Assessing Anaerobic Biodegradation of
Weathered Petroleum Hydrocarbons Using
Electron Acceptor Amendments
A Master’s Thesis Presented to the Faculty of
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
Meghann Frances Chell
October 2007
AUTHORIZATION FOR REPRODUCTION OF MASTER’S THESIS
I hereby grant permission for the reproduction of this thesis in its entirety or any portion
without further authorization, provided appropriate acknowledgement is made to the
author(s) and advisor(s).
______________________________
Meghann Frances Chell
______________________________
Date
ii
APPROVAL PAGE
TITLE:
ASSESSING ANAEROBIC BIODEGRADATION OF
WEATHERED PETROLEUM HYDROCARBONS USING
ELECTRON ACCEPTOR AMENDMENTS
AUTHOR:
MEGHANN FRANCES CHELL
SUBMITTED:
NOVEMBER 2007
THESIS COMMITTEE MEMBERS:
___________________________________
Dr. Yarrow Nelson
Advisor/Committee Chair
__________________
Date
___________________________________
Dr. Oscar Daza
Committee Member
__________________
Date
___________________________________
Dr. Christopher Kitts
Committee Member
__________________
Date
___________________________________
Dr. Tryg Lundquist
Committee Member
__________________
Date
___________________________________
Dr. Thomas Ruehr
Committee Member
__________________
Date
iii
ABSTRACT
A method was established to examine weathered petroleum hydrocarbon natural
attenuation by anaerobic bacteria in a laboratory setting to determine the potential
contribution of anaerobic biodegradation to in situ remediation. A former oil field located
on the Central California Coast was used as a test site to assess the efficacy of this
bioremediation method. Previous research on hydrocarbon biodegradation at this site
focused on aerobic microbial activity as it contributes to natural attenuation. In the
current study, anaerobic microcosms were established using ground water from the site to
investigate the role of specific anaerobic processes on biodegradation of dissolved
hydrocarbons.
Groundwater was collected from a monitoring well in an anoxic aquifer at the field site.
Microcosms were prepared in custom-made 2-L serum bottles with 100-mL gas
headspaces. Four separate electron acceptors – nitrate, sulfate, manganese(IV) and
iron(III) – were added separately to microcosms to test for their promotion of anaerobic
biodegradation. One set of microcosms utilized a mixture of nitrate, sulfate, and iron(III)
to examine the interaction of bacterial species on biodegradation. A set of unamended
microcosms was run to examine hydrocarbon biodegradation under natural attenuation
conditions. For comparison of biodegradation rates, aerobic microcosms were prepared
and operated side-by-side with the anaerobic microcosms. A set of killed controls was
prepared with 1 % sodium azide to inhibit microbial activity.
Microorganisms were supplied by site groundwater and inoculum from anaerobic soil
collected at the field site. The experiment was conducted inside of a glovebox purged
iv
with nitrogen gas, with testing performed on 80 sacrificial microcosms after 0, 26, 134,
and 407 days of incubation. The total petroleum hydrocarbon (TPH) concentration in
groundwater was determined using gas chromatography. Utilization of electron acceptor
amendments was monitored using the following methods: ion chromatography for
decrease in nitrate and sulfate concentrations, phenanthroline method for increase in
ferrous iron, and formaldoxime method for increase in aqueous manganese. Bacterial
communities were characterized using terminal restriction fragment length polymorphism
(TRFL-P) analysis. Gas headspaces in the microcosms were monitored for methane,
oxygen, helium, nitrogen, carbon dioxide, nitrous oxide, and hydrogen. Helium was
added to the headspace gas to serve as a tracer for gas leaks. Microtox® analysis was
used to determine toxicity changes in anaerobic and aerobic microcosms.
Results show no appreciable evidence of TPH reduction in any of the anaerobic
microcosms after 407 days. Aerobic microcosm TPH concentrations were reduced 53%
at the 26-day sampling period. Lack of microbial activity in anaerobic microcosms was
confirmed by the lack of appreciable change in electron acceptor concentrations.
However, methane was detected in the headspace gases for three microcosm conditions –
unamended, sulfate-amended, and manganese-amended – suggesting possible
methanogenic activity. Nitrous oxide was detected in two microcosms conditions –
nitrate amendment and mixed amendment – suggesting possible denitrification activity.
Results of TRFL-P analysis suggest that microcosms communities were significantly
different from initial conditions after 407 days incubation, that the communities were
most similar amongst the most anaerobic conditions (unamended and sulfate-amended
microcosms), which are also most similar to the field conditions, and that the aerobic and
v
iron-amended microcosms were the least similar. Despite evidence of anaerobic activity
in these microcosms, lack of detectable petroleum hydrocarbon reduction suggests that
anaerobic biodegradation contributions to natural attenuation of the weathered petroleum
hydrocarbons at this field site is negligible in comparison to aerobic biodegradation.
vi
ACKNOWLEDGEMENTS
To Laleh
I can’t imagine working on this with anyone else but you. You’ve changed me for the
better. I miss you already.
To Dr. Nelson
You are an amazing instructor, a considerate advisor, and a good friend. Thank you for
both your praise and criticism. You ask everyone for their best and give them nothing
less than yours.
To My Labmates
Thank you for your friendship, your assistance, your support. Thank you taking part in
all the fun and all the frustration. But most of all, thank you for being considerate and
trying not to wake me when I was sleeping on the floor. EPEL 2006 FOREVER!!!
To Bob Pease, Don Eley, Gonzalo Garcia, Kim Tulledge, Sheldon Nelson and the
Unocal Crowd
Thank you for making this experiment possible and for giving students at Cal Poly, San
Luis Obispo a chance to experience the Guadalupe Dunes.
To Dr. Kitts, Alice Hamrick, Tiffany Glaven
Thanks for being such good teachers, for putting up with all of my questions, and for
revealing the super scary secrets of TRF analysis.
To My Family
Thank you for your love and support. Thank you for calling me repeatedly to make sure I
didn’t fall off the face of the earth. I would not be who I am without you. Let’s not
discuss whether that is a good thing or a bad thing.
To My Wonderful Friends
Thank you for shaping my life at Cal Poly and helping me become the woman I am.
Thank you for helping me make and escape trouble. Thank you for all of the delicious
dinner parties, for the painful hangovers, for all the sweaty dance parties and mellow
movie nights. I can’t begin to express how much I will miss all of you. KIT 4RELZ!!!
To KCPR and EWB
Thank you for reminding me that work and fun exist outside of my laboratory, for putting
me on the airwaves and taking me to Thailand, for changing my life forever.
To Natasha
You are the best cat ever. You’ve seen me through a lot in our 17 years together.
vii
TABLE OF CONTENTS
LIST OF TABLES ........................................................................................................... X
LIST OF FIGURES ...................................................................................................... XII
CHAPTER 1 . INTRODUCTION .................................................................................. 1
CHAPTER 2 . BACKGROUND..................................................................................... 7
2.1 NATURAL ATTENUATION ........................................................................................ 7
2.2 BIOENERGETICS ..................................................................................................... 10
2.2.1 Electron Acceptors for Biological Reactions ................................................. 11
2.2.2 Bioenergetics: Microbial Growth Formulas.................................................. 13
2.2.3 Bioenergtics Calculations .............................................................................. 14
2.3 ANAEROBIC BIODEGRADATION OF PETROLEUM COMPOUNDS ............................... 15
2.3.1 Nitrate Reduction ........................................................................................... 17
2.3.2 Iron Reduction............................................................................................... 18
2.3.3 Manganese Reduction .................................................................................... 20
2.3.4 Sulfate Reduction............................................................................................ 21
2.3.5 Carbon Dioxide Fermentation ....................................................................... 22
2.3.6 Microbial Consortia....................................................................................... 25
2.4 GUADALUPE RESTORATION PROJECT .................................................................... 26
2.4.1 Site History .................................................................................................... 26
2.4.2 Natural Attenuation Studies at the Guadalupe Restoration Project .............. 30
CHAPTER 3 . MATERIALS AND METHODS......................................................... 36
3.1 EXPERIMENTAL DESIGN......................................................................................... 36
3.2 GROUNDWATER COLLECTION ................................................................................ 37
3.2.1 Groundwater Well Selection ......................................................................... 37
3.2.2 Anaerobic Groundwater Collection Barrel.................................................... 39
3.2.3 Groundwater Collection Procedure............................................................... 41
3.3 SOIL COLLECTION .................................................................................................. 43
3.4 MICROCOSMS ........................................................................................................ 45
3.4.1 Microcosm Bottles.......................................................................................... 45
3.4.2 Anaerobic Glovebox....................................................................................... 47
3.4.3 Anaerobic Microcosms Establishment ........................................................... 48
3.4.4 Aerobic Microcosm Establishment................................................................. 54
3.4.5 Adjusting Microcosm pH................................................................................ 55
3.5 GAS HEADSPACE ANALYSES ................................................................................. 57
3.6 TPH ANALYSIS ...................................................................................................... 60
3.5.1 Solvent Extraction (EPA Method 3510C) ...................................................... 60
3.6.2 Concentrating Extract Solution...................................................................... 62
3.6.3 Total Petroleum Hydrocarbon Analysis (EPA Method 8015c)...................... 63
3.7 ELECTRON ACCEPTOR ANALYSES ......................................................................... 70
3.7.1 Sulfate and Nitrate Analysis by Ion Chromatography ................................... 70
3.7.2 Iron Analysis by the Phenanthroline Method (Standard Method 3500-Fe)... 75
3.7.3 Manganese Analysis by the Formaldoxime Method ...................................... 79
viii
3.8 MICROTOX® TOXICITY ANALYSIS ........................................................................ 84
3.9 TERMINAL RESTRICTION FRAGMENT ANALYSIS .................................................... 89
3.9.1 Sample Filtration............................................................................................ 91
3.9.2 DNA Extraction .............................................................................................. 92
3.9.3 PCR Using Fluorescently-labeled Primers.................................................... 93
3.9.4 Production of Labeled Fragments by Enzyme Digestion............................... 97
3.9.5 TRF Pattern Generation by CEQ-8000 ......................................................... 99
3.9.6 TRF Pattern Analysis ..................................................................................... 99
CHAPTER 4. RESULTS AND DISCUSSION.......................................................... 101
4.1 MICROCOSM INTEGRITY ...................................................................................... 101
4.1.1 Redox Indicator Color.................................................................................. 101
4.1.3 Acidic pH in Iron-Amended Microcosms ..................................................... 104
4.1.3 Nitrogen-to-Helium Ratios for Leak Detection............................................ 105
4.2 TOTAL PETROLEUM HYDROCARBON RESULTS .................................................... 108
4.2 HEADSPACE AND AQUEOUS GAS CONCENTRATIONS ........................................... 113
4.2.1 Oxygen Concentration in Microcosm Bottles .............................................. 115
4.2.2 Methane Production..................................................................................... 115
4.2.4 Nitrous Oxide Production ............................................................................ 119
4.2.5 Carbon Dioxide Production ......................................................................... 121
4.4 NITRATE AND SULFATE CONCENTRATIONS (ION CHROMATOGRAPHY RESULTS) 123
4.4.1 Nitrate Concentration in Microcosms.......................................................... 124
4.4.2 Sulfate Concentration in Microcosms .......................................................... 127
4.5 FERROUS IRON CONCENTRATIONS IN MICROCOSMS ............................................ 130
4.6 MANGANESE(II) CONCENTRATIONS .................................................................... 131
4.6 MICROTOX® TOXICITY RESULTS ......................................................................... 134
4.7 TERMINAL RESTRICTION FRAGMENT ANALYSIS RESULTS ................................... 137
4.7.1 16S DNA Digests.......................................................................................... 137
CHAPTER 5. CONCLUSIONS.................................................................................. 147
REFERENCES.............................................................................................................. 152
APPENDIX A: BIOENERGETICS FORMULAS AND CALCULATIONS ........ 161
APPENDIX B: EPA METHOD 3510C...................................................................... 172
APPENDIX C: EPA METHOD 8015C...................................................................... 181
APPENDIX D: IRON ANALYSIS BY PHENANTHROLINE METHOD............ 215
APPENDIX E: MANGANESE(II) ANALYSIS BY THE FORMALDOXIME
METHOD ...................................................................................................................... 223
APPENDIX F: TERMINAL RESTRICTION FRAGMENT ANALYSIS
PROTOCOL.................................................................................................................. 228
ix
LIST OF TABLES
Table 2.1 Terminal Electron Acceptor Half-Reactions and their Gibb's Free Energy in
terms of kilojoules per electron equivalent (kJ/eeq) (Rittmann and McCarty, 2001) ...... 12
Table 2.2 Boiling Point Distribution of Diluent (Lundegard and Garcia, 2001)............ 28
Table 3.1 Microcosm Amendment Concentrations Based on Initial TPH Concentration of
6 ppm and Using a Safety Factor of 3 .............................................................................. 52
Table 3.2 Amendment Source and Form.......................................................................... 53
Table 3.3 Pilot-Scale Test Results for pH Adjustment Using Different NaOH Normalities
........................................................................................................................................... 55
Table 3.4 Henry's Law Constants and Temperature Conversion Factors for Gaseous
Solutes (National Institute of Standards and Technology, 2005) ..................................... 60
Table 3.5 GC Oven Specifications ................................................................................... 64
Table 3.6 GC Operating Conditions................................................................................ 65
Table 3.7 GC Calibration Standard Set for TPH Analysis.............................................. 66
Table 3.8 Diluent Standard Concentrations and GC Output for Calibration Curve ...... 67
Table 3.9 Hexacosane Concentrations in Diluent Standards.......................................... 68
Table 3.10 Hexacosane Concentration and GC Output for Calibration Curve .............. 68
Table 3.11 Calibration Curve Data and Elution Times for Nitrate, Nitrite, and Sulfate as
Monitored by Ion Chromatography .................................................................................. 72
Table 3.12 Fe(II)-Phenanthroline Dilution Series for Calibration Curve ...................... 77
Table 3.13 Fe(II)-Phen Concentrations and Absorbances .............................................. 77
Table 3.14 Mn(II) Dilution Series for Calibration Curve................................................ 81
Table 3.15 Mn(II)-Formaldoxime Absorbance as a Function of Concentration and
Development Time ............................................................................................................ 82
Table 4.1 Microcosm Water Color Due to Redox Indicator........................................... 102
Table 4.2 Average Microcosm pH at Sampling Dates................................................... 104
x
Table 4.3 Nitrogen-to-Helium Ratios for All Microcosms at Sampling Each Date ...... 106
Table 4.4 TPH Concentrations in Microcosm Replicates ............................................. 109
Table 4.5 Average TPH Concentrations and Standard Deviations............................... 110
Table 4.6 Carbon Dioxide and Oxygen Concentrations in All Microcosms and Replicates
at Sampling Dates ........................................................................................................... 113
Table 4.7 Methane, Hydrogen and Nitrous Oxide Concentrations in All Microcosm
Replicates at Sampling Dates ......................................................................................... 114
Table 4.8 Methane Generation in Gas Headspace and Calculation of Amount of
Headspace and Dissolved Methane in Unamended, Sulfate, and Manganese(IV)
Microcosms ..................................................................................................................... 116
Table 4.9 Bioenergetic Stoichiometry for Hexane Consumed Due to Methane Production
......................................................................................................................................... 118
Table 4.10 Nitrous Oxide Production Observed in Gas Headspace and Calculated in
Aqueous Solution in Nitrate and Mixed Amendment Microcosms.................................. 120
Table 4.11 Bioenergetics Stoichiometry for Hexane Consumed Calculated from Nitrous
Oxide Production ............................................................................................................ 121
Table 4.15 Nitrate Concentrations in All Microcosms and Replicates ......................... 125
Table 4.13 Nitrate Consumed Based on Nitrous Oxide Produced and Bioenergetic Molar
Ratios .............................................................................................................................. 127
Table 4.14 Sulfate Concentrations in All Microcosm Replicates .................................. 128
Table 4.15 Iron(II) Concentration in Iron-Amended and Unamended Microcosms ..... 130
Table 4.16 Manganese(II) Concentration in Manganese and Unamended Microcosms
......................................................................................................................................... 132
Table 4.17 Hexane Consumption Based on Bioenergetics Calculations and
Manganese(II) Concentration, Corrected to Exclude Manganese(II) Present at Day 0
Sampling Event ............................................................................................................... 134
Table 4.18 Percent Effect of Microcosm Samples on Bioluminescent Bacteria,
Calculated Using Microtox Omni Software.................................................................... 135
Table 4.19 Effective Concentration of Microcosm Sample that caused 50% Reduction in
Bacterial Bioluminescence, Determined Using Microtox Omni Software ..................... 136
CHAPTER 1
xi
LIST OF FIGURES
Figure 2.1 Processes Involved in the Natural Attenuation of Petroleum Hydrocarbons
Released into the Environment (EPA, 1999) ...................................................................... 8
Figure 2.2 Electron Donor Chemical Energy Partitioning into Energy Production and
Cell Synthesis (Rittmann and McCarty, 2001) ................................................................. 11
Figure 2.3 Terminal Electron Acceptor Utilization and Plume Dispersion (USGS 2005)
........................................................................................................................................... 13
Figure 2.4 Methanogenic Metabolism (Madigan, 2006)................................................. 24
Figure 2.5 Location of Guadalupe Restoration Project Site on the Central California
Coast (Diaz 2006) ............................................................................................................. 27
Figure 2.6 Diluent Source Zones and Groundwater Monitoring Wells at the Guadalupe
Restoration Project Site .................................................................................................... 31
Figure 3.1 Location of Monitoring Well J8-11 at the GRP Site, Source of Anaerobic
Groundwater ..................................................................................................................... 38
Figure 3.2 Grundfos Pumphead with Dissolved Oxygen Meter and Flow Meter ........... 39
Figure 3.3 Collection Barrel Design ............................................................................... 41
Figure 3.4 Collection Barrel Lid with Fill Line, Purge Line, and Air Release Valve..... 42
Figure 3.5 Drill Rig and Soil Collection at H-2 .............................................................. 44
Figure 3.6 Location of Well Pad H2 at the GRP site, Location of Anaerobic Soil
Collection.......................................................................................................................... 45
Figure 3.7 Microcosm Bottle Design............................................................................... 46
Figure 3.8 Anaerobic Glovebox....................................................................................... 49
Figure 3.9 Laleh and Meghann Establishing Anaerobic Microcosms Inside the N2/He
Purged Glovebox .............................................................................................................. 50
Figure 3.10 Greg Ouellette Conducting Gas Headspace Analysis ................................. 58
Figure 3.11 Glassware Set-Up in Chemical Fume Hood for TPH Extraction................ 61
Figure 3.12 Hewlett Packard 6890 GC/FID with Auto Sampler......Error! Bookmark not
defined.
xii
Figure 3.13 TPH Calibration Curve................................................................................ 67
Figure 3.14 Hexacosane Standard Curve........................................................................ 69
Figure 3.15 GC Output for 834 ppm Diluent Standard. Large Peak at 22.5 Minutes is
Hexacosane. ...................................................................................................................... 69
Figure 3.16 Dionex Ion Chromatogram with Auto Sampler ............Error! Bookmark not
defined.
Figure 3.17 7-Anion Standard Output from DX-190 Ion Chromatogram....................... 71
Figure 3.18 Ion Chromatograph for 50 ppm Nitrate and Sulfate Standard.................... 73
Figure 3.19 Calibration Curve for Nitrate, Nitrite, and Sulfate, 1 - 20 ppm Range ....... 74
Figure 3.20 Calibration Curve for Nitrate, Nitrite and Sulfate, 20 - 200 ppm Range .... 74
Figure 3.21 Hitachi U-3010 UV/VIS Spectrophotometer...Error! Bookmark not defined.
Figure 3.22 Fe(II)-Phen Calibration Curve .................................................................... 78
Figure 3.23 Change in Mn(II)-Formaldoxime Development with Time.......................... 82
Figure 3.24 Mn(II)-Formaldoxime Absorbances as a Function of Concentration,
Measured at Four Development Times............................................................................. 83
Figure 3.25 TRF Pattern Gernerated from Initial Groundwater Sample Collected from
the GRP Site at J8-11........................................................................................................ 90
Figure 3.26 Basics of Creating TRF patterns – DNA Labeling, Enzyme Digestion, and
Fragment Analysis ............................................................................................................ 91
Figure 3.27 GeneAmp Thermal Cycler Used for PCR and Enzyme Digestion ........ Error!
Bookmark not defined.
Figure 3.28 Polymerase Chain Reaction Stages - Denaturation, Annealing, and
Elongation......................................................................................................................... 95
Figure 4.1 Microcosm Color Spectrum at Day 26 Sampling Event. From Left to Right:
Mixed Amendment, Iron, Manganese, and Unamended Microcosms. ........................... 103
Figure 4.2 GC/FID Generated Chromatogram for Unamended Microcosm at the 407th
Day Sampling Event........................................................................................................ 108
Figure 4.3 Change in TPH Concentrations in Groundwater Microcosms During 407Day Incubation................................................................................................................ 111
xiii
Figure 4.4 Comparative TPH Concentrations in All Groundwater Microcosms,
Arranged by Microcosm Condition ................................................................................ 112
Figure 4.5 Methane Molarity in Microcosm Headspace Gas ....................................... 117
Figure 4.6 Gaseous Nitrous Oxide Concentration in Nitrate and Mixed Microcosms . 119
Figure 4.7 Gaseous Carbon Dioxide Molarity in Microcosm Headspace, Corrected to
Remove Outliers in Iron-Amended Microcosms ............................................................. 122
Figure 4.8 Change in Average Nitrate Concentration in Microcosms.......................... 126
Figure 4.9 Change in Average Sulfate Concentration in Groundwater Microcosms ... 129
Figure 4.10 Change in Ferrous Iron Concentration in Iron-Amended and Unamended
Microcosms ..................................................................................................................... 131
Figure 4.11 Change in Manganese(II) Concentration in Manganese and Unamended
Microcosms ..................................................................................................................... 132
Figure 4.12 % Effect of Microcosm Sample on Bacterial Bioluminescence, Microcosm
Comparison..................................................................................................................... 135
Figure 4.13 Concentration of Microcosm Sample Required to Reduce Bacterial
Bioluminescence by 50%, Microcosm Comparison........................................................ 136
Figure 4.14 Electropherograms and Dendrogram Produced from Bray-Curtis
Similarity for 16S DNA Fragments Produced by Dpn III Restriction Enzyme............... 140
Figure 4.15 Electropherograms and Dendrogram Produced from Bray-Curtis Similarity
for 16S DNA Fragments Produced by Hae III Restriction Enzyme................................ 141
Figure 4.16 Electropherograms and Dendrogram Produced from Bray-Curtis Similarity
for 16S DNA Fragments Produced by Hha I Restriction Enzyme .................................. 142
Figure 4.17 Electropherograms and Dendrogram Produced from Bray-Curtis
Similarity for Methanogen DNA Fragments Produced by Sau I96 Restriction Enzyme 145
Figure 4.18 Electropherograms and Dendrogram Produced from Bray-Curtis
Similarity for Archaea DNA Fragments Produced by Hae III Restriction Enzyme ....... 146
xiv
CHAPTER 2 . INTRODUCTION
Approximately half of the drinking water used in the United States is comprised of
groundwater, comprising 95 % of the freshwater on the continent (Kota et al, 2004). Due
to poor waste management and storage practices, much of the nation’s freshwater has
been contaminated or is in danger of contamination by xenobiotic chemicals. Petroleum
hydrocarbons and oxygenates are common groundwater contaminants due to leaking
underground storage tanks and accidental spills, with several hundred petroleum
contaminated sites in the continental United States (Boopathy, 1994, Kota et al, 2004).
Remediation processes are necessary to preserve the natural environment and to ensure
that drinking water is not compromised by anthropogenic pollution.
Natural attenuation is a passive remediation technique encompassing chemical and
physical processes that remove contaminants from soil and groundwater ecosystems (Kao
and Wang, 2000). The method is particularly valuable in relationship to groundwater
remediation, as it does not disturb fragile aquifer sediments and does not introduce
foreign chemicals into the groundwater (Kota et al, 2004). Of these processes, aerobic
and anaerobic bioremediation are the largest contributors to hydrocarbon natural
attenuation (Curtis and Lammey, 1998). Both processes consist of contaminant
degradation by native microorganisms, oxidizing contaminants while consuming
available electron acceptors. Aerobic processes utilize oxygen as the terminal electron
acceptor (TEA), while anaerobic processes utilize a variety of TEA, depending on the
specific contaminant and microorganism, including ferric iron, sulfate, nitrate,
manganese oxide, and carbon dioxide (Heldrich et al, 2004).
Anaerobic and aerobic bioremediation processes can occur simultaneously or in tandem
as the plume migrates, as dictated by environmental conditions and microbial competition
(Cho et al, 1997). Until recently, anaerobic processes were thought to contribute
negligibly to hydrocarbon biodegradation and were often discounted (Townsend et al,
2003), but recent research has brought to light the role anaerobic processes play in natural
attenuation, specifically in saturated and aquatic systems where oxygen supply is limited.
In these systems, anaerobic processes dominate once oxygen is consumed, with TEAs
utilized in the order of metabolic favorability (Spormann and Widdel, 2000). The order of
favorability generally follows from the free energy available from the electron acceptors.
This is seen in sites with long-term contamination; in these sites, dissolved oxygen is
depleted and areas of highly reduced sediment appear as anaerobic processes
predominate (Anderson and Lovley, 2000a). Numerous petroleum hydrocarbons are
susceptible to biodegradation in anaerobic environments, including benzene, toluene,
ethyl benzene and xylene (BTEX) (Burland and Edwards, 1998, Kao and Wang, 2000,
Cho et al, 1997, Lovley et al, 1994), methyl tert-butyl ether (MTBE) (Bradley et al, 2001,
Finneran and Lovley, 2001), n-alkanes (Chayabutra and Ju, 2000), Number 2 diesel fuel
(Boopathy, 1994), polycyclic aromatic hydrocarbons (PAH) (Schmitt et al, 1996), and
crude oil (Bekins et al, 2001).
2
In this study, anaerobic biodegradation of weathered petroleum hydrocarbons was
investigated using groundwater collected from the former Guadalupe Oil Field, a 2,700acre coastal dune ecosystem approximately 30 miles south of San Luis Obispo, CA. Due
to the high viscosity of oil produced at the site, a thinning agent referred to as “diluent”
was historically added to the crude oil in the wells to facilitate pumping (Lundegard and
Garcia, 2001). Diluent consisted of mid-range petroleum distillate from nearby refineries
and is similar to diesel fuel in equivalent carbon chain length. The diluent was stored on
the site in underground tanks and distributed to oil wells through a large network of
pipes. During its usage, both diluent and diluent-crude oil mixtures contaminated the site
through leaky storage tanks and corroded pipelines, impacting surface and groundwater
quality. Approximately 9 million gallons of hydrocarbons were leaked, creating 90
documented contaminant plumes (Lundegard and Garcia, 2001).
Previous research on weathered petroleum contaminants is limited, as most research has
focused on the fate and transport of petroleum-contaminants such as MTBE and BTEX.
Loehr and Webster (1996) showed that hydrocarbon bioavailability decreased with aging
and weathering, attributed to increased contact between the soil and the contaminants.
Since the nature of weathering is site specific, the bioavailability of weathered petroleum
contaminants will vary with site conditions. Loehr et al (2001) found that, among nine
soils from different petroleum and related industry sites, three had high potential, four
had low potential, and one had very low potential for further biodegradation. All three
with high potential were from diesel oil contaminated sites, analogous to the
contamination at the Guadalupe Restoration Project (GRP). Additionally, many studies
3
on anaerobic biodegradation have been performed using pure bacterial cultures or
enrichment cultures (Chayabutra and Ju, 2000), with evidence of weathered hydrocarbon
biodegradation in environmental samples reported in few cases (Salminen et al, 2004).
For environmental samples taken from weathered hydrocarbon sites, well-adapted
microbial consortia are more effective at degrading hydrocarbons than are those obtained
from recently contaminated soils (Trindade et al, 2005).
The fragile nature of the dune ecosystem at GPR and the numerous state and national
endangered species it hosts precludes the use of invasive remediation techniques. Less
disruptive techniques, including biosparging, phytoremediation, and natural attenuation
studies are currently underway at the Guadalupe site. Despite numerous natural
attenuation studies, anaerobic processes are poorly quantified at the former Guadalupe oil
field and other petroleum-contaminated sites. Though the kinetics of anaerobic processes
are less favorable than aerobic, oxygen is often limited in groundwater environments,
proving insufficient for full remediation of a contaminant spill. Due to the low solubility
of oxygen in water, oxygen concentrations in groundwater are very low, with a maximum
concentration of 9 mg/L at ambient temperature and pressure. As oxygen is consumed in
a contaminant plume, it can only be recharged at the water-air interface at the
groundwater surface, limiting the flux of oxygen to the plume fringe. Anaerobic
processes, using electron acceptors such as nitrate, sulfate, manganese and iron, and
methanogenesis by a microbial consortium, may contribute more to diluent degradation
than previously considered due to their presence in groundwater at concentrations greater
than oxygen.
