.2 % of saturation at approximately 14 oC. These DO levels

corresponded to an average DO concentration of 10.7 mg/L. A final measurement of the volume of D.I. water remaining in bottle 1 (D.I. Control) revealed the loss of approximately 14 mL of water to evaporation over the duration of the experiment. The amount of water lost to evaporation was less than one percent (1 %) of the total groundwater sample. Therefore, the final TPH concentrations were not adjusted to reflect decreases due to evaporation.

78

120.0

100.0
Percent of Initial TPH Remaining (%)

Non-amended Inoculum Added

80.0

60.0

40.0

20.0

0.0 C10-C12 C12-C14 C14-C16 C16-C18 C18-C20 Carbon Chain Range C20-C24 C24-C28 C28-C32

Figure 4.7: Effect of inoculation on the biodegradation of TPH fractions in groundwater samples with no nutrients added. A comparison of the percent of initial TPH remaining in nonamended TB8 groundwater samples and TB8 groundwater with added inoculum.

79

100.0 90.0 Nutrients Percent of Initial TPH Reamining (%) 80.0 Nutrients/Inoculum 70.0 60.0 50.0 40.0 30.0 20.0 10.0 0.0 C10-C12 C12-C14 C14-C16 C16-C18 C18-C20 Carbon Chain Range C20-C24 C24-C28 C28-C32

Figure 4.8: Effect of inoculum addition on biodegradation of TPH fractions for groundwater with nutrients added. A comparison of the percent of initial TPH remaining in TB8 groundwater samples with added Nutrients and TB8 groundwater samples with added Nutrients and Inoculum.

80

4.2 Effect of Dissolved Oxygen Concentration on Respiration Rates of TPHContaminated Groundwater

Decreasing the oxygen concentrations in the respirometer refresh air reduced the average DO levels in the sample waters as expected providing a range of 0.5 to 10.7 mg O2/L (Table 4.9). Reducing the dissolved oxygen in the sample waters significantly decreased the CO2 production (Figure 4.9 and Table 4.9). However, significant biodegradation was observed even with only 0.5 mg/L-dissolved oxygen (Figure 4.9). These samples had only 5 % as much DO as the samples purged with air, but the CO2 production rate was 50 % of the value for samples purged with air (Table 4.9). Although, not all the CO2 produced in the respirometric experiments can be attributed to the biodegradation of TPH, this may indicate the concentration of dissolved oxygen need not be high for biodegradation. A similar experiment by Yerushalmi et al (2001) reported the recovery of 65.9 ± 1.6 % of
14

C labeled CO2 from benzene biodegradation, following two

additions of DO at the initial concentration of 0.5 mg/L. However, the CO2 production rate may have remained high due to the evolution of CO2 from the same chemical reactions responsible for the excessive CO2 production (in excess of stoichiometric estimates) discussed in section 4.1.4.

81

Table 4.9: Oxygen Percentages in Respirometer Refresh Air, Average Dissolved Oxygen Concentrations, and Corresponding 48-Hour Average Carbon Dioxide Production Values. Standard Avg. CO2 Standard Avg. 48- Standard Experiment Oxygen in Avg. Deviation hour CO2 Deviation Rate Refresh Final DO Deviation (mg/L) (µg/ml-hr) (µg/ml-hr) (µg/ml) (µg/ml) (mg/L) Air (% v/v) EXP1 EXP2 EXP3 EXP4 20.9 8.02 1.00 0.25 10.7 3.9 1.9 0.5 0.1 0.2 0.1 0.1 2.6 2.0 1.5 1.3 0.1 0.1 0.0 0.0 122.8 94.4 74.3 64.7 2.4 6.5 0.9 0.6

82

3.0

2.5 Average CO2 Production Rate (µg/ml-hr)

2.0

1.5

1.0

0.5

0.0 0.0 2.0 4.0 6.0 8.0 10.0 12.0 Average Dissolved Oxygen Concentration (mg/L)

Figure 4.9: Effect of DO concentration on CO2 production rates during TPH biodegradation for TB8 groundwater. Rates are averaged over the 48-hour runs. Two-dimensional error bars indicate standard deviation of both the CO2 production rates and the measured DO concentrations.

