impact of thermal remediation on the degradation of ......degradation of naphthalene by indigenous...
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Impact of Thermal Remediation on the Degradation of Naphthalene by Indigenous Anaerobic Bacteria in
Hydrocarbon Contaminated Soil
by
Kirstin Newfield
A thesis submitted in conformity with the requirements for the degree of Master of Applied Science Graduate Department of Civil Engineering
University of Toronto
© Copyright by Kirstin Newfield 2014
ii
Impact of Thermal Remediation on the Degradation of
Naphthalene by Indigenous Anaerobic Bacteria in Hydrocarbon
Contaminated Soil
Kirstin Newfield
Master of Applied Science, Civil Engineering University of Toronto
2014
Abstract
Thermal remediation is an efficient and cost effective method for the removal of organic
compounds from the subsurface. However, complete removal of these compounds cannot be
achieved by this technology alone. It is generally assumed that bioremediation will provide the
polishing steps at thermally treated sites. In this study, soil was collected from a hydrocarbon
contaminated site that previously underwent thermal remediation. A microcosm batch study was
conducted to determine the impacts of thermal remediation on indigenous microorganisms and
their ability to degrade naphthalene. Soils that reached varying peak temperatures were set up in
microcosms at temperatures experienced along their respective cooling profiles. Naphthalene
degradation was not detected within any of the unamended microcosms within a 6 month time
frame, although, archaea growth was detected in the microcosms after 2 months of acclimation,
accompanied by iron reduction and significant methane production assumed to have arisen from
degradation of methanol.
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Table of Contents Table of Contents ........................................................................................................................... iii
List of Tables .................................................................................................................................. v
List of Figures ................................................................................................................................ vi
Chapter 1 Introduction .................................................................................................................... 1
1.1 Introduction ......................................................................................................................... 1
1.2 Research Objectives ............................................................................................................ 3
Chapter 2 Literature Review ........................................................................................................... 4
2.1 Biodegradation of Naphthalene .......................................................................................... 4
2.1.1 Impacts of Increased Temperature on Microbial Activity ...................................... 5
2.2 Bioavailability of Naphthalene ........................................................................................... 8
2.2.1 Impacts of Increased Temperature on Bioavailability .......................................... 10
2.3 Combined Treatment Technologies .................................................................................. 11
2.4 Field Site History .............................................................................................................. 12
Chapter 3 Materials and Methods ................................................................................................. 15
3.1 Soil Preparation ................................................................................................................. 15
3.1.1 Soil pH .................................................................................................................. 15
3.1.2 Soil Bulk Density and Particle Density ................................................................ 16
3.2 Synthetic Groundwater Preparation .................................................................................. 16
3.3 Solution Preparation .......................................................................................................... 17
3.4 Microcosm Batch Experiment .......................................................................................... 18
3.4.1 Microcosm Preparation ......................................................................................... 19
3.4.2 Microcosm Analysis ............................................................................................. 20
3.4.3 Molecular Analyses for Total Bacteria and Total Archaea .................................. 25
Chapter 4 Results and Discussion ................................................................................................. 30
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4.1 Soil Properties ................................................................................................................... 30
4.2 Adsorption Isotherm Batch Study ..................................................................................... 34
4.2.1 Impact of Temperature on Adsorption .................................................................. 38
4.2.2 Impact of Soil Type on Adsorption ...................................................................... 42
4.3 Degradation Batch Experiment ......................................................................................... 43
4.3.1 Naphthalene Degradation ...................................................................................... 43
4.3.2 Microbial Analysis ................................................................................................ 46
4.3.3 Qualitative Observations ....................................................................................... 53
4.3.4 Microscopy ........................................................................................................... 54
4.3.5 Ions Analysis ......................................................................................................... 55
4.3.6 pH, Specific Conductivity and ORP ..................................................................... 59
4.3.7 Methane Production .............................................................................................. 60
Chapter 5 Conclusions and Recommendations ............................................................................. 62
5.1 Conclusions ....................................................................................................................... 62
5.2 Further Work ..................................................................................................................... 63
5.3 Recommendations ............................................................................................................. 64
References ..................................................................................................................................... 65
Appendix A ................................................................................................................................... 75
Appendix B ................................................................................................................................... 76
Appendix C ................................................................................................................................... 78
Appendix D ................................................................................................................................... 80
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List of Tables Table 1 Half reaction processes for varying redox conditions ....................................................... 4
Table 2 Physical and Chemical Properties of Naphthalene [35] .................................................... 8
Table 3 Composition of synthetic groundwater ............................................................................ 17
Table 4 Scenarios Tested in Microcosm Batch Experiment ......................................................... 18
Table 5 Microcosm Distribution for Degradation Study .............................................................. 20
Table 6 Microcosm Distribution for Adsorption Study ................................................................ 21
Table 7: Henry's constants adjusted for temperature .................................................................... 23
Table 8 qPCR Primers and References ......................................................................................... 27
Table 9 Physical and Chemical Properties of Each Soil Type ...................................................... 30
Table 10 Concentrations of contaminants remaining in each soil post thermal remediation ....... 33
Table 11 Experimentally determined Freundlich adsorption isotherm coefficients for Soil 1and 3
....................................................................................................................................................... 37
Table 12 Concentration of Metals in µg/L at 5 months of microcosms running, where *BDL is
below the detection limit ............................................................................................................... 56
Table 13 Concentration of anions in synthetic groundwater and Soil 3 supernatant at 155 days.
Concentrations in mg/L ................................................................................................................. 58
Table 14 Supernatant parameters at 149 days ............................................................................... 59
Table 15 Groundwater Chemistry at North Carolina Bulk Fuel Terminal ................................... 76
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List of Figures Figure 1 Naphthalene Structure, C10H8 ........................................................................................... 4
Figure 2 Relationship of temperature and growth rate showing varying classes of
microorganisms with respect to temperature .................................................................................. 6
Figure 3 The solubility of naphthalene in water and the diffusion coefficient with increasing
temperature [12] ............................................................................................................................ 10
Figure 4 Cross Section Representative of Site Subsurface (ARCADIS, 2005) [45] .................... 13
Figure 5 Optimization of SPME extraction time for naphthalene using a 50/30µm
DVB/CAR/PDMS fiber ................................................................................................................ 24
Figure 6 Adsorption Isotherms for Soil 1 at A) 30°C B) 45°C C) 60°C ....................................... 35
Figure 7 Adsorption Isotherms for Soil 3 at A) 30°C B) 45°C ..................................................... 36
Figure 8 Soil 1 adsorption isotherms for naphthalene at 30, 45 and 60ºC .................................... 38
Figure 9 Soil 3 adsorption isotherms for naphthalene at 30 and 45ºC .......................................... 38
Figure 10 Adsorption Isotherms for Naphthalene on Soil 1 and 3 at 30, 45 and 60ºC ................. 42
Figure 11 Change in Naphthalene Concentration at 30°C for Soil 1, 2 and 3 .............................. 44
Figure 12 Change in Naphthalene Concentration at 45°C for Soil 1, 2 and 3 .............................. 44
Figure 13 Change in Naphthalene Concentration at 60°C for Soil 1 and 2 .................................. 45
Figure 14 Total Bacteria Concentration in Soil Prior to Experimentation. Representing time = 0
in the microcosms ......................................................................................................................... 47
Figure 15 Concentration of Total Bacteria and Total Archaea in Soil 1, 2 and 3 at 30C at 8
weeks. Grey and black dashed lines represent the detection limits for total bacteria and archaea
respectively. .................................................................................................................................. 50
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Figure 16: qPCR results for total bacteria and total archaea at 149 days for each soil and
temperature scenario. S1, 2, 3-30, 45, 60 represent the soil number followed by the temperature
of that scenario. For example, S1-30 represents a triplicate microcosm for Soil 1 at 30ºC. ........ 52
Figure 17 Colour transformation of microcosms. A) Soil at time = 0 B) Active microcosm at
time = 5 months C) Control microcosm at time = 5 months. ....................................................... 54
Figure 18: Images from a Leica DMI 3000 Inverted Microscope, slides stained with Acridine
Orange and viewed under a red filter. A and B are fluorescence images of sample from active
microcosm of Soil 3 at 30C. C is fluorescence image of sample from control microcosm of Soil 3
at 30C ............................................................................................................................................ 55
Figure 19 Cation analysis at 119 days of microcosm monitoring ................................................ 56
Figure 20 Temperature profile for Soil 1 during thermal remediation ......................................... 78
Figure 21 Temperature profile of Soil 2 during thermal remediation .......................................... 78
Figure 22 Temperature profile of Soil 3 during thermal remediation .......................................... 79
1
Chapter 1 Introduction
1.1 Introduction Thermal remediation is an efficient and cost effective in-situ method for the removal of volatile
and semi-volatile organic compounds from the subsurface; however, complete removal of these
compounds, particularly low volatility compounds, cannot be achieved by this technology alone.
This is evident at a former bulk fuel terminal in Greensboro, North Carolina where 85% of the
initial total petroleum hydrocarbon concentrations were removed using the Electro-Thermal
Dynamic Stripping Process TM (ET-DSPTM) developed by McMillan-McGee Corp [1] [2]. It is
generally assumed that natural attenuation processes, such as biodegradation, will provide the
polishing steps to reduce the residual contaminant concentrations post thermal remediation [2].
As biodegradation is the only in-situ process that can completely eliminate parent hydrocarbons
[3], it is imperative to understand how thermal remediation will impact the indigenous microbial
communities in the target treatment area.
Typical bioremediation processes stimulate the growth of microbial populations that break down
the target contaminant to less harmful byproducts. Although these processes occur naturally, the
remediation of high concentration source zones usually requires conditions to stimulate this
growth. This can be accomplished by the addition of an electron acceptor (oxygen, sulfate,
nitrate), micronutrients, an energy source (carbon) or changing other parameters such as
temperature in the soil [4]. Bioaugmentation is an additional approach to stimulating
biodegradation; introducing a microbial population known to degrade the target contaminants
into the subsurface. This study focuses on the impact of elevated temperatures on the ability of
indigenous microbial communities to degrade low volatility hydrocarbons typically remaining
post thermal remediation. Although the impact on increased temperatures has been studied on
bacteria isolated from high temperature environments [5] [6], limited research is available
regarding these impacts on indigenous microbial communities in the subsurface.
Naphthalene is a semi-volatile, low weight polycyclic aromatic hydrocarbon (PAH). Due to the
low vapor pressure, low solubility and high sorption coefficients, naphthalene is not easily
remediated using thermal treatment and remediation is reliant on biodegradation [2], providing
an ideal compound for this study. Naphthalene originates from the incomplete combustion of
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hydrocarbons and is a constituent of crude oil. It is widespread in the environment due to both
anthropogenic and natural sources, although the problematic source zones typically stem from
the release of petroleum products and coal-derived products [7]. Within the past decade a range
of 67,000-121,000 kgs of naphthalene have been released by industry into the Canadian
environment each year [8]. The USEPA and the Agency for Toxic Substances and Disease
Registry listed naphthalene as one of the most commonly found substances at hazardous waste
sites on the National Priorities List. It is important to understand the fate of naphthalene in the
environment as it poses a risk to human health, the environment and wildlife [7]. Naphthalene is
categorized by the International Agency for Research on Cancer (IARC) as a possible carcinogen
to humans [9].