4
For this experiment, microcosms were established with anaerobic conditions using
anaerobic groundwater collected from an aquifer at the Guadalupe site and using
anaerobic soil from the Guadalupe site as microbial inoculum. Eight distinct conditions
were established in the microcosms: iron reducing, manganese reducing, denitrification,
sulfate reducing, mixed-amendment (nitrate, sulfate, and iron), unamended
(methanogenic), and aerobic microcosms as well as azide-killed controls. Four
samplings were conducted after 0, 26, 134, and 407 days of incubation. At each
sampling date, duplicate microcosms were sacrificed for analysis. The 26-day sampling
date was chosen to provide important aerobic data for comparison with anaerobic data as
previous research at this site indicated over 50 % reduction in total petroleum
hydrocarbons (TPH) after approximately 20-days incubation under aerobic conditions
(Dreyer, 2004, Lassen, 2005, Waudby, 2003). Microcosms were sacrificial to prevent
loss of gases due to breaching Teflon-lined septa at sampling dates. At each sampling
date, microcosm water was analyzed for changes in total petroleum hydrocarbons (TPH)
and TEAs, and headspace gas was monitored for headspace gas composition. One
additional sampling date remains, tentatively scheduled to take place after 800-days of
incubation.
This project was designed as a joint research project between Meghann Chell and Laleh
Rastegarzadeh. The TPH and gas headspace analyses were conducted together, Laleh
Rastegarzadeh conducted ion chromatography analyses, and Meghann Chell conducted
iron and manganese analyses. For completion of her Masters of Science degree in
5
Environmental Engineering, Laleh Rastegarzadeh opted to conduct an additional study
for her thesis.
6
CHAPTER 3 . BACKGROUND
2.1
Natural Attenuation
Natural attenuation is a passive process involving physical and chemical changes
occurring over time, reducing the mass, toxicity and mobility of contaminant chemicals
(EPA, 2006). A non-invasive process, natural attenuation has gained considerable interest
and acceptance as a viable method for remediating petroleum contaminated sites (Wang
and Fingas, 1998). Though often negatively regarded as a “sit back and watch”
technique, natural attenuation methods require extensive monitoring, strategic planning
and management, site characterization, and data analysis (Cho et al, 1997). Additionally,
monitoring of compounds associated with microbial activity, such as oxygen, nitrate,
sulfate, carbon dioxide, and ferric iron is necessary to determine which microbial
processes are dominant at a particular site (EPA, 1999).
Five processes dominate natural attenuation, as depicted in Figure 2.1: biodegradation,
sorption, dispersion/dilution, volatilization, and chemical reactions (EPA, 1999). Of
these five processes, the most environmentally significant is biodegradation (Chapelle,
1999), encompassing all of the changes in chemical contaminants performed by
microorganisms, such as bacteria and fungi. Though biodegradation processes may
produce harmful end products depending on the contaminants at the site, numerous
studies indicate petroleum hydrocarbons appear to be less toxic after biodegradation in
almost all cases (EPA, 1999).
7
Figure 3.1 Processes Involved in the Natural Attenuation of Petroleum
Hydrocarbons Released into the Environment (EPA, 1999)
One of the central concerns regarding the effectiveness of biodegradation is the length of
time required by the method, especially in comparison with other more invasive
microbial techniques (Bento et al, 2004). Other bioremediation techniques of interest
include biostimulation – nutrient addition to a contaminated site to boost degradation
rates – and bioaugmentation – addition of nutrients and microorganisms known to
degrade the contaminant of concern to the contaminated area (Bento et al, 2004). In a
study comparing biodegradation to other bioremediation methods, Bento et al (2004)
found natural attenuation out-performed biostimulated and bioaugmented soil columns
containing diesel-contaminated sediments, despite increased dehydrogenase activity in
the bioaugmented columns. This observation was attributed to indigenous organisms
being adapted to their surroundings. Coates et al (1996) suggests that indigenous
8
microorganisms can out-compete introduced species despite bioaugmentation,
demonstrated by native sulfate-reducing microorganisms’ ability to out-compete
introduced iron-reducing bacteria despite stimulation by Fe(III) oxides. Simoni et al
(2001) cite that degradation rates are more controlled by nutrient supply to
microorganisms rather than microbial degradation capacity; nutrient supply is heavily
influenced by soil grain size and microbial biomass distribution. Additionally,
microorganisms are concentration sensitive, making them incapable of degrading very
low concentrations, which cannot support sufficient biomass to promote microbial
growth, or high concentrations that are too toxic to microbial cells (EPA, 1999).
Biodegradation by bacteria is divided into two types: aerobic and anaerobic. Aerobic
biodegradation is the destruction of contaminants in the presence of oxygen, whereas
anaerobic biodegradation occurs in environments devoid of oxygen. During preliminary
bioremediation studies, only aerobic organisms were widely regarded as capable of
degrading environmental contaminants (Cookson, 1995). Bailey et al (1973) reported
that molecular oxygen was necessary for the remediation of petroleum hydrocarbons.
Cookson (1995) stated absolutely that “oxygen is required” for bioremediation of
aliphatic petroleum hydrocarbons since anaerobic biodegradation is too uncertain and ill
defined, making aerobic processes necessary. During the time elapsed since these earlier
suppositions, extensive evidence has been offered in recent years supporting the
remediation of petroleum contaminants under a variety of anaerobic conditions. Lovley
(1997) established the importance of anaerobic microorganisms in biodegradation of the
aromatic petroleum compounds benzene, toluene, ethyl benzene and xylene (BTEX).
9
Aliphatic petroleum hydrocarbons are known to degrade by way of numerous anaerobic
processes, including iron reduction, denitrification, sulfate reduction, and
methanogenesis, though the specific mechanisms are poorly understood (Salminen et al,
2004).
In petroleum-contaminated environments, there is generally no limit of carbon source or
electron donor; therefore, biodegradation is generally limited by the availability of
terminal electron acceptors (TEAs). The thermodynamic favorability of potential TEAs,
in order from most to least energetically favorable, is oxygen > nitrate > ferric iron >
manganese(IV) > sulfate > carbon dioxide (Chapelle 1999).
2.2
Bioenergetics
Bioenergetics is a thermodynamic approach to studying biologically mediated redox
reactions incorporating the conversion of carbon and energy sources to cell mass and
cellular energy (Rittmann and McCarty, 2001). Bioenergetics uses the free-energy of
respiration and synthesis reactions to predict the theoretical stoichiometry of microbial
growth requirements, yields, and waste products. The end result is a balanced
stoichiometric reaction relating substrate utilization to electron acceptor concentrations
and biomass production.
One of the fundamental bases of bioenergetics is the partitioning of substrates into the
chemical energy harnessed during respiration and for usage in building cell mass and
developing cellular energy. Figure 2.2 depicts the partitioning of chemical energy
10
occurring during cellular respiration. The partitioning of energy into these two processes
is calculated based on energy created during cellular respiration – more efficient
metabolism permits more energy dedicated to cell growth rather than cellular energy.
When calculating carbon partitioning, fe represents the fraction of carbon utilized for
cellular energy and fs the fraction for cell synthesis (Rittmann and McCarty, 2001).
Energy fractions are calculated in terms of electron equivalents, the same as redox
balancing, because electron flow is the basis for cellular energy
Figure 3.2 Electron Donor Chemical Energy Partitioning into Energy Production
and Cell Synthesis (Rittmann and McCarty, 2001)
2.2.1 Electron Acceptors for Biological Reactions
Contaminant biodegradation in the environment can be described in terms of redox
reactions. Typical electron acceptors in soil and groundwater systems include oxygen,
nitrate, ferric iron [Fe(III)], manganese oxide, sulfate, and carbon dioxide. Half reactions
and reduction potential for these electron acceptors are listed in Table 2.1, in order of
11
decreasing reduction potential. The more energy yielded by the total reaction, the more
energy that can be used to create biomass. Thus, organisms that are capable of using
oxygen as a terminal electron acceptor are capable of out-competing organisms that are
not able to use them. Once oxygen is consumed, microorganisms utilizing less favorable
TEAs are able to grow. This cycle of TEA thermodynamic favorability, referred to as the
Electron Tower Theory, predicts how electron acceptors will be consumed in a
contaminant plume, as depicted in Figure 2.3, where consumption of terminal electron
acceptors proceeds in order of reduction potential as the contaminant plume is dispersed
in the water table, as depicted in Figure 2.3 (USGS, 2005).
Table 3.1 Terminal Electron Acceptor Half-Reactions and their Gibb's Free Energy
in terms of kilojoules per electron equivalent (kJ/eeq) (Rittmann and McCarty,
2001)
Reduced
TEA
TEA Half-Reaction
Species
O2
¼ O2 + H+ + e½ H2O
Oxygen
Fe(III)
Fe3+ + eFe2+
Iron
NO31/5 NO3- + 6/5 H+ + e1/10 N2 + 3/5 H2O
Nitrogen
Mn(IV)
½ MnO2 + e- + 2H+
½ Mn2+ + H2O
Manganese
SO42- 1/8 SO42- + 19/16 H+ + e1/16 H2S + 1/16 HS- + ½ H2O
Sulfate
+
CO2
1/8 CO2 + H + e
1/8 CH4 + ¼ H2O
Methanogenesis
∆G
(kJ/eeq)
-78.72
-74.39
-72.20
-59.04
20.85
23.53
Though the half reactions for sulfate reduction and methanogenesis are positive,
indicating that these reactions are not thermodynamically favorable, they are able to
proceed whenever the electron donor supplies sufficient energy for the total reaction to be
energetically favorable. As illustrated in Figure 2.3, TEA utilization in hydrocarbon
plumes typically follows Gibbs free energy with the more thermodynamically favorable
reactions first utilized (Curtis and Lammey, 1998).
12
Figure 3.3 Terminal Electron Acceptor Utilization and Plume Dispersion (USGS
2005)
2.2.2 Bioenergetics: Microbial Growth Formulas
Once free energy of respiration is determined, the next step in bioenergetics is to link
substrate utilization to electron acceptor utilization and cell synthesis. Half-reactions for
all three steps are required in order to create a representative equation for microbial
growth. Once half-reactions for substrate (electron donors) and electron acceptors are
determined (as listed in Section 2.2.1), they are combined with half-reactions for cell
synthesis. Synthesis reactions are based upon the nitrogen source utilized by the
microorganism, because nitrogen is required for cell growth, protein and nucleic acid
synthesis.
When the fe and fs are calculated, the impact of reducing potential becomes apparent. For
example, using acetate as electron donor and ammonium as nitrogen source, aerobic
organisms would have fs = 0.59, whereas methanogenic organisms (carbon dioxide as
13
electron acceptor) would have fs = 0.05. Capable of dedicating 11.8 times more energy to
cell synthesis, aerobic organisms would dominate in this environment so long as
conditions were suitable for their growth.
2.2.3 Bioenergetics Calculations
Reaction stoichiometry for nitrate, sulfate, ferric iron, manganese oxide, oxygen and
carbon dioxide reduction were developed using bioenergetics and are listed below. The
bioenergetics equations used and calculations specific to reducing conditions of interest
in this experiment are included in Appendix A. For the purposes of these calculations,
hexane (C6H14) was used as electron donor to represent petroleum compounds and
ammonium was used as the nitrogen source for all synthesis reactions. ∆Gpc, ∆Gr, and
∆Ga values were taken from Rittmann and McCarty (2001) for all TEAs except
manganese, which was calculated using the mathematical relationship between reduction
potential and Gibb’s Free Energy as described in Rittmann and McCarty (2001). Using
the balanced equations, stoichiometric ratios are calculated to correlate changes in
changes in substrate to changes in electron acceptor or end product concentrations.
Manganese Reduction
0.0263 C6 H14 + 0.222 MnO2 + 0.444 H + + 0.278 HCO3− + 0.0278 NH 4+
→ 0.378 H 2O + 0.0278 C5 H 7O2 N + 0.222 Mn 2+ + 0.0467 CO2
Iron Reduction
0.0263 C6 H14 + 0.405 Fe 3+ + 0.378 H 2O + 0.298 HCO3− + 0.0298 NH 4+
→ 0.0298 C5 H 7O2 N + 0.405 Fe 2+ + 0.0389 CO2 + 0.405 H +
14
Sulfate Reduction
0.0263 C6 H14 + 0.1125 SO42− + 0.344 H + + 0.0005 NH 4+ + 0.0005 HCO3−
→ 0.0005 C5 H 7O2 N + 0.0563 H 2 S + 0.0563 HS − + 0.138 CO2 + 0.179 H 2O
Nitrate Reduction
0.0263 C6 H14 + 0.0939 NO3− + 0.0939 H +
→ +0.0230 C5 H 7O2 N + 0.0354 N 2 + 0.0525 CO2 + 0.150 H 2O
Carbon Dioxide Reduction
0.0263 C6 H14 + 0.00034 NH 4+ + 0.00034 HCO3− + 0.0522 H 2O
→ 0.00034 C5 H 7O2 N + 0.0563 H 2 S + 0.117 CH 4 + 0.0278 CO2
Oxygen Reduction
0.0263 C6 H14 + 0.0988 O2 + 0.0303 NH 4+ + 0.0303 HCO3−
→ 0.0303 C5 H 7O2 N + 0.0369 CO2 + 0.154 H 2O
Though nitrogen gas is reported as the end product for nitrification, Zeng et al (2003)
reported that the majority of gas produced as a byproduct of nitrate reduction was nitrous
oxide, not nitrogen gas. Based on this conclusion, changes in nitrous oxide concentration
were attributed to nitrate reduction using the same molar ratios developed for nitrogen
gas as the end product. Additionally, nitrate was used as nitrogen source for nitrate
reducing bacteria (rather than ammonium, which was used for all other bioenergetics
calculations), as studies have shown nitrate reductase activity is reduced when
ammonium is used as the nitrogen source for denitrifying bacteria (Morris and Syrett,
1963).
2.3
Anaerobic Biodegradation of Petroleum Compounds
Originally thought to contribute marginally to overall biodegradation (Cookson 1995),
anaerobic biodegradation mechanisms have been gaining more attention in recent years
15
due to increased information regarding contaminant site conditions and rapid oxygen
depletion (Burland and Edwards, 1999). In Section 2.2, the energetic favorability of
aerobic biodegradation over that of anaerobic biodegradation was shown mathematically
using bioenergetic calculations, as oxygen has a greater reduction potential than do other
terminal electron acceptors. Despite this reasoning, numerous reasons exist for
considering the contribution of anaerobic microorganisms to biodegradation. In saturated
groundwater systems, microorganisms rapidly deplete oxygen when a contaminant
present is (Bregnard et al, 1995). Though aerobic processes can continue to dominate at
the air-groundwater interface, anoxic conditions dominate in most contaminant plumes
(Lovley et al, 1989). Additionally, oxygen is sparingly soluble in groundwater, with an
optimistic maximum at approximately 9 mg/L at typical ambient temperature and
pressure. Limited oxygen solubility in water becomes problematic when considering the
large oxygen demand required in heavily contaminated systems (Kazumi et al, 1997) due
to the large stoichiometric amount of oxygen necessary to mineralize the contaminants
completely.
Anaerobic biodegradation follows different biochemical pathways dependent on the
electron acceptor utilized by the microorganism. Petroleum-based contaminants have
been shown to degrade under various anaerobic conditions, including nitrate reduction,
sulfate reduction, ferric iron reduction, manganese reduction and methanogenic
conditions.
16
2.3.1 Nitrate Reduction
Nitrate reduction, more commonly referred to as denitrification, is a common
phenomenon in saturated soil systems (Payne, 1981). Many of the denitrifying organisms
are facultative anaerobes, capable of using alternate electron acceptors when oxygen is
not available (Knowles, 1982). As a fertilizer, a component of secondary-treated
wastewater, and a septic system effluent, nitrate is ubiquitous in shallow groundwater
(Knowles, 1982). The availability, high solubility in aqueous systems, and large reducing
potential make nitrate an ideal terminal electron acceptor in contaminated systems.
Studies using nitrate as a terminal electron acceptor document its success in aiding
biodegradation of numerous petroleum-based contaminants. Burland et al (1998)
documented success using nitrate as the primary electron acceptor in benzene
biodegradation in enriched soil and groundwater microcosms. Bradley et al (2001)
demonstrated greater than 43 % reduction of methyl tert-butyl ether (MTBE) by
denitrification in sediment microcosms. Chayabutra et al (2000) demonstrated 40 %
reduction of n-hexadecane by a pure culture of Pseudomonas aeruginosa under
denitrifying conditions following an initial oxic period. Boopathy (1994) reported 47 %
reduction in No. 2 diesel fuel from contaminated soils using nitrate amendment and
native microorganisms.
Concerns associated with wide scale application of denitrification are numerous. From
an analytical perspective, in situ application of nitrate as an electron acceptor is difficult
to monitor due to dilution effects and other hydrogeologic phenomena from biological
17
utilization (Chapelle, 1999). This is worsened by uncertain stoichiometry; since nitrate
can act as a nitrogen source for many microorganisms, it may be difficult to limit its
usage to biodegradation, yielding a higher ratio of electron acceptor to electron donor
consumption than can be accounted for by contaminant reduction (Hutchins et al, 1991).
Many denitrifying organisms are sensitive to their own by-product – nitrite – and are
unable to reduce it further. Under strict anaerobic conditions, nitrite completely stopped
biodegradation at concentrations as low as 0.1 g/L NO2- – N (Chayaburta et al, 2000).
Biodegradation of benzene was tied more to reduction of nitrate to nitrite rather than
complete reduction to nitrogen gas (Burland 1999), thus trading one toxic groundwater
contaminant for another. Zeng et al (2003) demonstrated that denitrification typically
results in the formation of nitrous oxide rather than nitrogen gas. Nitrous oxide is a
greenhouse gas with approximately 296 times more impact on global warming than
carbon dioxide (Albritton, 2001). Nitrate addition to groundwater is not considered
acceptable by water governance agencies due to toxicity disruption of oxygen utilization
in infants at concentrations exceeding 10 mg/L NO3- - N, termed methanoglobanemia
(Tchobanoglous and Schroeder, 1985).
2.3.2 Iron Reduction
Ferric iron reduction has been called “the most important chemical change that takes
place in the development of anaerobic soils and sediments” (Ponnamperuma 1972)
because it is responsible for the regulation of many soil and groundwater systems,
including the oxidation of organic matter and the distribution of phosphate and trace
metals (Lovley 1991). Unlike nitrate, an ion dissolved in solution, Fe(III) reacts with
18
water to form amorphous ferric hydroxide solids [Fe(OH)3 or FeOOH], which do not
move with groundwater flow and therefore are not subject to hydrogeologic influences
(Coates et al, 1996). Ferric iron reducing microorganisms were the first anaerobic
bacteria identified capable of degrading petroleum contaminants in laboratory studies
(Lovley and Phillips, 1988, Chapelle, 1999), providing the basis for future research
focusing on anoxic systems.
Iron-reducing bacteria have been linked to the successful biodegradation of numerous
petroleum-based contaminants. Finneran and Lovley (2001) demonstrated that ferric iron
utilizing microorganisms were capable of complete mineralization of MTBE, often
thought to be recalcitrant in anaerobic systems. This mineralization was completed
without the formation of tert-butyl alcohol (TBA), a toxic byproduct of incomplete
MTBE degradation. Kao and Wang (2000) demonstrated 93.1 % reduction of BTEX
within the iron-reducing zone of a gasoline-contaminated aquifer during an in situ
bioremediation study. Coates et al (1996) postulated that iron-reducing bacteria are more
effective at degrading certain contaminants than sulfate-reducing bacteria, as they
demonstrated that iron-reduction became the predominant method of biodegradation
when iron was added to contaminated sediments with sulfate-reducing conditions. Based
on computer simulations, Bekins et al (2001) estimated that sufficient ferric iron oxides
existed in crude oil contaminated sediments to allow for iron-reduction to continue for 10
– 15 years in areas of low hydraulic conductivity that support iron-reduction conditions.
19
As shown in the bioenergetics calculation, the ratio of iron required for hydrocarbon
degradation is very high. Curtis and Lammey (1998) calculated a highly unfavorable
molar ratio of 36:1 electron acceptor to contaminant for toluene biodegradation via iron
reduction. Unlike dissolved electron acceptors (NO3-, SO42-, and O2) that are subject to
lateral or areal groundwater recharge, elemental electron acceptors (Fe(III), Mn(IV)) can
be depleted due to slow recharge dependent on mineral weathering (McMahon and
Bruce, 1996).
2.3.3 Manganese Reduction
Though not extensively studied, significant evidence supports the utilization of
manganese oxide (MnO2) as a terminal electron acceptor for petroleum-related
contaminants. Shewanella putrefaciens, an isolate from anoxic sediments, is capable of
degrading petroleum hydrocarbons using MnO2 as a terminal electron acceptor (Burdige
and Dhakar, 1992). Bradley et al (2001) report MTBE degradation rates with manganese
oxides as terminal electron acceptor similar to those measured for ferric iron and sulfate.
Baedecker et al (1993) reported manganese and ferric iron, along with methanogenesis,
as being the most important electron acceptors in anaerobic petroleum hydrocarbon
remediation. Though MnO2 is thermodynamically favorable as a terminal electron
acceptor, the bio-available fractions account for only a small percentage of the total
manganese present in the soil system; most manganese in soil is tied up in insoluble
oxide-metal mineral systems (Huling et al, 2002). Cycling through the two common
forms (Mn(IV) and Mn(II)) keeps the relatively small percentage of free Mn available in
soil systems. Though it is only 10 % as abundant as Fe(III), a higher proportion of
20
Mn(IV) is available to act as an oxidant, adsorbant, and terminal electron acceptor
(Lovley 1991). This reactivity is due to Mn recycling in the environment, high surface
area for bioremediation and metal adsorption, and thermodynamic favorability.
Arguments that available manganese in soil systems is incapable of supporting
bioremediation are countered by statements that manganese cycling makes it more
bioavailable than Fe3+ or other terminal electron acceptors (Huling, 2002, Lovley, 1991).
This cycling is due to the rather high redox state of manganese (similar to nitrate),
promoting prompt cycling by aerobic bacteria and the formation of additional biogenic
oxides (Prescott et al, 2002).
Numerous concerns exist with using manganese(IV) for biostimulation. Limiting Mn2+ in
drinking water is of high importance because of the potential physiological side effects of
excess Mn consumption (Madrid et al, 2003). Although Mn is a micronutrient found in
small quantities in all living cells and activator of enzymes used in the TCA cycle, it is
toxic in large quantities, causing Parkinson’s-like syndrome, reproductive and immune
abnormalities, and hepatic cirrhosis (Xue et al, 2004). Water contaminated with Mn2+ can
also cause aesthetic and economic damage to pipes and fixtures (Madrid et al, 2003).
2.3.4 Sulfate Reduction
Although sulfate reduction in natural attenuation is theoretically limited by its low
thermodynamic favorability (∆G = 20.85 kJ/eeq), the relatively high oxidation state of
petroleum hydrocarbons makes the reactions possible by creating an overall positive free
energy value for the reaction (Rittmann and McCarty, 2001). Townsend et al (2003)
21
reported the ability of sulfate-reducing bacteria to mineralize polycyclic aromatic
hydrocarbons (PAH), classifying their remediation as “sulfate-dependent”. High PAH
biodegradation rates by sulfate-reducing bacteria were reported by Schmitt et al (1996) in
a shallow sand and gravel aquifer study and by Rothermich et al (2002) in petroleumcontaminated harbor sediments. Lovley et al (1994) documented the complete
mineralization of benzene by unamended sulfate reduction, the first report of a
successful, unamended anaerobic study at that time (Chapelle, 1999). Additionally,
sulfate is generally more abundant in groundwater systems than nitrate since sulfate is a
micronutrient, required only in small amounts for cell biomass. Relatively high
concentrations at typical field sites make sulfate a likely electron acceptor, as reported by
Cho et al (1997), who stoichiometrically linked it to approximately 66 % of BTEX
biodegradation in a jet fuel contaminated site. Despite inefficiencies, sulfate-reducing
organisms are thus capable of mineralizing numerous petroleum contaminants.
Though capable of mineralizing hydrocarbons, sulfate reduction has a multitude of
undesirable byproducts, including hydrogen sulfide gas and ionic sulfide (Mancini et al,
2003). Hydrogen sulfide gas is flammable and is extremely toxic at low concentrations,
causing pulmonary paralysis. Additionally, biodegradation can be limited by bisulfide
(HS-) accumulation, as reported by Edwards et al (1992).
2.3.5 Carbon Dioxide Fermentation
A distinct class of organisms called Archaea (Madigan and Martinko, 2006) performs
carbon dioxide reduction (methanogenesis). Methanogens are poorly classified, their
22
metabolism is poorly classified, and their true extent in the environment is unknown,
though Archaea are now believed to constitute approximately 20 % of all biomass on the
planet. Methanogens are obligate anaerobes, extremely sensitive to oxygen, pH, and
osmotic changes.
Methanogens are not thought to be responsible for hydrocarbon biodegradation, but work
in a consortium of microorganisms to degrade complex polymers completely, as shown
in Figure 2.4. Methanogens utilize numerous oxidized carbon sources, including carbon
dioxide and monoxide, formate, methyl compounds and organic acids (Madigan, 2006),
many of which are byproducts of other microbial metabolisms.
Methanogenic metabolism is the least thermodynamically favored of those considered in
this project (∆G = 23.53 kJ/eeq). Despite this, Wiedemeier et al (1999) estimated that
methanogenesis was responsible for 16 % of all petroleum hydrocarbon biodegradation at
a site under anaerobic conditions. In a review article, Wiedemeier et al (1999) stated that
methanogenesis is likely to be the most sustainable method of anaerobic natural
attenuation since the fermentation reactions driving it are limited only by the availability
of petroleum hydrocarbons. Salminen et al (2004) demonstrated that methanogenic
conditions become dominant once other TEAs are depleted. Grbic-Galic and Vogel
(1987) demonstrated 50 % reduction in benzene and toluene under methanogenic
conditions. Methane gas produced by this microbial pathway can be monitored in gas
headspace because the gas is fairly insoluble (Amos et al, 2005). Methanogenesis may
produce undesirable end products; for example under methanogenic conditions, MTBE is
23
oxidized to the more toxic TBA due to lack of sufficient energy to mineralize the
contaminant (Bradley et al, 2001).
Figure 3.4 Methanogenic Metabolism (Madigan, 2006)
24
2.3.6 Microbial Consortia
In environmental systems, microorganisms do not exist as pure cultures, but rather as
complex and interdependent ecological systems (Madigan and Martinko, 2006). In these
systems, microbial communities tend to be interdependent, contributing to maintaining
the nutrient cycles (nitrogen, carbon, sulfur) within close proximity, as can be
demonstrated within a Winogradsky Column (Madigan and Martinko, 2006). In
contaminated systems, a consortium of microorganisms with different metabolisms may
be more successful in managing a contaminant plume than one group of microorganisms
with the same or similar metabolic pathways (Boopathy, 2004).
Microorganisms can metabolize syntrophically, where the consortia to combine
metabolic abilities to degrade a substance they are incapable of degrading individually.
Townsend et al (2003) demonstrated syntrophic metabolisms between sulfate-reducers
and methanogens in the absence of sulfate. Meckenstock (1999) created a syntrophic coculture where a toluene-consuming sulfate-reducer and a hydrogen-consuming nitratereducer grew with nitrate as the sole electron acceptor.
In a consortium, the metabolic byproducts of one microbial species may enhance the
metabolism of another species. Schmitt et al (1996) demonstrated that biodegradation of
aromatic hydrocarbons by iron-reducing bacteria was stimulated by nitrate and sulfate
reducers. Though the exact mechanism was unknown, Schmitt et al postulated organic
acids from aromatic biodegradation complexed insoluble Fe(III) oxides, increasing their
bioavailability. Shelobolina et al (2003) demonstrated that nitrate-reducing
25
chemolithotrophs oxidized Fe(II) to Fe(III) while reducing nitrate to nitrite, promoting
the iron recycling. Though the specific cause was unknown, Boopathy (2004)
demonstrated that the mixed electron acceptor condition, amended with nitrate, sulfate
and ferric iron, outperformed the individually amended soil columns, reducing diesel
contamination by 88 % in 310 days. Kao and Wang (2000) determined that a mixed
amendment process prevented the further spread of a BTEX plume in an aquifer.
Consortia can be inhibitory. Coates et al (1996) demonstrated that native sulfate reducing
bacteria were able to out-compete biostimulated iron-reducing bacteria due to larger
numbers, regardless of the increased thermodynamic favorability of iron reduction.
Lovley (1991) indicates that iron-reduction can prevent manganese-reduction, since
Fe(II) will reduce manganese(IV) oxides.
2.4
Guadalupe Restoration Project
2.4.1 Site History
The former Guadalupe Oil Field, now titled the Guadalupe Restoration Project (GRP), is
located on the central California coast. The site straddles the San Luis Obispo – Santa
Barbara County line, with the bulk of the site in San Luis Obispo County (Figure 2.5).
Oil exploration at the site began in 1947, undertaken by the Sand Dune Oil Company,
which was largely unsuccessful in their efforts (Levine-Fricke Recon, 1996). Union Oil
of California
26
Figure 3.5 Location of Guadalupe Restoration Project Site on the Central
California Coast (Diaz 2006)
27
(Unocal) purchased 49 % interest in the field in 1951 and purchased the remaining shares
in mid-1953. By March 1953, production had increased to 2,000 barrels per day from 34
wells. By 1990, Unocal had increased production to 3,500 barrels per day from 218
production wells.