83

4.3 Implications of Stirring on Respiration During the 48-hour EXP 2 respirometry run (8.02 % O2 in the respirometer refresh air) a reduction in CO2 production was observed when one of the triplicate TB8 groundwater samples was found with a failed magnetic stirrer. A significant decrease in the CO2 production from this sample (Series 3) was observed beginning at hour 24 (Figure 4.10). The loss of stirring resulted in a 30 % reduction in the total CO2 produced over the 48-hour experiment. The observed benefit of stirring likely stemmed from the increased transport of DO in the fluid, as well as an increase in the bioavailability of the TPH in the groundwater. The stirring may better

approximate the processes occurring in the field because it simulates the mixing involved with water filtering between soil particles in the subsurface. 120
Control (D.I. Water)

Cummulative CO2 Production (µg/mL)

100 80 60 40 20 0 0 -20

Series2 Series3 Series4

10

20

30 Time (hours)

40

50

60

Figure 4.10 Cumulative CO2 production versus time with a reduction of CO2 produced by the Series 3 triplicate sample after the magnetic stirrer failed at approximately 24-hours. 84

The identification of changes in CO2 production due to a loss of stirring during EXP2 encouraged more investigation into the phenomenon. Therefore, two additional groundwater samples, containing inactive stir bars, were included in the EXP4 respirometer experiment. The experiment was allowed to run the normal 48-hours with three groundwater samples stirred and two not stirred. After 48-hours, the two inactive magnetic stirrers were activated and the CO2 production measurements were continued (Figure 4.11). Over the first 48-hours virtually no difference existed in CO2 production among the three stirred samples, which produced an average of 64.6 µg/mL. However, the non-stirred samples produced an average of 80 % less CO2 over the 48-hour interval with an average cumulative CO2 production of 12.2 µg/mL. After the stirrers were activated, the CO2 production in one of the samples increased sharply, while the production in the other sample was not affected (Figure 4.11).

120 110 Cumulative CO2 Production (µg/ml) 100 90 80 70 60 50 40 30 20 10 0 -10 0

Control (D.I. Water) Stirred 1 Stirred 2 Stirred 3 Nonstirred 1 Nonstirred 2

20

40

60

80 100 Time (hours)

120

140

160

180

Figure 4.11: Cumulative CO2 production vs. time showing a reduction of CO2 production in duplicate samples containing inactive magnetic stirrers over 48-hours.

85

4.4 Long-term Biodegradability Experiment

To examine the long-term biodegradability of the hydrocarbons in the diluent-contaminated groundwater, two large samples were aerated for a total of 138 days while monitoring TPH concentration. The dissolved oxygen concentrations in these sample bottles consistently

remained at or near saturation (between 8 – 10 mg/L), indicating the aeration was adequate throughout the experiment (Table 4.10). The TPH data for the long-term experiment were somewhat sporadic revealing some initial biodegradation followed by very slow degradation (Table 4.11, Figure 4.12). The TPH concentrations of Bottle 1 were sporadic for the first week and then leveled off at 5200 µg/L for the remainder of the experiment (Table 4.11). A steady decline in TPH concentration was observed in the first three samples of Bottle 2, but it also leveled off at about 5000 µg/L (Figure 4.12). Similar fluctuations of TPH concentrations were observed throughout the individual TPH carbon chain distributions for all three experimental bottles (data not shown).