Microcosm batch studies were conducted to simulate the in-situ anaerobic degradation of
naphthalene by indigenous microorganisms at the ET-DSPTM remediated, former bulk terminal
previously contaminated with hydrocarbons in Greensboro, North Carolina. Soil samples came
from three locations that experienced different peak temperatures through the remediation
process: 37, 70 and 101ºC. Each soil was tested for sorption capacity and microbial activity at
temperatures representative of what each soil experienced post thermal remediation: 30, 45 and
60ºC. The variations in microbial communities at each peak temperature were monitored to
determine the resilience of these microbes at high temperatures. Improving the understanding of
the potential for bioremediation to treat naphthalene remaining after thermal remediation may
lead to the development of a cost-effective strategy for a thermal treatment – bioremediation
treatment train approach.
All work and writing presented in this thesis was completed by the author with the exception of
the qPCR method which was completed by Dr. Pulin Mundal and Dr. Simone Larcher. Review
and editing was conducted by Dr. Brent Sleep. Some routine laboratory preparations were
performed by Viviane Malveira Cavalcanti and Mateus Xavier De Lima. qPCR method
development and analysis was completed by Dr. Pulin Mundal and Dr. Simone Larcher.
3
1.2 Research Objectives A microcosm scale study was conducted to determine the impacts of thermal remediation on the
degradation of naphthalene by indigenous microbial communities at a hydrocarbon contaminated
site. The following describes the detailed objectives.
• To develop adsorption isotherms for each soil (from locations with peak site temperatures
of 101, 70, and 37 C) at the temperatures of 30, 45 and 60ºC, to determine the availability
of naphthalene in the aqueous phase for each condition.
• To monitor microbial activity for each soil in anaerobic conditions with naphthalene as
the carbon source.
• To determine the potential for degradation for each soil using microcosm batch studies
maintained at 30, 45 and 60°C.
4
Chapter 2 Literature Review
2.1 Biodegradation of Naphthalene Naphthalene is a polycyclic aromatic hydrocarbon composed of two fused benzene rings, shown
in Figure 1.
Figure 1 Naphthalene Structure, C10H8
Many bacteria, fungi, cyanobacteria and algae capable of degrading naphthalene have been
isolated from soils [10] in a variety of reducing conditions. Aerobic studies have been
extensively studied [11] [10] [12] [13] although many contaminated source zones occur in
anaerobic environments [10]. This has led to the study of naphthalene degradation in a variety of
anaerobic environments. It has been shown that naphthalene degradation may occur under nitrate
reducing [14] [13] [15], sulfate reducing [16] [17] [18] [19], methanogenic [20] and iron
reducing conditions [21] [22] [23]. The half reactions for these processes are shown in Table 1.
Table 1 Half reaction processes for varying redox conditions
Oxidation Process Half Reaction Aerobic Respiration 0.25O2 + H+ + e- = 0.5 H2O Iron Reduction Fe3+ + e- = Fe 2+ Denitrification 0.2NO3
- + 1.2 H+ + e- = 0.1N2 +0.6H2O Sulfate Reduction 0.125SO4
2- + 1.1875H+ +e- + 0.0625H2S + 0.0625HS- + 0.5H2O Methanogenesis 0.125CO2 + H+ + e- = 0.125CH4 + 0.25H2O Reduction Process Half Reaction Naphthalene 1/48C10H8 + 5/12H2O = 5/24CO2 + H+ + e-
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Anaerobic processes typically have lower degradation rates than aerobic degradation [24] and
usually are not a significant pathway for naphthalene degradation in natural environments [25].
Anaerobic processes, however, can be stimulated with amendments to the soil such as
macronutrients (nitrogen, phosphorous, potassium), micronutrients (trace metals) and
physical/chemical parameters such as temperature and pH [26].
Microorganisms require the proper nutrients such as nitrogen, potassium, and phosphate and
micronutrients such as trace metals for growth to occur. Generally, microorganisms will not
suffer from nutrient depletion if the substrate concentration is less than ppm levels [27]. In the
case of the former bulk fuel terminal, concentrations ranged in the ppm levels for naphthalene
along with many other hydrocarbons at high concentrations. Due to the complexity of factors
required to satisfy biodegradation, an efficient bioremediation process is typically only possible
by the amendment of the soils to meet all of the conditions, however, this study will focus on the
impacts of temperature on indigenous microbial communities capable of degrading naphthalene
in an unamended subsurface.
2.1.1 Impacts of Increased Temperature on Microbial Activity
Temperature is a critical factor impacting the growth and survival of microorganisms [26]. Each
microorganism has a minimum temperature at which they can function and as temperatures are
increased, the chemical and enzymatic reaction rates in the cell increase up to the optimal
temperature for growth. As the temperature increases above this level, inhibitory temperatures
can be reached for microorganisms where all molecules lose their structure and functionality in a
process referred to as denaturation [26]. This is temperature and time dependent; it takes longer
at lower temperatures to sterilize the same microbial population than it would at higher
temperatures. This is also impacted by the moisture content as moist heat has a better penetrating
power than dry heat and results in a higher rate of destruction of living organisms at a given
temperature [26]. This phenomenon is often used to sterilize water and soil when microbial
communities are not performing a desirable function. The temperature range varies for each
particular microbial species with general classifications of microorganisms shown in Figure 2.
Microorganisms have the potential to survive these unfavourable conditions through the
formation of non-vegetative structures such as spores which are less metabolically active and
have the potential to persist in these harsh environments [26].
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Figure 2 Relationship of temperature and growth rate showing varying classes of
microorganisms with respect to temperature [28]
Thermophiles can be isolated from a variety of environments such as soils, sewage, compost,
rivers, lakes and seawaters, sediments, geothermal and hydrothermal environments [29]. Many
of the thermophilic microorganisms that consume naphthalene as a sole carbon source have been
isolated from compost [30]. Hyperthermophilic microorganisms, however, are typically
restricted to extreme environments such as hot springs, geysers and deep-sea hydrothermal vents.
Many naphthalene and other hydrocarbon degrading microorganisms have been isolated from
such areas [6] [31].
Figure 2 indicates that thermophiles typically have higher growth rates at their optimal
temperatures than do mesophiles. PAH biodegradation typically occurs slowly in mesophilic
conditions, between 20ºC and 40ºC [32] [6], however, studies have shown that enhanced
biodegradation can occur at elevated temperatures [33] [32].
Hemalatha and VeeraManikandan (2011) found that the optimal temperatures for PAH
degradation by aerobic Flavobacterium spp l, Pseudomonas spp 1, and Pseudomonas spp 2
isolated from hydrocarbon contaminated sites were 40ºC, 40ºC and 45ºC, respectively. However,
these results were representative of hydrocarbon utilization over a 24 hour testing period and did
not experience the peak temperatures or duration of heating typically experienced at thermal
remediation sites [33]. Annweiler et al. (2000) found that Bacillus thermoleovorans, an aerobic
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thermophilic bacteria isolated from contaminated compost, metabolized naphthalene at an
optimal temperature of 60ºC [11]. Lin et al. (2012) found thermophilic Pseudoxanthomonas
grew at 70ºC and survived during a decrease in temperature to 55ºC during thermophilic
biodegradation of diesel oil in food waste composting [34]. Feitkenhauer et al. (2003) isolated
several aerobic microorganisms capable of degrading aromatic hydrocarbons from hot springs,
compost piles and industrial wastewater which had an optimal growth temperature between 60-
70ºC [6]. These studies indicate that hydrocarbon biodegradation can be enhanced at a range of
elevated temperatures, although limited information is available regarding the potential for in-
situ, indigenous microorganisms to degrade naphthalene at the elevated temperatures
experienced during thermal remediation. There is also limited information on anaerobic
thermophilic microorganisms and their ability to degrade PAH.
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2.2 Bioavailability of Naphthalene Biodegradation can be accomplished only when the substrate, in this case naphthalene, is in a
state where the microorganisms can access it. Typically, microorganisms exist in the aqueous
phase, thus, the bioavailability of the hydrocarbons is dependent on mass transfer from the
NAPL or sorbed phase to the aqueous phase [35] [12]. This is driven by both the properties of
the compound, naphthalene, and the properties of the soil.
Naphthalene is a semi-volatile, compound with low aqueous solubility and high sorption
coefficients, suggesting it is not readily available in the aqueous phase at typical subsurface
temperatures. Physical and chemical properties for naphthalene are shown in Table 2.
Table 2 Physical and Chemical Properties of Naphthalene [36]
Property Naphthalene
Molecular Weight 128.19 g/mol
Melting Point 80.5 ºC
Boiling Point 218ºC
Density at 20ºC 1.145 g/ml
Water Solubility at 25ºC 31.7 mg/L
log Kow 3.29
log Koc 2.97-3.27
Vapour Pressure 0.087mmHg
Henry's Law Constant 4.6x10-4 atm-m3/mol
In addition to the compound’s properties, the availability is also dependent on the fraction of
organic content of the soil, foc [37, 38]. A linear relationship has been demonstrated for the
9
adsorption isotherms of naphthalene with a directly proportional relationship between Kd and foc
[37]. The adsorption can be predicted by the following relationship:
𝑲𝒐𝒄 = 𝑲𝒅𝒇𝒐𝒄
(1)
Where, Koc is the soil-water partition coefficient for a specific organic compound (L/kg), in this
case naphthalene, and is independent of the soil or sediment type, and Kd is the partition
coefficient. As the organic carbon content in the soil increases the sorption also increases.
As previously mentioned, the soil properties can also impact the availability of naphthalene in
the aqueous phase. Owabor et al. (2012), found a significant difference in the amount of
naphthalene adsorbed to two different soil types, clay and sand. The clay adsorbed significantly
higher amounts of naphthalene than the sand while the sand was able to retain more naphthalene
during desorption than the clay [39]. It has also been found that residual contaminant
concentrations increase with increasing organic matter and clay content in the soil [25]. ERH is a
desirable remediation technology for heterogeneous sites with complex subsurfaces containing a
range of soil compositions. These variations in adsorption and desorption with varying soil types
may significantly impact the bioavailability of naphthalene within the varying layers.
The contact time between the contaminant and the soil also impacts the potential for
biodegradation. Increasing the contact time has been found to decrease the effectiveness of
biodegradation [25]. Studies have shown that freshly spiked contamination could be completely
degraded while contamination that had been present in soil over a longer period of time was
susceptible to residual contamination remaining post biodegradation [25]. This may be due to
adsorption processes that occur over a longer period of time (months to years) such as micropore
diffusion and intra-organic matter diffusion [40]. Once these adsorptive processes take place, in
subsurfaces exposed to contamination for a long period of time, it is more difficult for desorption
to occur.
Additionally, factors such as temperature and pH can impact the adsorption of a compound. The
impacts of temperature will be the focus of this study and described in more detail in the
subsequent section.