Oil produced at GRP was extremely viscous and dense (API Gravity 8 – 12, Lundegard
and Garcia, 2001), tending to behave similar to asphalt at ambient conditions (Levine
Fricke Recon, 1996). A lower viscosity petroleum mixture – called “diluent” – was
added to the crude oil to lower its viscosity in production wells and pipelines (Lundegard
and Johnson, 2003). Diluent was a mid-cut range of petroleum distillate, typically
coming from Unocal’s Santa Maria Refinery (Unocal, 1994). Diluent is chemically
similar to a mixture of kerosene and motor oil, with carbon ranges as displayed in Table
2.2.
Table 3.2 Boiling Point Distribution of Diluent (Lundegard and Garcia, 2001)
Approximate
Carbon
Range
<nC11
nC11 – nC14
nC14 – nC22
nC22 – nC30
>nC30
% of Diluent
Separate-Phase
Product
1
9
65
20
5
Diluent was distributed to the site by via pipeline constructed in 1955 (Unocal, 1994). It
was then stored in diluent tanks until pumped to individual wells at minimum pressure.
Once at the well, diluent flowed by gravity to the oil wells and was pumped out once the
diluent became well mixed with the underground petroleum. The mixture was then
28
transported to the refinery for distillation, where diluent could be fractioned, removed,
and returned to the site. Diluent was utilized for this purpose for 35 years (Lundegard
and Garcia, 2001).
During the 45 year time period when diluent was utilized at the site, diluent was
inadvertently released from the distribution system at multiple times and locations
(Lundegard and Johnson, 2003). Though the total volume of the released diluent is not
known, estimations range from 8.5 million (Lundegard and Garcia, 2001) to over 20
million gallons (Sneed, 2002). Many of the releases were sufficient for diluent to reach
the groundwater table and spread laterally, forming a light non-aqueous phase liquid (LNAPL), referred to as a source zone (Lundegard and Johnson, 2003). Source zone
contaminants are able to release toxic substances into the aquifer continuously as the
groundwater moves along its gradient. There are approximately 90 source zones at the
site, ranging in volume from 93 to 231,000 m3 (Board, 2005, Lundegard and Garcia,
2003). Figure 2.6 depicts the numerous source zones at the GRP site.
The first report of oil on the beach adjacent to the oil field occurred in 1988, but this
release did not match the fingerprint of diluent used at the field. A second release was
reported in 1990, and its proximity to the oil field and volume of the release resulted in
the foregone conclusion that the release was from GRP. A bentonite clay wall was
installed to prevent subsequent releases, but failed to prevent another release in 1994. At
this point, the Central Coast Regional Water Quality Control Board and the California
Department of Fish and Game ordered Unocal to develop a method to prevent further
29
releases. Later the same year, the U. S. Coast Guard filed a Notice of Federal Intent,
exerting jurisdiction over the beach to ensure the beach ecosystem was preserved. The
Central Coast Regional Water Quality Control Board issued a Cleanup or Abatement
Order 98-38 (CAO 98-38) in 1998, mandating site characterization, excavation of any
source zones posing an imminent threat to surface water quality, product recovery, pilot
testing, and treatment of affected soils (Board, 1998). Pilot tests performed at the site
include the following:
•
Land treatment units, where oxygen and nutrients were added to affected soils to
promote biodegradation;
•
Biosparging;
•
Dual pump recovery systems;
•
Hot water and steam flooding, and;
•
Full-scale phytoremediation using Arroyo willow trees.
2.4.2 Natural Attenuation Studies at the Guadalupe Restoration Project
Numerous studies have been performed to determine the extent and efficiency of
bioremediation at the GRP site.
Cal Poly graduate student Eileen Mick conducted a study to determine the chemical
changes occurring in dissolved-phase diluent during aerobic biodegradation processes
(Mick, 2006). Mick monitored TPH changes in on-site mesocosms – 4 ft cubes filled
with non-contaminated sand and approximately 100-gallons of diluent-contaminated
groundwater. Samples were collected during two runs, conducting infrared analysis to
30
Figure 3.6 Diluent Source Zones and Groundwater Monitoring Wells at the Guadalupe Restoration Project Site
31
determine chemical structure, column fractionation to determine changes in chemical
composition, TPH biodegradation potential measured over 20-day intervals and
equivalent carbon chain lengths using simulated distillation. Her results indicated that
the initial material was extremely polar, with non-detectable concentrations in the
aromatic and aliphatic fractions.
Stewart Lehman and Brian Dragich deduced a method to determine the efficiency of
natural attenuation by monitoring electron acceptors. This method was preferred due to
issues in accurately measuring TPH degradation due to competing natural attenuation
reactions, namely sorption, as well as unreliability of TPH quantification techniques at
the time. Lehman and Dragich developed theoretical stoichiometric ratios using
bioenergetics, and then TPH biodegradation in aqueous microcosms was assumed based
on the changes in electron acceptor concentration. However, since change in TPH
concentration was not monitored, the changes in TEA concentrations could not be
conclusively attributed to TPH biodegradation.
Marie Dreyer examined the effects of hydrocarbon weathering on their biodegradability
and toxicity in groundwater (Dreyer, 2004). This study was conducted in support of
natural attenuation as a means of reducing the toxicity of dissolved-phase diluent at GRP.
Thirty-four samples were collected at different wells along plume transects and were
incubated for 20-days to test short-term biodegradation. Sample toxicity was measured
before and after the 20-day incubation period. Dreyer reported that TPH degradation
rates decreased with increasing distance from the plume source, with first-order rate
32
constants decreasing from 5 to 46 % along the plume transects, suggesting that
biodegradability decreased with increased weathering. Though toxicity decreased with
decreasing TPH concentration, samples with low initial TPH had little change in toxicity,
indicating the presence of a toxicity threshold for biodegradation processes. Though this
study supported the reliability of natural attenuation for decreasing TPH and toxicity, it
also demonstrates that a threshold limit exists for both, possibly due to the weathering
processes.
Robin Cunningham studied the biodegradation rates of weathered hydrocarbons using
microcosms and soil columns (Cunningham 2004). The study focused on determining
biodegradation kinetics and accessing the ability of native species under field-modeled
conditions. Soil columns in triplicate and groundwater microcosms in duplicate were
established using materials collected from the GRP site. TPH changes in columns and
microcosms were monitored for 150 days. Cunningham demonstrated that
biodegradation was slightly faster in soil columns than in microcosms without soil.
Though biodegradation was observed in both systems, the increased rate and overall
degradation in the soil columns suggests that the fixed-surface and/or added inoculum
provided by the soil columns stimulated microbial activity.
Barbara Orchard (2005) and Lynne Maloney (2003) observed methane production in
microcosms containing soil from the GRP site. Kirk Gonzalez established microcosms
using anaerobic groundwater and soil to investigate biodegradation via methanogenic
pathways and connect this with observed TPH degradation and methanogenesis observed
33
at the site (Gonzalez, 2006). Microcosms containing either soil and groundwater or only
soil were prepared in bottles with minert valves to allow gas headspace sampling. During
the first phase, groundwater was not purged prior to microcosm establishment, thus
methane produced during the incubation period (240 days) could not be conclusively
attributed to methanogenic activity. During the second phase, the groundwater was
purged, but methane generation was measured in the control microcosm, indicating the
control was not sufficiently inhibited or methane generation was abiotic.
Paul Lundegard and Paul Johnson conducted an investigation into natural attenuation
occurring at the source zone (Lundegard and Johnson, 2003). A source zone (SZ) is an
area of petroleum-impacted soil potentially contributing to contamination in water or
vapor phases. In this study, source zone natural attenuation (SZNA) was investigated
using nested groundwater wells and soil gas probes in an attempt to confirm SZNA is
occurring at the GRP site and to evaluate SZNA rates and sustainability. Data collected
during the study confirmed natural attenuation is occurring along the source zone,
including:
•
Increasing hydrocarbon concentrations along groundwater flow paths between SZ
and down-gradient wells is evidence that the SZ contamination is dissolving into
the groundwater;
•
Decreases in electron acceptor concentrations along groundwater flow path from
up to down gradient are evidence of anaerobic biodegradation;
•
Increasing methane concentrations from up to down gradient provides evidence of
methanogenic activity;
34
•
Decreasing oxygen concentration and increasing carbon dioxide concentration
provides evidence of aerobic biodegradation, and;
•
Changes in hydrocarbon composition relative to SZ composition are evidence of
natural attenuation.
35
CHAPTER 4 . MATERIALS AND METHODS
3.1
Experimental Design
Anaerobic microcosms were established using anoxic groundwater and soil from the
Guadalupe Restoration Project site (GRP). Five anaerobic reducing conditions were
created by addition of electron acceptor amendments: nitrate, sulfate, ferric iron,
manganese oxide, and mixed amendment (nitrate, sulfate, and ferric iron). Unamended
anaerobic microcosms were established to compare biodegradation between microcosms
stimulated with electron acceptors and microcosms that represented natural GRP field
conditions. Aerobic microcosms were established by addition of oxygen to compare
anaerobic and aerobic biodegradation kinetics. Killed controls were established to ensure
that any changes in total petroleum hydrocarbon (TPH) concentrations observed in the
microcosms was the result of biodegradation rather than other abiotic methods of natural
attenuation.
Anoxic microcosms were incubated in an anaerobic chamber designed to simulate
groundwater conditions. Samples were analyzed after 0, 26, 134, and 407 days of
incubation. At each sampling date, microcosms were sacrificed in duplicate for chemical
analyses. Gas headspace was monitored for changes in helium, hydrogen, methane,
nitrogen, oxygen, and nitrous oxide concentrations. Groundwater was analyzed for
changes in TPH, electron acceptor amendments, and toxicity. Changes in the microbial
community were assessed at the 0 and 407-day sampling events.
36
3.2
Groundwater Collection
3.2.1 Groundwater Well Selection
Groundwater was collected at the Guadalupe Restoration Project site for use in
establishing the aqueous microcosms. Based on our experimental design, requiring
establishment of eighty-nine 2-Liter microcosms, approximately 50-gallons of
groundwater would be necessary to establish our microcosms. Selection of an
appropriate groundwater monitoring well (MW) for sample collection was made on the
basis of several conditions:
•
Groundwater at the well must be anaerobic. Anaerobic conditions were identified
by non-detectable dissolved oxygen concentration, presence of reduced iron,
presence of ammonium and absence of nitrate;
•
Groundwater should have low to non-detect concentrations of sulfate and nitrate
so that these electron acceptors could be added individually without interference
due to naturally-occurring electron acceptors;
•
Groundwater at the well should be subject to a single source-plume, not from comingled plumes;
•
Groundwater must have TPH concentrations less than 10 mg/L to ensure
homogeneity and would preferably have aliphatic, aromatic, and polar fractions
present;
•
The well must be able to accommodate a Grundfos submersible pump and
continuous nitrogen purging to maintain anaerobic conditions and minimize
disturbance during groundwater pumping;
37
•
The well must be able to supply the necessary volume, eliminating most nested
wells at the GRP site.
Based on these considerations, Monitoring Well (MW) J8-11 was chosen for
groundwater collection, pictured in Figure 3.1. Historically, MW J8-11 showed low
concentrations of dissolved oxygen (less than 0.60 mg/L since 2001), total iron (nondetect to 1.9 mg/L), sulfate (non-detect to 8.8 mg/L), and nitrate (non-detect since 2001)
(LFR, 2005).
J8-11
Figure 4.1 Location of Monitoring Well J8-11 at the GRP Site, Source of Anaerobic
Groundwater
At the time of the sampling, the sulfate concentration was 3.1 mg/L and ferrous iron was
7.5 mg/L. During the groundwater collection, dissolved oxygen was monitored using a
38
dissolved oxygen probe inserted in the pump, as seen in Figure 3.2. Dissolved oxygen at
time of collection was 0.14 mg/L.
Figure 4.2 Grundfos Pumphead with Dissolved Oxygen Meter and Flow Meter
3.2.2 Anaerobic Groundwater Collection Barrel
The need to maintain anaerobic conditions during groundwater collection, transport, and
microcosm establishment warranted the design of a collection barrel. Given the large
volume of water required, a large barrel was preferable over several small containers to
reduce the surface area to volume ratio for the containers, reducing the potential for
oxygen exposure. Additionally, multiple sample containers could introduce variability
into the experiment due to differences in purging efficiency or groundwater variability,
whereas a large container ensured homogenous initial groundwater samples.
39
The collection barrel, pictured in Figure 3.3, was designed to ensure effective nitrogen
purging and prevent oxygen infiltration before collection, during collection, transport,
and microcosm establishment. The barrel was constructed of polypropylene and had an
airtight lid with two threaded plugs, pictured in Figure 3.4. These plugs were modified to
allow development and maintenance of an anaerobic environment. Modifications
included the following:
•
Nitrogen Sparge Line: Continuous nitrogen purge was used before collection and
during groundwater collection and microcosm establishment to remove oxygen
within the barrel. The purge line is ¼-in diameter stainless steel tubing extending
to 2-in above the barrel base. Four diffusing stones were connected to the end of
the purge line to reduce nitrogen bubble size, increasing mass-transfer of nitrogen
into the water. This purge line helped to remove dissolved oxygen and methane.
•
Air Exhaust Outlet: An air outlet was necessary to prevent pressure build-up
within the barrel during the nitrogen purge. The outlet was ¼-in stainless steel
tubing, connected in series to a check valve and shut-off valve. The valves were
necessary to prevent re-entry of atmospheric oxygen during transport.
•
Fill Line: A ¾-in PVC fill-line extended inside the barrel to 2-in above the base.
This line is used to fill the barrel with groundwater and for filling the microcosms
when in the laboratory. The fill-line was connected to two valves and a ¾-in hose
fitting attached to the barrel lid. These two valves were used to purge the barrel
and fill-line prior to collection. After approximately 30-minutes of purging with
nitrogen gas, the exhaust line and main valve on the fill line were closed and the
second valve opened to ensure the fill line was completely purged of oxygen.
40
Fill line
Air exhaust
N2 sparge
Air-tight lid
Bulk head
Diffuser
Figure 4.3 Collection Barrel Design
3.2.3 Groundwater Collection Procedure
The collection barrel was transported to the GRP site on the day of groundwater
sampling. Prior to transport, resazurin (C12H6NNaO4, Acros Organics, #418900050) and
a spin bar were added to the empty barrel. Resazurin is a redox indicator used to
determine establishment of anaerobic conditions. Under anaerobic conditions, with a
reduction potential less than E = -0.42 mV, resazurin remains colorless (Gleason and
Gordon, 1989). In the presence of oxygen, resazurin turns bright pink, making detection
of more oxidized conditions unmistakable. Though research indicated resazurin
concentrations up to 0.1 % are common in anaerobic studies, 1 mg/L resazurin was
41
preferable to keep it below the initial TPH concentration. Since resazurin is a
hydrocarbon salt, the concentrations were kept below initial TPH concentrations to
prevent interference with future TPH analyses. To obtain this concentration, 0.19 g of
resazurin was necessary for 50 gal of groundwater.
Figure 4.4 Collection Barrel Lid with Fill Line, Purge Line, and Air Release Valve
To facilitate barrel transport, a drum dolly with locking casters was used. The barrel was
strapped into the truck to prevent spilling contaminated water or injuring the GRP site
staff. The GRP site and Levine Fricke Recon staff provided two 300-psi nitrogen gas
tanks necessary to purge the barrel and well during the collection process. The barrel,
fill-lines, and wellhead were all purged for approximately two hours. The well was
developed prior to collection by discarding the first 5-gallons extracted, collected in a
second 50-gallon polypropylene tank for proper disposal. The collection barrel was filled
by connecting the pump outlet to the nitrogen-purged fill-line. Nitrogen sparging was
continued during filling; excess nitrogen gas escaped through the exhaust line. The
42
volume collected was monitored using a flowmeter attached to the pumphead to prevent
overfilling the barrel. The collection barrel was purged with nitrogen during the entire
collection process and for approximately 20 minutes after pumping was ceased.
Bob Pease, a Levine Fricke Recon employee, conducted the barrel filling and transported
the filled barrel to the Environmental Protection Engineering Lab (EPEL) on the
California Polytechnic State University, San Luis Obispo campus. The groundwater was
purged with nitrogen gas for approximately 24 hours to remove any dissolved methane
prior to microcosm establishment. This was necessary to ensure methane detected in the
microcosms could be attributed to methanogenic activity during the experiment rather
than formed by evaporation of dissolved methane present in the groundwater at the time
of collection.
3.3
Soil Collection
Soil was collected from site H2 well pad area the GRP site, as depicted in Figure 3.5.
The location of site H2 is depicted in Figure 3.6. Site H2 was selected for selected for the
following reasons:
•
Low sulfate, iron, dissolved oxygen and nitrate concentrations existed in pore
water;
•
Anaerobic conditions were confirmed, as previously defined, and;
•
The proximity to the road to permit drill rig access.
MW H2-2 was originally selected for groundwater collection, but this well was inside of
a perched aquifer within the dune sand aquifer at the GRP site. For this reason, the GRP
43
site and Levine Fricke Recon staff members were uncertain whether MW H2-2 could
provide the full groundwater volume requested.
Figure 4.5 Drill Rig and Soil Collection at H-2
Soil was collected during the week prior to groundwater collection. The soil was
excavated using a hollow-stem auger drill rig under nitrogen purge and stored frozen in
eight soil cores under nitrogen gas to maintain anaerobic conditions. The soil was
transported to the Environmental Biotechnology Institute lab at California Polytechnic
State University, San Luis Obispo, campus, where the eight cores were mixed together in
an anaerobic glovebox. The cores were blended to help maintain soil sample uniformity
minimize differences among microcosms due to soil heterogeneity. Once the soil was
mixed in a 1-gal glass jar, the jar was closed, and sealed with parafilm, the soil was
transported to the EPEL and stored in the purged glovebox (described in Section 3.4.2)
until the microcosms were established.
44
H2
Figure 4.6 Location of Well Pad H2 at the GRP site, Location of Anaerobic Soil
Collection
3.4
Microcosms
3.4.1 Microcosm Bottles
Microcosm bottles used in this experiment were of a novel design to provide adequate
volume for duplicate TPH analyses while providing an airtight seal and minimizing gas
leakage. The bottles were constructed by Research and Development Glass Products and
Equipment of Berkeley, CA, and were constructed by welding the body of a 2-liter glass
media bottle (Fisher Scientific 06-414) to the neck of a standard 160-mL serum bottle
with 20 mm outer-diameter mouth (Figure 3.7). This modification allowed the use of 22mm Teflon-lined crimp seals to close the bottles once they were filled, thereby preventing
the loss of gases generated during the experiment from the microcosm headspace. Since
45
microcosms were custom-welded rather than manufactured, their total volume varied
individually. Average microcosm total volume was measured at 2.64 ± 0.02 L based on
the volumes of 6 randomly selected microcosms.
Figure 4.7 Microcosm Bottle Design
46
3.4.2 Anaerobic Glovebox
A custom-constructed glovebox, approximately 2.5 m3 in volume, was utilized for
microcosm establishment and storage during the entire incubation period. Robin
Cunningham and Daniel Gutierrez constructed the glovebox for use in an earlier project
using anaerobic soil columns completed in Summer 2005. To accommodate the
groundwater microcosm study, several modifications were made to the existing glovebox.
The glovebox characteristics are described below.
•
Structural Characteristics:
o Two gas lines were drilled to facilitate nitrogen gas circulation and to
allow for a direct purge line with diffusing stone, used to purge the
microcosms bottles with nitrogen gas before filling;
o To fill the bottles without exposing the groundwater to oxygen, the filling
occurred inside the glovebox. To accomplish this, a ¾-in hose barb was
added to the glovebox face, allowing the tubing to connect to the inside
and outside of the glovebox;
o To ensure proper sealing, all glovebox joints were re-secured using
silicone gel and weather-stripping tape was added to all removable
portions to aid in leak prevention;
•
Storage Capacity: To accommodate 89 microcosm bottles, three shelves were
constructed and placed into the glovebox prior to microcosm establishment;
•
Temperature and Light Control: In order to maintain GRP site groundwater
conditions, the glovebox interior was kept dark and at a temperature of
approximately 19 °C.
47
o A glovebox cover was designed to maintain dark conditions and prevent
absorption of additional heat. The cover was constructed of a layer of ¼in black felt to prevent light entry and a layer of thin, off-white cotton to
prevent heat absorption.
o To maintain groundwater temperature conditions, cooling water was
circulated through approximately 250-ft of ¾-in beverage-grade tubing
looped around the storage shelving, providing approximately 50-ft2 of heat
transfer area. Cooling water temperature was controlled using a Fisher
Scientific IsoTemp® 1006S Refrigerated Circulator. A thermometer was
placed inside an Erlenmeyer flask filled with deionized water to monitor
the temperature inside the glovebox. To maintain local groundwater
temperatures, the IsoTemp was adjusted to approximately 14 °C in the
winter and 22 °C in the summer.
With these modifications, the glovebox could accommodate the anaerobic microcosms
and maintain desired conditions. Figure 3.8 depicts the glovebox after the Day 0
sampling event, when 14 microcosms and our two initial TRF samples were removed.
3.4.3 Anaerobic Microcosms Establishment
3.4.3.1 Preparation
Prior to microcosm establishment, multiple steps were taken to prevent oxygen
contamination. Approximately 48 hours before establishment, all materials needed to fill
the microcosms were placed inside the glovebox, the glovebox was sealed, and a gas
48
mixture comprised of 98 % nitrogen and 2 % helium was pumped into the box at 20
L/min. This gas mixture was used to purge the microcosm bottles of residual oxygen by
inserting a diffusing stone connected to the gas line into each bottle for approximately
two minutes to ensure no oxygen was trapped inside the bottles.
Figure 4.8 Anaerobic Glovebox
3.4.3.2 Establishment
Anaerobic microcosms were established inside the nitrogen and helium purged glovebox,
as depicted in Figure 3.9. Each microcosm was established using 20 mL soil
(approximately 50 g dry weight) and approximately 2.3 L groundwater. Sand from the
GRP site was measured using a Pyrex beaker and added to the microcosm bottles using a
stainless steel funnel with tubing added to prevent spilling the measured volume of sand
49
and preventing sand from collecting on the lip of the microcosm, reducing the
effectiveness of the crimp seal. The sand was rinsed into the bottle using a wash bottle
filled with anaerobic groundwater from the collection barrel. Microcosm bottles were
filled to the neck with groundwater, and then 100 mL was removed using a 100 mL
Pyrex pipet. This was done to ensure all bottles had 100 mL headspace for gas analysis
since bottle volumes varied slightly due to custom construction. The groundwater in the
microcosms was then purged with the nitrogen/helium mixture for approximately 1
minute before the amendments were added.
Figure 4.9 Laleh and Meghann Establishing Anaerobic Microcosms Inside the
N2/He Purged Glovebox
For amended microcosms and controls, the electron acceptors were added as a
concentrated liquid. The amount of electron acceptor needed was determined using
bioenergetics (see Section 2.2) and assuming an initial TPH concentration of 6 ppm. A
50
safety factor of 3 was used to ensure electron acceptors would not be rate limiting. This
is important due to the presence of natural organic matter possibly consuming electron
acceptor amendments. Bioenergetics calculations yield a ratio (Y) of electron acceptor to
carbon source as determined based on theoretical electron acceptor energy yield.
Y=
moles electron acceptor
moles carbon source
(1)
Multiplying yield by the TPH concentration gives the stoichiometric concentration of the
electron acceptor required to mineralize the specific amount of TPH. Multiplying this
number by the safety factor (SF) increases the amount of the electron acceptor to ensure
the electron acceptor was not rate limiting.
Electron Acceptor Concentration = SF (Y × [TPH(mg /L)])
(2)
This is the molar concentration of the electron acceptor in ionic form. To determine the
concentration of the bound form, the molar electron acceptor concentration was
multiplied by the molar ratio of electron acceptor present in solid, salt form and the
weight of the solid amendment. For example, ferric iron was added as FeCl3, thus the
calculated concentration was multiplied by the ratio of Fe(III) ions in the solid form and
by the weight of ferric chloride solid. The mass of compound added to each bottle was
determined by multiplying this final concentration by the microcosm bottle volume.
Electron acceptor amendment concentrations are listed in Table 3.1. Unamended
microcosms did not have any amendments added. The aerobic microcosms were
established outside of the anaerobic glovebox, as described in Section 3.3.4. Note the
sodium azide concentration in the control was not multiplied by a safety factor, but was
51
determined based on success when used in previous experiments. Control microcosms
had no electron acceptor amendments.
Table 4.1 Microcosm Amendment Concentrations Based on Initial TPH
Concentration of 6 ppm and Using a Safety Factor of 3
Theoretical
Electron
Amendment
TEA
Safety Factor
Acceptor
Mass in
Concentration Concentration
Amendment Microcosm
Needed
(ppm)
Form
(mg)
(ppm)
Microcosm
Condition
Electron
Acceptor
Iron
Fe(III)
92
280
FeCl3
1,700
Manganese
Mn(IV)
51
150
MnO2
500
Sulfate
SO42-
26
77
Na2SO4
320
Nitrate
NO3-
20
59
KNO3
170
Oxygen
O2
19
59
O2
120
Unamended
CO2
N/A
N/A
CO2
N/A
Azide
Control
N3-
1,000
1,000
NaN3
2,100
Fe(III)
92
280
FeCl3
1,700
SO42-
26
77
Na2SO4
320
NO3-
20
59
NaNO3
170
Mixed
Amendments were added to the microcosms inside of the glovebox after microcosms
were inoculated with soil and purged briefly with a mixture of nitrogen and helium gas.
Amendments were added after gas purge to prevent contamination of the diffusion stone
on the gas purge line, potentially contaminating other microcosms. Amendments were
made into concentrated liquids to ensure the amendments dissolved in the microcosm, to
make the amendments easier to add while working within the glovebox, and to ensure all
52
microcosms received the same mass of amendment by way of a volume with known
concentration.
All amendments were dissolved in deionized water with the exception of manganese
oxide (MnO2), which was added as a precipitated solid suspension. Table 3.2 lists the
salt or solid form of the amendments and their sources.
Table 4.2 Amendment Source and Form
Amendment
Nitrate
Sulfate
Ferric Iron
Manganese (IV)
Azide
Chemical Formula
KNO3
Na2SO4
FeCl3
MnO2
NaN3
Source
Fisher Scientific
Fisher Scientific
Fisher Scientific
Amorphous Solid
Fisher Scientific
Cat #
P263
ACS35425
F1010
S2271
Since MnO2 is not an ionic salt, it does not dissolve in solution; instead, it remained solid
in the microcosms. Since biotic manganese reduction involves close contact between the
oxide and the bacteria (Burdije and Dhakar, 1992), manganese oxide surface area should
be maximized to ensure the solid is bioavailable. Fresh abiotic manganese oxide
precipitate was used rather than a crystalline solid to improve bioavailability.
As detailed in Nelson et al (1999), amorphous MnO2 was created by combining KMnO4
and MnCl2 in a basic solution at 90 °C with constant stirring. According to Murray et al
(1984), this method produces solid-phase manganese with a low degree of crystallinity
and an X-ray diffraction pattern attributed to δ-MnO2. The concentration of manganese
solid in suspension was determined by extracting a known volume from suspension and
53
heating at 100 °C until all water was evaporated, then weighing the solid using an
electronic balance.
Once the amendment solutions were added to the groundwater in the microcosms, the
microcosm bottles were crimp-sealed using 22-mm Teflon® seals. Once all bottles were
purged, amended, labeled, and sealed, the glovebox was shut. When the microcosms
were sealed, the helium gas was removed from the gas flow and the nitrogen gas flow
was reduced to approximately 3 L/min. This flow was maintained for the remainder of
the experiment unless the glovebox was opened, after which the flow was increased to
approximately 20 L/min overnight.
3.4.4 Aerobic Microcosm Establishment
Aerobic microcosms were established outside of the glove box as oxygen contamination
was not a concern. Microcosms were established using 20 mL soil (approximately 50 g
dry weight) and approximately 2.3 L groundwater. As with the anaerobic microcosms,
the aerobic microcosm bottles were filled to the neck with groundwater, and then 100 mL
was removed using a 100-mL Pyrex pipet. This was done to ensure all bottles had 100
mL headspace for gas analysis. To ensure that oxygen was not the limiting reagent for
TPH degradation, the microcosms were purged using a mixture of approximately 98 %
oxygen and 2 % helium for approximately two minutes. Helium was added as a quality
control measure to monitor gas leakage from the bottle during the experiment. Bottles
were then crimp-sealed with 22-mm Teflon® seals and placed in a Fisher Iso-Temp
54
incubator set to 19 °C. Establishing aerobic microcosms in sealed bottles permitted gas
headspace analysis and prevented interference from potential hydrocarbon volatilization.
3.4.5 Adjusting Microcosm pH
During the experiment, iron-amended microcosms required pH correction due to the
decrease in pH after ferric chloride addition. A method to test the reason for pH
reduction and subsequent pH correct was determined.