Table 4.10: Measured Dissolved Oxygen Concentrations and Percent of DO Saturation for the Long-term Experimental Sample Bottles (1), (2), (2A)

DO (mg/L) Date 3/22/2003 3/25/2003 3/26/2003 5/7/2003 7/7/2003 8/4/2003 Average Std. DEV

DO (% Saturation)

Bottle (1) Bottle (2)/(2A*) Bottle (1) Bottle (2)/(2A*) 8.64 8.50 8.46 9.66 9.50 8.64 8.90 0.5 8.82 9.99 9.17 10.20 8.78 8.50 9.24 0.7 91.70 88.20 91.30 102.50 97.20 98.30 94.87 5.3 94.90 104.30 96.60 109.50 91.90 91.50 98.12 7.3

86

The TPH values shown in Table 4.11 and Figure 4.12 do not consider the likely increases in TPH concentration due to evaporative water loss. The sample bottles were open to the atmosphere and aerated so they likely lost considerable amounts of water to evaporation. If significant quantities of water were lost to evaporation over the experimental time period, the reported concentrations of TPH in the samples would be higher than the actual concentration of TPH. To Account for the possible evaporation, evaporation rates were measured and used to correct the TPH concentrations. These corrections are described in the next section (Section 4.4.2).

Consideration of the surrogate recovery values for the individual samples may offer some explanation of the sporadic TPH results (Table 4.11). The surrogate recoveries varied from only 66 % to as high as 97 %. The jump in surrogate recoveries observed between the initial and second TPH samples of Bottle 1, likely contributed to the increase in TPH observed over this sample interval (Table 4.11). Although, the surrogate recovery information may not be used as a direct correction factor, it may explain some of the variability in the TPH data.

87

Table 4.11: TPH Carbon Chain Distribution and Total TPH for Bottles (1), (2,) and (2A) from the Long-term Biodegradability Study. Values were not Adjusted for Water Loss Due to Evaporation. Fifth (t = 110 d) Initial (t = 0) Second (t = 1.6 d) Third (t = 6.6 d) Initial 2A Fourth (t = 48.8 d) (t2 = 28.2 d) (t2 = 89.4 d) (t2 = 0)

Bottle Number Surrogate Recovery C Range C10-C12 C12-C14 C14-C16 C16-C18 C18-C20 C20-C24 C24-C28 C28-C32 C32-C36 C36-C40 Total

Bottle (1) Bottle (2) Bottle (1) Bottle (2) Bottle (1) Bottle (2)

Bottle (2A)

Bottle (1)

Bottle (2A)

Bottle (1)

Bottle (2A)

66% 87 349 989 1357 1312 1122 291 93 ND ND 5600

81% 99 428 1205 1630 1592 1335 371 140 ND ND 6800

96% 79 395 1156 1665 1532 1184 331 158 ND ND 6500

91% 71 368 1085 1580 1434 1115 309 138 ND ND 6100

90% 55 288 917 1437 1336 972 295 100 ND ND 5400

85% 51 277 889 1404 1312 951 312 104 ND ND 5300

79% 30 203 869 1494 1478 1105 250 71 ND ND 5500

89% 18 189 880 1550 1357 979 194 33 ND ND 5200

87% 24 202 954 1679 1526 1137 234 44 ND ND 5800

97% 11 185 796 1488 1303 1014 284 119 ND ND 5200

87% 16 178 800 1442 1240 945 272 107 ND ND 5000

Hydrocarbon Concentration (µg/L)

88

8000 7000 6000 TPH Concentration(ug/L) 5000 4000 3000 2000 1000 0 0 20 40 60 Time (days) Figure 4.12: TPH concentrations vs. time during the long-term biodegradation of TB8 groundwater without nutrient or inoculum addition. Concentrations were not adjusted to account for the evaporation of water from the sample bottles. 80 100 120 Bottle (1) Bottle (2) Bottle (2A)

89

4.4.1 Evaporation Effects on TPH Concentrations During The Long-term Biodegradability Experiment Towards the end of the long-term experiment, measurements of water loss indicated substantial evaporation from the long-term sample waters (Table 4.12). Average

evaporation rates were calculated using the measurements of the water lost over two 14day periods. The resulting average evaporation rates were 14.2 ± 1 mg/L and 17.9 mg/L for Bottles 1 and 2A, respectively. The difference in evaporation rates for the two bottles may be attributed to the size of the bubbles pumped through the sample water. The aeration system in Bottle 2A delivered the air into the groundwater in fine bubbles. The larger surface area of the smaller bubbles would likely entrain water vapor more readily than would the larger bubbles produced by the air stones in Bottle 1.