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2.2.1 Impacts of Increased Temperature on Bioavailability
The bioremediation of PAHs can proceed slowly at low temperatures as PAHs have limited
water solubility and low mass transfer rates from the adsorbed phase to the aqueous phase [6]
[32]. Elevating temperatures can increase the bioavailability of the PAH compounds by
increasing the solubility and mass transfer rates of PAH to aqueous solutions [32] [12] [29].
Feitkenhauer and Markl (2003) found that the mass transfer coefficient and solubility of
naphthalene increased by factors of approximately 5 and 10, respectively, with an increase in
temperature from 20ºC to 70ºC [12]. This relationship is shown in Figure 3.
Figure 3 The solubility of naphthalene in water and the diffusion coefficient with increasing
temperature [12]
Viamajala et al. (2007) also found an increase in mass transfer rates and equilibrium solubility
concentrations of PAHs at increased temperatures. However, it was determined that although the
mass transfer rates are significantly increased with increased temperature, the degradation of
PAH is still mass transfer rate limited at these increased temperatures [32].
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2.3 Combined Treatment Technologies Thermal remediation technologies are typically designed to optimize the vapour extraction of
contaminants, with the assumption that there is an added benefit of increased biological activity
resulting from the increased temperatures. Many industries refer to the rule of thumb that for
every 10°C increase in temperature the microbial population doubles [41], often omitting that
this occurs within the limited temperature range for each particular class of microorganisms [42].
Many of these assumptions lack substantive evidence, especially regarding the compounds that
are most likely to remain post thermal remediation, (example: low-volatility PAH). There is also
disregard for the conditions required to support the microbial communities. As stated earlier,
bioremediation requires amendments to the soil to enhance the degradation of the target
compounds and has temperature dependent limits in which biodegradation can take place. If the
temperature profile of thermal remediation sites can be designed to optimize the degradation of
low-volatility contaminants while maintaining efficient rates of vapour extraction of the higher
volatility compounds, the costs and efficiency of a combined thermal and bioremediation
technology can be improved.
Although there has been little research conducted on a combined technology for PAHs, there has
been research conducted for chlorinated solvents. Friis et al. (2006) studied amended and
unamended soils for the potential of TCE dechlorination during thermal remediation. The soils
were taken from a contaminated site that had not undergone prior treatment. Thermal
remediation conditions were simulated at the microcosm scale and concluded that complete
dechlorination was not observed in any of the unamended soils [43], suggesting that some form
of bioaugmentation would be required in combination with thermal treatment to obtain
conditions favourable to biodegradation.
TRS Group, Inc. has documented research into combining thermal and bioremediation at
chlorinated solvent contaminated sites. The thermal treatment reached an optimal temperature of
37ºC increasing the amount of microorganisms 46 to 100 fold over a time period of 3 months
[44] [45]. In order to achieve these results, the subsurface was amended with nutrients to
enhance the degradation rates through injection ports located along the electrodes [45]. It was not
stated whether these conditions were aerobic or anaerobic. This indicates the need to augment
the subsurface to achieve efficient biodegradation, especially during thermal treatment. It also
12
suggests a significantly lower optimal temperature than was is typically experienced at thermal
remediation sites (75-100°C).
These studies show the potential for a combined remediation technology, however, amendments
to the subsurface and alterations to the typical temperature profiles are required for post-thermal
bioremediation to take place.
2.4 Field Site History Soil samples were taken from a former bulk fuel terminal in Greensboro, North Carolina which
was operational between 1973 and 1990. During this time it was heavily contaminated with
leaded and unleaded gasoline, #2 diesel fuel and kerosene. An estimated 1.7 million liters of
hydrocarbons, mainly comprised of gasoline and diesel fuel, were released on the site. The
LNAPL plume was comprised of 7.4 weight% naphthalene. In 1994, the excess fuel products
were removed and the facilities were decommissioned, removing above ground and subsurface
structures. In 1996 a dual-phase extraction system was installed, removing approximately 42,000
liter equivalents of hydrocarbons by 2003. In 2004, a pilot study investigated 3 different
remediation technologies; 1. Dewatering and high-vacuum extraction 2. Air sparge and high-
vacuum extraction and 3. ET-DSPTM. Air sparging combined with vapour extraction was found
to promote aerobic biodegradation in the soil with an estimated degradation rate between 606
and 1,985 mg total hydrocarbons per kg soil per year, however, the estimated remedial time
frame was estimated at 15-20 years compared to the ET-DSPTM method which had an estimated
remedial time frame of 3 years. Due to the complex geology of the site and the efficiency
determined by the pilot-scale test, ET-DSPTM was implemented and by 2007 85% of the initial
total petroleum hydrocarbon concentrations were removed in one of the four contaminated zones
on the site [1].
Electrical-resistance heating requires the installation of electrodes throughout the subsurface of
the contaminated area. A three-phase voltage is applied to the electrodes, heating the subsurface
soil and groundwater to temperatures up to 100ºC. At the North Carolina site, in addition to
conventional electrical resistance heating, heated groundwater was added at the electrodes,
preventing desiccation of the surrounding soil, maintaining subsurface moisture levels, and
electrical conductivity. The contaminants were either vapor extracted or flushed to the extraction
wells, driven by the influx of heated water [1]. ERH effectively targets compounds with vapour
13
pressures greater than 5mmHg at 10ºC [41], however, compounds that do not meet this threshold
with vapour pressures less than 5mmHg at 10 ºC rely on biodegradation to remove any residual
contamination, including PAHs.
The subsurface is composed of sandy silt and clayey silt saprolites which are weathered soils
from underlying metamorphosed granitic to dioritic parent bedrock. The consolidated bedrock is
located 60-80ft bgs. Figure 4 shows a cross section obtained approximately 150m north of where
the soil samples for this study were extracted. The saprolite contains relic structures of mineral
veins, fractures and metamorphic foliations which create a secondary porosity and permeable
micro-conduit in the soil. The water table undergoes fluctuations between 8ft and 20ft bgs. All of
the samples for this study were taken >20ft bgs to ensure saturated conditions. The site is
situated on regional and local divides and is also located in a groundwater recharge zone causing
dynamic shifts in the groundwater movement within the site. Concentrations of naphthalene vary
throughout the site reaching concentrations of 3900µg/L, however the concentrations were very
low in the region where the soil samples for this study were taken (~1µg/L). A concentration
gradient map is shown in Appendix A.
Figure 4 Cross Section Representative of Site Subsurface (ARCADIS, 2005) [46]
14
The subsurface at the field site in Greensboro, North Carolina has low dissolved oxygen
concentrations, < 0.05 mg/L and high iron and sulfate concentrations of approximately 95mg/L
and 4mg/L respectively. A detailed water chemistry analysis is found in Appendix B.
15
Chapter 3 Materials and Methods
3.1 Soil Preparation Soil samples were obtained from three locations at varying depths and peak temperatures from
the outer perimeter of the thermally remediated zone at the former bulk fuel terminal in
Greensboro, North Carolina; 37 ºC at 36.2 ft. bgs, 70 ºC at 26.5 ft. bgs and 101 ºC at 27.5 ft. bgs.
Sample extraction was organized by McMillan McGee and samples were extracted by a
subcontractor, GES. All samples were taken from the saturated zone where biodegradation is
more favourable. The cores were taken using a geoprobe rig and were 2’ by 2” diameter. The
samples reached peak temperatures in 2011-2012, temperature profiles can be found in Appendix
C. Each soil core was sampled and sent for analysis by Maxxam Analytics for polyaromatic
hydrocarbons, BTEX and F1 hydrocarbon fractions, F2-F4 hydrocarbon fractions and moisture
content on December 20, 2012. Samples of homogenized soil were sent for cation exchange
capacity and organic content analysis by University of Guelph, Laboratory Services, Agriculture
and Food Laboratory on July 18, 2013. The samples were stored at 4°C.
3.1.1 Soil pH
The pH of the soil was determined using the Standard Test Method for pH of Soils, ASTM
D4972-01(2007) [47]. The soil pH is tested using a sensitive electrode system (Radiometer
Analytical PHM 92 pH meter) in both a suspension of water and a suspension in 0.01M calcium
chloride solution.
The calcium chloride solution was prepared in a volumetric flask combining 14.7g of
CaCl22H2O in 100ml MilliQ® water to obtain a stock concentration of 1.0M CaCl2. An
intermediate stock solution was prepared combining 10ml of 1.0M stock CaCl2 1000ml MilliQ®
water to obtain a concentration of 0.01M.
10grams of the soil were placed into a glass container and 10ml of MilliQ® water was added.
The same procedure was completed for the CaCl2 solution, placing 10grams of the soil in a glass
container and adding 10ml of 0.1M CaCl2 solution. The suspensions were mixed thoroughly and
16
left to stand for 1hr. Once complete the pH was tested in both suspensions at room temperature
(15-25°).
3.1.2 Soil Bulk Density and Particle Density
Bulk density of the homogenized soil core was taken to determine the exact volume of soil added
to each microcosm. The bulk density was calculated using the following formula:
𝜌𝑏 = 𝑀𝑠𝑉𝑡
(2)
A hollow glass tube, open at both ends was used to measure the mass, Ms (g), and volume, Vs
(cm3), of the soil. The glass tube was measured for weight and inner diameter. It was then driven
straight into the soil. The height of the soil and the weight of the soil in the tube were then taken.
The particle density was determined to classify the soil and calculate the porosity of the
homogenized soil. The following formula was used, where Msolids is the mass of the solid
particles (g), Vs is the volume of the solids without pore space (cm3):
𝜌𝑝 = 𝑀𝑠𝑜𝑙𝑖𝑑𝑠𝑉𝑠
(3)
A sample of soil was dried and weighed in a 25ml volumetric flask. The flask was filled part way
with distilled water, stoppered, agitated and left to stand for 48 hours. This step was done to fully
saturate the pore space within the soil and remove any air pockets. Once saturated the flask was
filled to the graduation mark and weighed again. Using this data the volume of the displaced
water was calculated and the exact volume of the solid particles was determined.
The porosity was then calculated from the previously determined data as follows:
𝑛 = 1 − 𝜌𝑏𝜌𝑝
(4)
3.2 Synthetic Groundwater Preparation A mineral salt medium was prepared to simulate the groundwater conditions at the site,
amenable to anaerobic biodegradation of naphthalene, consisting of the following: (stock
solutions per liter of autoclaved distilled, Milli-Q® water): 1ml of calcium solution (80.2g/L
17
CaCO2), 1ml of iron solution (11.6g/L iron(II) sulfate), 1ml of magnesium solution (268.5 g/L
MgCl2) and 1ml of nitrate solution (4.2g/L concentration NaNO3). The solutions were prepared
in autoclaved MilliQ® water and filtered with a 250 ml Nalgene Sterile Filter Unit to ensure
sterility. Final concentrations of ions are shown in Table 3.The mineral salt medium was purged
for 2 hours using nitrogen gas to remove oxygen. Either 0.1N hydrochloric acid or 0.1N sodium
hydroxide was added to the medium to obtain a pH of 7. The solution was measured for specific
conductivity and redox potential.