The main concern in adjusting microcosm pH was to ensure the amount of base added
required a minimum amount of microcosm gas headspace. Tests were performed using
0.1 N, 1.0 N, and 2.5 N sodium hydroxide solution (NaOH). The results of these tests are
listed in Table 3.3. Following this experiment, 5 N NaOH was chosen to correct
microcosm pH because it would only require approximately 5 % of headspace.
Table 4.3 Pilot-Scale Test Results for pH Adjustment Using Different NaOH
Normalities
Groundwater + Sand pH
pH After FeCl3 Addition
Test 1
8.20
2.98
Test 2
8.38
3.00
Test 3
8.00
3.32
Test 4
8.23
3.16
Normality NaOH Used
0.1 N
0.5 N
1.0 N
2.5 N
Final Volume Base Added
10 mL
4 mL
2.4 mL
0.9 mL
Final pH
5.2
8.7
8.2
8.5
Headspace Required
100 %
40 %
24 %
9%
Note: Since experiment was conducted at 1/10 scale, 10 times the volume listed would be
necessary to correct the pH in the microcosms.
55
The pH adjustment was conducted in the glovebox to ensure anaerobic conditions were
not compromised. One day prior to pH correction, helium was added to the gas flow into
the glovebox at approximately 4 mL/min. Both the helium and nitrogen flow rates were
increased at the time of pH correction. However, the helium flow meter was not
accurately calibrated prior to this usage, and due to this oversight, helium concentrations
in the re-established microcosms were much lower than other microcosms. To correct
pH, 5.0 N NaOH was added dropwise to iron-containing microcosms using a 10-mL pipet
while monitoring pH. Microcosms required approximately 4.5 mL of 5.0 N NaOH to
reach a final pH of approximately 8.0. The NaOH solution was mixed in the microcosm
using the N2/He purge line, which was inserted into the bottle and allowed to bubble for
30 seconds. This also removed any oxygen dissolved in the basic solution. After mixing,
the pH was tested and adjusted further with dropwise addition of NaOH followed by a
few seconds of gas bubbling to mix. If the target pH was exceeded, 1 N HCl was added
dropwise to raise pH to the desired range.
Once pH was corrected, microcosms were re-inoculated with 50 g fresh anaerobic sand,
since the acidic pH was assumed to kill all microorganisms in the iron-amended
microcosms. After sand was added, the gas headspace was re-established by removing
approximately 24 mL from the microcosm once pH was corrected (20 mL is the
approximate volume necessary for 50 g of sand, and the volume of NaOH required
ranged from 3.5 – 4.5 mL). The water was then purged for 1 min using the N2/He
mixture, and microcosms were resealed with Teflon-lined 22 mm crimp seals. Ironamended microcosms were adjusted before mixed amendment microcosms to prevent
56
NO3- and SO42- contamination by the gas diffusion stone. The glovebox was sealed, He
gas line was closed, and the N2 gas was maintained at approximately 27 L/min for 24
hours to purge any potential O2 contamination. After 24 hours, the N2 flow rate was
reduced to 3 L/min.
3.5
Gas Headspace Analyses
On each sampling date, gas headspace analyses were the first analyses performed.
Monitoring changes in headspace gas concentration was used to indicate microbial
activity and microcosm integrity. Gases monitored were nitrogen (N2), helium (He),
hydrogen (H2), oxygen (O2), carbon dioxide (CO2), methane (CH4), and nitrous oxide
(N2O). The nitrogen to helium ratio was monitored as an indicator of microcosm
integrity, since microcosms were established and purged with a 98 % / 2 % nitrogenhelium mixture. Oxygen concentration was monitored in anaerobic microcosms to
ensure anaerobic conditions were maintained. Carbon dioxide was monitored as an endpoint of biodegradation, indicating hydrocarbons (or other organic compounds) were
mineralized within the microcosms. Methane and nitrous oxide are indicators of specific
microbial metabolisms – methanogenesis and nitrate reduction, respectively. Hydrogen
gas served as an indicator of fermentation as a possible precursor to methanogenesis.
Gas headspace analysis was performed by Greg Ouellette, owner and operator of Inland
Empire Analytical (Figure 3.10). A 10-mL gas-tight syringe, stopcock and needle
assembly was first purged three times with nitrogen. The headspace sample was then
collected by inserting the needle through the septa on the microcosm and collecting a 10
57
mL headspace gas sample. The syringe was pumped three times to collect a
representative sample of the headspace gas. The gas sample was then analyzed for the
fixed gases on an Agilent 3000A Micro Gas Chromatograph equipped with a sample
inlet, three sample loop injectors, three columns and three detectors. For this analysis,
two of the columns were used. The first split was injected onto a 10-meter Molesieve 5A
PLOT column. This column separated helium, hydrogen, oxygen, nitrogen and methane.
The second split was injected onto an 8-meter poraPlot Q column. This column
separated carbon dioxide from the gas mixture. Components were detected by solid-state
thermal conductivity detectors. Both columns are maintained at 45 °C. The total
analysis time was 2 minutes.
Figure 4.10 Greg Ouellette Conducting Gas Headspace Analysis
For some of the gases, it was necessary to account for dissolved gases to calculate the
total amount of gas produced (or consumed). Aqueous concentrations were calculated
from headspace concentrations using Henry’s Law. Henry’s law specifies that, at
58
constant temperature, the amount of a given gas dissolved in a given type and volume of
liquid is directly proportional to the partial pressure of the gas in equilibrium with the
liquid. The formula for Henry’s Law is:
pi = k H ,T Ci
(3)
where pi is the partial pressure of the solute gas above the solution, Ci is the concentration
of the solute gas in the solution, and kH,T is the Henry’s Law Constant for the solute gas
at temperature T. When kH,T is used in this manner, it has the following units:
k H ,T =
Lso ln atm
molgas
(4)
However, Henry’s Law takes many mathematical forms; therefore care must be taken to
check the units and make proper conversions when using this method.
The value of kH,T is temperature dependent and most values are indexed at standard
temperature, 298 K (25 °C). When using this method at another temperature, the
constant must be converted using the van’t Hoff Equation:
k (T ) = k (TΘ ) × e
⎡ ⎛ 1 1 ⎞⎤
⎢−C ⎜ −
⎟⎥
⎣ ⎝ T TΘ ⎠⎦
(5)
where T is the temperature in Kelvin, k(TΘ) is the Henry’s Constant at standard
temperature and pressure, and C is a constant specific for the type of gas in solution.
Values of kH,T and C used in this experiment are included in Table 3.4.
Gases were typically measured in units of parts per million on a volume basis (ppmv), or
percent. To determine aqueous concentrations, gaseous concentrations were converted to
mg/m3, then to atmospheres (atm) using the ideal gas law. Once the gaseous species are
59
converted to partial pressure units, the molarity in solution was calculated using Henry’s
Law. The gaseous concentration was determined using the molecular mass of the
species.
Table 4.4 Henry's Law Constants and Temperature Conversion Factors for
Gaseous Solutes (National Institute of Standards and Technology, 2005)
Gaseous
Solute
CH4
O2
CO2
N2
He
N20
H2
3.6
kH,T
[(L-atm)mol-1]
725.2
781.0
29.01
1692
2672
40.61
1302
C
1600
1700
2400
1300
230
2600
500
TPH Analysis
3.5.1 Solvent Extraction (EPA Method 3510C)
EPA Method 3510c (described in Appendix B) was used in extracting petroleum
hydrocarbons from the microcosm water samples into methylene chloride (MeCl) for
further TPH analyses. Extractions were started the day of sampling and completed within
three days of sampling. Glassware was prepared by washing with Alconox soap, rinsing
with tap water followed by three nominal rinses with deionized water, rinsing with
methanol, then rinsing with MeCl. After preparation, glassware was dried in the fume
hood until used.
Extractions were performed using 2-Liter separatory funnels supported by ring stands in a
chemical fume hood, as seen in Figures 3.10. Pyrex funnels were placed below the
separatory funnels (Figure 3.11), each of which contained approximately 50 mL of
60
anhydrous sodium sulfate (Fisher Scientific S415-212), dried at 100 °C for two hours
prior to use. The sodium sulfate was utilized as a drying agent to remove any water
dissolved in the methylene chloride during the extraction process. Glass wool (Supelco
2-0411) was used to plug the stem of the Pyrex funnel to hold the anhydrous sodium
sulfate while allowing the solvent to pass through the funnel into the TurboVap
Concentration Tube below the funnel. The TurboVap Concentration Tube (Zymark
Corporation) was 200 mL in volume with a 1 mL endpoint stem, designed to fit into the
TurboVap II Concentrator (Zymark Corporation) following the extraction. The
workstation used a water bath for sample heating and directed gas nozzles for directed
gas-stream evaporation. The sodium sulfate, glass wool, and all glassware were rinsed
twice with MeCl prior to starting the extraction.
Figure 4.11 Glassware Set-Up in Chemical Fume Hood for TPH Extraction
Each 2-Liter microcosm bottle provided two one-liter water samples for extraction. The
1-L aliquots were measured using a Kimax 1-L graduated cylinder. Each extracted
volume was spiked with 1 mL of an internal standard (hexacosane dissolved in MeCl) at
61
a concentration of approximately 1 mg/L, which would aid in monitoring extraction and
analytical efficiency. As a high molecular weight compound, hexacosane would elute at
the upper range of the diluent-range peaks in the chromatogram produced by the gas
chromatograph (GC). The area of this peak was used to monitor extraction efficiency.
Approximately 60 mL of pure GC-grade MeCl was added to the separatory funnel using
a Kontes autopourer. The separatory funnel was capped, removed from the ring-stand,
and gently shaken several times to mix the water and MeCl. The funnel was inverted and
the stopcock opened to relieve accumulated gas pressure. This was repeated until gas
pressure was equalized. At this point, the sample was vigorously shaken for one minute
(approximately 100 shakes). The separatory funnel was returned to the ring stand,
uncapped, and allowed to sit for ten minutes. During this time, the MeCl (specific
gravity = 1.3255) settled to the bottom of the separatory funnel, below the less dense
aqueous layer. After 10 minutes, the MeCl extract was slowly drained from the
separatory funnel through the Pyrex funnel filled with anhydrous sodium sulfate and into
the TurboVap Concentration Tube. This process was repeated three times for each oneliter sample, using a total of 180 mL MeCl. After the final draining, the sodium sulfate
was rinsed with approximately 30 mL of MeCl to remove any TPH residual in the drying
agent and glass wool.
3.6.2 Concentrating Extract Solution
Since the TPH concentrations in the samples were relatively low (approximately 5 mg/L
in the initial groundwater), the extracts were concentrated to increase detection
sensitivity. To concentrate the extract, the TurboVap Concentrator Tubes (the Tubes),
62
each containing approximately 200 mL of MeCl and extracted petroleum hydrocarbons,
were placed into a TurboVap II Concentrator (the Concentrator). The Tubes were placed
into the Concentrator’s water bath at 35 °C with ultra-high purity nitrogen gas directed at
the solvent surface with a constant pressure of 16 bar. The extract was concentrated to an
endpoint volume of approximately 1 mL.
After evaporation, the Tubes were removed from the Concentrator and returned to the
fume hood. The extract was transferred to a 10-mL Kimax graduated cylinder using a 2mL glass Pasteur pipet and a silicon pipet bulb. Once the extract was transferred, the
Tube was rinsed with approximately 1 mL of MeCl. The rinsed MeCl was then
transferred to the graduated cylinder. This was repeated until the total volume of extract
in the graduated cylinder measured 5 mL. The final extract volume was important
because it was used to calculate the amount of TPH initially present in the water sample,
which can be calculated using a concentration factor. For this extraction, 1 L of water
concentrated to 5 mL of methylene chloride resulted in a concentration factor of 200. The
extract was then transferred into two 2-mL crimp-top vials and stored until analyzed
using gas chromatography/flame ionization detection.
3.6.3 Total Petroleum Hydrocarbon Analysis (EPA Method 8015c)
Each extracted sample was analyzed for total petroleum hydrocarbons (TPH) using a
Hewlett Packard 6890 Gas Chromatograph with Flame Ionization Detection (GC/FID)
and a 6890 Series Auto Sampler. A Supelco SBP-1 16892-02B capillary column was
used in the GC. The GC/FID was remotely controlled using Agilent Technologies
63
ChemStation® software (rev A.08.03). The GC-based TPH analysis method was based
on EPA Method 8015c, included in Appendix C.
Standards were run periodically between sample analyses and with each sample run as a
quality control measure to ensure consistency. Tables 3.5 and 3.6 summarize the GC
oven specifications and operating conditions used during sample analyses.
Table 4.5 GC Oven Specifications
Initial Temperature:
45 °C
Final Temperature:
275 °C
Oven Ramp Specifications
Rate
Final
Final Time
Temperature
(min)
(°C/min)
(°C)
0
45
3.00
12
275
19.17
0
275
12.00
Routine maintenance was performed on the GC/FID throughout the experiment. The
injection septum was changed after approximately 50 injections, the liner was
periodically inspected and was changed when necessary (approximately every 100
injections), and the injection needle was changed whenever alignment concerns occurred.
An average lower detection limit of 38 mg/L TPH in MeCl was established by finding the
peak area average from ten methylene chloride blanks and converting the peak area to a
concentration using the calibration curve. This was equivalent to 0.19 mg/L TPH in H2O
using the concentration factor. At least one vial of pure methylene chloride was run as
blanks with every set of samples. Monitoring peak area curves in the methylene chloride
blanks gave an indication of the column cleanliness and the reliability of data produced
64
by the GC. If the peak area was greater than the average of the standard deviation for
blanks, additional blanks were run and the column was inspected or manually purged. If
the methylene chloride blanks continued to result in peak areas above the average, the
column was kept at 400 °C overnight to volatilize and purge residual hydrocarbons in the
column.
Table 4.6 GC Operating Conditions
INLET
Mode
Initial Temperature
Purge Flow
Total Flow
Splitless
200 °C
50.0 mL/min
59.8 mL/min
COLUMN
Capillary Column
Model Number
Maximum Temperature
Nominal Diameter
Nominal Inlet Pressure
Outlet Pressure
SBP-1
Supelco 16892-02B
320 °C
530.00 µm
4.0 psi
Ambient
DETECTOR
Detector Type:
Temperature
Air Flow
Make-up Gas Type
Flame Ionization Detector (FID)
Hydrogen Flow
340 °C
On
Make-up Flow
Nitrogen, UHP
Carrier Gas
Gas Saver
Pressure
Purge Time
Off
9.9 psi
0.20 min
Nominal Length
Nominal Film Thickness
Mode
Initial Flow
Average Velocity
30.0 m
1.00 µm
Constant Flow
7.0 mL/min
48 cm/s
On
On
Helium
Sample concentrations were determined by direct comparison to calibration curves
created using known concentrations of pure diluent source material collected from the
GRP site. A 10,000-ppm diluent in MeCl stock solution was made gravimetrically from
pure diluent source material. Stock solutions were created gravimetrically due to
difficulties pipetting the diluent. Six or seven dilutions were made from this stock to
construct a calibration curve, with the lowest dilution near practical quantitation limits.
65
Typical concentrations used were approximately 40, 80, 240, 400, 800, 1,000 and 2,000
mg diluent per liter of MeCl.
A diluent concentration standard was created by adding 0.5036 grams of pure diluent to a
volumetric flask and diluting to 50 mL total volume with MeCl, yielding a final stock
concentration of 10,130 mg/L diluent in MeCl. The dilutions were prepared from this
stock solution by first creating a 2,026 mg/L diluent in MeCl solution from the stock,
then using this solution to create additional dilutions. Calibration standards made with
this stock solution are listed in Table 3.7
Table 4.7 GC Calibration Standard Set for TPH Analysis
Volume Stock
(mL)
Volume MeCl
(mL)
1
1
3
5
10
10
5
24
24
22
20
15
10
20
Stock
Concentration
(mg/L)
405.2
2,026
2,026
2,026
2,026
2,026
10,130
Total Volume
(mL)
Concentration
(mg/L)
25
25
25
25
25
20
25
40.52
81.04
243.1
405.2
810.4
1,013
2,026
Samples of each standard were run on the GC, whereby peak areas were obtained. A
calibration curve was generated using the concentration of standards and their
corresponding peak areas. Linear regression was applied to the points, correlating peak
areas and concentrations. The calibration curve obtained from this set of standards had
an R2 value of 0.9996 and is displayed in Table 3.8 and Figure 3.12.
66
Table 4.8 Diluent Standard Concentrations and GC Output for Calibration Curve
TPH Concentration
(mg/L in MeCl)
40.52
81.04
243.1
405.2
810.4
1013
2026
Peak Area
3462.5
6702.6
20542.7
34990.9
69558.7
87407
168444.5
180000
160000
140000
Peak Area
120000
100000
y = 83.46x + 798.12
R2 = 0.9996
80000
60000
40000
20000
0
0
400
800
1200
1600
2000
TPH Concentration in MeCl (mg/L)
Figure 4.12 TPH Calibration Curve
To calculate extraction efficiency and recovery when extracting TPH from water
samples, a calibration curve was developed for hexacosane for use as an internal quality
control standard. To accomplish this, 106.5 mg of hexacosane was added to 50 mL of the
10,130 mg/L diluent in MeCl standard before the remaining dilutions were created. This
created a set of hexacosane standards within the diluent standards, as each of the
67
standards had proportionately less hexacosane with subsequent dilutions. Hexacosane
concentrations within the diluent standards are listed in Table 3.9.
Table 4.9 Hexacosane Concentrations in Diluent Standards
TPH Concentration
(mg/L in MeCl)
40.52
81.04
243.1
405.2
810.4
1013
2026
10,130
Hexacosane
Volume (mL)
1
1
3
5
10
13
5
5
Total Volume
(mL)
25
25
25
25
25
25
25
50
Hexacosane
Concentration
(mg/L)
0.443
0.852
2.556
4.26
8.52
11.076
21.3
106.5
Hexacosane peaks were integrated separately from the diluent peaks in the GC
chromatograms and subtracted from the total peak area. Hexacosane peak area was then
related to hexacosane concentration by linear regression. The hexacosane calibration
curve obtained from this set of standards with an R2 value of 0.9999 is displayed in Table
3.10 and Figure 3.13. An example chromatogram is included in Figure 3.14.
Table 4.10 Hexacosane Concentration and GC Output for Calibration Curve
Hexacosane
Concentration (mg/L)
0.443
0.852
2.56
4.26
8.52
11.1
21.3
Peak Area
34.7
61.9
184.3
318.3
631.6
808.9
1571.6
68
1800
1600
y = 73.698x 0.0595
1400
R2 = 0.9999
Peak Area
1200
1000
800
600
400
200
0
0
5
10
15
20
Hexacosane Concentration in MeCl (mg/L)
Figure 4.13 Hexacosane Standard Curve
Figure 4.14 GC Output for 2014 ppm Diluent Standard. Large Peak at 22.5
Minutes is Hexacosane.
69
3.7
Electron Acceptor Analyses
3.7.1 Sulfate and Nitrate Analysis by Ion Chromatography
Changes in sulfate and nitrate concentrations were monitored using a Dionex DX-120 Ion
Chromograph (IC) equipped with an AS40 Autosampler, AS-9 analytical column and
AS-9 guard column. The IC is controlled using the Dionex Chromeleon®
Chromatography Management System.
Ion chromatography functions on the basis of ion selectivity. Based on their selectivity,
ions will elute from the column at different time intervals. The elution time is controlled
by the ion, the eluent strength and flow rate, and system backpressure. During operation,
an electrical conductivity (EC) detector is continuously monitoring solution EC as time
elapses. When only the eluent is leaving the column, the EC records a flat line.
However, when an ionic species is eluted, the detector records a peak in EC proportional
to the strength of the ion detected. The area under the peak at the time the ion is eluted is
relative to its concentration in the sample.
For the AS-9 4 mm analytical column, 9 mM sodium carbonate was used as eluent.
Eluent is degassed using helium prior to usage. To determine the elution times of nitrate
and sulfate, a Dionex 7-anion standard was used. This is necessary since the samples
being analyzed were from a complex groundwater system and can have multiple anions
present. An example of 7-anion standard output is presented in Figure 3.15 with the
species of interest labeled on the accompanying table.
70
iI
140 n
µS
1
7
D
C
E
2 - 5.767
100
75
3 - 7.017
50
5 - 10.283
1 - 3.500
4 - 9.033
7 - 15.250
6 - 14.417
25
0
min
-20
0.0
No.
1
2
3
4
5
6
7
Total:
2.0
Ret.Time
min
3.50
5.77
7.02
9.03
10.28
14.42
15.25
4.0
6.0
Peak Name
n.a.
n.a.
n.a.
n.a.
NO3
n.a.
SO4
8.0
10.0
Height
µS
34.186
126.964
52.667
33.387
37.766
25.821
29.062
339.854
12.0
Area
µS*min
5.913
25.364
13.420
9.429
12.436
12.351
16.255
95.167
14.0
Rel.Area
%
6.21
26.65
14.10
9.91
13.07
12.98
17.08
100.00
16.0
Amount
PPM
n.a.
n.a.
n.a.
n.a.
n.a.
n.a.
n.a.
0.000
18.5
Type
MB
BMb
bMB
BMb
bMB
BM
MB
Figure 4.15 7-Anion Standard Output from DX-190 Ion Chromatogram
Sulfate and nitrate concentrations were determined by comparing sample peak areas to a
standard curve. Since the ionic species will not react with each other, standards were
prepared containing nitrate, nitrite, and sulfate. A standard stock solution was first made
containing 1000 mg/L each of KNO3, KNO2, and Na2SO4. This standard stock solution
was diluted to construct a dilution series. A typical dilution series, elution times and peak
areas for the three anionic species observed are listed in Table 3.11. Once the peak areas
were obtained, they were used to create a calibration curve mathematically relating
71
concentration to IC peak area. To ensure linearity at high and low concentrations, two
standard curves are made: 1 – 20 ppm for low concentration range and 20 – 200 ppm for
high concentration range. Typical IC output is seen in Figure 3.16. Calibration curves
are shown in Figures 3.17 and 3.18.
Table 4.11 Calibration Curve Data and Elution Times for Nitrate, Nitrite, and
Sulfate as Monitored by Ion Chromatography
Species
Concentration
(ppm)
1
2
5
10
20
40
60
80
100
200
NO2
(Elutes at 6.15 s)
NO3
(Elutes at 8.58 s)
SO4
(Elutes at 13.4 s)
0.27
0.38
0.48
0.99
1.95
4.2
6.87
9.42
11.9
24.17
0.22
0.31
0.39
0.82
1.6
3.52
5.72
8.09
10.51
23.58
0.29
0.41
0.52
1.033
1.95
4.16
8.09
11.36
14.66
32.39
Duplicate IC analyses were conducted for each of the samples from each microcosm
condition, making a total of four samples per electron acceptor amendment per sampling
date. Nitrate and sulfate concentrations were monitored for all microcosm conditions. On
each sampling date, all microcosm samples were vacuum filtered using 0.2 µm
nitrocellulose fiber filters (Fisher Scientific). Once an adequate volume was filtered,
samples were frozen until IC analysis. To analyze them, samples were loaded into
Dionex 5-mL PolyVials and loaded in the auto sampler. One deionized water blank was
loaded for each sample to clean the column between samples. One standard was run per
set of samples to monitor IC performance. If samples were too concentrated for
72
interpretation using standard curves, the samples were diluted appropriately and the
analysis was repeated using the diluted sample.
IS
M
D
20.0 T
µS
M
T
S
D
C
E
17.5
1 - 10.633
15.0
12.5
10.0
7.5
µS
2 - 15.350
5.0
2.5
0.0
-2.5
-5.0
-7.5
min
-10.0
0.0
2.0
4.0
6.0
8.0
10.0
12.0
14.0
16.0
18.5
Minutes
Figure 4.16 Ion Chromatograph for 50 ppm Nitrate and Sulfate Standard
73
2.5
y = 0.0875x + 0.1758
R2 = 0.9926
SO4
2
1.5
y = 0.0887x + 0.1398
Peak Area
R2 = 0.9901
NO2
1
y = 0.073x + 0.1133
R2 = 0.9902
NO3
0.5
0
0
5
10
15
20
25
Concentration (mg/L)
NO2
NO3
SO4
Figure 4.17 Calibration Curve for Nitrate, Nitrite, and Sulfate, 1 - 20 ppm Range
35
30
y = 0.1718x - 2.213
R2 = 0.9984
SO4
Peak Area
25
20
y = 0.124x - 0.5802
R2 = 0.9998
NO2
15
y = 0.1235x - 1.4579
R2 = 0.9975
NO3
10
5
0
0
50
100
150
200
250
Concentration (mg/L)
NO2
NO3
SO4
Figure 4.18 Calibration Curve for Nitrate, Nitrite and Sulfate, 20 - 200 ppm Range
74
3.7.2 Iron Analysis by the Phenanthroline Method (Standard Method 3500-Fe)
Possible iron reduction from ferric iron (Fe3+) to ferrous iron (Fe2+) was monitored by the
concentration of ferrous iron in the microcosms. This was accomplished using the
phenanthroline method as described in Standard Methods for the Examination of Water
and Wastewater (American Public Health Association, 1998) 3500-Fe, an EPA-approved
method for iron analysis added to Standard Methods in 1997. Method 3500-Fe is
included in Appendix D.
The phenanthroline method differentiates between ferrous and ferric iron by taking
advantage of the dissolved form of ferrous iron, as opposed to the colloidal form of ferric
iron. 1,10-Phenanthroline chelates ferrous iron cations at a 3-to-1 ratio, forming an
orange-red complex. The color formed by the complexation follows Beer’s Law – the
intensity of the color is proportional to the complex concentration, which is independent
of pH from 3 to 9. Dissolved iron concentration can be detected as low as 10 µg/L using
a spectrophotometer at 510 nm.
A Hitachi U-3010 UV/VIS Spectrophotometer was used for this colorimetric method,
remotely controlled by Hitachi UV Solutions 2.0 software. Iron concentration in
microcosms was determined by comparing them to a calibration curve created using
known concentrations of complexed Fe(II)-phenanthroline. The standards were created
in accordance with Standard Methods. Reagents used included concentrated
hydrochloric (HCl) acid, hydroxylamine solution, ammonium acetate buffer and
75
phenanthroline solution, both made in laboratory. All reagents used were certified for
use in metal detection and had low to non-detectable iron concentrations.
A 10 mg/L Fe(II) solution was prepared by diluting 200 mg/L Fe(II) stock solution in a
volumetric flask. A 1,000 mg/L phenanthroline solution was made by adding
approximately 100 mg 1,10-phenanthroline monohydrate (C12H8N2⋅H2O, Fisher
Scientific P-70) to a 100-mL volumetric flask and diluting with deionized water (DI
water) by stirring and heating to 80 °C. Ammonium acetate buffer was made by
dissolving 250 g ammonium acetate (NH4C2H3O2) in 150 mL water and 700 mL glacial
acetic acid.
To create the standard dilutions, 50 mL of 10 mg/L Fe(II) solution, 2 mL concentrated
HCl, and 1 mL hydroxylamine hydrochloride (NH2OHHCl, Lab Chem Inc, LC15530-1)
were added to a 250-mL Erlenmeyer flask and boiled to reduce the volume by 50 %. The
mixture was cooled to room temperature. Once cooled, 10 mL ammonium acetate buffer
and 4 mL phenanthroline solution were added. The mixture was diluted to 100 mL using
a volumetric flask, yielding a 10-ppm solution of complexed ferrous iron and
phenanthroline [Fe(II)-Phen]. After thorough mixing, the solution was allowed to stand
for 10 minutes for full color development. Once the color fully developed, the mixture
was diluted using a 50-mL volumetric flask as described in Table 3.12 to create the
standards used in the calibration curve.
76
Table 4.12 Fe(II)-Phenanthroline Dilution Series for Calibration Curve
Volume of 10 ppm
Fe(II)-Phen Solution
(mL)
1
5
10
20
30
40
50
Final Fe(II)-Phen
Concentration
(ppm)
0.5
1
2
4
6
8
10
Total Volume
(mL)
50
50
50
50
50
50
50
The standards were read against a DI water blank set at zero absorbance. The absorbance
of each standard was recorded and related to its concentration by linear regression. An
example set of concentrations and respective absorbances and the calibration curve are
included in Table 3.13 and Figure 3.19.
Table 4.13 Fe(II)-Phenanthroline Concentrations and Absorbances
Fe(II)-Phen
Concentration
(mg/L)
0.5
1
2
4
6
8
10
Absorbance at
510 nm
0.056
0.117
0.231
0.467
0.700
0.932
1.190
When analyzing samples from microcosm bottles, samples were filtered with a 0.20 µm
filter to remove organic contaminants and ferric iron colloids. Since Standard Methods
warns that other dissolved metals present in the sample may interfere with the color
development by complexing with phenanthroline, an excess of phenanthroline was used
to ensure that color development was not impeded by interfering metals and that
phenanthroline was not the limiting reagent in the complexation reaction. Since the
77
sample itself is a pink color due to the presence of resazurin, the blank used to zero the
spectrophotometer is a sample of water from the microcosms diluted in the same manner
as the test sample, but without the phenanthroline. This compensated for the color
present due to the resazurin and other the reagents used to promote the complexation
reaction.