Table 4.12 Evaporation Measurements Over Two 14-Day Periods and the Resulting Calculated Evaporation Rates Start Date Final Date 7/7/2003 7/21/2003 7/21/2003 8/4/2003 Time (days) 14 14 Average Std. DEV Evaporation (mL) 188 210 199 16 250 250 250 0 Evaporation Rate (mL/day) Bottle (2A) 17.9 17.9 17.9 0 13.4 15.0 14.2 1

Bottle (1) Bottle (2A) Bottle (1)

The evaporation rates were applied to the TPH data to correct the concentrations for evaporation loss (Table 4.13). First, the total water loss at the time of sampling was estimated by multiplying the evaporation rate (Er) by the number of days at which the

90

sample was taken (td). Then this quantity of water evaporated, was subtracted from the quantity of water remaining in the sample bottle if no water was lost to evaporation (R) to determine the actual volume. Finally, the corrected TPH concentration (Ccorrected) was calculated by multiplying the measured TPH concentration in each carbon range (Cobs) by the actual volume th [R – (Er * td)] and dividing by the volume expected without evaporation (R). In Summary, Ccorrected = {Cobs * [R – (Er * td)]} / R (Equation 4.1)

Because evaporation rates were low, the corrections for evaporation did not reduce the concentration of TPH in the samples taken over a short period of time. However, concentrations of TPH from samples taken over large time periods were reduced considerably (Figure 4.13). Using TPH concentrations corrected for evaporation, it

appears the TPH biodegradation in the TB8 groundwater samples was relatively rapid over the initial six days of the experiment, and then slow and steady over the long-term Figure 4.13).

The levels of TPH for the 110 day analyses were similar to the final concentration of TPH observed in the non-amended sample of EXP1 at 28 days (4400 µg/L). This result demonstrates similar degradation rates occurred over the first 28 days of the respective experiments. The respiration observed in EXP1 within the first 7 days complements the TPH degradation in the long-term samples within the same time period. Both

experiments were characterized by early rapid degradation/respiration, followed by

91

considerable slowing of the degradative processes. Of the total TPH degradation observed in Bottle 1 over 110 days, 47 % was degraded in the first 6.6 days of the experiment. The 47 % is based on a comparison of the maximum TPH concentration (for Bottle 1, the sample taken at 1.6 days was higher than the initial, see respective surrogate recoveries) and the concentrations observed at 6.6 days and 110 days. Similarly, an average of 76 % of the total CO2 produced in the nonamended samples of EXP1, evolved over the first 7 days of the 28-day experiment.

One difference between EXP1 and the long-term biodegradability experiment stands out as a possible explanation for the decreased rate of biodegradation observed in the longterm experiment. The samples used in EXP1 received constant stirring where the longterm samples were not mechanically stirred (minimal stirring would have been provided by the bubbling of air through the samples). Given the significant reductions in CO2 production observed without strirring (Section 4.3), a lack of thorough mixing within the long-term samples possibly limited the rates of biodegradation. This stirring effect was further investigated by adding stirring at the end of the long-term experiment. The results are described in the next section.

92

Table 4.13 TPH Carbon Chain Distribution and Total TPH Concentrations for Bottles (1), (2,) and (2A) of the Long-term Biodegradability Experiment Adjusted for Water Loses Due to Evaporation Initial (t = 0) Second (t = 1.6 d) Third (t = 6.6 d) Initial 2A (t2 = 0) Bottle (2A) Fourth (t = 48.8 d) (t2 = 28.2 d) Bottle (1) Bottle (2A) Fifth (t = 110 d) (t2 = 89.4 d) Bottle (1) Bottle (2A)

Bottle Number Surrogate Recovery C Range C10-C12 C12-C14 C14-C16 C16-C18 C18-C20 C20-C24 C24-C28 C28-C32 C32-C36 C36-C40 Total

Bottle (1) Bottle (2) Bottle (1) Bottle (2) Bottle (1) Bottle (2)