Table 3 Composition of synthetic groundwater
3.3 Solution Preparation Naphthalene solutions were prepared to calibrate the GC and spike the microcosms. Methanol
(Sigma Aldrich Chromosolv ® for HPLC, ≥99%) was used to prepare stocks with concentrations
>10mg/L and distilled, Milli-Q® water was used to prepare standards with concentrations
<10mg/L. The solid naphthalene (Aldrich, >99%) was measured gravimetrically to create the
initial stock in HPLC grade methanol. Once in the aqueous phase the standards were prepared
Salts
Final Concentration of Ion in Synthetic Groundwater
(mg/L)
Calcium Chloride Di hydrate
Calcium 21.9 Chloride 38.7
Ferrous Sulfate Heptahydrate
Iron 2.3 Sulfate 4.0
Magnesium Chloride Hexahydrate
Magnesium 32.2 Chloride 93.8
Sodium Nitrate
Sodium 1.1
Nitrate 0.7
Total Chloride 132.5
18
volumetrically to the desired concentrations and stored in 20ml glass vials with septum screw
caps. The microcosm degradation batch study spiking solution was prepared to 610 mg/L in
methanol and stored in a 250ml amber bottle with a septum cap. 1ml of spiking solution was
used in each microcosm to obtain a concentration of 10mg/L naphthalene.
3.4 Microcosm Batch Experiment A microcosm batch experiment was conducted to determine the impacts of increased
temperatures experienced during thermal remediation on the potential for biodegradation of
naphthalene by indigenous microbial communities. Soil samples from a thermally remediated
site, described previously, were used in this study. Each soil type was tested at varying
temperatures that would be experienced along the cooling profile to determine the optimal
temperature for degradation to take place: 30, 45 and 60ºC. A total of 8 scenarios were tested,
shown in Table 4.The three soil types at varying peak temperatures were used to determine how
the peak temperatures impact the microbial communities in the soil and their potential for
biodegradation post thermal remediation.
Table 4 Scenarios Tested in Microcosm Batch Experiment
Scenario Peak Temperature
Reached in the Field
Temperature Tested in the
Laboratory
1 37 30
2 37 45
3 70 30
4 70 45
5 70 60
6 101 30
7 101 45
8 101 60
19
3.4.1 Microcosm Preparation
The microcosms were prepared in a polyethylene anaerobic glove bag (Atmosbag, Sigma-
Aldrich) filled with 20% CO2 and 80% Nitrogen gas mix (Praxair) to maintain anaerobic
conditions in the microcosms. All of the glassware and stainless steel items were washed with
Fisherbrand, Sparkleen detergent, rinsed with Milli-Q® water and autoclaved prior to use. All
other items were washed and cleaned with 70% ethanol prior to use.
The casing of the soil core was cut partially around the circumference of the casing at 2 inch
intervals using a sterilized hacksaw blade in an aerobic environment in the fumehood. Each
incision point was sealed with parafilm to ensure limited oxygen entered the core. Once cut, the
entire core was handled within the anaerobic glovebag. The glovebag was filled and purged
twice before the preparation. The soil core was split into smaller cylinders along the incision
points described above. The soil from the inner diameter of the core was extracted and
homogenized to ensure consistency in each microcosm and to avoid using the outer edge of the
soil that may have been exposed to oxygen. Any of the homogenized soil that was not used in the
initial setup was stored in autoclaved mason jars and refrigerated at 4ºC for future use.
Each microcosm scenario was prepared in triplicate in a 100 ml serum bottle (total volume of
120ml) sealed with a 20mm blue chlorobutyl septum stopper (Bellco Glass) and capped with a
stainless steel crimp cap to secure the stopper. Each serum bottle was filled with approximately
10-15g of soil and 60 ml of synthetic groundwater (as prepared previously) before being sealed.
The bottles were weighed upon each addition. Each microcosm was spiked to achieve a total
aqueous concentration of 10mg/L (1ml of 610 mg/L naphthalene in methanol solution). The
control bottles were prepared in duplicate and spiked to obtain a concentration of 0.2 g/L sodium
azide and 0.5 g/L mercuric chloride to inactivate any microorganisms that may have been present
in the soil. Resazurin (50uL of 1 g/L stock solution) was injected into selected microcosms to
ensure anaerobic conditions were maintained.
The increased temperatures were obtained through a series of water baths held at 60ºC, 45ºC and
30ºC. The microcosms were placed up to the neck in deionized water. Water was replenished
regularly with heated DI water when levels approached the water level in the microcosm bottle.
The water baths were covered to prevent photodegradation of the microcosms. The series of
microcosms at each condition are shown in Table 5. Each soil was tested at temperatures
20
experienced along the in-situ cooling profiles and was not tested at temperatures far exceeding
the peak temperature that was experienced in-situ.
Table 5 Microcosm Distribution for Degradation Study
Number of Microcosms
Temperature (ºC)
Soil 1 Controls Soil 1 Soil 2
Controls Soil 2 Soil 3 Controls Soil3
60 2 3 2 3
45 2 3 2 3 2 3
30 2 3 2 3 2 3
3.4.2 Microcosm Analysis
Although the naphthalene was directly injected into the aqueous phase in each microcosm,
overtime the compound partitions into the gaseous and sorbed phases. In order to determine the
actual amount that was degraded due to microbial activity a mass balance was performed for all
three of these phases: aqueous, sorbed and gaseous phases. The mass balance is given by the
following equation:
𝑴𝑻 = 𝑴𝒂𝒒 + 𝑴𝒈 + 𝑴𝑺 + 𝑴𝒅 (5)
Where, MT, g, aq, S, d are the total mass, mass in the gaseous, aqueous and sorbed phase and mass
degraded respectively in µg. The total mass of naphthalene is added to the microcosms in a
known amount. The mass in the aqueous phase is determined by the analytically determined
concentration in the aqueous phase (µg/L), Caq, and the known volume of the aqueous phase, Vaq
(L), expressed by equation 6. The mass in the gaseous phase is determined theoretically, as
described in a subsequent section, using the theoretically determined gaseous concentration
(µg/L), Cg, and the volume of the gaseous phase, Vg (L), expressed by equation 7. The adsorbed
mass is calculated using the sorbed concentration (µg/g), determined experimentally and
expressed as adsorption isotherms, and the known mass of the soil in each microcosm, expressed
by equation 8.
𝑴𝒂𝒒 = 𝑪𝒂𝒒𝑽𝒂𝒒 (6)
21
𝑴𝒈 = 𝑪𝒈𝑽𝒈 (7)
𝑴𝑺 = 𝑺𝑴𝒔𝒐𝒊𝒍 (8)
The change in the total mass over time will give the rate of degradation of naphthalene, rd,
expressed by equation 9.
𝒓𝒅 = ∆𝑴𝑻𝒕
(9)
3.4.2.1 Adsorption Isotherm Preparation and Modeling
A batch experiment was conducted for each soil type to determine the adsorption isotherms for
varying concentrations at each temperature. Microcosm bottles were prepared as described
previously; approximately 10-15g of soil and 60ml of synthetic groundwater were added to the
serum bottle in an anaerobic glovebag and sealed with a chlorobutyl stopper and stainless steel
crimp cap. Each microcosm was spiked to a concentration of 2 g/L sodium azide and 0.5 g/L
mercuric chloride to inactivate any microorganisms that may have been present in the soil. The
microcosms were then agitated and left for a period of 24 hours to ensure the sterilization of
microorganisms. This was to ensure that all reductions in naphthalene concentration were a result
of adsorption and not a result of biological activity. The microcosms were then spiked with
varying concentrations of naphthalene in methanol solution with a range of 0.5ml – 2.5ml total
solution added. The distribution of microcosms is shown in Table 6. Control bottles were
prepared without the addition of soil.
Table 6 Microcosm Distribution for Adsorption Study
Concentration of Naphthalene (µg/L)
Number of Microcosms with Soil
Number of Control Microcosms without Soil
100 2 2
500 2 2
2000 2 2
6000 2 2
12000 2 2
The microcosms were first tested at 30ºC. Aqueous samples were taken at time 0, 24 and 48 hr to
determine the equilibration time. Previous studies show that equilibration time typically occurs
22
within 24 hours [37] [39]. Once sampled the microcosms were transferred to 48ºC and allowed
to equilibrate for 48 hours before sampling. This was further completed at 60ºC.
The Freundlich adsorption model has been previously demonstrated to represent PAH adsorption
in water/sediment and water/soil systems [38].
𝑺 = 𝑲𝑪𝒂𝒒𝑵 (10)
where, S is the sorbed concentration of naphthalene to soil (µg/g) determined through mass
balance as previously described, K and N are empirical constants determined through batch
experiments and Caq is the equilibrium concentration in the aqueous phase (µg/L).
This can be displayed in the form of a linear equation on a logarithmic scale as follows:
𝒍𝒐𝒈𝑺 = 𝟏𝒏�𝒍𝒐𝒈𝑪𝒂𝒒� + 𝒍𝒐𝒈𝑲 (11)
The amount of adsorption was determined using this relationship for the degradation study.
3.4.2.2 Gaseous Analysis
The mass in the gaseous phase was determined using Henry’s Law, equation 12, and the Ideal
Gas Law, equation 14.
𝒑 = 𝒌𝑯 ∗ 𝑪𝒂𝒒 (12)
Where, the partial pressure, p, is dependent on the Henry’s constant for naphthalene, kH (𝑎𝑡𝑚∗𝑚3
𝑚𝑜𝑙),
and the aqueous concentration, Caq (µg/L), in the microcosm. Henry’s constant is temperature
dependent and is adjusted using the Van’t Hoff equation as follows:
𝒌𝑯(𝑻) = 𝒌𝑯(𝑻°)𝐞𝐱𝐩 [−𝑪�𝟏𝑻− 𝟏
𝑻°�] (13)
Where, T is the target temperature, Tº is standard temperature, 298K and C is a temperature
dependence constant. C for naphthalene is 3600K [48].
The Henry’s Constants calculated for the temperatures of interest in this study are found in Table
7.
23
Table 7: Henry's constants adjusted for temperature
Temperature (ºC) Henry’s Constant (𝒂𝒕𝒎∗𝒎𝟑
𝒎𝒐𝒍)
30 0.00056
45 0.00098
60 0.00164
The amount of naphthalene in the gas phase can then be determined using the Dalton’s Law as
follows:
𝒏𝑨 = 𝒑𝑨𝑽𝑹𝑻
(14)
where, n is the number of mol of naphthalene in the gas phase of volume, V (ml), R is the
universal gas constant, 8.3145 𝐽𝑚𝑜𝑙∗𝐾
and T is the target temperature (K).
3.4.2.3 Aqueous Sampling and Analysis
The microcosm stopper was wiped with 70% ethanol prior to extracting a 1ml sample through
the stopper using a 1ml glass syringe (gastight #1001 Hamilton) with a disposable 22 gauge
needle (BD PrecisionGlideTM Needle). The sample was dispensed through a 0.22 micron filter
(Millex Syringe driven filter unit with PVDF membrane) in combination with a 25 gauge needle
(BD PrecisionGlideTM Needle) into a crimp sealed, 2ml amber GC vial containing 0.3ml of
Milli-Q® water.