1.2
1.0
A bsorbance @ 510 nm
y = 0.1184x - 0.0052
R2 = 0.9998
0.8
0.6
0.4
0.2
0.0
0
2
4
6
8
10
Conc Fe(II)-Phen (ppm)
Figure 4.19 Fe(II)-Phen Calibration Curve
During sampling periods, the iron, mixed, and unamended microcosms were analyzed for
increases in Fe(II) concentration. The iron and mixed microcosms were both amended
with ferric iron because iron reduction was anticipated for these conditions. The
unamended microcosm acted as an iron control; analyzing these microcosms would
indicate if an increase in ferrous iron occurred in a microcosms not amended with ferric
iron, indicating that increased Fe(II) concentration was due to naturally-occurring ferric
iron in the soil or groundwater rather than iron addition.
78
For sample analysis, 10-mL aliquots from each microcosm were acidified with 0.2 mL of
concentrated HCl. The sample was pipetted up and down briefly to mix. Once mixed, 5
mL of acidified sample was removed and added to a 10-mL volumetric flask, to which 2mL phenanthroline solution and 1-mL ammonium acetate buffer were added and then
diluted to 10 mL. The flask was inverted ten times to mix. Since ferrous iron is oxygen
sensitive and will oxidize to ferric iron with extended exposure, the samples had to be
read within 5 – 10 minutes of mixing. To ensure the samples were all within the range of
the calibration curves at sampling time, several dilutions were made to the initial 10-mL
samples before acidification. Typically 10 % and 50 % dilutions were made, but were
only read if the undiluted sample was outside of the quantitation limits of the calibration
curve.
3.7.3 Manganese Analysis by the Formaldoxime Method
The formaldoxime method, as outlined by Brewer and Spencer (1971), allows for
detection of Mn(II) without interference from other ionic manganese species in solution.
Brewer and Spencer’s article describing the method is included in Appendix D. The
method is stable and sensitive compared to other methods of analyzing Mn(II) and is
suitable for environmental samples with numerous dissolved constituents.
Formaldoxime, a solution comprised of formaldehyde and hydrochloride, reacts with
Mn(II) to form a red-purple colored complex. Beer’s Law linearly relates the intensity of
the color formed to the concentration of Mn(II). The complex can be detected using a
UV spectrophotometer at 450 nm with detection limits at 10 µg/L Mn(II).
79
All materials used in the formaldoxime method were low in iron content and of
appropriate grade for metal analysis. The three reagents required were 10 %
hydroxylamine hydrochloride, 37 % formaldehyde (Fisher Scientific, F79), and
ammonium hydroxide. Formaldoxime solution was made by combining hydroxylamine
hydrochloride and formaldehyde in a 20:1 ratio. Ammonium hydroxide was diluted 1:10
with deionized water (DI water), then this was combined with the formaldoxime in a 2:1
ratio of formaldoxime to dilute ammonium hydroxide. 0.5 mL of this mixture was added
to 3.5 mL of sample. Since pH needed to be within the range of 8.8 – 8.9 for proper color
development, ammonium hydroxide diluted to a 1:5 ratio with DI water was added by
dropwise addition while monitoring pH with an digital pH meter (Cole Palmer, Model
59003-00). At this point, color would begin to develop; once color development was
stable, the color development was measured using a colorimeter at 450 nm. Sample color
is stable for a period of 60 minutes after full color development.
Mn(II) concentrations in laboratory microcosms were determined by comparison to a
calibration curve created prior to the sampling date. The standards for the calibration
curve were made using a stock 2000-ppm Mn(II) standard reagent (VMR Scientific, CV
8537). The 2000-ppm standard reagent was used to create a 10-ppm Mn(II) solution by
adding 5 mL of 2000-ppm Mn(II) standard reagent to a 100-mL volumetric flask, then
diluting with DI water. This 10-ppm Mn(II) solution was then used to create additional
standard solutions used for the calibration curve. A typical dilution scheme is seen in
Table 3.14.
80
Table 4.14 Mn(II) Dilution Series for Calibration Curve
Volume of 10 ppm
Mn(II) Solution
(mL)
5
10
20
30
40
50
Total Volume
(mL)
50
50
50
50
50
50
Final Mn(II)
Concentration
(ppm)
1
2
4
6
8
10
Once an appropriate dilution series was created, it was necessary to determine an
appropriate incubation time for full, stable color development. Visual inspection
revealed that color intensity increased with time, thus it was necessary to determine if the
color development remained linear with time as well as to determine the time when the
color development was most stable. To accomplish this, the Mn(II) dilutions were
developed with the formaldoxime-ammonium hydroxide mixture and then allowed to
develop for four different time periods – 0, 2, 10, and 30 minutes, measured with a
laboratory stop watch. After appropriate time development, the sample absorbance of
450 nm wavelength light was determined with a Hitachi U-3010 UV/VIS
spectrophotometer. The results, shown in Table 3.15 and Figure 3.20, indicate that color
development was rapid at first and began to level out for all concentrations with increased
incubation time. Though all development times had acceptable R2 values, ranging from
0.9073 to 0.9925, 10 minute development time appeared to have the most stable color
development, as shown in Figure 3.21. Thus a 10-minute development time was utilized
for all sample analyses during this experiment.
81
Table 4.15 Mn(II)-Formaldoxime Absorbance as a Function of Concentration and
Development Time
Concentration
(ppm)
1
2
3
4
5
0 Minutes
0.021
0.032
0.075
0.121
0.22
Absorbance
2 Minutes 10 Minutes
0.088
0.198
0.135
0.321
0.202
0.442
0.32
0.618
0.43
0.707
30 Minutes
0.202
0.387
0.503
0.745
0.842
1
Absorbance @ 450 nm
0.8
0.6
0.4
0.2
0
0
5
10
15
20
Time (min)
5 ppm
25
4 ppm
3 ppm
30
2 ppm
35
1 ppm
Figure 4.20 Change in Mn(II)-Formaldoxime Development with Time
At each sampling event, the manganese and unamended microcosms were analyzed for
Mn(II) concentration by the formaldoxime method. The manganese microcosms were
amended with MnO2, and Mn(II) concentrations were expected to increase with time if
manganese reduction were occurring. The unamended microcosms act as an manganese
control; analyzing these microcosms indicated whether an increase in Mn(II) occurred
without manganese addition, demonstrating that increased Mn(II) concentration was due
82
to naturally occurring manganese oxides present in the soil or groundwater rather than
MnO2 addition.
Absorbance @ 450 nm
1
0.9
30 min
y = 0.1638x +
0.0444
0.8
R2 = 0.9858
0.7
0.6
10 min
y = 0.1315x +
0.0627
0.5
R2 = 0.9925
0.4
0.3
2 min
y = 0.0869x 0.0257
0.2
R2 = 0.9688
0.1
0 min
y = 0.0487x 0.0523
0
0
1
2
3
4
5
6 R2 = 0.9073
Concentration Mn(II) [ppm]
0 min
2 min
10 min
30 min
Figure 4.21 Mn(II)-Formaldoxime Absorbances as a Function of Concentration,
Measured at Four Development Times
All samples were filtered with a 0.2 µm filter before beginning analyses. Filtration
removed any suspended organic material or solids that could potentially interfere with the
formaldoxime method. To ensure that sample concentrations did not exceed the limits of
the calibration curve, two dilutions were made using DI water – 1:1 and 1:4 ratio of
sample to DI water. The diluted samples were recorded only if the non-diluted sample
was out of range of the standard curve. At each sampling event, 0.5 mL of formaldoxime
mixture is added to 3.5 mL of sample. The pH was adjusted to 8.8 – 8.9 immediately
using dropwise addition of 1:5 dilution of ammonium hydroxide in DI water while
monitoring pH with a digital pH meter (Cole Palmer). Once sample pH was corrected,
83
samples were incubated in open air at room temperature for 10 minutes to allow color to
develop. Once developed, samples were read using a UV/VIS Spectrophotometer at 450
nm.
3.8
Microtox® Toxicity Analysis
Microtox® is a biosensor-linked assay used for rapid determination of chemical toxicity
for a variety of organic and inorganic contaminants. Microtox® is favored over more
traditional animal-based testing procedures because it is fast, reliable, inexpensive, and
humane. The method allows toxicity testing to be performed in by modest laboratories,
eliminating the need to send samples out to labs for further toxicity analysis. Within an
experimental setting, Microtox® provides a simple, effective method to monitor changes
in sample toxicity during the duration of an experiment.
The main drawback associated with the Microtox® method is the lack of correlation
between toxicity to microorganisms and toxicity to species of concern due to the
differences between prokaryotic and eukaryotic metabolic pathways. However, linear
regression relating LC50 data from studies using fathead minnow and Microtox® studies
using Photobacterium phosphoreum had a reasonable correlation (r = 0.81) when
compared for over 200 individual compounds (Kaiser and Palabrica,1991). Kaiser and
Palabrica (1991) reported that Photobacterium phosphoreum results had poor correlation
with rat toxicity data, but stated that rat toxicity data rarely has good correlation with
other parameters, such as biological endpoint or physiochemical coefficients. In support
of the method, Chang et al (1981) state that Microtox® can detect the effects of a variety
84
of biologically important or toxic molecules with more sensitivity than other techniques
due to the involvement of enzymes and proteins required for biolumniscence.
The method has received recognition in the scientific community, becoming a US EPA
approved method for Whole Effluent Toxicity Testing under the National Primary and
Secondary Drinking Water Standards and addition to the ASTM Standard Methods for
determining toxicity of aqueous wastes before and after treatment. The Microtox®
method utilizes a fluorometer (M500 Analyzer, SDI) remotely controlled by Microtox
Omni software.
The Microtox® suite of testing methods is extensive, but for this experiment, only the
Basic Test was used. The Basic Test necessitated four samples of decreasing sample
concentration (45 %, 22.5 %, 11.25 %, and 5.625 % sample) and a control. The control
was used to determine the change in luminescence without exposure to sample. The
change in the sample luminescence over time is used to correct the experimental samples
thereby assuring only the change in metabolic rate due to chemical exposure is
considered when calculating the effect of exposure on the bacteria. The Basic Test
measures light intensity of the bacteria at three time periods – before exposure, after 5
min exposure, and after 15 min exposure. If desired, the user can alter these exposure
times and the sample concentrations.
The Microtox® Method involves analyzing the change in light emission by a
bioluminescent bacterial species, Vibrio fischeri, after exposure to a sample of known
85
concentration. Since the bioluminescense is a byproduct of their metabolism, intensity of
light produced is directly proportional to their metabolic rate, which is directly linked to
the toxicity of their environment. Change in luminescence was reported as percent effect
(% effect). The % effect is mathematically defined as:
% Effect =
Iot − It
It
(6)
where Iot is the corrected baseline light intensity and It is the measured light intensity after
t minutes exposure to sample. Iot is corrected for each test sample by comparison to a
change in the control under the assumption that any change in control luminescence
indicates a change in metabolic activity unrelated to the sample. The baseline light
intensity was calculated by:
⎡I ⎤
Iot = Io⎢ ct ⎥
⎣ Ico ⎦
(7)
where Ict is the luminescence of the control sample at time t, Ico is the measured
luminescence of the control sample at time t = 0 min.
The measured light intensities are used to calculate an EC50 for the sample. The EC50 is
analogous to LD50 used in chemical toxicity determinations, and indicates the
concentration at which a 50 % reduction in activity was noted in a measured quantity – in
this case, bioluminescence. To calculate the EC50, Microtox® Omni software introduces
a parameter called gamma (Γt), mathematically defined as:
86
⎡I ⎤
Γt = ⎢ ot ⎥ −1
⎣ It ⎦
(8)
By algebraic manipulation, % effect can be calculated in terms of Γt rather than light
intensity:
⎛ Γ ⎞
% effect = ⎜ t ⎟ ×100%
⎝ Γt + 1⎠
(9)
When the light intensity is reduced by 50%, the factor Iot/It = 2 since the sample
concentration is half of its baseline intensity. Therefore, the EC50 concentration can be
determined by setting Γt = 1, or by setting Iot = 2It.
To determine EC50, Microtox® Omni software plots the log10 of Γt versus log10 of
concentration on a log-log plot for each sample time. The points are automatically fitted
to a linear regression. Once this regression is created, the EC50 can be extrapolated or
interpolated from the graph by setting Γt = 1 and determining the concentration where
this occurs. The linear regression produced by Microtox® Omni is of the form:
logC = m logΓt + b
( 10 )
Where Γt is the gamma value produced by the sample concentration at t minutes exposure
and m and b are the slope and y-intercept, respectively, determined by the linear
regression.
The EC50 can be determined by the linear regression equation by setting Γt = 1. Since
log10(1) = 0, this reduces the linear equation to:
87
log EC50 = b
EC50 = 10 b
( 11 )
It is important to note at this point that C is the percent concentration, not the total
concentration. Since the sample was diluted in order to perform the basic test, the C
generated by the software was the percent concentration relative to the sample used. The
initial concentration of your sample must be known to report the final EC50. For example,
if the initial sample contained 6 mg/L TPH in water and the EC50 determined was 25 %,
then the EC50 concentration was 1.5 mg/L TPH in water.
This linear regression equation can also be used to calculate the percent effect of an
undiluted sample. To do this, set C = 100. Then log10(C) = log10 (100) or 2. The linear
regression equation can then be modified to solve for Γt.
log10 C = m log10 Γt + b
2 − b = m log10 Γt
log10 Γt =
Γt = 10
2−b
m
( 12 )
2−b
m
This equation can then be used to calculate the percent effect of an undiluted sample
using equation (7).
Microtox® Omni software calculates percent effect and EC50 for samples after 5 and 15
minutes exposure. For the purpose of this experiment, the greater of the two values was
recorded.
88
3.9
Terminal Restriction Fragment Analysis
Terminal restriction fragment (TRF) analysis, also known as Terminal Restriction
Fragment Length Polymorphism (T-RFLP), is a molecular method of monitoring the
dynamics of microbial populations (Kitts 2001). This method uses DNA extraction,
polymerase chain reaction (PCR), capillary and gel electrophoresis, enzymatic digestion,
and fluorimetry. One of the primary advantages of the method is the digital data
produced, which is easily imported into a spreadsheet and statistical software, thereby
eliminating the need for manual data reduction. This is especially helpful when dealing
with complex microbial communities present in contaminated soil systems, where the
microbial populations are extremely diverse. It also helps reduce the impact of culture
bias where certain bacterial species are easier to cultivate than others. This is particularly
important in an anaerobic experiment since many of the target organisms may be difficult
to culture, fastidious in their nutrient and carbon-source requirements, or extremely slowgrowing, reducing the likelihood of growing them in a laboratory setting for further
identification. A TRF pattern generated from groundwater is included in Figure 3.22.
To produce TRF patterns, the DNA was extracted from the cells and purified. Once
extracted, the DNA was amplified by PCR using primers specific to a range of targeted
genes. One primer was fluorescently labeled on the 5’ end of the DNA, allowing further
analysis. After amplification, the DNA was cut by a restriction enzyme targeting a four
base-pair sequence in the DNA. Depending on the enzyme used and the species present,
the DNA may be cut once, multiple times, or not at all, creating DNA fragments of
multiple lengths. These fragments were then analyzed using capillary electrophoresis.
89
Similar to gel electrophoresis, capillary electrophoresis utilized the negative-ionic charge
associated with DNA to move the fragments along an electrical gradient. The fragments
were moved through the gel medium at rates relative to their size-to-charge ratio: smaller
fragments eluted faster than larger fragments. The relative abundance of a particular
fragment length was resolved by differences in peak area in the generated TRF patterns.
Once completed, these patterns were used to compare initial and final microbial
populations by noting changes in relative abundance with time as well as emergence or
disappearance of certain peaks. An example of the process is shown in Figure 3.23.
14000
135.81
13000
12000
11000
10000
9000
74.15
8000
7000
151.19
6000
176.76
5000
343.13
248.43
4000
76.59
3000
2000
226.90
277.02
493.71
532.82
1000
-0
50
100
150
200
250
300
350
400
450
500
550
Size in Base pairs
Figure 4.22 TRF Pattern Generated from Initial Groundwater Sample Collected
from the GRP Site at J8-11
90
Figure 4.23 Basics of Creating TRF patterns – DNA Labeling, Enzyme Digestion,
and Fragment Analysis
3.9.1 Sample Filtration
Groundwater samples were first concentrated by filtration before conducting any further
analyses. Water samples in the microcosms were filtered using a vacuum pump
(M100GX, Fisher Scientific) and 0.2-µm nylon membrane filters (Schleicher and
91
Schuell). The 0.2-µm filters were placed atop Watman paper backing filter to ensure that
the suction produced by the vacuum pump did not tear the finer filter. The entire 2-L
sample was filtered. Soil present was allowed to collect on the sample. Once the entire
sample was passed through the filter, the filter and any filtrate trapped on it were wrapped
in aluminum foil and stored in a -20 °C freezer until further analyses.
3.9.2 DNA Extraction
The first step associated with creating TRF patterns was DNA extraction and purification.
This was accomplished using MoBio Power Soil DNA Extraction Kit, a suite of
proprietary chemicals and equipment used in series to remove DNA from the cells and
separate DNA from other cellular and extracellular material present in the
microorganisms or the soil sample itself. MoBio Power Soil DNA Kit provides media
necessary for bead beating to homogenize and lyse the cells in the sample, followed by a
series of reagents to purify the DNA by precipitating contaminants. The specific steps
and reagents used are explained in the TRF protocol developed by the Environmental
Biotechnology Institute, California Polytechnic State University, San Luis Obispo, CA,
included in Appendix E.
Once extracted, DNA was visualized to ensure the extraction was successful, which was
accomplished using gel electrophoresis.
92
3.9.3 PCR Using Fluorescently-labeled Primers
Polymerase Chain Reaction (PCR) is a technique used in molecular biology to amplify
the DNA of interest present in a sample. PCR is conducted using an Applied Biosystems
GeneAmp® PCR System 9700 thermal cycler, a device that heats and cools samples to
specific temperatures for specified time intervals. The cycling temperatures control DNA
denaturation, elongation, and annealing.
The initial step was DNA denaturation, during which the thermal cycler raised the
temperature to denature the double-stranded DNA helix, resulting in two strands of DNA.
Annealing followed denaturation, during which the temperature was lowered. At this
temperature, the primers adhere to the two DNA strands. The types of primers used
selected for specific genes, gene fragments, or sequences, controlling the section of DNA
amplified during the process. Additionally, the forward (5’-to-3’) primers utilized in
TRF analysis had a Cy5 fluorescent tag used to detect DNA fragments and create the
TRF pattern. The final stage, elongation, was the step when new DNA was synthesized
by complementary base pairing to the template strand. DNA polymerase mediated
synthesis, adhering to the single-stranded DNA at the primer and continuing along the
strand in the 5’-to-3’ direction. This process of denaturation, annealing and elongation
consisted of one cycle; the thermal cycler repeated this process for 25 – 30 cycles,
depending on the parameter of interest. This process is depicted in Figure 3.24.
Mathematically, this translates into 2n fluorescently labeled double-stranded DNA
segments, referred to as the “short product” because they are only the desired segment of
the full DNA strand, where n is the number of cycles utilized in the thermal cycler.
93
To conduct PCR, several reagents are needed. These are as follows:
•
Primers. These are short segments of DNA recognizing the specific codons of
interest and allowing replication to proceed with the specific enzymes. Primers
used in developing TRF patterns utilize fluorescently labeled forward primers.
Different primers are used depending on the bacterial species of interest. In our
experiment, four different types of primers were used: a general microbial count
was gathered by focusing on the DNA region that codes for 16S RNA, and
specific primers are used to enumerate methanogens, Archaea, and chloroflexi.
•
DNA Template. This contains the region of DNA to be amplified. Similar to the
primers, the DNA template was varied depending on the region of DNA or target
organism.
•
Enzyme. The enzyme mediates DNA replication during the elongation stage.
Taq polymerase was utilized for all PCR runs conducted in this experiment.
•
Deoxynucleotide triphosphates (dNTP). These are the nucleotides Taq
polymerase uses to form the complementary DNA strands. This mixture contains
all four DNA nucleotides – adenine, guanine, cytosine, and thiamine – in equal
proportion.
•
BSA. This additive aids PCR in various ways. BSA binds proteins and other
compounds in solution that otherwise interfere with the process. BSA coats the
walls of the PCR tube, preventing the DNA template from being adsorbed and
reducing “primer-dimer,” a phenomenon where non-target amplifications occur
due to low template concentration. Finally, BSA can help to stabilize the Taq
polymerase.
94
Figure 4.24 Polymerase Chain Reaction Stages - Denaturation, Annealing, and
Elongation
95
•
Divalent cations. Typically manganese or magnesium, divalent cations aid Taq
polymerase attachment to DNA. Divalent cation concentration is altered to
optimize PCR depending on the cleanliness of the product desired. In general, the
less concentrated the divalent cations are, the more specific the PCR result is.
MgCl2 was used as a source of divalent cations for all PCR runs conducted.
•
Buffer. The buffer helps maintain ideal pH for optimal Taq polymerase activity
and stability. A 10X gold buffer was used in all PCR runs conducted.
•
PCR Water. Reagents were diluted to a final volume with PCR water. The
volume used depends on the amount of DNA added. The total volume was 40 µL
in all PCR runs conducted.
For each run conducted, PCR reagents were combined in a UV-sterilized clean room.
Positive and negative controls were utilized. The positive control, which varied
depending on the target DNA, utilized known species DNA to confirm that the PCR was
successful. For example, 16S primers used E. coli as a positive control. The negative
control did not have any DNA added; if any bands were visible in a confirmation gel run
after the PCR is complete, then PCR reagents were contaminated and had to be repeated.
Similar to the primers, the thermal cycler’s program depended on the target DNA
segment or species. Once the program was completed, gel electrophoresis was conducted
to determine if the PCR was successful. If successful, all PCR products, including the
positive control, had bright bands under visualization, while the negative control did not
have any bands.
96
To ensure that enough DNA was present to produce TRF patterns and to minimize
variation, PCR was run in triplicate for each sample. The triplicates were combined
during PCR cleanup. The cleanup removed excess divalent cations, primer, template, and
dNTPs from the solution, as these interfered with the TRF patterns. PCR cleanup is
conducted using MoBio PCR Ultra-Clean kit. The method used for the cleanup is
included in Appendix E.
Once cleaned, the PCR product was quantitated using a FLx 800 Microplate Fluorescence
Reader (Fluorometer, Bio-Tek Instruments), controlled remotely using KC4 Fluorometer
software. The fluorometer measured the luminescence of the Cy5-labeled forward
primer. Once measured, the fluorometer software estimated the DNA concentration in
the PCR product by comparison of readings to standard curves. This estimation was used
to determine the volume of DNA added to the enzyme digest.
3.9.4 Production of Labeled Fragments by Enzyme Digestion
Once amplified DNA segments were cleaned and quantified, the segments were digested
into smaller fragments by a restriction enzyme. The restriction enzyme fractured the
segments based upon a four-nucleotide sequence; the enzyme cut the DNA segment
wherever the specific sequence was located, creating a broad variety of fragments due to
genetic diversity. DNA segments can be fragmented once, twice, multiple times, or not
at all depending on how many times the sequence is present.
97
Numerous enzymes are available for use, each having a different tetranucleotide
recognition sequence. For example, one of the restriction enzymes used, Dpn III,
recognizes the basepair sequence guanine-cytosine-guanine-cytosine (GCGC). Since the
fragments produced are not species specific, performing multiple digests with different
restriction enzymes can aid in species identification. For this experiment, 16S product
was digested with Dpn III, Hae III, and Hha, Methanogen product was digested with
SauI96, and Archaea product was digested with Hae III. Digestion products were stored
at -20 °C until further analyses.
Before proceeding with generating TRF patterns, the digested DNA was cleaned of
excess salt through ethanol precipitation. The cleaning agent combined cold 95 %
ethanol, 3 M sodium acetate solution, and glycogen (20 mg/mL) in a 100:2:1 ratio. 100
µL of this mixture was added to the digest, inverted to mix, and incubated in a freezer at 20 °C for 30 min. Tubes were then centrifuged at 5,300 rpm for 15 minutes, causing the
DNA to form a pellet in the base of the tube. The ethanol solution was removed by
inverting the tubes and lightly tapping out the fluid. Once most of the solution was
removed, an additional 100 µL of cold 70 % ethanol is added to the tubes and centrifuged
for 5 min at 5,300 rpm. The ethanol was removed by inverting the tubes and lightly
tapping out the fluid. Tubes were placed into a fume hood for 30 minutes to allow the
remaining ethanol to evaporate. Precipitated DNA fragments were stored at -20 °C until
further analyses.
98
3.9.5 TRF Pattern Generation by CEQ-8000
TRF patterns were generated by capillary electrophoresis using the Beckman Coulter
CEQ-8000 Genetic Analysis System, comprised of capillary electrophoresis hardware
remotely controlled by system software. The precipitated DNA was resuspended by
adding 2 µL formamide and 0.25 µL of 600 base pair standard, an internal standard used
by the CEQ-8000. Once mixed, a drop of mineral oil was placed on the solution to
ensure that the samples did not dry out during the analysis. Samples were then run in a
CEQ-8000. Raw data generated from the CEQ-8000 consisted of peaks at the fragment
lengths that represent the relative abundance of microorganisms present in the sample
(Kitts, 2001). The resulting patterns were statistically analyzed to determine which peaks
contribute to similarities and dissimilarities between the samples.
3.9.6 TRF Pattern Analysis
Once these fragments were generated, the data was realigned using Microsoft Excel to
align peaks and to set a new data threshold. Patterns developed were compared sets to
group similar fragment lengths generated by the different restriction enzymes.
The CEQ-800 Genetic Analysis System was used to define the data threshold. Results
were then analyzed using macros in MS Excel and statistically analyzed using Primer-5
software (Primer-E Ltd). The data threshold was 1 %, eliminating all peaks that
contributed less that 1% of the total peak area. This removed background noise from the
analysis, since these small peaks did not sizably contribute to the total community (Rees
99
et al, 2004). Once the data were aligned and noise reduced, the similarity was calculated
using Bray-Curtis coefficient:
n
⎧
⎫
y ij − y ik ⎪
⎪
∑
⎪
⎪
S jk = 100 ⎨1− ni=1
⎬
⎪
(y ij + y ik )⎪⎪⎭
⎪⎩ ∑
i=1
( 13 )
where i was the peak function compared across j and k samples (Rees, 2004) and yij and
yjk is one set of n attributes. The degree of similarity calculated between the different
microcosm conditions was then used to create a dendrogram. A dendrogram is a treecluster facilitating the visualization of clusters produced by the Bray-Curtis similarity
calculation. The connection point between two samples is related to the average percent
similarity between them; grouping of clusters represents the relative similarity of a group
of samples. For final presentation, the dendrograms produced by Primer V and the
electropherograms produced by the CEQ 8000 were combined to facilitate the
comparison of individual TRF patterns as they relate to similarity between the
microcosms.
100
CHAPTER 5 .
4.1
RESULTS AND DISCUSSION
Microcosm Integrity
4.1.1 Redox Indicator Color
The redox indicator resazurin was added to the water used to establish all microcosms in
order to test for oxygen exposure during the duration of the experiment. A color change
from colorless to pink would indicate an increase in redox potential and could indicate
that oxygen contamination had occurred. Table 4.1 lists the colors of the microcosm
bottles during the establishment period and each sampling event. The groundwater was
colorless when pumped out of the barrel and along the entire length of tubing into the
glovebox, indicating that the groundwater was successfully transported under anaerobic
conditions and the mass transfer into the tubing had no measurable effect on oxygen
concentration.
Aerobic microcosms turned pink as expected due to oxygen purging during their
establishment. Conversely, unamended microcosms remained colorless during the length
of the experiment, indicating that the groundwater and soil used were anaerobic during
microcosm establishment and were not exposed to oxygen. The nitrate and manganese
oxide amended microcosms turned pink when established, most likely due to the high
redox potential of these amendments. Since the reduction potential for resazurin to
change color (E = – 0.42 mV) is more reduced than the reduction potential for manganese
or nitrate (E = 0.612 mV and E = – 0.30 mV, respectively), microcosms amended with
101
nitrate or manganese were expected to develop a pink color. The sulfate and ironamended microcosms turned slightly pink when established but because colorless before
the 26-day sampling date. The initial color change in sulfate and iron amended
microcosms was not expected, since these microcosms should have been sufficiently
reduced to prevent the color change to pink. A selection of microcosms is illustrated in
Figure 4.1.