66% 87 349 989 1357 1312 1122 291 93 ND ND 5600

81% 99 428 1205 1630 1592 1335 371 140 ND ND 6800

96% 79 394 1153 1661 1529 1181 330 158 ND ND 6485

91% 71 367 1083 1577 1431 1113 308 138 ND ND 6087

90% 54 285 907 1422 1322 962 292 99 ND ND 5344

85% 50 274 880 1390 1299 942 309 103 ND ND 5248

79% 30 203 869 1494 1478 1105 250 71 ND ND 5500

89% 16 173 804 1416 1239 894 177 30 ND ND 4750

87% 23 191 903 1589 1444 1076 221 42 ND ND 5489

97% 9 144 618 1156 1012 788 221 92 ND ND 4040

87% 13 144 648 1167 1004 765 220 87 ND ND 4047

Hydrocarbon Concentration (ug/L)

93

8000 7000 6000 TPH Concentration(ug/L) 5000 4000 3000 2000 1000 0 0 20 40 60 Time (days) Figure 4.13: TPH concentrations vs. time adjusted for water loss due to evaporation during the long-term biodegradation of TB8 groundwater without nutrient or inoculum addition. 80 100 120 Bottle (1) Bottle (2) Bottle (2A)

94

4.4.2 Results for the Investigation of the Effects of Stirring and Nutrient Addition on the Long-term TPH Biodegradation in TB8 Groundwater Because the observed long-term TPH biodegradation was slow, two experiments were initiated at the end of the long-term run, to determine whether a lack of nutrients or stirring was limiting biodegradation. At day 124, Bottle 1 received nutrients (N, P, K) and Bottle 2A received constant stirring. After 14 days, the TPH concentration in Bottle 2A had decreased slightly (0.2 mg/L) with the addition of stirring (Table 4.14 and Figure 4.14). However, the biodegradation of TPH in Bottles 1 and 2A, over the previous 14day period (days 110 – 124, without stirring or nutrients) was considerably higher (0.94 mg/L and 1.15 mg/L, respectively). An increase in TPH was observed for the nutrient amended Bottle 1, over the 14 days since adding nutrients (surrogate recoveries of 74 and 72 %, respectively) (Table 4.14). This increase in TPH over the 14-day period may have been due to sampling error or analytical variability. Further examination of the sample analyses of Bottle 1 at days 110 – 124 revealed a 23 % reduction in surrogate recovery corresponded to a 23 % reduction in TPH (Table 4.14). A similar relationship existed between TPH analyses and surrogate recovery for Bottle 2A over the same time period. Considering the difference in surrogate recovery values between the samples, the quantities of TPH reduction observed between days 110 and 124 are likely due to inherent variation in the analysis technique (Table 4.14). Inorganic nutrient analysis of

Bottle 2A (stirring without nutrient addition) at 117.4 days resulted in non-detectable readings for nitrogen and ortho phosphate (sulfate was 200 mg/L). Nevertheless, no other evidence was observed over the 14-day period to support the required addition of nutrients to TB8 groundwater to enhance biodegradation at the biosparge site.

95

No conclusions were made from these experiments because they were conducted over a single, relatively short time interval, and no duplicate samples were available. Subsequent TPH measurements are recommended for the continued investigation of the effects of nutrient addition and stirring on the biodegradation of TPH contaminants in TB8 groundwater. However, further experimentation on this matter is beyond the scope of this thesis.