The analysis of naphthalene was conducted using the Agilent Technologies 7890A GC system
with flame ionization (GC-FID) and a DB 624 capillary column. An automated CTC CombiPal
was used in combination with the GC-FID to conduct solid phase micro extraction (SPME).
SPME was conducted by inserting a 24 gauge, 50/30µm DVB/CAR/PDMS fiber (Supelco)
through the septa (thermogreen LB-2, 11mm, Supelco) and immersing it in the liquid sample of
the 2ml GC vials. The sample was extracted for 25 minutes prior to desorbing directly into a
0.75mm straight inlet liner of the GC injection port for 5 minutes at 260 ºC and splitless flow.
The extraction time was optimized to ensure an efficient GC run time, shown in Figure 5.
24
Figure 5 Optimization of SPME extraction time for naphthalene using a 50/30µm
DVB/CAR/PDMS fiber
Prior to running a sequence of samples, the SPME fiber required a conditioning step. The fiber
was set to desorb for 3600s in the GC inlet set at 250°C and splitless flow set at 8ml/minute. The
oven was set to 65°C for 1 minute and further ramped at 25°C/minute up to 260°C and held for
50.8 minutes. The FID was set to 250°C.
Following the conditioning step the sample sequence was analyzed as follows. The GC-FID
method uses ultra high purity 5.0 helium gas (Praxair) as the carrier gas with a flow rate of 8.0
mL/min. An oven temperature profile of 40ºC for 1min, 25ºC/min ramp to 200ºC for 1.5 min
was used. The injector temperature was set to 260ºC and the FID detector is set at 250ºC. The
ultra high purity 5.0 helium, zero air and ultra high purity 5.0 hydrogen (Praxair) flow rates for
the FID were set at 30ml/min, 400ml/min and 30ml/min respectively. The runtime was a total of
8.9 minutes.
The analytical method has a method detection limit of 100µg/L.
In the case that a sample had a concentration greater than 500µg/L, a desorption method was run
between samples in order to clean the fiber. The following method was used. The SPME fiber
0
50
100
150
200
250
0 5 10 15 20 25 30 35
Area
Cou
nt
Time (minutes)
25
was desorbed for 600s with the GC inlet set to 260°C and a split ratio of 3:1. Oven was set to
40°C for 1 minute and ramped at 35°C/min up to 200°C and held for 5 minutes. The FID was set
to 250°C. This method had a total run time of 10.6 minutes.
3.4.3 Molecular Analyses for Total Bacteria and Total Archaea
16S rDNA gene copy numbers of General Bacteria and Total Archaea were determined with a
quantitative polymerase chain reaction (qPCR) technique. qPCR assays were performed in DNA
templates obtained from soil samples, supernatant of active microcosms, and homogenized
aliquots of active microcosms (soil + water). The DNA extraction, plasmid extraction, and qPCR
to determine General Bacteria, and Total Archaea were performed at BioZone facilities at the
University of Toronto.
3.4.3.1 Sampling and DNA Extraction
The soil samples (~ 10 g) were collected in Falcon tubes at the beginning of the microcosm start-
up. The supernatant (0.5 ml) from microcosms were collected in Eppendorf tubes during the
middle of the microcosm incubation (58 days) and homogenized soil slurries were collected in
Falcon tubes (150 days).
DNA was extracted from these samples using PowerSoil® DNA Isolation Kit (catalog no.
12888-50) from MO BIO Laboratories, Inc. (CA, USA). For soil samples, approximately 0.25 or
0.4 g of soil was used for DNA extraction. Supernatant liquid samples (0.5mL) were collected
using disposable syringes and DNA was extracted from these samples without further
processing. For both cases, the collected samples were stored in the freezer at or lower than -
20oC prior to DNA extraction. Homogenized samples (5mL) for DNA extractions were obtained
within an anaerobic glovebag with a 20%CO2 and 80%N mixture, bottles were vigorously
agitated and 5ml of the soil suspension was immediately removed. Aliquots were centrifuged at
maximum speed at 4oC for 30 minutes, and DNA was extracted from the pellets. The final DNA
extracts in UV treated Ultra Pure Distilled Water (100 µL) were stored in the freezer at or lower
that -20oC prior to NanoDrop and qPCR analysis.
26
3.4.3.2 Standard Plasmid Preparation
To quantify the 16S rDNA gene copy numbers of total General Bacteria and total Archaea in
DNA extracts using NanoDrop or qPCR analysis, plasmids extracted from dehalobactor (for
general bacteria) and archaea (for total archaea) were used as standards. The sequences are found
in Table 8. Recombinant E. coli with dehalobactor or archaea plasmid were grown in LB media
in two 17 hour cycles in an incubator shaker (37oC and 180 rpm). Plasmid was extracted from
2mL of this culture using GenEluteTM Plasmid Miniprep Kit (catalog no. PLN350) from Sigma-
Aldrich. This provides approximately 1010 gene copies/µL of extract. To prepare the calibration
curves, nine serial dilutions (10 times dilution in each step) were prepared for the plasmids using
UV treated Ultra Pure Distilled Water. The extracted plasmids were stored in fridge at 4oC for
less than 7 days before analysis by qPCR. For longer period of storage, plasmid extracts were
stored in the freezer at or lower that -20oC prior to NanoDrop and qPCR analysis.
3.4.3.3 NanoDrop Analysis
Concentrated DNA extracts of the samples, concentrated plasmid extracts, and 10 and 100 times
diluted plasmid extracts were analyzed using a NanoDrop 1000 Spectrophotometer (Thermo
Fisher Scientific) to quantify DNA concentrations in ng/µL. UV-treated Ultrapure Distilled
Water was used to initialize the instrument (2 µL) and as a blank (1.5 µL). These concentrations
were converted to gene copy number/µL using the molecular weight of each base pair (1bp = 660
g/mol) and the total number of base pairs in general bacteria (5400bp) or in total archaea
(5000bp). The plasmid concentration values were used to prepare the calibration curve for 16S
rDNA copy numbers quantification by qPCR.
3.4.3.4 qPCR Analysis
The sample DNA extracts were diluted 10, 50, and 100 times with UV treated Ultra Pure
Distilled Water, and the diluted extracts underwent qPCR analysis in duplicate or triplicate. The
calibration standards (101 to 108 times dilutions of plasmid extracts) were analyzed in duplicate.
In each reaction 2µL of diluted sample/standard was added to 18µL of Master Mix. The Master
Mix contained 10µL of EvaGreen® Supermix (catalog no. 172-5200, BIO RAD), 0.5µL of
forward primer (10µM), 0.5µL of reverse primer (10µM), and 7µL of UV treated Ultra Pure
Distilled Water. At least two blanks (18µL Master Mix only) were included in each batch of
27
qPCR analysis. Table 8 shows the forward and reverse primers used in the qPCR and their
reference protocols. The sample and standard plasmid dilutions were performed on a clean lab
bench (sterilized using 70% ethanol) and the qPCR strip preparation was performed in a clean
qPCR cabinet (sterilized using UV light for ~30mins). The qPCR analysis was performed with a
CFX96™ Real-Time System (C1000™ Thermal Cycler, BIO-RAD). BIO-RAD qPCR CFX
Manager 3.0 software was used to operate the qPCR system and to quantify the gene copy
number concentration of each sample.
Table 8 qPCR Primers and References
Target Primer 5’3’ Sequence Reference
Total Archaea Arch 787f ATTAGATACCCGBGTAGTCC (Yu et al. 2005) [49]
Arch 1059r
GCCATGCACCWCCTCT
Total Bacteria Bac 1055f ATGGCTGTCGTCAGCT (Amann et al. 1995) [50]
Bac 1392r ACGGGCGGTGTGTAC (Stahl et al. 1988) [51]
3.4.3.5 Metals Analysis
Microcosms were sampled after 119 days to analyze the concentration of dissolved metals in the
supernatant. This was done to further understand the reduction processes occurring in the
microcosms over time.
1ml samples were extracted from the microcosm and filtered through a 0.22μm syringe filter to
remove any sediment from the samples. The samples were placed in 15ml Falcon tubes and 9ml
of MilliQ® water was added to dilute the sample. These samples were further acidified with
nitric acid to ensure the metals remain in the aqueous phase. The samples were capped and
placed at 4°C prior to analysis.
The samples were analyzed by the Analytical Lab for Environmental Science Research and
Training (ANALEST) in the Department of Chemistry at the University of Toronto using
Inductively Couples Plasma Atomic Emission Spectroscopy (ICP-AES). A Perkin Elmer Model
Optima 7300DVICP AEOS was used to conduct the analysis.
28
3.4.3.6 Anion Analysis
Anion Analysis was conducted at 161 days to further characterize the re-dox chemistry within
the microcosms.
1ml samples were extracted from the microcosm and filtered through a 0.22μm syringe filter to
remove sediment from the samples. Samples were placed in screw cap vial to transport to the
BioZone analytical facilities at the University of Toronto. Dilutions were prepared directly into
the analytical vials using pipettes. The IC sample vials held a total of 0.5ml sample for analysis.
Vials were sealed with IC filter caps. Samples were run undiluted as well as at ten times dilution
to ensure all of the anions fit within the range of calibration standards. Ten time dilutions were
prepared with 50µL of sample in 450µL deionized water. Blanks were run in between each
sample to ensure no carry over was detected.
A Thermo Scientific Dionex ICS-2100 fitted with a Dionex IonPacTM AS18 was used in
combination with a Thermo Scientific Dionex AS-DV autosampler to conduct the analysis.
Potassium Hydroxide was used as the eluent with a concentration of 23mM. Standards prepared
in the BioZone were used to analyze for chloride, nitrate, nitrite, sulfate and phosphate.
3.4.3.7 Microscopy Analysis
Microscopy analysis was conducted to observe the variations in microbial communities in the
microcosms. A Leica DMI 3000 inverted microscope was used conduct the analysis.
To prepare the samples, 0.5ml of supernatant was extracted from the microcosms and injected
into a 2ml vial containing 1.3ml of phosphate buffer of pH 7.4. The vial was then sealed and
sonicated for 1 minute. 0.2 ml of 100µg/ml Acridine Orange solution (Sigma Aldrich) was added
to the sample vial and vortexed for 5 seconds to thoroughly mix. The sample was then covered
with tinfoil and allowed to react for 3 minutes in the absence of light.
The sample was filtered onto a 0.1µm, 25mm black membrane polycarbonate filter (GE Water
and Process Technologies), backed by an 8µm nitrocellulose support membrane (Millipore) and
further backed by a glass filter (Whatman GF/F). The sample was washed with the buffer
solution using a low pressure pump. The polycarbonate filter was placed on a 76x26mm
GoldLine microscope slide containing a drop of ProLong Gold Antifade reagent. An additional
29
drop of this antifade gel was placed on the top surface of the membrane to mount the coverslip
(25x25mm, No.1) and maintain the fluorescence of the slide. The coverslip was sealed with a
clear nail polish around the edges.