Table 5.1 Microcosm Water Color Due to Redox Indicator
Microcosm
Day 0
Day 26
Day 134
Condition
Mn(IV)-1
Pink
Pink
Pink
Mn(IV)-2
Pink
Pink
Pink
Fe(III)-1
Pink-Orange
Yellow/Brown Colorless
Fe(III)-2
Pink-Orange
Yellow/Brown Colorless
Mix-1
Layered Pink/Rust
Pale Yellow
Bright Pink
Mix-2
Layered Pink/Rust
Pale Yellow
Bright Pink
SO4-1
Light Pink
Colorless
Colorless
SO4-2
Light Pink
Colorless
Colorless
NO3-1
Bright Pink
Bright Pink
Bright Pink
NO3-2
Bright Pink
Bright Pink
Bright Pink
Unamended-1 Colorless
Colorless
Light Pink
Unamended-2 Colorless
Colorless
Colorless
O2-1
Bright Pink
Bright Pink
Bright Pink
O2-2
Bright Pink
Bright Pink
Bright Pink
Control-1
Pink
Pink
Pink
Control-2
Layered Pink/Colorless Pink
Pink
** Final Oxygen microcosms observed on 298 day, not 407 day.
Day 407
Pink
Pink
Colorless
Colorless
Light Pink
Light Pink
Colorless
Slight Pink Tint
Bright Pink
Bright Pink
Colorless
Colorless
Bright Pink**
Bright Pink**
Slight Pink Tint
Light Pink
Though oxygen contamination was an obvious culprit, the gas headspace data indicated
that oxygen concentration was less than 0.01 % in anaerobic microcosms during the
sampling events (Section 4.2 Gas Headspace Data). Furthermore, since the unamended
microcosms remained colorless, the groundwater used to establish the microcosms was
102
not contaminated with oxygen at the time microcosms were established. Another source
of oxygen may have been the electron acceptor amendments.
Figure 5.1 Microcosm Color Spectrum at Day 26 Sampling Event. From Left to
Right: Mixed Amendment, Iron, Manganese, and Unamended Microcosms.
The electron acceptor amendment solutions were made with DI water that was not
anoxic, contributing trace amounts of oxygen. It is possible the oxygen content of the DI
water was sufficient to induce color change in all amended microcosms and offers
possible explanation why only the unamended microcosms remained clear during
establishment. Since all microcosms amendment volumes were minimal in comparison
103
with the bottle volume, the amount of oxygen contributed should not have been sufficient
to compromise anaerobic conditions.
4.1.3 Acidic pH in Iron-Amended Microcosms
The pH of all microcosms during the course of this experiment are listed in Table 4.2.
Microcosm pH was not measured at the initial sampling date.
Table 5.2 Average Microcosm pH at Sampling Dates
Microcosm
Condition
Mn
Fe
Mix
NO3
SO4
Unamended
0 Day**
pH
NA
NA
NA
NA
NA
NA
NA
26 Day
pH
7.77
2.67
2.68
8.24
8.19
8.15
134 Day
pH
7.65
7.58
7.55
8.22
8.28
8.09
407 Day
pH
7.9
7.47
7.51
8.03
7.78
8.23
-
O2
7.60
7.53
NA
Control
8.27
8.17
7.9
** Samples were not analyzed for pH on Day 0
At the 26-day sampling event, the pH of iron-containing microcosms was measured at
approximately 2.6. The pH reduction was attributed to the formation of ferric hydroxide,
producing 3 moles of hydronium cations for each mole of ferric iron, thereby exceeding
the buffering capacity of the groundwater.
Fe 3+ + 3H 2O → Fe(OH) 3 + 3H +
( 14 )
Once the source of the acidity was determined, a side experiment was conducted to
determine the volume and concentration of alkaline solution needed to correct the pH.
The pilot-scale test was performed at 1:10 scale using 0.2 L groundwater, 5 g sand, and
104
0.169 g FeCl3 dissolved in DI water. Groundwater pH was recorded initially, after sand
addition, and at time intervals after iron addition. In four trials, the pH was reduced to 3
when FeCl3 was added at the same concentration as used in the iron-amended
microcosms. The acidic pH in the iron-amended microcosms was adjusted as described
in Section 3.4.5.
4.1.3 Nitrogen-to-Helium Ratios for Leak Detection
The nitrogen to helium ratio in the microcosm gas headspace reflects the integrity of the
seal provided by the crimp seal and its ability to contain headspace gases. Nitrogen and
helium headspace data for all microcosm replicates are listed in Table 4.3. The initial
N2/He ratio in the microcosm was approximately 20:1. In most microcosms, N2/He ratios
were maintained near 20:1 for the duration of the experiment. One unamended
microcosm had a very high ratio at the 134-day sampling event (725:1), indicating that
the microcosm leaked during incubation. This microcosm was excluded from subsequent
gas headspace analyses. Oxygen microcosms had very low N2 content due to the oxygen
purge used during their establishment. In these microcosms, the N2:He ratio ranged from
0.24:1 to 0.68:1 during the course of the experiment. When the iron-containing
microcosms were opened to correct the pH drop, their N2/He ratios were re-established at
a higher nitrogen to helium ratio because a lower He flow rate was used. After the pH
correction, the N2/He ratios in the iron-containing microcosms were approximately 156:1.
Maintaining a stable N2:He ratio indicates that leakage from the crimp seals was minimal
in most microcosms during the 407-day incubation period.
105
Table 5.3 Nitrogen-to-Helium Ratios for All Microcosms at Sampling Each Date
Microcosm
Condition
Mn(IV)
Fe
Mixed
SO4
NO3
Unamended
O2
Control
Mn(IV)
Fe
Mixed
SO4
NO3
Unamended
O2
Control
He
ppm
0 Day
46601
43359
48991
34581
46688
N2
%
N2:He
91.2
91.6
85.1
87.7
89.6
19.6
21.1
17.4
25.3
19.2
51243
35369
45209
12926
69531
69048
372097
253009
44628
90.3
91.4
91.6
87.1
90.7
90.0
10.6
6.8
91.8
17.6
25.8
20.3
67.4
13.0
13.0
0.3
0.3
20.6
26 Day
48517
46185
42788
44434
44356
43107
51243
35369
47163
46377
69295
71915
402442
409896
46758
48116
92.4
92.9
85.3
85.4
85.6
85.3
91.7
92.5
92.4
93.0
90.4
89.8
21.1
22.1
92.3
92.8
19.0
20.1
19.9
19.2
19.3
19.8
17.9
26.2
19.6
20.0
13.0
12.5
0.5
0.5
19.7
19.3
106
Table 4.3 Nitrogen to Helium Ratios for All Microcosms
at Sampling Dates (Continued)
He
N2
Microcosm
N2:He
Condition
ppmv
%
Mn(IV)
Fe
Mixed
SO4
NO3
Unamended
O2
Control
Mn(IV)
Fe
Mixed
SO4
NO3
Unamended
134 Day
43283
42624
4730
5323
87356
7219
54118
53808
42814
43929
695
59157
300989
460411
42954
42633
407 Day
42397
41070
5067
7505
7173
6396
52914
46247
41722
41165
61745
48696
94.8
94.8
98.8
99.1
98.4
98.7
93.7
93.7
94.8
94.4
99.8
94.5
24.0
26.4
95.1
95.2
21.9
22.2
208.8
186.2
112.6
136.7
17.3
17.4
22.1
21.5
1434.7
16.0
0.8
0.6
22.1
22.3
96.6
94.4
97.1
96.9
95.4
97.0
92.6
93.0
96.5
93.3
93.7
94.6
22.8
23.0
191.7
129.0
133.0
151.6
17.5
20.1
23.1
22.7
15.2
19.4
94.6
95.7
23.5
40.1
O2
Control
40211
23889
107
4.2
Total Petroleum Hydrocarbon Results
Figure 4.2 shows a GC/FID generated chromatogram from the 407th Day sample of one
unamended microcosm. The large spike eluted at approximately 22.5 minutes was
hexacosane. The “humpogram” from 2 to 22 minutes indicates that this is an unresolved
mixture of hydrocarbons, as expected since the contamination at GRP consisted of
weathered diesel-range petroleum distillate. The lack of spikes in the humpogram
indicates that few n-alkanes were present in the mixture. The elution time of the
humpogram suggested that most of the remaining weathered petroleum compounds are
equivalent to C-20 in carbon length.
Figure 5.2 GC/FID Generated Chromatogram for Mixed-Amendment Microcosm
at the 26th Day Sampling Event
Table 4.4 lists the TPH concentrations in the microcosms on Days 0, 26, 134, and 407 for
all microcosms and replicates. With few exceptions, extractions show good replication.
Averages and standard deviations are listed in Table 4.5. Statistical analyses were based
upon four extractions – two 1-liter extractions were made from each microcosm, and two
microcosms of each condition were sacrificed at each sampling date. TPH fluctuations in
108
all microcosms are depicted in Figure 4.3. Comparative TPH concentrations are depicted
in Figure 4.4, representing the change in TPH concentration for each microcosm
condition as the experiment progressed.
Table 5.4 TPH Concentrations in Microcosm Replicates
Microcosm
Condition
A
Mn(IV)
B
A
Fe(III)
B
A
Mixed
B
A
SO4
B
A
NO3
B
A
Unamend
B
A
O2
B
A
Control
B
TPH Concentration (mg/L)
0 Day 26 Day 134 Day 407 Day
4.39
3.98
4.46
4.87
4.67
4.83
4.37
4.86
5.16
3.66
4.46
4.79
4.02
4.31
4.29
5.89
3.91
5.57
1.92
6.38
3.71
4.52
2.01
5.67
4.91
5.64
4.77
5.50
5.01
5.06
4.28
5.26
5.38
4.81
3.79
6.06
4.94
6.07
4.16
6.21
5.76
3.80
6.30
6.03
3.79
6.33
4.21
4.18
4.96
5.86
4.24
4.72
4.92
6.50
4.67
4.11
4.69
6.01
4.11
4.32
4.76
6.23
4.05
4.20
4.50
5.19
4.29
4.12
4.53
5.17
4.45
4.24
4.71
5.33
4.59
3.84
4.52
5.49
4.23
4.75
4.74
6.08
4.52
4.63
4.69
6.17
4.65
4.19
4.70
6.24
4.54
4.46
4.76
6.10
4.28
1.94
1.37
1.27
4.59
1.94
1.35
1.24
4.93
2.21
1.20
1.26
4.17
2.09
1.19
1.23
4.18
4.64
4.76
6.54
4.49
4.66
4.93
6.23
4.88
4.61
5.01
6.13
4.73
4.30
5.13
6.13
109
Table 5.5 Average TPH Concentrations and Standard Deviations
0 Day
Microcosm Concentration
std dev
(mg/L)
Condition
Mn(IV)
4.56
0.48
Fe(III)
4.38
0.67
Mix
5.53
0.47
SO4
4.31
0.25
NO3
4.34
0.23
Unamended
4.48
0.18
O2
4.49
0.34
Control
4.57
0.31
134 Day
Microcosm Concentration
std dev
(mg/L)
Condition
Mn(IV)
4.39
0.08
Fe(III)
3.25
1.49
Mix
3.89
0.18
SO4
4.83
0.13
NO3
4.56
0.09
Unamended
4.72
0.03
O2
1.28
0.09
Control
4.96
0.15
* This sample was extracted at 298 Days
26 Day
Concentration
(mg/L)
4.20
5.20
5.44
4.33
4.10
4.51
2.05
4.55
std dev
0.50
0.52
0.89
0.27
0.18
0.24
0.13
0.17
407 Day
Concentration
std dev
(mg/L)
5.10
0.53
5.83
0.48
6.22
0.12
6.15
0.28
5.30
0.15
6.12
0.06
*
1.25
0.02
6.26
0.20
Rapid TPH decrease was observed in the aerobic microcosms: TPH decreased 54.3 % at
Day 26 sampling date and 72.2 % at Day 298. In contrast, TPH decreases were not
observed in any anaerobic microcosms or the controls. No statistically relevant
difference in TPH concentrations occurred between the six anaerobic conditions in the
experiment with the exception of the 134-day iron-amended microcosms. However, this
TPH reduction was most likely due to loss of methylene chloride in the iron-colloid
emulsion layer formed during the extraction rather than TPH biodegradation. Ironamended microcosms developed extensive flocs due to the addition of FeCl3, a
commercial coagulant, making extraction very difficult due to the formation of a large
110
emulsion layer between the methylene chloride (MeCl) and aqueous layers. This made
collection of MeCl very difficult – after addition of three 60-mL aliquots of MeCl, only
approximately 30 mL was recovered. This is supported by hexacosane recovery from
these iron-amended microcosms, which was approximately 30 % of that retrieved from
the other microcosms. All other iron-containing microcosms were passed through a
coarse filter prior to extraction with MeCl to prevent loss of TPH in the emulsion layer.
For the 407-day sampling event, the samples from the iron-amended microcosms were
filtered prior to methylene chloride extraction. This minimized the emulsion layer and
resulted in higher TPH concentrations than were recorded during the 134-day samples
(Figure 4.3). Based upon similar hexacosane recoveries in filtered and unfiltered
microcosms, the filtration did not affect TPH concentration.
7.0
TPH Concentration (mg/L)
6.0
5.0
4.0
3.0
2.0
1.0
0.0
0
50
100
150
200
250
300
350
400
Time (day)
Mn(IV)
Fe(III)
Mix
SO4
NO3
Unamended
O2
Control
Figure 5.3 Change in TPH Concentrations in Groundwater Microcosms During
407-Day Incubation
111
7
TPH Concentration (mg/L)
6
5
4
3
2
1
0
Mn(IV)
Fe(III)
Mix
SO4
NO3
Unamend
O2
Control
Conditions
0 Day
26 Day
134 Day
407 Day
Figure 5.4 Comparative TPH Concentrations in All Groundwater Microcosms,
Arranged by Microcosm Condition
Despite minute fluctuations in TPH concentrations at different sampling dates, there was
little statistically significant variation between the samples at the onset of the experiment.
Observed TPH concentrations were higher at 407-days than observed at any previous
sampling date for both the anaerobic microcosms and controls. Since the increase was
consistent for both experimental and control microcosms, this increase is likely due to an
analytical anomaly. High TPH readings across a range of samples measured at a
particular time are common with TPH analyses, and are sometimes referred to as “TPH
inflation”. The cause of this anomaly is unknown. Despite this anomaly, it is clear by
comparing TPH concentrations in the anaerobic microcosms to that of the controls that
TPH concentrations have not changed significantly for any of the anaerobic microcosms
during the course of the 407-day experiment.
112
4.2
Headspace and Aqueous Gas Concentrations
All gas data generated during this experiment are listed in Tables 4.6 and 4.7. Gas datarelated calculations are described in Sections 4.2.1 through 4.2.4.
Carbon Dioxide (PPM)
0
Day
26
Day
134
Day
407
Day
134
Day
407
Day
47
0.01
0.02
0.02 0.151 0.38 0.573
0.4
0.93
0.05
0.02
0.02 0.161 0.42 0.588
0.4
0.35
0.02
0.02
0.02
8.19
9.49 0.438
0.1
1.74
0.05
0.02
0.02
6.67
9.56 0.169
0.3
0.36
0.02
0.01
0.02
3.09
9.88 1.336
1.6
0.02
0.02
0.699
1.1
Control
O2
Unamend
NO3
SO4
Mn(IV)
0
Day
Fe(III)
Oxygen (%)
Mixed
Condition
Table 5.6 Carbon Dioxide and Oxygen Concentrations in All Microcosms and
Replicates at Sampling Dates
0.06
26
Day
0.6
0.02
0.03
0.02
0.07 0.108
0.1
2.43
0.02
0.02
0.02 0.062 0.07 0.114
0.1
0.64
0.01
0.18
0.02
0.07
0.07 0.093
0.1
8.9
0.01
0.02
0.02
0.06
0.07 0.086
0.1
0.75
0.01
0.03
0.03
0.07
0.09 0.154
0.1
0.69
0.02
0.03
0.03
0.07
0.09 0.132
0.1
38.11 23.62 34.81
0.057 0.43 1.774
55.79 23.13 11.93
0.056 0.58 1.766
1.32
0.01
0.02
0.02
0.06
-
0.01
0.02
0.02
-
113
0.07 0.093
0.1
0.06
0.1
0.09
Methane
(ppm)
Hydrogen
(ppm)
26
Day
134
Day
407
Day
0
26 134 407
Day Day Day Day
<5
<5
<5
13
nd
nd
nd
nd
<10 <10 <10
<10
<5
<5
<5
10
nd
nd
nd
nd
<10 <10 <10
<10
<5
<5
<5
<5
nd
150
nd
nd
<10 <10 <10
<10
<5
<5
<5
<5
nd
125
nd
nd
<10 <10 <10
<10
<5
<5
<5
<5
nd
125
nd
nd
<10 <10 <10
542
<5
<5
nd
nd
<10
490
Control Unamend
O2
NO3
SO4
Mn(IV)
0
Day
Fe(III)
0
26 134 407
Day Day Day Day
Nitrous Oxide
(ppm)
Mixed
Condition
Table 5.7 Methane, Hydrogen and Nitrous Oxide Concentrations in All Microcosm
Replicates at Sampling Dates
<5
<5
<5
16
nd
nd
nd
nd
<10 <10 <10
<10
<5
<5
10
19
nd
nd
nd
nd
<10 <10 <10
<10
<5
<5
<5
<5
nd
<10
nd
nd
<10 <10
35
18
<5
<5
<5
<5
nd
<10
nd
nd
<10 <10
55
18
<5
<5
<5
nd
nd
nd
<10 <10 <10
<5
<5
<5
nd
nd
nd
<10 <10 <10
<5
9
<5
94
nd
nd
nd
nd
<10 <10 <10
<10
<5
12
5
48
nd
nd
nd
nd
<10 <10 <10
<10
<5
<5
<5
<5
nd
nd
nd
nd
<10 <10 <10
<10
-
<5
<5
<5
-
nd
nd
nd
-
<10 <10
<10
Note that only one mixed amendment microcosm was analyzed for headspace gases on
the 0-day and 26-day sampling events, only one control microcosm was analyzed for
headspace gas on the 0-day sampling event, and no oxygen microcosms were analyzed on
114
the 407-day sampling event. Additionally, the headspace data for the second unamended
microcosm was omitted from headspace data due to a headspace leak, as noted in Section
4.1.3.
4.2.1 Oxygen Concentration in Microcosm Bottles
Table 4.6 lists the results of oxygen analyses during the microcosm experiment. The
results indicate that anaerobic conditions were maintained in the anaerobic microcosms
during the full extent of the incubation period. Though there were some samples with
high O2 detected at the 0 Day sampling event, Greg Ouellette of Inland Empire
Analytical attributed them to poor sample storage. Since the O2 sensor on the gas
chromatogram used for gas headspace analysis was not operational at the time of
sampling, the samples had to be stored until the sensor could be replaced. This was only
a problem for the Day 0 samples. Overall oxygen levels were extremely low in all
samples and replicates had low standard deviations.
4.2.2 Methane Production
No methane was detected in any microcosms at 0 day sampling event. This confirms that
there was no dissolved methane in the water samples when the microcosms were
established, allowing any future methane detection to be attributed to methanogenic
activity in the microcosms. Methane production was observed in unamended
microcosms at 26, 134 and 407 days, in sulfate-amended microcosms at 134 and 407
days, and in manganese(IV)-amended microcosms at 407 days (listed in Table 4.7 and
115
depicted in Figures 4.5). Methane detection limits in gas headspace were 5.0 ppmv.
Using Henry’s Law, aqueous methane concentrations were calculated from measured
gaseous concentrations to determine the total amount of methane generated in the
microcosm, as displayed in Table 4.8.
Table 5.8 Methane Generation in Gas Headspace and Calculation of Amount of
Headspace and Dissolved Methane in Unamended, Sulfate, and Manganese(IV)
Microcosms
Microcosm
Condition
Unamended
Gaseous Concentration
Aqueous Concentration
mol/L (g) Average
Std Dev mol/L (aq) Average Std Dev
26 Day Samples
3.76E-07
1.22E-08
4.4E-07 8.85E-08
1.4E-08 2.9E-09
5.01E-07
1.62E-08
Sulfate
Manganese(IV)
134 Day Samples
Unamended
Sulfate
2.09E-07
4.17E-07
2.1E-07
6.77E-09
4.2E-07
1.35E-08
6.8E-09
1.4E-08
Manganese(IV)
407 Day Samples
Unamended
Sulfate
Manganese(IV)
9.40E-05
1.60E-05
1.90E-05
1.30E-05
1.00E-05
1.27E-07
9.4E-05
1.8E-05
2.12E-06
1.2E-05
2.12E-06
116
2.17E-08
2.57E-08
1.76E-08
1.35E-08
1.3E-07
2.4E-08
2.9E-09
1.6E-08
2.9E-09
8E-05
Methane Concentration (mol/L)
7E-05
6E-05
5E-05
4E-05
3E-05
2E-05
1E-05
0E+00
0
50
100
150
200
250
300
350
400
450
Time (Day)
Unamended
Sulfate
Manganese(IV)
Figure 5.5 Methane Molarity in Microcosm Headspace Gas
Using bioenergetics (see Section 2.2), the amount of methane produced can be
mathematically related to the amount of TPH that would be consumed if all observed
methane production was attributed to anaerobic fermentation of TPH, as displayed in the
equation below:
0.0263 C6 H14 + 0.00034 NH 4+ + 0.00034 HCO3− + 0.0522 H 2O
→ 0.00034 C5 H 7O2 N + 0.0563 H 2 S + 0.117 CH 4 + 0.0278 CO2
Based on stoichiometry generated using bioenergetics and using hexane as the carbon
source, 4.43 moles of methane would be produced for every mole of hydrocarbon
consumed. This theoretical molar ratio was used to calculate the moles of hydrocarbon
consumed to produce the moles of methane measured in the microcosm gas and dissolved
in the microcosm water. These calculations are listed in Table 4.9.
117
Table 5.9 Bioenergetic Stoichiometry for Hexane Consumed Due to Methane
Production
0.0263
mol/L
Hexane Consumed
Methane Produced
0.1165
mol/L
Ratio
4.430
Microcosm
Condition
Unamended
Sulfate
Manganese(IV)
Unamended
Sulfate
Manganese(IV)
Unamended
Sulfate
Manganese(IV)
Average Concentration Methane Produced
Gaseous
Aqueous
(mol)
(mg)
(mol/L)
(mol/L)
26 Day Samples
4.4E-07
1.4E-08
7.6E-08 1.2E-03
Calculated Hexane
Consumed
(mol/L)
(mg/L)
7.6E-09
6.5E-04
2.1E-07
4.2E-07
134 Day Samples
6.8E-09
3.6E-08
1.4E-08
7.2E-08
5.8E-04
1.2E-03
3.6E-09
7.2E-09
3.1E-04
6.2E-04
9.4E-05
1.8E-05
1.2E-05
407 Day Samples
1.3E-07
9.7E-06
2.4E-08
1.8E-06
1.6E-08
1.2E-06
1.6E-01
2.9E-02
1.9E-02
9.7E-07
1.8E-07
1.2E-07
8.3E-02
1.5E-02
1.0E-02
Based on bioenergetics calculations, approximately 0.083 mg/L, 0.015 mg/L, and 0.010
mg/L of hexane would be necessary to produce the amount of methane measured in the
unamended, sulfate-amended, and manganese(IV)-amended microcosm, respectively,
after 407 days of incubation. These numbers are near or below the practical quantitation
limits of the GC/FID (0.05 mg/L) and would not be considered detectable. Thus, it is
possible that anaerobic biodegradation is resulting in methane production but that the
magnitude of this biodegradation is too small to be detected by GC/FID within 407 days.
118
4.2.4 Nitrous Oxide Production
No nitrous oxide was measured at 0 day sampling in any microcosms. Nitrous oxide
production was measured in nitrate-amended microcosms beginning at 134 days and in
mixed-amendment microcosms at 407 days. All nitrous oxide data are listed in Table 4.7
and depicted in Figures 4.6. The nitrous oxide detection limit in gas headspace was 10.0
ppmv. Nitrous oxide concentrations in aqueous solution were calculated from observed
gaseous concentrations using Henry’s Law, included in Table 4.10.
Nitrious Oxide Concentration(mol/L)
3E-05
2E-05
2E-05
1E-05
5E-06
0E+00
0
50
100
150
200
250
300
350
400
450
Time (day)
Nitrate
Mixed
Figure 5.6 Gaseous Nitrous Oxide Concentration in Nitrate and Mixed Microcosms
119
Table 5.10 Nitrous Oxide Production Observed in Gas Headspace and Calculated in
Aqueous Solution in Nitrate and Mixed Amendment Microcosms
Microcosm
Condition
Nitrate
Mixed
Nitrate
Mixed
Nitrate
Mixed
Gaseous Concentration
Aqueous Concentration
mol/L (g) Average Std Dev mol/L (aq) Average Std Dev
26 Day Samples
8.76E-07
5.08E-07
6.5E-07 3.25E-07
4.5E-07 8.55E-08
4.17E-07
3.87E-07
ND
ND
134 Day Samples
1.46E-06
8.46E-07
1.9E-06 5.90E-07
1.1E-06 3.42E-07
2.30E-06
1.33E-06
ND
ND
407 Day Samples
7.51E-07
4.35E-07
7.5E-07 0.00E+00
4.4E-07 0.00E+00
7.51E-07
4.35E-07
2.26E-05
1.31E-05
2.2E-05 1.53E-06
1.2E-05 8.89E-07
2.05E-05
1.18E-05
Using bioenergetics (see Section 2.2), the amount of nitrous oxide produced can be
mathematically related to the amount of TPH that would be consumed if all of the
observed nitrous dioxide production was attributed to denitrification, as displayed in the
stoichiometric equation below:
0.0263 C6 H14 + 0.0939 NO3− + 0.0939 H +
→ +0.0230 C5 H 7O2 N + 0.0354 N 2 + 0.0525 CO2 + 0.150 H 2O
Based on bioenergetics using hexane as the carbon source, 1.35 moles of nitrous oxide
would be produced for every mole of hydrocarbon consumed. This theoretical molar
ratio was used to calculate the moles of hydrocarbon that would be consumed to produce
the moles of nitrous oxide measured in the microcosm gas as well as dissolved in the
microcosm water. These calculations are listed in Table 4.11. Based on bioenergetics
calculations, approximately 0.086 mg/L and 0.0030 mg/L of hexane would be consumed
to produce the amount of nitrous oxide measured in the mixed-amendment and nitrate
120
amended microcosms, respectively, after 407 days incubation. These numbers are near or
below the practical quantitation limits of the GC/FID (0.05 mg/L) and would not be
considered detectable. Thus, it is possible that anaerobic biodegradation is resulting in
methane production but that the magnitude of this biodegradation is too small to be
detected by GC/FID within 407 days.
Table 5.11 Bioenergetics Stoichiometry for Hexane Consumed Calculated from
Nitrous Oxide Production
Hexane Consumed
Nitrous Oxide Produced
Ratio
Microcosm
Condition
Nitrate
Mixed
0.0263
0.0354
1.3
mol/L
mol/L
Average Concentration
N2O Produced
Gaseous
Aqueous
(mol)
(mg)
(mol/L)
(mol/L)
26 Day Samples
1.5E-06
8.5E-07
2.1E-06
9.1E-05
Hexane Consumed
(mol)
(mg/L)
1.5E-06
5.82E-02
Nitrate
Mixed
1.9E-06
134 Day Samples
1.1E-06
2.7E-06
1.2E-04
2.0E-06
7.48E-02
Nitrate
Mixed
7.5E-07
2.2E-05
407 Day Samples
4.4E-07
1.1E-06
4.7E-05
1.2E-05
3.0E-05
1.3E-03
7.9E-07
2.3E-05
2.99E-02
8.58E-01
4.2.5 Carbon Dioxide Production
Carbon dioxide was measured in all microcosms as an indicator of microbial
mineralization of petroleum hydrocarbons. As shown in bioenergetics calculations
(Section 2.2), carbon dioxide is an expected by-product of TPH biodegradation in all
microcosms, even in the unamended microcosm where it acts as an electron acceptor.
Thus, an increase in carbon dioxide concentration was expected to correlate to a decrease
121
in TPH concentration. Carbon dioxide concentrations in all microcosms are listed in
Table 4.6. Changes in gaseous carbon dioxide concentration are plotted in Figures 4.7.
Large increases in carbon dioxide concentration were observed in both iron-amended
microcosms at the 0 Day and 26-Day sampling events. This increase in carbon dioxide
concentration was attributed to carbonate protonation and formation of carbonic acid due
to the acidic pH from the ferric chloride addition. Due to its instability, carbonic acid
decomposes to form water and carbon dioxide. Carbon dioxide concentrations in all
microcosms except the 0-day and 26-day data points for the two iron-amended
microcosms are displayed in Figure 4.7.
Carbon Dioxide Concentration(mol/L)
8E-04
7E-04
6E-04
5E-04
4E-04
3E-04
2E-04
1E-04
0E+00
0
50
100
150
200
250
300
350
400
450
Time (day)
Mn(IV)
Fe(III)
Mixed
SO4
NO3
Unamended
Oxygen
Control
Figure 5.7 Gaseous Carbon Dioxide Molarity in Microcosm Headspace, Corrected
to Remove Outliers in Iron-Amended Microcosms
122
Carbon dioxide concentrations in the oxygen-amended microcosms increased rapidly
from the 0-day to 26-day sampling events and continued to rise until the 134-day
sampling event. This increase in carbon dioxide concentration was supported as evidence
of biodegradation by the subsequent decrease in TPH concentration at those sampling
events. The increase in carbon dioxide concentration in the mixed-amendment
microcosm between the 134 and 407-day sampling dates supported the assertion that
denitrification is taking place in these microcosms. However, this assertion was not
supported by the lack of TPH biodegradation, which is near the quantitation limit for
GC/FID. The lack of change in carbon dioxide concentration in the other microcosms
supported the assertion that TPH biodegradation did not take place in these microcosms
during the 407-day incubation period.