Table 4.14 TPH Carbon Chain Distribution and Total TPH Concentrations for Bottles (1) and (2A) Comparing the Biodegradation in TB8 Groundwater With and Without Added Nutrients (1) and With and Without Stirring (2A). All Values are Adjusted for Water Loses Due to Evaporation (t = 110 d) (t2 = 89.4 d) (t = 124 d) (t2 = 103.4 d) Bottle (2A) 72% 10 126 516 904 687 502 124 31 ND ND 2900 (t = 138 d) (t2 = 117.4 d) Bottle (1) Bottle (2A) 73% ND 170 792 1420 974 348 ND ND ND ND 3704 67% ND 112 529 892 661 433 72 ND ND ND 2700

Bottle Bottle (1) Bottle Bottle (1) Numbers (2A) Surrogate Recovery C Range C10-C12 C12-C14 C14-C16 C16-C18 C18-C20 C20-C24 C24-C28 C28-C32 C32-C36 C36-C40 Total 9 144 618 1156 1012 788 221 92 ND ND 4040 13 144 648 1167 1004 765 220 87 ND ND 4047 97% 87% 74% 11 134 543 956 753 540 131 27 ND ND 3096

Result (ug/L)

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8000 Bottle (1) 7000 6000 TPH Concentration(ug/L) 5000 4000 3000 2000 1000 0 0 20 40 60 80 Time (days) Figure 4.14: TPH concentrations versus time for all TPH sample events (including those for the investigation of the effects of stirring and nutrient addition on TPH biodegradation. All values are adjusted for evaporation. 100 120 140 160 Bottle (2) Bottle (2A)

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4.4.3 Kinetics of the Long-term Biodegradation of TPH in TB8 Groundwater Zero and first order rate constants were calculated for the observed biodegradation of TPH compounds in experimental sample bottles 1, 2 and 2A. The TPH data collected after nutrient and stirring addition (bottles 1 and 2A, respectively) were not used in the calculation of kinetic constants. The biodegradation observed in bottle 2, over the first 6.6 days, was found to fit first order kinetics with a rate constant of 0.0357 day-1 (R2 = 0.9523) (Figure 4.15). Due to the increase in TPH in bottle 1, between the initial sample and the sample at 1.6 days, only the TPH concentrations from the samples taken at 1.6 days and 6.6 days were used to estimate the initial rate (0 to 6.6 days) of TPH biodegradation in Bottle 1 (Figure 4.15). The first order rate constant for the initial TPH biodegradation in Bottle 1 was 0.0385 day-1 (Figure 4.15). No meaningful r-squared value could be determined for initial Bottle 1 biodegradation because only two points were used for the determination. However, the first order rate constants were similar for both bottles, which adds certainty to these first order rate constants. Zero order kinetics better describes the overall TPH biodegradation observed in the non-amended (no nutrient addition and no stirring) TB8 groundwater. The zero order rate constants for Bottles 1 and 2A over the experimental periods of non-amended biodegradation (124 days for Bottle 1 and 103 days for Bottle 2A) were 19.8 µg/L-day (R2 = 0.87) and 24.0 µg/L-day (R2 = 0.88), respectively (Figure 4.16). The previously reported first order rate constant for the biodegradation of TPH at the TB8 site was 0.0023 day-1 (Nelson, 2003), which is approximately half of the overall rates observed in these laboratory experiments. The overall, nonamended, first order rate constants for Bottles 1 and 2A have slightly smaller R2 values (0.85 and 0.84, respectively) than those for the zero order constants

98

(data not shown). The overall first order rate constants were (0.0042 and 0.0057 day-1 for Bottles 1 and 2, respectively.

99

8.85

Bottle (1)
8.8

Natural Log of TPH Concentration

y = -0.0385x + 8.8391 R2 = 1 (1)

Bottle (2)

8.75

8.7

8.65

y = -0.0357x + 8.8016 R2 = 0.9523 (2)
8.6

8.55 0 1 2 3 4 5 6 7

Time (days)

Figure 4.15: The Natural Log of TPH Concentration vs. Time for experimental bottles (1) and (2) over the first 6.6 days of the Long-term Biodegradability experiment. Linear trendlines for bottles (1) and (2A) showing first order rate constants (day-1) and respective R2 values.