The prepared slides were analyzed using the Leica DMI 3000 inverted microscope with a
standard Texas red filter and fluorescence with 100x magnification.
3.4.3.8 Methane Analysis
The production of methane was analyzed by gas chromatography using a Hewlett Packard 5890
Series II Gas Chromatograph. A headspace sample of each microcosm was taken by a locking
gas tight syringe and directly injected into the inlet of the GC. A GS-Q ® Agilent Technologies
column, 30m x 0.53mm I.D. was used in combination with FID. The injector and detector were
set to 250°C with an oven temperatures set isothermally at 35°C for 5 minutes. The helium, air
and hydrogen were set at flow rates of 4.4, 300 and 30 ml/minute with a septum purge of 2.2
ml/minute. Splitless flow was used.
3.4.3.9 pH, Specific Conductivity, ORP Analysis
The pH, specific conductivity and ORP were measured in the synthetic groundwater prior to
microcosm setup as well as the supernatant of the microcosms at 149 days following their setup.
pH and ORP were measured using a Radiometer PHM92 pH Meter. Specific conductivity was
measured using an Orion Model 150 meter.
In an anaerobic glovebag filled with 20% CO2 and 80% N, microcosms were destructed. The
supernatant and soil were agitated to homogenize the microcosm into a slurry prior to pouring a
portion of the slurry into a falcon tube for analysis. Analysis was conducted within the anaerobic
glovebag by promptly inserting the probes into the slurry. The ORP was measured first, followed
by the specific conductivity and the pH.
30
Chapter 4 Results and Discussion
This study was conducted in three distinct phases:
1. Characterization of the three soils that reached varying peak temperatures during thermal
remediation, including physical and chemical parameters.
2. Adsorption isotherm batch studies analyzing the interactions between naphthalene and
the soil samples.
3. Microcosm degradation batch studies, monitoring naphthalene degradation in each soil at
varying temperatures. This included the analysis of naphthalene concentrations, microbial
populations, chemical parameters of the aqueous phase and methane production over a 6
month period.
The results for each of these phases will be discussed sequentially in the following sections.
4.1 Soil Properties
Each soil sample was homogenized prior to the analysis of physical and chemical parameters.
The results are shown in Table 9.
Table 9 Physical and Chemical Properties of Each Soil Type
Soil 1 Soil 2 Soil 3
Depth (ft. bgs) 27.5 26.5 36.2
Peak Temperature (ºC)1 101.2 69.5 36.9
Grain Size Classification Sandy Silt Sandy Silt Sandy Clay
Moisture Content 22.2 36.4 45.7
1 Complete temperature profiles can be found in Appendix C
31
Cation Exchange Capacity
(cmol+/kg)2 31.4
30.8 38.9
Organic Content (%dry) 3 0.1 0.1 0.1
pH4 6.1-6.7 6.2-6.5 6.0-6.5
Bulk Density (g/cm3) 1.55 1.25 1.79
Particle Density (g/cm3) 2.36 2.43 2.41
Porosity 0.34 0.48 0.26
The most critical difference between the three soils is that they reached different peak
temperatures at varying depths. Apart from temperature and depth, all three soils had similar
properties. This makes it possible to compare the differences based on peak temperature while
leaving other parameters constant. The soils had differing moisture contents, however, once the
soil is in the microcosms it is fully saturated with synthetic groundwater thereby eliminating this
variability.
There was a slight variation in cation exchange capacity (CEC), although each soil fell between
30 and 40 cmol+/kg. Soil 3 had a slightly higher CEC as it had higher clay content than Soils 1
and 2. As clay develops a negatively charged surface it has the capacity to host more positively
charged cations. A higher CEC value results in a soil that can hold increased amounts of
nutrients, such as potassium, calcium, sodium and trace metals that will leach out of the soil over
time and become available for microbial communities. A low CEC results in a soil that cannot
easily maintain the nutrients as they are more likely to be flushed through as the groundwater
moves through the system. This is an important factor in the field as it determines the amount of
nutrients that can be held in the soil until they are required for biodegradation. The CEC also
2 Values obtained from Guelph Laboratories 3 Values obtained from Guelph Laboratories 4 Values obtained over a range using the CaCl2 and MilliQ® water values described in the section 3.1.1
32
impacts the buffering capacity of the soil as it controls the number of hydrogen ions retained in
the soil available for replenishing the groundwater or buffering the groundwater in order to
maintain equilibrium. This has a significant impact on the overall pH of the system.
The pH can significantly impact the microbial communities in the soil as it controls microbial
enzyme activity, transport processes and nutrient solubility [52]. Many microbes cannot survive
in extreme pH environments, with many naphthalene degrading bacteria performing optimally
around neutral pH [5] [53] [52] [33]. The pH of the soil at the site in this study falls between 6.0
and 7.0, providing suitable conditions for many naphthalene degrading bacteria. Degradation
processes, however, can lead to fluctuations in the pH. For example, during the degradation of
naphthalene by sulfate reduction, sulfuric acid can be formed. This would result in a slight
decrease in the pH which could potentially hinder microbial growth, however, soils have a
tendency to act as a buffer for the groundwater and prevent these variations. As mentioned
above, a higher CEC value and increased presence of organic matter increase the buffering
capacity of the soil.
pH also impacts the sorption of naphthalene. It has been found that naphthalene adsorption
increases from pH 1.5 to pH 4.0 and slightly decreases from pH 4.0 up to pH 7 [54].This
indicates that a fluctuating pH may result in a change in adsorption of naphthalene, ultimately
changing the bioavailability. The organic carbon (OC) content also significantly impacts the
adsorption of naphthalene as it is mainly responsible for the sorption of hydrophobic organics to
soils when the OC content in the soil is greater than 0.1% [55]. Each soil type in this study
contained 0.1 dry% organic matter, falling within this threshold, so it is likely that OC was the
driving factor in adsorption for these soils. Further, the OC value was consistent for each soil,
therefore, it was anticipated that each soil would have similar adsorption results with respect to
OC. This is investigated further in the subsequent section, Adsorption Isotherms.
In addition, the soil was analyzed by Maxxam Analytics for F1-F4 hydrocarbon fractions, PAH
and BTEX prior to experimental setup. The results for the constituents that had concentrations
above the detection limit are shown in Table 10.
33
Table 10 Concentrations of contaminants remaining in each soil post thermal remediation
Units Soil 3 Soil 2 Soil 1
Polyaromatic Hydrocarbons
Acenaphthene µg/g 0.059 0.17 ND
Acenaphthylene µg/g 0.016 0.017 ND
Fluorene µg/g 0.078 0.035 ND
1-Methylnaphthalene µg/g 2.0 0.74 ND
2-Methylnaphthalene µg/g 5.2 1.5 ND
Naphthalene µg/g 2.5 0.19 ND
Phenanthrene µg/g 0.077 0.051 ND
BTEX & F1 Hydrocarbons
Benzene µg/g 2.0 ND ND
Toluene µg/g 1.3 ND ND
Ethylbenzene µg/g 3.0 ND ND
o-Xylene µg/g 3.0 ND ND
p+m-Xylene µg/g 5.5 ND ND
Total Xylenes µg/g 8.5 ND ND
F1 (C6-C10) µg/g 260 11 ND
F1 (C6-C10) - BTEX µg/g 250 11 ND
F2-F4 Hydrocarbons
F2 (C10-C16 Hydrocarbons) µg/g 910 390 ND
F3 (C16-C34 Hydrocarbons) µg/g 81 55 ND
F4 (C34-C50 Hydrocarbons) µg/g ND ND ND
It was expected that the soil that reached the lowest peak temperature would have the largest
concentrations of residual contamination. This is consistent with the results as Soil 3, which only
reached 37ºC, had the highest concentrations of PAH, BTEX and F1-F3 hydrocarbon fractions.
This was followed by Soil 2, with a peak temperature of 70ºC and finally Soil 1, with a peak
34
temperature of 101ºC. This variability in residual compounds adds an additional factor
complicating the potential to compare the biodegrading ability of each soil at each temperature.
The presence of additional compounds complicates the biological processes occurring within the
microcosm as they provide additional substrate for the microbial communities to sustain growth.
Competing substrates have the ability to both hinder and promote degradation depending on the
types of microorganisms and compounds present. Contaminants can reach inhibitory
concentrations, becoming toxic to microbial communities. For example, Shuttleworth and
Cerniglia (1995) determined that concentrations of 5mg/L naphthalene would inhibit the growth
of many strains of phenanthrene degrading bacteria [56]. Furthermore, microbial communities
can preferentially acclimate to different substrates in a complex system depending on the
substrates, nutrients, and physical and chemical conditions of the subsurface. Although
naphthalene is the target contaminant in this study, the presence of other substrates may hinder
naphthalene degradation. This is discussed further along with the qPCR results.
This preliminary assessment of the soils provided a basis upon which the adsorption isotherm
batch study and degradation batch study were conducted. It is essential to understand the
variations in each soil’s parameters in order to assess the results of the adsorption isotherms and
degradation studies.
4.2 Adsorption Isotherm Batch Study
Adsorption isotherms were developed for each soil type to determine the sorption of naphthalene
at each temperature: 30, 45 and 60ºC. The Freundlich adsorption isotherm model was used to
model the data on the logarithmic scale. The results for Soil 1 at 30, 45 and 60°C are presented in
Figure 6 and Soil 3 at 30 and 45°C in Figure 7. Due to the similarities of Soil 1 and 2 adsorption
isotherms were not conducted for Soil 2.
35
Figure 6 Adsorption Isotherms for Soil 1 at A) 30°C B) 45°C C) 60°C
y = 0.0012x1.3724 R² = 0.9643
0.10
1.00
10.00
100.00
1.00 10.00 100.00 1000.00 10000.00
y = 0.0024x1.2603 R² = 0.9408
0.10
1.00
10.00
100.00
1.00 10.00 100.00 1000.00 10000.00
y = 0.0062x1.2172 R² = 0.9827
0.10
1.00
10.00
100.00
1.00 10.00 100.00 1000.00 10000.00
Aqueous Concentration of Naphthalene (µg/L)
A)
B)
C)
Sorb
ed N
apht
hale
ne C
once
ntra
tion
(µg/
g)
36
Figure 7 Adsorption Isotherms for Soil 3 at A) 30°C B) 45°C
Adsorption isotherms for Soil 1 and 3 demonstrated a close correlation to the Freundlich
adsorption model with the relationship expressed by equation 10on a logarithmic scale. At low
concentrations the sorption data approaches a linear relationship. Linear adsorption isotherms are
typical for many non-polar organic compounds up to 60 - 80% of their water solubility [40] [55].