4.4
Nitrate and Sulfate Concentrations (Ion Chromatography Results)
Samples were collected, filtered, and analyzed for nitrate, nitrite and sulfate
concentrations on all four sampling dates. Nitrite was not detected above 1 ppm in any
microcosms any of the four sampling dates and therefore was not subjected to further
analysis. Nitrate and sulfate concentrations results for all microcosms are listed in
Sections 4.4.1 and 4.4.2. Sulfate and nitrate samples taken at the Day 0 sampling event
were diluted, as the concentrations used to amend the microcosms were believed to be
too high for IC analysis. However, it was observed for these initial samples that the
concentrations were within the IC detection range, so future samples were not diluted
after the 0 Day event.
123
4.4.1 Nitrate Concentration in Microcosms
Nitrate data from all microcosms and replicates are listed in Table 4.15. Changes in
average nitrate concentrations and standard deviations during the course of the
experiment are visualized in Figure 4.8.
Microcosms not amended with nitrate were generally low to non-detectable in nitrate
concentrations throughout the experiment. The only exception to this was the control
microcosm, which exhibited larger nitrate concentrations than the nitrate-amended
microcosms throughout the experiment. This was surprising, since the control
microcosms were not amended with nitrate. Potential explanations for this are that azide
interferes with nitrate readings by the IC or azide was converted to nitrate due to a
chemical reaction in the microcosms (Cotton and Wilkinson, 1976). In the nitrateamended microcosms, no statistically significant difference occurred between the
samples taken at the different time intervals, indicating that nitrate was not utilized as an
electron acceptor in these microcosms at a detectable level.
Nitrate concentration in the nitrate amended microcosm increased at the 407-day
sampling event, which is the opposite of what would be expected if nitrate was utilized as
an electron acceptor in biodegradation. Though the final concentration of nitrate
increased in comparison with the other two concentrations, the nitrate concentrations at
the four sampling dates are no significantly different from each other.
124
Table 5.12 Nitrate Concentrations in All Microcosms and Replicates
Microcosm
Condition
A
Manganese
B
A
Iron
B
A
Mix
B
A
Sulfate
B
A
Nitrate
B
A
Unamended
B
A
Oxygen
B
A
Control
B
Nitrate Concentration (mg/L)
0 Day
26 Day
134 Day
407 Day
0.20
0.19
0.00
0.00
0.20
0.16
0.00
0.00
0.36
0.16
0.00
0.00
0.33
0.16
0.00
0.00
0.33
0.08
0.00
0.00
0.20
0.00
0.00
0.00
0.43
0.16
0.00
0.00
0.45
0.00
0.00
0.00
53.88
61.63
66.21
62.06
69.01
66.61
57.05
59.32
68.42
49.75
58.17
49.67
0.20
0.00
0.00
0.00
0.20
0.00
0.00
0.00
0.00
0.00
0.00
0.00
0.00
0.00
0.00
0.00
85.84
58.02
65.60
150.46
86.16
58.13
65.68
151.43
60.52
65.83
65.65
150.38
62.54
66.22
65.58
150.22
0.20
0.00
0.00
0.00
0.20
0.00
0.00
0.00
0.47
0.16
0.00
0.00
0.45
0.16
0.00
0.00
0.20
0.52
1.01
0.20
0.50
1.01
0.41
0.65
0.96
0.38
0.60
0.99
156.65
0.00
148.67
2.03
157.60
0.00
148.99
2.15
190.98
152.37
148.72
1.92
191.53
152.10
149.87
2.03
125
200
180
NitrateConcentration (PPM)
160
140
120
100
80
60
40
20
0
Mn(IV)
Fe(III)
Mix
SO4
Initial
NO3
Unamend
Day 26
O2
Day 134
Control
Day 407
Figure 5.8 Change in Average Nitrate Concentration in Microcosms
Using bioenergetics stoichiometry, the nitrate consumed to produce the nitrous oxide
measured in gas headspace can be calculated. Theoretical results are listed in Table 4.13.
Based on results of these calculations, the nitrate consumed to produce the highest
measured nitrous oxide concentrations (500 ppm in Mixed microcosms at 407 days) is
1.8 mg/L. Based on these calculations, the amount of nitrate consumed in order to
produce the observed amount of nitrous oxide is small relative to the amount of nitrate
added as amendment.
126
Table 5.13 Nitrate Consumed Based on Nitrous Oxide Produced and Bioenergetic
Molar Ratios
Microcosm
Condition
Nitrate
Mixed
407 Day Samples
Average N2O Concentration
Gaseous
Aqueous
(mol/L)
(mol/L)
7.5E-07
4.4E-07
2.2E-05
1.2E-05
NO3 Consumed
(mol/L)
(mg/L)
2.8E-06
8.1E-05
6.21E-02
1.78E+00
4.4.2 Sulfate Concentration in Microcosms
Sulfate data from all microcosms and replicates are listed in Table 4.14. Changes in
average sulfate concentrations and standard deviations during the course of the
experiment are visualized in Figure 4.11.
Sulfate concentrations were generally low in the microcosms not amended with sulfate,
approximately 3 mg/L, which was the initial sulfate concentration in the groundwater at
J8-11. The two final iron microcosms exhibited high sulfate concentrations in one of the
microcosms and low-to-non-detect concentrations in the other, yielding a high standard
deviation. The source of this sulfate is not known. It is possible this sulfate was due to
an overall analytical error in sulfate analysis, since other anomalies were observed.
Sulfate microcosms exhibited an increase, but it was not statistically significant. Overall,
sulfate does not appear to be a statistically significant difference in sulfate concentration
in any microcosm condition, indicating sulfate was not utilized as an electron acceptor.
127
Table 5.14 Sulfate Concentrations in All Microcosm Replicates
Microcosm
Condition
Manganese
Iron
Mix
Sulfate
Oxygen
Nitrate
Unamended
Control
Replicates
Mn-1A
Mn-1B
Mn-2A
Mn-2B
Fe-1A
Fe-1B
Fe-2A
Fe-2B
Mix-1A
Mix-1B
Mix-2A
Mix-2B
SO4-1A
SO4-1B
SO4-2A
SO4-2B
O2-1A
O2-1B
O2-2A
O2-2B
NO3-1A
NO3-1B
NO3-2A
NO3-2B
CH4-1A
CH4-1B
CH4-2A
CH4-2B
N3-1A
N3-1B
N3-2A
N3-2B
0 Day
2.17
1.82
2.76
2.78
2.55
2.63
1.97
2.06
42.06
49.98
43.37
46.29
50.38
51.02
23.90
24.22
1.96
2.56
2.99
2.60
2.93
1.59
2.33
2.07
3.65
3.26
22.12
21.95
2.75
2.68
2.97
2.93
Sulfate Concentration (mg/L)
26 Day
134 Day
407 Day
1.33
2.46
2.37
1.31
2.93
2.15
1.74
3.33
3.17
1.50
3.16
2.94
5.23
4.08
91.09
4.86
4.26
91.61
0.00
61.72
19.97
0.00
62.12
20.08
104.71
76.04
93.85
76.27
54.31
93.73
93.18
93.47
76.22
81.69
73.31
76.59
82.35
73.83
60.28
77.54
80.05
60.29
77.83
79.93
7.11
3.89
7.03
4.06
6.00
3.90
5.84
4.03
6.37
2.73
2.49
6.24
2.69
2.49
5.84
2.85
2.60
5.98
2.77
2.49
1.32
2.49
0.90
1.24
2.42
0.78
1.21
2.16
1.24
1.92
2.42
1.35
6.42
2.34
2.03
6.39
2.34
2.15
5.29
2.33
1.92
5.42
2.66
2.03
Some anomalous changes occurred in sulfate concentration. One 407-day iron
microcosms and one 134-day iron microcosm had sulfate concentrations similar to the
128
sulfate-amended microcosms, as did one of the 0-day unamended microcosms. These
anomalous concentrations were observed in both microcosm replicates, as shown in
Table 4.14.
120
Sulfate Concentration (PPM)
100
80
60
40
20
0
Mn(IV)
Fe(III)
Mix
SO4
NO3
Initial
Unamend
O2
Day 26
Day 134
Control
Day 407
Figure 5.9 Change in Average Sulfate Concentration in Groundwater Microcosms
Unfortunately, no additional end product was monitored that could be correlated to
sulfate reduction (unlike nitrous oxide and nitrate). Based on the sulfate concentrations
and the lack of TPH reduction, it was assumed that the sulfate addition had no effect on
TPH biodegradation in this experiment.
129
4.5
Ferrous Iron Concentrations in Microcosms
Ferrous iron analyses were conducted on iron-amended, mixed amendment, and
unamended microcosms at each sampling date. The unamended microcosms were tested
to monitor potential iron reduction taking place in microcosms without iron-amendment,
representing the changes in iron concentration in field conditions. Ferrous iron data is
listed in Table 4.15. Changes in average ferrous iron concentration and standard
deviations are depicted graphically in Figure 4.12.
Table 5.15 Iron(II) Concentration in Iron-Amended and Unamended Microcosms
Microcosm
Condition
A
Iron
0 Day
26 Day
134 Day
407 Day
5.05
9.65
10.7
10.7
5.62
5.90
9.72
9.68
0.03
0.06
1.02
0.83
1.57
1.29
3.68
3.72
1.08
1.06
0.211
0.215
0.510
0.501
0.622
0.640
0.900
0.900
0.416
0.417
2.25
2.16
0.197
0.189
0.248
0.248
0.611
0.594
0.619
0.501
B
A
Mixed
B
A
Unamended
B
Initial Fe(II) concentrations in the unamended microcosms were approximately 2 ppm.
In the iron-amended microcosms, initial Fe(II) concentrations were higher. In all
microcosms, the Fe(II) concentration decreased during incubation.. These results
indicate that there is no evidence of iron reduction the iron-amended microcosms.
130
6
Fe(II) Concentration (ppm)
5
4
3
2
1
0
Fe
Mix
Initial
26 Day
Unamend
134 Day
407 Day
Figure 5.10 Change in Ferrous Iron Concentration in Iron-Amended and
Unamended Microcosms
According to these analyses, there is no evidence to support iron reduction as a
mechanism for biodegradation in this experiment. The degree that the initial pH decrease
contributed to this result cannot be determined; however, comparing the bacteriological
results in the TRF Data (see Section 4.7) indicated that the pH change considerably
altered the microbial community.
4.6
Manganese(II) Concentrations
Aqueous manganese (Mn(II)) was analyzed using the formaldoxime method for the
manganese-amended microcosms and unamended microcosms at each sampling date.
The unamended microcosm was tested to assess whether or not naturally present Mn(IV)
131
was contributing to biodegradation. Mn(II) concentrations, averages, and standard
deviations are listed in Table 4.16. Changes in average Mn(II) concentration and
standard deviations are depicted graphically in Figures 4.11.
Table 5.16 Manganese(II) Concentration in Manganese and Unamended
Microcosms
Manganese(II) Concentration (mg/L)
0 Day
26 Day
134 Day
407 Day
2.37
1.07
1.00
2.89
2.15
1.83
0.98
3.87
1.41
0.85
4.00
0.08
0.91
4.19
1.05
4.73
2.29
2.17
0.78
3.61
2.48
2.81
4.59
2.63
2.90
5.00
2.53
1.99
Microcosm
Condition
A
Manganese
B
A
Unamended
B
6
Mn(II) Concentration (ppm)
5
4
3
2
1
0
Mn
Unamend
0 Day
26 Day
134 Day
407 Day
Figure 5.11 Change in Manganese(II) Concentration in Manganese and
Unamended Microcosms
132
Manganese(II) concentrations fluctuated between 1 ppm and 4 ppm in manganeseamended microcosms and between 1 ppm and 5 ppm in unamended microcosms (Figure
4.11). For the manganese-amended microcosms, a statistically significant increase in
Mn(II) concentrations occurred during the experiment. However, this increase was not
significantly different from the increase measured in the unamended microcosms. The
increase in concentration of Mn(II) in the amended and unamended microcosm is
intriguing, and suggests naturally-occurring MnO2 may have been present in the
groundwater or soil and was more bioavailable than the MnO2-precipitates used as
amendments. However, there was no consistent increase in Mn(II) to suggest that native
microorganisms utilized the manganese-oxide amendment.
Bioenergetics (see Section 2.2) was utilized to developed balanced stoichiometric
equations for the biodegradation of hexane using MnO2 as the terminal electron acceptor,
as seen in the following equation:
0.0263 C6 H14 + 0.222 MnO2 + 0.444 H + + 0.278 HCO3− + 0.0278 NH 4+
→ 0.378 H 2O + 0.0278 C5 H 7O2 N + 0.222 Mn 2+ + 0.0467 CO2
Based on bioenergetics using hexane as the carbon source, 8.44 moles of Mn(II) would
be produced for every mole of hydrocarbon consumed. This theoretical molar ratio was
used to calculate the moles of hydrocarbon that would be consumed to produce the
amount of Mn(II) measured in the microcosm. Since there was some Mn(II) present in
the initial samples, additional calculations were performed to remove initial Mn(II) so
that only the net increase of Mn(II) was contributed to biodegradation. The results of
these calculations are listed in Table 4.17. Based on these calculations, approximately
0.27 mg/L and 0.66 mg/L of hexane would be consumed to produce the amount of Mn(II)
133
measured in the manganese oxide amended and unamended microcosms, respectively,
after 407 days incubation. These decreases in TPH concentration are above the detection
limits, but do not correlate with TPH data obtained from the GC/FID. It is possible that
there is a lack of correlation between theoretical and actual TPH degradation due to the
difference in carbon length between hexane (C-6) and the unresolved petroleum mixture
present at the GRP site (C-20) or that natural organic matter present in the microcosms
was used as an alternative carbon source. Regardless, there is insufficient evidence to
support the use of manganese oxide as a terminal electron acceptor for petroleum
hydrocarbon biodegradation during the 407-day incubation period.
Table 5.17 Hexane Consumption Based on Bioenergetics Calculations and
Manganese(II) Concentration, Corrected to Exclude Manganese(II) Present at Day
0 Sampling Event
Mn(II) Produced
Microcosm
Condition
Manganese
Unamended
4.6
Concentration
(mg/L)
1.48
3.57
Molarity
(mol/L)
2.69E-05
6.49E-05
Hexane Consumed
Molarity
(mol/L)
3.18E-06
7.69E-06
Concentration
(mg/L)
0.27
0.66
Microtox® Toxicity Results
Toxicity analyses were conducted on samples from one of the two microcosms sacrificed
per amendment condition on each sampling date. Table 4.18 summarizes Microtox
toxicity testing results in terms of % effect for all microcosms during this experiment. %
Effect is calculated from variables generated in a linear regression shown in Figure 4.12.
Table 4.19 lists the toxicity results in terms of EC50. Note that for EC50, lower number
indicates greater toxicity, since EC50 indicates the amount of a sample required to reduce
134
bacterial bioluminescence by 50 percent. The results are represented visually in Figure
4.13, comparing microcosm types during the length of the experiment.
Table 5.18 Percent Effect of Microcosm Samples on Bioluminescent Bacteria,
Calculated Using Microtox Omni Software
Microcosm
Condition
Mn(IV)
Fe(III)
Mix
NO3
SO4
Unamend
O2
Control
Day 0
81.07
100.00
100.00
82.76
77.14
82.07
81.65
82.51
% Effect
Day 26 Day 134 Day 407
78.09
84.34
75.75
100.00
83.06
82.95
100.00
85.18
86.13
69.85
81.17
94.87
65.21
89.57
86.67
80.62
90.07
91.98
31.28
45.87
74.41
71.02
85.64
100
90
80
% Effect
70
60
50
40
30
20
10
0
Mn(IV)
Fe(III)
Mix
NO3
SO4
Day 0
Unamend
Day 26
O2
Day 134
Control
Day 407
Figure 5.12 % Effect of Microcosm Sample on Bacterial Bioluminescence,
Microcosm Comparison
135
Table 5.19 Effective Concentration of Microcosm Sample that caused 50%
Reduction in Bacterial Bioluminescence, Determined Using Microtox Omni
Software
Microcosm
Condition
Mn(IV)
Fe(III)
Mix
NO3
SO4
Unamend
O2
Control
Day 0
EC50
29.53
4.45
7.03
32.81
36.76
29.30
28.11
20.96
Day 26
EC50
33.19
4.40
5.51
44.91
63.14
30.67
114.00
40.75
Day 134 Day 407
EC50
EC50
30.72
37.14
33.55
28.21
26.15
25.05
28.03
17.62
25.34
22.52
25.34
18.36
114.10
36.38
25.07
120
EC50 (% of Sample)
100
80
60
40
20
0
Mn(IV)
Fe(III)
Day 0
Mix
Day 26
NO3
SO4
Day 134
Unamend
O2
Control
Day 407
Figure 5.13 Concentration of Microcosm Sample Required to Reduce Bacterial
Bioluminescence by 50%, Microcosm Comparison
By comparing % Effect and EC50 results for all microcosms, it is evident that very little
overall change occurred in sample toxicity for the anaerobic microcosms during the
experiment. The initial toxicity of the iron-amended microcosms was reduced when the
136
pH was corrected, as seen in the change from 26 to 134 day samples for iron and mixed
microcosms (Figures 4.12 and 4.13). With only two exceptions (nitrate and sulfate
microcosms on 26th Day sampling), toxicity of the anaerobic microcosms did not vary
appreciably from the value of the control microcosm. By comparison, the aerobic
microcosms experienced a large drop in toxicity from 0th to 26th day and remained below
toxicity detection limits for the rest of the experiment. Very little difference occurred
between the unamended microcosms and the amended microcosms throughout the
experiment, indicating that the amendments had no effect on sample toxicity. However,
little difference occurred between the control microcosm, with 1,000 mg/L sodium azide,
the amended microcosms, and the unamended microcosms. The toxicity cannot be due to
the resazurin indicator, since the aerobic microcosms reduced in toxicity to detection
limits.
4.7
Terminal Restriction Fragment Analysis Results
TRF patterns were compared to observe differences in microbial species within the
different microcosms. Dendrograms were produced using Bray-Curtis similarity grouped
by group average. No complex statistical analysis could be conducted since samples
were not analyzed in duplicate and only initial and final samples were taken.
4.7.1 16S DNA Digests
Three enzyme digests were performed on the 16S portion of the DNA strand. This
method is equivalent to a heterotrophic plate count, as it is non-specific in bacterial types.
The three restriction enzymes used were Dpn III, Hae III, and Hha I. Figures 4.14
137
through 4.16 combine the electropherograms produced by the CEQ 8000 and the with
dendrograms produced by Primer software using Bray-Curtis similarity grouped by group
average.
The dendrograms created using the Brae-Curtis similarity analysis highlighted many of
the differences observed between the different microcosm conditions. The iron
microcosm was always an outlier with only 30 % similarity to the other microcosms.
Nitrate-amended microcosm and initial sample were grouped together in dendrograms
produced from each of the three restriction enzymes with a relative similarity ranging
from 50 – 70 %. The three microcosms that produced methane gas – unamended, sulfate,
and manganese – were all grouped together. Oxygen and mixed were always either
grouped together or branched off of each other.
Although no appreciable logic is apparent in the grouping of initial and nitrate-amended
microcosms, it is possible that the addition of nitrate did not disturb the microbial
communities, and did not serve as a nitrogen source or terminal electron acceptor for
existing bacteria. Grouping manganese, sulfate and unamended together is logical on the
basis since all of these microcosms produced methane gas during the course of the
experiment. Though manganese is a more oxidized system than the other two methaneproducing microcosms, it is possible that the manganese oxide amendment did not
disturb methanogenic microorganisms. This is likely since manganese reduction was not
appreciable, so the amendment did not stimulate manganese-reducing organisms during
the 407-day incubation period.
138
The iron-amended microcosms were extremely different from all other microcosms in
each digest, most likely due to the pH change during microcosm establishment. Though
the mixed-amendment microcosm analyzed was not as dissimilar as was the ironamended microcosms, the mixed microcosm was grouped with the aerobic microcosm on
each dendrogram. This could be due to the iron-induced pH reduction killing off of
sensitive organisms, allowing other organisms to flourish. Given the production of
nitrous oxide in the mixed microcosm, these organisms were likely to be denitrifiers.
Nitrous oxide was produced in the nitrate-amended microcosm as well, but at 1/25 the
amount detected in the mixed amendment microcosm.
Shifts were observed in the microcosms between the 0-day and 407-day sampling events.
Methane-producing microcosms were grouped together in dendrograms produced by all
three restriction enzymes and iron-amended and aerobic microcosms were significantly
different from the other microcosm conditions.
139
Figure 5.14 Electropherograms and Dendrogram Produced from Bray-Curtis
Similarity for 16S DNA Fragments Produced by Dpn III Restriction Enzyme
140
Figure 5.15 Electropherograms and Dendrogram Produced from Bray-Curtis
Similarity for 16S DNA Fragments Produced by Hae III Restriction Enzyme
141
Figure 5.16 Electropherograms and Dendrogram Produced from Bray-Curtis
Similarity for 16S DNA Fragments Produced by Hha I Restriction Enzyme
142
4.7.2 Methanogen and Archaea Results
Methanogen and Archaea TRF electropherograms are grouped with the dendrograms
produced by Primer in Figures 4.17 and 4.18.
Each of microcosms provided sufficient DNA for TRF analysis of methanogenic and
Archaea populations, but there was not a readily apparent difference in the patterns
developed by the different microcosms. Each of the Archaea patterns is essentially
defined by three peaks: 205, 215 and 315. Each pattern has a substantial peak at 215 base
pairs long, contributing 29 – 69 % of the total relative abundance of all peaks, even in the
aerobic microcosm where the environment was expected to be lethal to Archaea. The
methanogenic patterns yielded similar results. There was no evidence of grouping by
either redox condition or gas production observed during the course of the experiment.
Each pattern had a significant peak at 405 base pairs comprising 35 – 64 % of the total
relative abundance. Similar to the Archaea pattern, the majority of the similarity
comparison was based upon peaks at three base pair lengths: 403, 405, and 505. This
result was not expected considering the different reducing environments in the seven
microcosms conditions. A likely reason is the presence of artifact DNA from dead cells.
Since the technique has no method of distinguishing between active biomass and
decaying cellular material, it is impossible to know if the DNA present came from active
bacteria.
When examining the dendrograms for the Archaea, the unamended microcosm was
significantly different from all other microcosms, sharing only 30 % similarity. This
143
results suggests that any amendment affected the Archaea community, regardless of the
redox state induced by the amendment. The unamended microcosm had 74.5 % of its
total peak area under peaks at 205 and 215 base pairs, with 39.5 % at 205 base pairs. No
other condition had more than 20 % relative abundance at this peak, and the initial
sample had no peak area at 205 base pairs. Thus, the unamended microcosm’s microbial
community changed significantly both from the initial sample and compared to all
amended microcosms.
144
Figure 5.17 Electropherograms and Dendrogram Produced from Bray-Curtis
Similarity for Methanogen DNA Fragments Produced by Sau I96 Restriction
Enzyme
145
Figure 5.18 Electropherograms and Dendrogram Produced from Bray-Curtis
Similarity for Archaea DNA Fragments Produced by Hae III Restriction Enzyme
146
CHAPTER 6
CONCLUSIONS
Based on the gas headspace results from the four sampling periods, anaerobic conditions
were maintained in the groundwater microcosms. Additionally, gas headspace data
confirmed that microcosms did not leak based on the N2:He ratios, which were
maintained in all un-altered microcosms for the length of the experiment.
No significant hydrocarbon biodegradation was observed in any of the anaerobic
microcosms during 407 days incubation based on TPH analyses. Similarly, no significant
changes were observed in electron acceptor concentrations in any amended microcosms.
In contrast, TPH concentration in the aerobic microcosms was reduced by 54.3 % after
26-days incubation and 72.2 % after 298-days incubation.
Methane and nitrous oxide concentrations in headspace gases increased in several
microcosms, suggesting fermentation and denitrification were occurring. However, the
volume of gases produced correlate to minimal decreases in hydrocarbon concentration
based on bioenergetics calculations. Anaerobic microcosms did not exhibit significant
changes in toxicity during the 407-day incubation period, remaining close to 80 % effect
throughout the experiment. By contrast, aerobic microcosms reduced from 80 % effect to
below detection limits by 134 days.
Terminal Restriction Fragment (TRF) analyses produced from 16S DNA digests revealed
a shift occurred in microbial communities between the initial and the final conditions.
147
The iron microcosms were significantly different from all other conditions, most likely
due to the pH reduction that occurred when the microcosms were initially established.
The aerobic microcosm was grouped with the mixed microcosm and the microcosms with
methanogenic activity (SO4, CH4, MnO2) were grouped together. For digests of
methanogenic DNA, the aerobic and iron-amended microcosms were significantly
different from all other microcosms, but no other grouping was indicative of redox
condition or observed metabolic activity. For digests of Archaea DNA, the unamended
microcosm was significantly different from all other microcosms, suggesting that
addition of any amendment to the microcosms significantly altered the Archaea
community present. Despite these groupings, no compelling statistical analyses could be
performed regarding similarity due to too few samples and lack of replicates.
The results of this experiment support the experimental findings of Loehr et al (2001),
suggesting that the biodegradability of weathered hydrocarbons tends to be site specific
due to specific biotic and abiotic weathering processes occurring. However, in the
anaerobic study published by Loehr et al (2001), all sites contaminated with diesel-range
hydrocarbons had high biodegradability. This was not the case in this study, where no
measurable anaerobic biodegradation was noted after 407-days incubation.
The results of the study do not correlate with the findings of Lundegard and Johnson
(2003), who observed extensive methanogenic activity in the source zone region of
plumes on the GRP site. Though methanogenic activity was detected in this experiment,
the amounts produced were not related to measurable TPH reduction; this suggests that
148
methane produced could be linked to TPH reduction below our detection limit (0.05
mg/L) or could be due to oxidation of natural organic matter present in the soil or
groundwater. While methanogenesis/fermentation may appreciably contribute to source
zone natural attenuation, the results of this experiment suggest it may not contribute
significantly to dissolved-phase diluent biodegradation.
Based on these results and experimental conditions after 407-days incubation, anaerobic
biodegradation may not contribute appreciably to natural attenuation at the Guadalupe
Restoration Project site.
In regards to future anaerobic studies at the GRP site, I wish to make the following
recommendations:
•
Conduct analyses on the remaining anaerobic microcosms after 800-days
incubation. Since the study was initially designed for 5 sampling periods in a
150-day incubation period, there remains one set of microcosms incubating in the
anaerobic glovebox. These samples should be analyzed for changes in TPH
concentration, amendment (or amendment by-product) concentrations, and
headspace gases to determine if anaerobic processes contribute appreciably to
hydrocarbon biodegradation within an 800-day period.
•
Quantify micronutrient availability at the site prior to well selection. Research
suggests that micronutrient concentrations (such as nickel and molybdenum) are
especially important to some anaerobic microorganisms, as they are necessary for
149
enzyme structures. Well selection should be based on nutrient availability as
well as TPH and electron acceptor concentrations.
•
Retrieve groundwater and sand from the same location on the site. Studies
suggest native microorganisms are more efficient at degrading contaminants than
are introduced microorganisms. It is reasonable to assume that the nature of the
contamination and microorganisms present vary from well to well. Therefore,
the acclimation period for organisms introduced to a foreign environment may
increase the lag phase.
•
Consider the costs and benefits of sacrificial microcosms before choosing to use
them over large microcosms. Though they may be simpler for gas analyses,
heterogeneities between individual microcosms can prove to be very frustrating.
•
Focus on chemical changes in the diluent during the incubation period as well as
the potential biodegradability of contaminants present at the site. Research
suggests that some compounds do not provide sufficient energy to promote
complete mineralization by native microorganisms (as is the case for the
biodegradation of MTBE by methanogens, resulting in TBA). Analysis of TPH
changes using gas chromatography with mass spectroscopy (GC/MS) could
indicate chemical changes in the unresolved petroleum mixture that are not
identified by GC/FID, indicating that TPH biodegradation is taking place, but the
compounds are not being completely mineralized.
•
Try a similar experiment where microcosms spiked with defined hydrocarbons
(such as BTEX, MTBE, octane, or No. 2 diesel fuel) to compare the
150
biodegradability of known compounds under anaerobic conditions to that of
weathered petroleum compounds.
151
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160
APPENDIX A: BIOENERGETICS FORMULAS AND
CALCULATIONS
161
A.1
Bioenergetics Introduction
Oxidation – reduction reactions, or redox reactions, are defined as reactions in which
electrons are passed between chemical species (Prescott, Microbiology). Redox reactions
are the basis of biochemical systems, as energy is released as chemicals transfer
electrons. The chemicals donating the electrons are oxidized, whereas the chemicals
receiving the electrons are reduced.
Redox reactions are often described in terms of half-reaction couples. These intermediate
steps are helpful since they permit the viewing of “electron transfer” between chemicals.
For the oxidation half-reaction, the electrons appear on the products side of the equation,
as depicted below:
1
1
C6 H12O6 → CO2 + e− + H +
24
4
For the reduction half, reaction, the electrons appear on the reactants side of the equation,
as depicted below:
1
1
O2 + e− + H + → H 2O
4
2
When the two reactions are combined, the electrons are cancelled and are not included in
the net reaction, since they are on both the products and reactants sides of the reaction, as
seen below:
1
1
1
1
C6 H12O6 + O2 → CO2 + H 2O
24
4
4
2
The fractions can be removed by multiplying the equation by the least common
denominator, yielding the familiar equation for aerobic respiration of glucose.