100

8000 7000 6000 TPH Concentration(ug/L) 5000 4000 3000 2000 1000 0 0 20 40 60 Time (days) Figure 4.16: TPH vs. Time for all nonamended (no nutrients or stirring) TB8 groundwater TPH sampling events during the Long-term Biodegradability experiment. Linear trendlines for bottles (1) and (2A) showing zero order rate constants (µg/L-d) and respective R2 values. 80 100 y = -19.768x + 5860.4 R2 = 0.8683 (1) y = -24.008x + 6305 R2 = 0.8835 (2A)

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CHAPTER 5 CONCLUSIONS

In the respirometry experiments, considerable carbon dioxide (CO2) production and concomitant TPH loss occurred in non-amended groundwater samples from the TB8 biosparge site, indicating hydrocarbon degrading microorganisms were present in the TB8 groundwater. Nutrient addition (N, P, K, & S) increased TPH degradation rates only slightly for both inoculated and non-inoculated groundwater (over 28 days of EXP1), but this increase was not statistically significant. Based on the recommended carbon to nutrient mole ratio range of 100: 1: 0.5 to 100: 10: 1 (C: N: P, respectively) necessary to enhance biodegradation, the bioremediation of TPH in TB8 groundwater is not expected to be limited by nitrogen availability. The C: N mole ratio was 100: 17 in the nonamended TB8 groundwater samples. However, ortho phosphate (P) was not detectable in the TB8 groundwater, and this lack of phosphate could significantly limit the biodegradation potential of TPH in TB8 groundwater. Since the phosphate detection limit was 0.5 mg/L, and phosphate concentrations < 0.5 mg/L would be sufficient for enhanced hydrocarbon biodegradation, phosphate should be measured with a more sensitive method. At the end of the long-term experiment, inorganic nutrient analysis of Bottle 2A (without nutrient addition) at 117.4 days resulted in non-detect (ND) readings for both nitrogen and ortho phosphate. Thus, the long-term biodegradation of

hydrocarbons in TB8 groundwater could be limited by a lack of inorganic nutrients.

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Decreasing dissolved oxygen concentrations in the groundwater samples decreased CO2 production. However, the observation of considerable CO2 production in samples with DO concentrations as low as 0.5 mg/L may indicate the concentration of dissolved oxygen need not be high for the biodegradation. DO concentrations less than 0.5 mg/L have been reported in three of six TB8 groundwater-monitoring wells (greater than 3 mg/L for the three remaining wells) (Coffey, 2003). Efforts should be made to increase the level of DO in monitoring wells containing microaerophilic concentrations (DO < 2 mg/L).

The evolution of CO2 in excess of stoichiometric estimates for the respective TPH degradation indicates the CO2 production during the respirometer experiments cannot be accounted for by TPH biodegradation alone. Other sources, possibly competing for dissolved oxygen and contributing to the evolution of CO2 in TB8 groundwater are, the degradation of dissolved soil organics, high groundwater alkalinity and reactions pertaining to chemical oxygen demand (COD). Competition for already low available DO concentrations is likely limiting the bioremediation of hydrocarbons at the TB8 site.

During the long-term biodegradability experiment, the initial biodegradation of TPH compounds was relatively rapid, with first order rate constants of 0.0385 day-1 and 0.0357 day-1 (R2 = 0.9523) over the first 6.6 days of experimentation. The TPH degradation beyond 6.6 days was considerably slower. Zero order rate constants of 19.8 µg/L-day (R2 = 0.87) and 24.0 µg/L-day (R2 = 0.88) were determined for the overall non-amended biodegradation of hydrocarbons in long-term experiments (124 and 103 days,

103

respectively). Although the overall rates were slow, TPH biodegradation was observed throughout the long-term experiment, and no minimum threshold TPH concentration was determined. However, after 124 days, 60 % of the initial TPH was still present.

Therefore, longer-term experiments need to be conducted in the future to confirm diluent contamination can be completely biodegraded.

104

CHAPTER 6 RECOMMENDATIONS

The following are recommendations are included for future evaluation of limitations on the biodegradability of hydrocarbon contaminants at the GRP TB8 biosparge site.
•

The use of a more precise analytical method (with a finer resolution than the 0.5

mg/L) for the presence of ortho phosphate would be beneficial in understanding the true potential phosphate limitation on the bioremediation at the TB8 site.