In this study, adsorption isotherms were conducted for naphthalene at concentrations less than
10mg/L which is approximately 30% of solubility in water, suggesting that a linear model is
likely at the lower concentrations. Although a linear relationship exists at lower concentrations,
y = 0.097x0.8754 R² = 0.9785
1.00
10.00
100.00
1.00 10.00 100.00 1000.00 10000.00
y = 0.3056x0.6968 R² = 0.9814
1.00
10.00
100.00
1.00 10.00 100.00 1000.00 10000.00
Aqueous Concentration of Naphthalene (µg/L)
Sorb
ed N
apht
hale
ne C
once
ntra
tion
(µg/
g)
A)
B)
37
the Freundlich model is the most appropriate fit and best represents the complete data set for this
study.
The experimentally determined Freundlich adsorption coefficients, expressed by equation 10, are
presented in Table 11 .
Table 11 Experimentally determined Freundlich adsorption isotherm coefficients for Soil
1and 3
Temperature (ºC) K experimental
(L/kg)
N R2
Soil 1
30 1.2 1.37 0.96
45 2.4 1.26 0.94
60 6.2 1.22 0.98
Soil 3 30 97.0 0.70 0.98
45 305.6 0.88 0.98
The partition coefficients (K) for Soil 1 are similar to the partition coefficients found in a study
conducted by Bayard et al. (1998). In the Bayard study K values ranged from 4.8-14.17 L/kg for
five different types of soil. Soils were typically a silty sand with total organic carbon ranging
from 1.4-3.4% which is comparable to the soils in this study. The sorption values for Soil 3,
however, far exceed these previously reported values. The impacts of both temperature and soil
type are discussed further in the subsequent sections.
As discussed previously, the partition coefficient can also be estimated using the linear
relationship found in equation 1, assuming that the fraction of organic carbon is linearly
correlated to the partition coefficient and is the main contributor to the adsorption of
naphthalene. The theoretical value for K, using equation 1 fell between 0.933-1.862 L/kg (over a
range of reported Koc values). The theoretical values are much lower than the values reported in
this study. This suggests that the adsorption is much greater than theoretically estimated. This
may be attributed to losses from the microcosm bottles which would have been accounted for in
the adsorption data.
38
4.2.1 Impact of Temperature on Adsorption
In order to show the relationship between the adsorption isotherms at each temperature, the
isotherms for each soil are plotted on one figure. Adsorption isotherms for Soil 1 and Soil 3 at
each temperature are shown in Figure 8 and Figure 9 respectively.
Figure 8 Soil 1 adsorption isotherms for naphthalene at 30, 45 and 60ºC
Figure 9 Soil 3 adsorption isotherms for naphthalene at 30 and 45ºC
0.10
1.00
10.00
100.00
1.00 10.00 100.00 1000.00 10000.00
Sorb
ed N
apht
hale
ne C
once
ntra
tion
(µg/
g)
Measure Naphthalene Aqueous Concentration (µg/L)
30C
45C
60C
0.10
1.00
10.00
100.00
1.00 10.00 100.00 1000.00 10000.00
Sorb
ed C
once
ntra
iton
(µg/
g)
Aqueous Concentration (µg/L)
30C
45
39
As shown in Figure 8 and Figure 9, this study indicates an increase in sorption with increasing
temperatures. This is an unexpected result. Many studies have shown a significant decrease in
the sorption of PAHs at increasing temperatures. Hiller et al. (2008) found a 24% reduction in
adsorption as temperatures increased from 4-27ºC, and similarly, He et al. (1995) found a 25%
reduction in adsorption as temperatures increased from 5-15 ºC [57]. The decrease in sorption is
a result of an increasing solubility at higher temperatures along with an increase in mass transfer
rates [12] [40]. Increased sorption at higher temperatures has been found for compounds that
experience a decrease in solubility at higher temperatures [40]. This, however, does not explain
the results found in this study as solubility of naphthalene increases with increasing temperature
[12].
There are two potential explanations for the results found in this study: 1. the method used was
dependent on an instantaneous desorption as the microcosms were transitioned from lower
temperatures to higher temperatures. 2. The co-solvent fraction, methanol, may have impacted
the solubility and therefore adsorption of naphthalene at varying temperatures. These
explanations are further addressed in the following sections.
1. Impacts of Batch Study
Adsorption isotherms for Soil 1 were first analyzed for equilibration time. Each concentration
was tested in triplicate. The microcosms were placed at 30ºC with supernatant samples taken at
24 and 48 hours. Previous studies conducted by other researchers determined that the time to
reach equilibrium was typically less than 24 hours [37] [39] [14]. Based on the literature and the
results indicating negligible change in concentration between 24 and 48 hours during this study it
was determined that equilibrium was reached within this timeframe. Therefore, the results for
Soil 1 are based on equilibrium reached within 48 hours.
Soil 1 microcosms were subsequently sampled at 45ºC at 48 hours and finally sampled at 60 ºC
at 48 hours. The resulting isotherms are shown in Figure 8. This process was dependent on the
assumption that desorption would not be a limiting factor in the microcosms reaching
equilibrium. It has since been determined that this assumption is not valid for this study. The
literature suggests there is a hysteresis effect which decreases the potential for desorption as the
contact time increases [57] [37] [25]. This could potentially prevent the microcosm from
40
reaching equilibrium at the higher temperatures due to the transfer of the microcosms from low
temperatures to high temperatures.
Following the adsorption isotherm test, the degradation study was conducted showing a much
longer equilibration time of approximately 500 hours required in the control microcosms
suggesting that the adsorption isotherms hadn’t yet reached equilibrium when analyzed
throughout the adsorption study. In this case, as the microcosms were transitioned from 30 to
60°C, it is possible that adsorption proceeded to increase throughout the transition period and
was never fully reached at 30 and 45ºC. Following this analysis, the isotherms were transitioned
back to 30ºC and left to equilibrate for 7 days prior to sampling. Again, the adsorption in the
isotherms was found to dramatically decrease, suggesting that the initial relationship was in fact
representing the naphthalene partitioning in the microcosms.
To investigate this further, the control microcosms from the degradation study were analyzed as
a single point on the adsorption isotherm. This was possible as the control microcosms in the
degradation study were set up in the same manner as the adsorption isotherms, however, only the
high concentration (10mg/L) condition was tested. This allowed for the analysis of this condition
over a longer period of time and was representative of a microcosm that was only exposed to one
temperature over this period. Control microcosms were analyzed for 30, 45 and 60°C. These
microcosms demonstrated the same trend as the adsorption isotherms, increasing temperatures
resulting in increasing adsorption. However, the degradation studies showed a slightly higher
adsorption as they were maintained over a longer period of time. This suggested that the use of
one batch of microcosms for the entire adsorption study was not the cause of the unexpected
trend.
Although the single batch adsorption experiment was not fully responsible for the trend of
increased sorption at increased temperature, the uncertainty could be mitigated by first testing the
batch of microcosms at 60ºC and subsequently decreasing the temperatures to test the soils at
lower temperatures. This, however, may increase the potential for losses to occur, further
impacting the subsequent analysis at lower temperatures. An alternative solution is to prepare
separate batches of microcosms for each temperature tested. As stated previously, this was not a
viable option for this study as there was limited soil available.
41
2. Effect of Cosolvent
The microcosms may have been showing an unexpected trend due to the presence of the
cosolvent, methanol. In order to obtain the range of naphthalene concentrations required for the
adsorption isotherms, 0.5-2.5ml of varying concentrations of naphthalene in methanol solution
were instilled into each microcosm. A high percentage of cosolvent may impact the validity of
the results. For this study, a range of 0.8-4% co-solvents were used in the adsorption isotherm.
Standard method ASTM E1195-01 recommends that the incorporation of co-solvent should not
exceed 0.1%.
Studies have been conducted to determine the impact of increased cosolvent fraction on
adsorption of organics. Studies show the sorption coefficients decrease log-linearly with an
increasing co-solvent fraction [58] [59]. This relationship has been found to correspond to the
increase in solubility of naphthalene with an increase in the cosolvent fraction [58]. Although the
ASTM E1195-01 recommended cosolvent fraction for adsorption studies is 0.001 or lower,
Nzengung et al. (1996) determined the impacts for fractions values between 0.008 and 0.042
proved to have minimal impacts on the sorption isotherms [59]. This, however, doesn’t explain
the higher K values reported compared to the estimated K values, as a higher cosolvent fraction
should decrease the K values as the solubility in the aqueous phase increases.
The higher K values reported in this study may be due to a decrease in the amount of cosolvent
in the aqueous phase at higher temperatures with an increasing fraction of methanol entering the
gas phase at increased temperatures. This could potentially create a higher solubility at the lower
temperatures as the fraction of methanol in the aqueous phase increases and a lower solubility at
the higher temperatures as the methanol enters the gaseous phase.
42
4.2.2 Impact of Soil Type on Adsorption
The impact of soil type on the adsorption of naphthalene was analyzed. The comparison of
adsorption isotherms of each soil type is shown in Figure 10.
Figure 10 Adsorption Isotherms for Naphthalene on Soil 1 and 3 at 30, 45 and 60ºC
Sorption of naphthalene was generally determined to be greater for Soil 3 than for Soil 1 at each
temperature tested. Given the soils had the same fraction of organic carbon (0.1%), it can be
estimated that both soils would have the same partitioning coefficient, governed by the equation
1. However, the partitioning coefficients were experimentally determined to be significantly
greater for Soil 3 than Soil 1. A difference in the adsorption for the two soils indicates that
organic carbon was not the main contributor to adsorption of naphthalene and the properties of
the soil themselves impacted the results. As stated earlier, Owabor et al. (2012), found a
significant difference in the amount of naphthalene adsorbed to two different soil types, clay and
sand. The clay adsorbed significantly higher amounts of naphthalene than the sand while the
sand was able to retain more naphthalene during desorption than the clay [39]. Soil 3 contained a
0.10
1.00
10.00
100.00
1.00 10.00 100.00 1000.00 10000.00
Sorb
ed C
once
ntra
tion
(µg/
g)
Aqueous Phase Concentration (µg/L)
Soil 1 30
Soil 1 45
Soil 1 60
Soil 3, 30
Soil 3 45
43
higher percentage of clay than Soil 1 and this may be the cause of the higher adsorption by Soil
3.
Soil 3 also contained an initial concentration of 2.5 µg/g naphthalene while Soil 1 did not contain
an initial concentration of naphthalene. This contamination had been residual in the soil prior to
site sampling. Residual contamination can be more difficult to desorb from the soil matrix as the
contact time increases. As discussed previously, there are two accepted types of adsorption
processes; those that occur in a short period of time (minutes to hours) and those that occur over
a longer period (months to years) [40]. Slow adsorption is a result of processes such as
micropore diffusion and intra-organic matter diffusion [40]. Once these adsorptive processes take
place it is more difficult for desorption to occur. It is unclear from the literature whether or not
the act of slow adsorption increases the adsorption capacity of the soil over time resulting in
greater partitioning coefficients as noted in this study.
Although the results of the adsorption study are not consistent with the literature, the degradation
batch experiment is not dependent on these results. This analysis was conducted for the purposes
of completing a mass balance for the degradation study; however, it is possible to monitor the
naphthalene concentration over time alongside control microcosms to determine if
biodegradation is taking place.