162
C6 H12O6 + 6O2 → 6CO2 + 12H 2O
Reactions are typically listed in tables, are balanced in terms of chemical species and
charge, and are written in terms of 1 electron (e-) to make them easier to combine. The
redox tables rank the reactions in orders of reduction potential (E'°, measured in volts),
best described as the chemical species’ ability to accept electrons or the likelihood of the
species being chemically reduced. The greater the value of E'°, the more likely the
species is to be reduced in a redox reaction. Chemicals with low or negative E'° are
generally not reduced, but rather act as electron donors. The E'° generated during the
oxidation-reduction reaction determines the overall energy released by the reaction, and
therefore the favorability of a specific reaction occurring.
To use a reduction potential table, select the species to be oxidize and to be reduced. The
reduction reaction should always have a higher reduction potential or else the net reaction
will have a negative reduction potential. For example, the oxidation of lactate by
reduction of nitrate would proceed according to the following:
−
RR : NO3 + 2H + + 2e− → NO2− + H 2O
E R = 0.421 V
RO : Pyruvate− + 2H + + 2e− → Lactate−
E O = (−0.185 V )
where RO is the oxidation half-reaction and RR is the reduction half-reaction. EO and ER
are the reduction potentials of the oxidation (electron donor) and reduction (electron
acceptor) half-reactions, respectively. The total reduction potential of the net reaction is
determined as depicted subtracting the oxidation potential from the reduction potential.
163
E T = E Electron Acceptor − E Electron Donor
The potential of the oxidation half of the reaction is subtracted because the equation is
listed in the table as a reduction; when it is used as an oxidation, the negative value is
taken. The net reaction is:
RT : Lactate− + NO3− → Pyruvate− + NO2−
E T = 0.421V − (−0.185 V )
E T = 0.606 V
In biochemical processes, redox reactions can become quite complex, particularly in
microbial systems. This is because the microbes are simultaneously the catalyst for the
reaction and the product of the reaction (Rittmann and McCarty, 2001). In general, when
dealing with biochemical systems, tabled values are used because the equations are too
complex for simple species and charge balancing.
Using ammonium as the nitrogen source yields the following synthesis reaction and
∆Gpc:
1
1
1
1
9
CO2 +
HCO3− +
NH 4+ + H + + e− → C5 H 7O2 N +
H 2O
5
20
20
20
20
∆G pc = −18.8
kJ
eeq
Using nitrate as nitrogen source yields the following synthesis reaction and ∆Gpc:
1
1
11
5
CO2 + NO3− + H + + e− → C5 H 7O2 N + H 2O
28
28
28
28
164
∆G pc = −13.5
kJ
eeq
The half-reaction and reduction potential for hexane reduction are:
1
12
6
CO2 + H + + e− → C6 H14 +
H 2O
38
38
38
∆Gr = 28.66
kJ
eeq
This value will be negative when hexane acts as an electron donor in our bioenergetics
calculations. Since the electron donor is the same for all calculations, the ∆Gp for
conversion of hexane to pyruvate will be the same. Additionally, since the ∆Gp is greater
than zero, n = 1 for all equations. Therefore, εn = ε
kJ
− ∆Gd
eeq
kJ
kJ
= 35.09
− 28.66
eeq
eeq
kJ
= 6.43
eeq
∆G p = 35.09
Manganese(IV) Reduction
The Gibbs Free Energy of manganese oxide reduction was obtained by converting the
reduction potential using the equation below:
1
1
MnO2 (s) + H + + e− → Mn 2+ (aq) + H 2O
2
2
∆E° = 0.612V
∆G° = −nℑ∆E°
∆G° = −(1)(96.48
∆G° = −59.04
kJ
)(0.612V )
Veeq
kJ
eeq
This value was then used for ∆Ga in all subsequent bioenergetics calculations.
165
1
12
6
H 2O → CO2 + H + + e−
−Rd : C6 H14 +
38
38
38
1
1
Ra : MnO2 + H + + e− → Mn 2+ + H 2O
2
2
∴∆Gr = −87.70
1
A= ε
(∆G
p
kJ
eeq
kJ
∆Ga = −59.04
eeq
− (∆Gd ) = −28.66
kJ
eeq
+ ∆G pc )
ε∆Gr
kJ
kJ ⎞
1 ⎛
+ 18.8
⎜6.43
⎟
eeq
eeq ⎠
0.6 ⎝
=
⎛
kJ ⎞
0.6⎜87.70
⎟
eeq ⎠
⎝
= 0.799
fs =
1
1
=
= 0.556
1+ A 1+ 0.799
fe =
A
0.799
=
= 0.444
1+ A 1+ 0.799
1
12
6
C6 H14 +
H 2O → CO2 + H + + e−
38
38
38
⎛1
⎞
1
+ f e Ra : (0.444 )⎜ MnO2 + H + + e− → Mn 2+ + H 2O⎟
⎝2
⎠
2
⎛1
⎞
1
1
1
9
+ f sRc : (0.556)⎜ CO2 + HCO3− + NH 4+ + H + + e− → C5 H 7O2 N + H 2O⎟
⎝5
⎠
20
20
20
20
−Rd :
R : 0.0263C6 H14 + 0.222MnO2 + 0.444H + + 0.278HCO3− + 0.0278NH 4+
→ 0.378H 2O + 0.0278C5 H 7O2 N + 0.222Mn 2+ + 0.0467CO2
166
Ferric Iron Reduction
1
12
6
−Rd : C6 H14 +
H 2O → CO2 + H + + e−
38
38
38
Ra : Fe 3+ + e− → Fe 2+
∴∆Gr = −102.93
1
A= ε
(∆G
p
kJ
eeq
kJ
∆Ga = −74.27
eeq
− (∆Gd ) = −28.66
kJ
eeq
+ ∆G pc )
ε∆Gr
kJ
kJ ⎞
1 ⎛
+ 18.8
⎜6.43
⎟
eeq
eeq ⎠
0.6 ⎝
=
⎛
kJ ⎞
0.6⎜102.93
⎟
eeq ⎠
⎝
= 0.680
fs =
1
1
=
= 0.595
1+ A 1+ 0.680
fe =
A
0.680
=
= 0.405
1+ A 1+ 0.680
1
12
6
C6 H14 +
H 2O → CO2 + H + + e−
38
38
38
3+
−
+ f e Ra : (0.405)(Fe + e → Fe 2+ )
−Rd :
⎛1
⎞
1
1
1
9
+ f sRc : (0.595)⎜ CO2 + HCO3− + NH 4+ + H + + e− → C5 H 7O2 N + H 2O⎟
⎝5
⎠
20
20
20
20
R : 0.0263C6 H14 + 0.405Fe 3+ + 0.378H 2O + 0.298HCO3− + 0.0298NH 4+
→ +0.0298C5 H 7O2 N + 0.405Fe 2+ + 0.0389CO2 + 0.405H +
167
Denitrification
1
12
6
−Rd : C6 H14 +
H 2O → CO2 + H + + e−
38
38
38
1
6
1
3
Ra : NO3− + H + + e− → N 2 + H 2O
5
5
10
5
∴∆Gr = −100.86
1
A= ε
(∆G
p
kJ
eeq
kJ
∆Ga = −72.20
eeq
− (∆Gd ) = −28.66
kJ
eeq
+ ∆G pc )
ε∆Gr
1 ⎛
kJ
kJ ⎞
+ 13.5
⎜6.43
⎟
0.6 ⎝
eeq
eeq ⎠
=
⎛
kJ ⎞
0.6⎜100.86
⎟
eeq ⎠
⎝
= 0.549
fs =
1
1
=
= 0.646
1+ A 1+ 0.549
fe =
A
0.549
=
= 0.354
1+ A 1+ 0.549
1
12
6
C6 H14 +
H 2O → CO2 + H + + e−
38
38
38
⎛1
⎞
6
1
3
+ f e Ra : (0.354 )⎜ NO3− + H + + e− → N 2 + H 2O⎟
⎝5
⎠
5
10
5
⎛5
⎞
1
1
11
+ f sRc : (0.646)⎜ CO2 + NO3− + H + + e− → C5 H 7O2 N + H 2O⎟
⎝ 28
⎠
28
28
28
−Rd :
R : 0.0263C6 H14 + 0.0939NO3− + 0.0939H +
→ +0.0230C5 H 7O2 N + 0.0354N 2 + 0.0525CO2 + 0.150H 2O
168
Sulfate Reduction
1
12
6
kJ
H 2O → CO2 + H + + e−
− (∆Gd ) = −28.66
−Rd : C6 H14 +
eeq
38
38
38
1
19
1
1
1
kJ
Ra : SO42− + H + + e− → H 2 S + HS − + H 2O
∆Ga = 20.08
8
16
16
16
2
eeq
∴∆Gr = −7.81
1
A= ε
(∆G
p
kJ
eeq
+ ∆G pc )
ε∆Gr
1 ⎛
kJ
kJ ⎞
+ 13.5
⎜6.43
⎟
0.6 ⎝
eeq
eeq ⎠
=
⎛
kJ ⎞
0.6⎜ 7.81
⎟
eeq ⎠
⎝
= 8.974
fs =
1
1
=
= 0.100
1+ A 1+ 8.974
fe =
A
8.974
=
= 0.900
1+ A 1+ 8.974
1
12
6
C6 H14 +
H 2O → CO2 + H + + e−
38
38
38
⎛ 1 2− 19 + −
⎞
1
1
1
+ f e Ra : (0.900)⎜ SO4 + H + e → H 2 S + HS − + H 2O⎟
⎝8
⎠
16
16
16
2
⎛1
⎞
1
1
1
9
+ f sRc : (0.100)⎜ CO2 + HCO3− + NH 4+ + H + + e− → C5 H 7O2 N + H 2O⎟
⎝5
⎠
20
20
20
20
−Rd :
R : 0.0263C6 H14 + 0.1125SO42− + 0.344H + + 0.0005NH 4+ + 0.0005HCO3−
→ 0.0005C5 H 7O2 N + 0.0563H 2 S + 0.0563HS − + 0.138CO2 + 0.179H 2O
169
Carbon Dioxide Reduction
1
12
6
H 2O → CO2 + H + + e−
−Rd : C6 H14 +
38
38
38
1
1
1
Ra : CO2 + H + + e− → CO2 + H 2O
8
8
4
∴∆Gr = −5.13
1
A= ε
(∆G
p
kJ
eeq
kJ
∆Ga = 23.53
eeq
− (∆Gd ) = −28.66
kJ
eeq
+ ∆G pc )
ε∆Gr
1 ⎛
kJ
kJ ⎞
+ 18.8
⎜6.43
⎟
0.6 ⎝
eeq
eeq ⎠
=
⎛
kJ ⎞
0.6⎜5.13
⎟
eeq ⎠
⎝
= 13.66
fs =
1
1
=
= 0.068
1+ A 1+ 13.66
fe =
A
13.66
=
= 0.932
1+ A 1+ 13.66
1
12
6
C6 H14 +
H 2O → CO2 + H + + e−
38
38
38
⎛1
⎞
1
1
+ f e Ra : (0.932)⎜ CO2 + H + + e− → CH 4 + H 2O⎟
⎝8
⎠
8
4
⎛1
⎞
1
1
1
9
+ f sRc : (0.068)⎜ CO2 + HCO3− + NH 4+ + H + + e− → C5 H 7O2 N + H 2O⎟
⎠
⎝5
20
20
20
20
−Rd :
R : 0.0263C6 H14 + 0.00034NH 4+ + 0.00034HCO3− + 0.0522H 2O
→ 0.00034C5 H 7O2 N + 0.0563H 2 S + 0.117CH 4 + 0.0278CO2
170
Aerobic Respiration (Oxygen Reduction)
1
12
6
H 2O → CO2 + H + + e−
−Rd : C6 H14 +
38
38
38
1
1
Ra : O2 + H + + e− → H 2O
4
2
∴∆Gr = −107.38
1
A= ε
(∆G
p
kJ
eeq
kJ
∆Ga = −78.72
eeq
− (∆Gd ) = −28.66
kJ
eeq
+ ∆G pc )
ε∆Gr
1 ⎛
kJ
kJ ⎞
+ 18.8
⎜6.43
⎟
0.6 ⎝
eeq
eeq ⎠
=
⎛
kJ ⎞
0.6⎜107.38
⎟
eeq ⎠
⎝
= 0.653
fs =
1
1
=
= 0.605
1+ A 1+ 0.653
fe =
A
0.653
=
= 0.395
1+ A 1+ 0.653
1
12
6
C6 H14 +
H 2O → CO2 + H + + e−
38
38
38
⎛1
⎞
1
+ f e Ra : (0.395)⎜ O2 + H + + e− → H 2O⎟
⎝4
⎠
2
⎛1
⎞
1
1
1
9
+ f sRc : (0.605)⎜ CO2 + HCO3− + NH 4+ + H + + e− → C5 H 7O2 N + H 2O⎟
⎠
⎝5
20
20
20
20
−Rd :
R : 0.0263C6 H14 + 0.0988O2 + 0.0303NH 4+ + 0.0303HCO3−
→ 0.0303C5 H 7O2 N + 0.0369CO2 + 0.154H 2O
171
APPENDIX B: EPA METHOD 3510C
172
173
174
175
176
177
178
179
180
APPENDIX C: EPA METHOD 8015C
181
182
183
184
185
186
187
188
189
190
191
192
193
194
195
196
197
198
199
200
201
202
203
204
205
206
207
208
209
210
211
212
213
214
APPENDIX D: IRON ANALYSIS BY PHENANTHROLINE METHOD
Standard Methods for Analysis of Water and Wastewater
215
216
217
218
219
220
221
222
APPENDIX E: MANGANESE(II) ANALYSIS BY THE
FORMALDOXIME METHOD
223
224
225
226
227
APPENDIX F: TERMINAL RESTRICTION FRAGMENT ANALYSIS
PROTOCOL
228
TRF Protocol
STEP 1- To extract DNA out of your environmental sample
Extraction
Using MoBio Power Soil DNA Extraction Kit
Please wear gloves at all times
1.
2.
3.
4.
5.
6.
7.
8.
9.
10.
11.
12.
13.
14.
15.
16.
17.
18.
19.
20.
21.
22.
To the 2ml PowerBead Tubes provided, add 0.25 gm of soil sample. For sand add 1.0 gram, for feces 0.1 gram
Gently vortex to mix.
Check Solution C1. If Solution C1 is precipitated, heat solution to 60°C until dissolved before use.
Add 60µl of Solution C1 and invert several times or vortex briefly.
Fast Prep for
Soil 5.0 m/s for 45 sec.
Feces 4.5 m/s for 30 sec.
Pure culture 4.5 m/s for 30 sec
Make sure the PowerBead Tubes rotate freely in your centrifuge without rubbing. Centrifuge tubes at 10,000 x g for 30 seconds.
CAUTION: Be sure not to exceed 10,000 x g or tubes may break.
Transfer the supernatant to a clean microcentrifuge tube (provided).
Note: Expect between 400 to 500µl of supernatant. Supernatant may still contain some soil particles.
Add 250µl of Solution C2 and vortex for 5 seconds. Incubate in the freezer for 10-15 minutes.
Centrifuge the tubes for 1 minute at 10,000 x g.
Avoiding the pellet, transfer up to, but no more than, 600µl of supernatant to a clean microcentrifuge tube (provided).
Add 200µl of Solution C3 and vortex briefly. Incubate in the freezer for 10-15 minutes.
Centrifuge the tubes for 1 minute at 10,000 x g.
Avoiding the pellet, transfer up to, but no more than, 750µl of supernatant into a clean microcentrifuge tube (provided).
Add 1200µl of Solution C4 to the supernatant and vortex for 5 seconds.
Load approximately 675µl onto a spin filter and centrifuge at 10,000 x g for 1 minute. Discard the flow through and add an
additional 675µl of supernatant to the spin filter and centrifuge at 10,000 x g for 1 minute. Load the remaining supernatant onto
the spin filter and centrifuge at 10,000 x g for 1 minute. Keep loading until all supernatant from all replicates has been filtered
through the same filter.
Add 500µl of Solution C5 and centrifuge for 30 seconds at 10,000 x g.
Discard the flow through.
Centrifuge again for 1 minute.
Carefully place spin filter in a new clean tube (provided). Avoid splashing any Solution C5 onto the spin filter.
Add 100µl of Solution C6 to the center of the white filter membrane. Let sit for 15 minutes. Alternatively, sterile DNA-Free
PCR Grade Water may be used for elution from the silica spin filter membrane at this step (Mo Bio Catalog No. 17000-10).
Centrifuge for 30 seconds.
Discard the spin filter. DNA in the tube is now application ready. No further steps are required.
We recommend storing DNA frozen (-20° to -80°C). Solution C6 contains no EDTA.
Detailed Extraction Protocol (This is the same protocol as above, with explanations for
each step)
1.
2.
3.
4.
5.
To the 2ml PowerBead Tubes provided, add 0.25 gm of soil sample. For sand add 1.0 gram, for feces 0.1 gram
After your sample has been loaded into the PowerBead Tube, the next step is a homogenization and lysis procedure. The
PowerBead Tube contains a buffer that will (a) help disperse the soil particles, (b) begin to dissolve humic acids and (c) protect
nucleic acids from degradation.
Gently vortex to mix.
Gentle vortexing mixes the components in the PowerBead Tube and begins to disperse the sample in the PowerBead Solution.
Check Solution C1. If Solution C1 is precipitated, heat solution to 60°C until dissolved before use.
Solution C1 contains SDS and other disruption agents required for complete cell lysis. In addition to aiding in cell lysis, SDS is
an anionic detergent that breaks down fatty acids and lipids associated with the cell membrane of several organisms. If it gets
cold, it will precipitate. Heating to 60°C will dissolve the SDS and will not harm the SDS or the other disruption agents. In
addition, Solution C1 can be used while it is still hot.
Add 60µl of Solution C1 and invert several times or vortex briefly.
Fast Prep for
Soil 5.0 m/s for 45 sec.
Feces 4.5 m/s for 30 sec.
Pure culture 4.5 m/s for 30 sec
Note: The vortexing step is critical for complete homogenization and cell lysis. Cells are lysed by a combination of chemical
agents from steps 1-4 and mechanical shaking introduced at this step. By randomly shaking the beads in the presence of
disruption agents, collision of the beads with one another and with microbial cells causes the cells to break open.
229
6.
7.
8.
9.
10.
11.
12.
13.
14.
15.
16.
17.
18.
19.
20.
21.
22.
We have designed the Mo Bio Vortex Adapter as a simple platform to facilitate keeping the tubes tightly attached to the vortex. It
should be noted that although you can attach tubes with tape, often the tape becomes loose and not all tubes will shake evenly or
efficiently. This may lead to inconsistent results or lower yields. Therefore, the use of the Mo Bio Vortex Adapter is a highly
recommended and cost effective way to obtain maximum DNA yields.
Make sure the 2 ml PowerBead Tubes rotate freely in your centrifuge without rubbing. Centrifuge tubes at 10,000 x g for 30 seconds.
CAUTION: Be sure not to exceed 10,000 x g or tubes may break.
Transfer the supernatant to a clean microcentrifuge tube (provided).
Note: Expect between 400 to 500µl of supernatant at this step. The exact recovered volume depends on the absorbancy of your
starting material and is not critical for the procedure to be effective. The supernatant may be dark in appearance and still
contain some soil particles. The presence of carry over soil or a dark color in the mixture is expected in many soil types at this
step. Subsequent steps in the protocol will remove both carry over soil and coloration of the mixture.
Add 250µl of Solution C2 and vortex for 5 seconds. Incubate in the freezer for 10-15 minutes.
Solution C2 contains a reagent to precipitate non-DNA organic and inorganic material including humic acid, cell debris, and
proteins. It is important to remove contaminating organic and inorganic matter that may reduce DNA purity and inhibit
downstream applications for the DNA.
Centrifuge the tubes for 1 minute at 10,000 x g.
Avoiding the pellet, transfer up to 600µl of supernatant to a clean microcentrifuge tube (provided).
The pellet at this point contains non-DNA organic and inorganic material including humic acid, cell debris, and proteins. For
the best DNA yields, and quality, avoid transferring any of the pellet.
Add 200µl of solution C3 and vortex briefly. Incubate in the freezer for 10-15 minutes.
Solution C3 is a second reagent to precipitate additional non-DNA organic and inorganic material including humic acid, cell
debris, and proteins. It is important to remove contaminating organic and inorganic matter that may reduce DNA purity and
inhibit downstream applications for the DNA.
Centrifuge the tubes for 1 minute at 10,000 x g.
Transfer up to 750µl of supernatant to a clean microcentrifuge tube (provided).
The pellet at this point contains additional non-DNA organic and inorganic material including humic acid, cell debris, and
proteins. For the best DNA yields, and quality, avoid transferring any of the pellet.
Add 1.2ml of Solution C4 to the supernatant (be careful solution doesn’t exceed rim of tube) and vortex for 5 seconds.
Solution C4 is a high concentration salt solution. Since DNA binds tightly to silica at high salt concentrations, this solution will
adjust the salt concentrations to allow binding of DNA, but not non-DNA organic and inorganic material that may still be
present at low levels, to the spin filters.
Load approximately 675µl onto a spin filter and centrifuge at 10,000 x g for 1 minute. Discard the flow through and add an
additional 675µl of supernatant to the spin filter and centrifuge at 10,000 x g for 1 minute. Load the remaining supernatant onto
the spin filter and centrifuge at 10,000 x g for 1 minute. Note: A total of three loads for each sample processed are required.
DNA is selectively bound to the silica membrane in the spin filter device in the high salt solution. Almost all contaminants pass
through the filter membrane, leaving only the desired DNA behind.
This is the step where you add your replicates together on the filter.
Add 500µl of Solution C5 and centrifuge for 30 seconds at 10,000 x g.
Solution C5 is an ethanol based wash solution used to further clean the DNA that is bound to the silica filter membrane in the
spin filter. This wash solution removes residues of salt, humic acid, and other contaminants while allowing the DNA to stay
bound to the silica membrane.
Discard the flow through from the collection tube.
This flow through fraction is just non-DNA organic and inorganic waste removed from the silica spin filter membrane by the
ethanol wash solution.
Centrifuge again for 1 minute.
This second spin removes residual Solution C5 (ethanol wash solution). It is critical to remove all traces of wash solution
because the ethanol in C5 can interfere with many downstream applications such as PCR, restriction digests and gel
electrophoresis.
Carefully place spin filter in a new clean tube (provided). Avoid splashing any Solution C5 onto the spin filter.
Note: It is important to avoid any traces of the ethanol based wash solution.
Add 100µl of Solution C6 to the center of the white filter membrane. Let sit for 15 minutes.
Note: Placing the Solution C6 (sterile elution buffer) in the center of the small white membrane will make sure the entire
membrane is wetted. This will result in a more efficient release of the DNA from the silica spin filter membrane.
As the Solution C6 (elution buffer) passes through the silica membrane, DNA is released because it only stays bound to the
silica spin filter membrane in the presence of high salt. Solution C6 is 10mM Tris pH 8 and does not contain salt. Alternatively,
sterile DNA-Free PCR Grade Water may be used for elution from the silica spin filter membrane at this step (Mo Bio Catalog
No. 17000-10).
Note: Solution C6 contains no EDTA. If DNA degradation is a concern, Sterile TE may also be used instead of buffer C6 for
elution of DNA from the spin filter.
Centrifuge for 30 seconds.
Discard the spin filter. DNA in the tube is now application ready. No further steps are required.
We recommend storing DNA frozen (-20° to -80°C).
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STEP 2 – To see if your DNA extraction worked. You have two choices
1. DNA Quantitation by A260 UV spectrophotometer
• Make a 1/10 dilution of DNA and PCR water in a UV plate
OR
2. Electrophoresis
• Use 10 uL of DNA from extraction. Run on a 1% gel for 20-25 minutes at ≈100V
STEP 3
PCR (Forward primer is labeled with a Cy5 fluorescent tag)
Now the question is how much DNA to add to the PCR.
If you have spec’ed the DNA use 10 ng of DNA.
If you have run a gel, how bright is the band? Usually 1 uL of the straight DNA or 1uL of a 1/10 dilution
works well. If the band is super bright dilute it.
•
Two control reactions are needed
1. a closed negative (master mix, no DNA, not opened outside PCR room),
2. a positive (DNA known to amplify with PCR conditions).
• Use E. coli for general 16S eukaryotes
• Use H. volcanii for archea
• Ask about other positive controls for other primers sets
•
Run three reactions for each sample. The three reactions will be combined in a later step.
•
The following volumes are to be used for a 50 uL 16S PCR
5 µL - 10X Buffer
3 µL – dNTPs (10mM, 2.5mM of each, A,T,C,G)
2 µL – BSA (20ug/mL)
7 µL - MgCl2 (25mM)
1 µL - Ba2F (10 uM)
1 µL - K2R- (10 uM)
0.3 µL - AmpliTaq Gold (5U/uL)
Water to bring final volume, after adding DNA, to 50 uL
Template DNA (this is added last outside the PCR room)
•
Use the following cycling parameters.
94°C for 10 minutes
30 cycles of (94°C for 1 minute, 46.5°C for 1 minute, 72°C for 2 minutes)
72°C for 10 minutes
4°C soak.
STEP 4- to see if your PCR worked
Electrophoresis
•
Use 3-5 uL of PCR product. Run on a 1.5% gel for 20-25 minutes at ≈80-100V
STEP 5- to combine the PCR replicates that worked and remove leftover salts, dNTPs, and
primer
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Using MoBio PCR Ultra-Clean kit
1.
2.
3.
4.
5.
6.
7.
8.
9.
Add 5 volumes SpinBind solution to each well and pipet up and down well
Transfer 750 µl to the spin filter unit. Centrifuge for 30 sec at 10 x kg. Discard eluate.
Repeat step 3 until all PCR SpinBind mixture is filtered. You are combining the three PCR replicates
at this point.
Add 300 µl of SpinClean buffer to spin filter and Centrifuge for 30 sec. At 10 x kg. Discard eluate.
Centrifuge spin filter for 120 sec. At 10 x kg to remove any remaining fluid.
Transfer spin filter to clean 2.0 ml collection tube.
Add 60 µl of PCR water to spin filter and incubate 10 min.
Centrifuge for 60 sec. At 10 x kg.
Discard spin filter and store at –20oC.
STEP 6 – Quantitate PCR product
Using the Bio-Tek Fluorometer determine the PCR product concentration by measuring the Cy5
incorporated fluorescent label from the forward primer
STEP 7 – produce the labeled fragments
Enzyme Digests (Amount of DNA digested varies depending on the samples being prepared. Ask for
instructions before proceeding further.)
1.
2.
3.
4.
5.
6.
Digest 75 ng - 300 ng DNA.
Digest 5-10 ng of a E. coli digest standard. Do not use the E. coli genomic DNA. Use the digest
standard which is E. coli PCR product with the fluorescent label.
For DpnII (10,000 U/mL) use 0.6 uL-1.0 uL enzyme and 4 uL buffer per reaction. Add DNA and
water to bring the volume 40 uL
For HaeIII (10,000 U/mL) use 0.4 uL enzyme and 4 uL buffer per reaction. Add DNA and water to
bring the volume to 40 uL
For HhaI (20,000 U/mL) use 0.5 uL enzyme, 0.4 uL BSA, and 4 uL buffer per reaction. Add DNA
and water to bring the volume to 40 uL
Place tubes in PCR machine for 4 hours @ 37°C then cycle to either 65°C for DpnII , 65°C for HhaI,
or 80°C for HaeIII for 20 minutes to deactivate the enzyme and finally to 4°C for infinity.
7. Store the digests in the -20°C freezer until ready for ethanol precipitation.
STEP 8 – remove excess salts
Ethanol Precipitation (Note: prompt removal of samples from centrifuge will ensure minimal loss of
sample.)
1.
2.
3.
4.
5.
6.
7.
8.
9.
To the digest, add 100 µl (2.5 x digest volume) of cold 95% ethanol and 2 µl 3M NaAc pH4.6 (5%
digest volume) and 1 uL glycogen (20 mg/mL)
Invert five times making sure the lids are securely on.
Place the tubes in the -20°C freezer for 30 minutes.
Centrifuge the tubes for 15 minutes at 5300 RPM to pellet DNA. (program 2)
Remove ethanol by inverting the PCR tray on a paper towel.
Add 100 µl of cold 70% ethanol.
Centrifuge the tubes for 5 minutes at 5300 RPM. (program 3)
Remove ethanol by inverting the PCR tray on a paper towel.
Centrifuge rack in inverted position on top of a paper towel for 1 min. @ 700 RPM to dry the pellet.
(program 4)
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10. Store the DNA in the -20°C freezer until ready to proceed to CEQ8000 preparation.
STEP 9 – Separates the labeled digested fragments
CEQ 8000-sample preparation
1.
2.
3.
4.
Make a master mix of 20 uL formamide and 0.25 uL 600 base pair standard per reaction. Add 20
µl of the master mix to each tube.
Add one drop of mineral oil to the top of each well to prevent sample evaporation.
Run in the CEQ 8000
Look at your beautiful TRF patterns!!
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