•

Any future experiments using the measurement of CO2 evolution to represent the

biodegradation of hydrocarbon contaminants in TB8 groundwater should include a set of biologically inactivated groundwater samples. The use of such samples, in addition to a D.I water control, would be useful in the determination of a baseline chemical CO2 production value for TB8 groundwater. Furthermore, the measurement of oxygen

consumption would be beneficial in determining the correlation between respirometry and TPH degradation.

•

A long-term biodegradability experiment with and without the amendment of inorganic nutrients would provide valuable insight on the possible nutrient limitations involved in the bioremediation of TB8 groundwater over the long-term.

A long-term biodegradability experiment should be run for a year to allow for observation of complete biodegradation. Due to relatively high concentrations of sulfate (SO42-), an investigation of sulfate inhibition on the metabolic processes of aerobic hydrocarbon degrading microorganisms is suggested.
•

•

105

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Bioremediation of petroleum pollutants.

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Dragun, J. 1998. The soil chemistry of hazardous materials. Publishers, Amherst MA. GOF. 2003 Guadalupe Oil Field: http://www.guaddunes.com. August 2003. Guadalupe

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Liebeg, Elizabeth Ward and Teresa J. Cutright. 1999. The investigation of enhanced bioremediation through the addition of macro and micro nutrients in a PAH contaminated soil. International Biodeterioration and Biodegradation 44:55-64. Miles, Rick A. and William J. Doucette. 2001. Assessing the aerobic biodegradability of 14 hydrocarbons in two soils using a simple microcosm/respiration method. Chemosphere 45:1085-1090. Marin, Mercedes, Ana Pedregosa, Santiago Rios, M. Luisa Ortiz, and Fernando Laborda. 1995 Degradation of diesel and heating oil by acinetobacter calcoaceticus MM5: It’s applications on bioremediation. International Biodeterioration and Biodegradation 1995:269-285. Mishra, Sanjeet, Jeevan Jyot, Ramesh C. Kuhad, and Banwari Lal. 2001. Evaluation of inoculum addition to stimulate in-situ bioremediation of oily-sludge-contaminated soil. Applied and Environmental Microbiology Apr. 2001:1675-1681. Namkoong, Wan, Eui-Young Hwang, Joon-Seok Park, and Jung-Young Choi. 2001. Bioremediation of diesel-contaminated soil with composting. Environmental Pollution 119:23-31. Ng, Michael S. 2003. Personal communication with the Assistant Laboratory Director (Ng) of ZymaX envirotechnology, Inc. on July 30, 2003. Nelson, Yarrow. 2003. Personal communication, CC. of email from Dr. Yarrow Nelson Associate Professor of Civil and Environmental Engineering, California Polytechnic State University to a Unocal employee (Coffey) on Febuary 5, 2003. Norris, R.D., Hinchee, R.E., Brown, R.A., McCarty, P.L., Semprini, L., Wilson, J.T., Kampbell, D.H., Reinhard, M., Bower, E.J., Borden, R.C., Vogel, T.M., Thomas, J.M., and C.H. Ward. 1994. Handbook of Bioremediation. Boca Raton, FL:CRC Press. Onwurah, Ikechukwu. 1999. Role of diazotrophic bacteria in the bioremediation of crude oil-polluted soil. Journal of Chemical Technology and Biotechnology 74:957-964. Rahman, Kaja Sheik Mujibur, Thahira Rahman, Perumalsamy Lakshmanaperumalsamy, and Ibrahim M. Banat. 2002. Occurrence of crude oil degrading bacteria in gasoline and diesel station soils. Journal of Basic Microbiology 42:284-291. Sharabi, Nel-Din and R. Bartha. 1993 Testing of some assumptions about biodegradability in soil as measured by carbon dioxide evolution. Appl. Envir. Microbiol. 1993 59: 1201-1205. Siddiqui, Samina, and W.A. Adams. 2001. The fate of diesel hydrocarbons and their effect on the germination of perennial rygrass. Inc. Environmental Toxicology 17:49-62.

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