4.3 Degradation Batch Experiment
A microcosm batch experiment was conducted to determine the potential for anaerobic
naphthalene degradation by indigenous microorganisms in the soil samples from the North
Carolina site that previously underwent thermal remediation. The microcosms were analyzed for
naphthalene degradation, microbial growth, ion concentrations and methane production over the
course of the study. These results are discussed in the following sections.
4.3.1 Naphthalene Degradation
The concentration of naphthalene throughout the microcosm study is displayed in Figure 11,
Figure 12, and Figure 13 for 30, 45 and 60ºC respectively. These figures demonstrate a rapid
decrease in naphthalene concentration between 0 and 120 hours with final equilibrium
stabilization around 500 hours. This occurs for both the biologically active microcosms as well
as the control microcosms. This suggests that this initial decrease is not due to biodegradation
44
and is likely due to adsorption processes which reach equilibrium around 500 hours. This is
unexpected as many studies indicate a typical equilibration time for naphthalene less than 24
hours, as discussed previously.
Figure 11 Change in Naphthalene Concentration at 30°C for Soil 1, 2 and 3
Figure 12 Change in Naphthalene Concentration at 45°C for Soil 1, 2 and 3
0.0
1000.0
2000.0
3000.0
4000.0
5000.0
6000.0
7000.0
8000.0
9000.0
-500 0 500 1000 1500 2000 2500 3000 3500
Nap
htha
lene
Con
cent
ratio
n (µ
g/L)
Time (hr)
Soil 1
Soil 1 Control
Soil 2
Soil 2 Control
Soil 3
Soil 3 Controls
0.0
1000.0
2000.0
3000.0
4000.0
5000.0
6000.0
7000.0
8000.0
9000.0
-500 0 500 1000 1500 2000 2500 3000 3500
Conc
entr
atio
n (µ
g/L)
Time (hrs)
Soil 1
Soil 1 Control
Soil 2
Soil 2 Control
Soil 3
Soil 3 Control
45
Figure 13 Change in Naphthalene Concentration at 60°C for Soil 1 and 2
The concentration at which naphthalene stabilizes is shown to depend on the temperature of the
microcosms and not the type of soil. Microcosms at 45 ºC and 60 ºC rapidly stabilized at
approximately 1000µg/L and microcosms at 30 ºC slowly stabilized at approximately 1500µg/L
for all soil types. This is consistent with the previous adsorption isotherm study which showed
decreased concentrations in the aqueous phase at increased temperatures. However, this result is
inconsistent with the adsorption study with respect to the varying soil type, insofar as the
absorption study displayed significant differences in the adsorption to Soil 3 than Soil 1. Soil 3
was found to adsorb larger concentrations of naphthalene than Soil 1in the adsorption study. This
variation may be due to the fact that the degradation study microcosms were monitored over a
longer period of time.
The lack of naphthalene degradation does not definitively indicate that there is a lack of
microbial communities in the soil that are capable of degrading naphthalene. Microbial
communities often require a period to acclimate to a new compound. Mihelcic and Luthy (1988),
found acclimation periods between 12 and 36 days for denitrifying organisms in soil that was
previously not exposed to naphthalene [14]. Once acclimated, naphthalene was degraded from
4mg/L to non-detectable levels within 47 days [14]. Other studies suggest anaerobic
0.0
1000.0
2000.0
3000.0
4000.0
5000.0
6000.0
7000.0
-500 0 500 1000 1500 2000 2500 3000 3500
Conc
entr
atio
n (µ
g/L)
Time (hrs)
Soil 1
Soil 1 Control
Soil 2
Soil 2 Control
46
microorganisms may require up to 18 months to acclimate [60]. The microcosms will require
further monitoring to determine the potential for naphthalene degradation as they may require
additional time to acclimate.
Although the naphthalene degradation study does not indicate microbial activity related to
naphthalene degradation at this time, there was evidence of microbial activity within the
microcosms. These analyses are discussed in the subsequent sections.
4.3.2 Microbial Analysis
DNA samples were extracted from Soil 1, Soil 2 and Soil 3 prior to any experimental setups to
determine the presence of microbes in the soil as sampled from the former bulk fuel terminal. A
qPCR analysis was conducted to determine the general bacteria and archaea count in the soil.
These results can be indicative of both active and inactive DNA in the soil or supernatant,
therefore, microbes that have been recently inactivated will be detected using the qPCR method.
The results are discussed in this section.
qPCR prior to microcosm setup:
Soil samples were extracted prior to experimental setup to analyze for initial microbial
populations in the soil. The results in Figure 14 show that the concentration of total bacterial
DNA in the soil is far below the detection limit of 190 DNA Copies/0.5g soil for Soils 1, 2 and 3.
This is not representative of what was expected for these samples. Many hydrocarbon degrading
bacteria thrive in conditions between 30- 40ºC [33]. It was anticipated that Soil 3, which only
reached a peak temperature of 37 ºC would have detectable bacteria populations.
47
Figure 14 Total Bacteria Concentration in Soil Prior to Experimentation. Representing
time = 0 in the microcosms
There are many reasons that may have resulted in such low bacteria counts in the soil initially:
• Sterilization by increased temperatures: The high temperatures experienced during
thermal remediation may have sterilized the soil, inhibiting any bacteria growth. As
discussed previously, microorganisms have a maximum temperature at which they can
survive, after which point they lose their structure and function [26]. This denaturation
process is also more likely as the time of peak temperature increases and is intensified in
the presence of increased moisture [26]. The soils at this site were maintained at their
peak temperatures for approximately 12 months (refer to Appendix C) and were also
subject to the injection of hot water throughout this process to prevent desiccation. Both
of these factors increase the potential for sterilization in each of the soils. This, however,
was unexpected for the samples of soil with relatively low peak temperatures (37°C and
70°C). Many microorganisms capable of degrading naphthalene thrive in these
temperatures [33] [11] [34] [6] . It has also been found that various organisms that cease
to function at high temperatures have the ability to form spores and re-acclimate when the
0
20
40
60
80
100
120
140
160
180
200
Soil 1 30C Soil 2 30C Soil 3 30C
Tota
l Bac
teria
DN
A Co
pies
/ 0.
5g s
oil
Total Bacteria Concentration in Soil Prior to Experimentation
48
temperatures have cooled back to acceptable levels [34]. This suggests that the peak
temperatures reached during thermal remediation should not have resulted in a complete
sterilization of the soil.
• Toxicity due to high concentrations of mixed contaminants: High concentrations of
contaminants can become inhibitory to microbial activity in soil. This site was heavily
contaminated with a wide array of hydrocarbons that may have reached inhibitory
concentrations, destroying the microbial communities. Pumphrey and Madsen (2007)
found the inhibition of certain strains of naphthalene degrading organisms at naphthalene
concentrations of 10mg/L and greater [61].
Although many bacteria are capable of degrading a variety of PAHs, varying
concentrations of contaminants can become inhibitory to the degradation of other
substrates present. Shuttleworth and Cerniglia (1995) determine that concentrations of
5mg/L naphthalene would inhibit the growth of many strains of phenanthrene degrading
bacteria [56]. Similarly, naphthalene inhibits phenanthrene degradation in the presence of
a variety of pseudomonads. However, when these substrates were present as the sole
carbon source, degradation was possible [62]. This suggests that the contaminants were
only inhibitory to the microorganisms when in the presence of additional substrates.
Naphthalene and other related compounds were also found to be inhibitory to
cyanobacterium Agmenellum quadruplicatum, a bacteria strain capable of degrading
hydrocarbons [63]. Given the low concentrations of total bacteria in the soil samples it is
possible that inhibition due to complex substrates may have occurred.
Alternatively, complex systems of varying substrates and cultures have the potential to
promote degradation. Horng et al. determined that the complex nature of cosubstrates and
cocultures has the potential to facilitate increased microbial activity and diversify the
types of substrates consumed by certain bacteria. For example, Pseudomonas putida
M2T14, a toluene degrader, was capable of degrading both monoaromatics as well as
PAHs when in the presence of toluene, representing a scenario where cosubstrates
promoted degradation. The use of cocultures, Pseudomonas putida M2T14 and P.
azelaica ND, a PAH degrader, stimulated the degradation rates of naphthalene and
toluene [64], representing a scenario where cocultures enhanced biodegradation rates.
49
Although the potential exists for enhanced degradation in complex systems, this was not
the case for the soils used in this study as the total initial bacteria count was much lower
than the detection limit.
With no information on the exact microbial populations present in these soils, it is
difficult to determine whether or not the contaminants would inhibit microbial activity.
This information could be obtained through pyrotag analysis but was not obtained for this
study.
• Impact of competitive cultures and substrates on nutrient availability: The complex
system of many substrates may also have a significant impact on the microbial
communities present. As previously mentioned, competing species may inhibit or
promote the growth of varying microbes [56]. If some species dominate the subsurface
they may create a high demand for nutrients and electron acceptors limiting the supply
for other less dominant species. With increased temperatures, the metabolisms of these
microorganisms can be enhanced, therefore consuming the nutrients at a more rapid rate
[26]. Due to a lack of augmentation or influx of nutrients in this thermal remediation
project, a nutrient depletion may have developed in the subsurface, therefore, preventing
any further degradation to take place. Recent groundwater analyses in the location of the
sampling were not available to confirm the concentrations of nutrients in the subsurface
at the time of sampling. Additionally, microbial analyses had not been conducted prior to
thermal remediation or throughout the remediation process, so it is difficult to determine
if microbial activity was occurring throughout the treatment period.
• Exposure to oxygen: These soils were sampled from anaerobic conditions where an
introduction of oxygen may have been toxic to the anaerobic microorganisms. The
sampling procedure was conducted by a third party and it is unknown if the soils were
exposed to oxygen. If, in the sampling process the soils were exposed to oxygen, the
anaerobic microorganisms may have been depleted.
• Extended cold storage period of samples: The soils were preserved at 4ºC from
December, 2012 until June 2013. Just as there are maximum temperatures that can be
reached that destroy microbial populations, there are also minimum temperatures at
50
which cell reproduction is not possible or even complete destruction can occur [26]. This
may have resulted in the inactivation and destruction of DNA in the soil.
Due to the complexity of the system and lack of site information it is difficult to determine the
cause of the undetectable concentrations of DNA in the soil. Further analyses were conducted
throughout the microcosm study to track the changes over time.
qPCR at 8 weeks:
Naphthalene degradation was undetected 8 weeks after setting up the microcosms. It was
unexpected that microbial communities would be flourishing within the microcosms due to the
lack of degradation and the low bacteria counts found in the initial soil samples. Supernatant
samples of each soil and temperature scenario were extracted at 8 weeks. Each scenario at 30ºC
was analyzed for total bacteria and total archaea as these were scenarios anticipated to be the
most microbially active. Scenarios not analyzed at this time were placed in falcon tubes and
stored at -20ºC for potential future analysis. The results are shown in Figure 15.
Figure 15 Concentration of Total Bacteria and Total Archaea in Soil 1, 2 and 3 at 30C at 8
weeks. Grey and black dashed lines represent the detection limits for total bacteria and
archaea respectively.
1.00E+00
1.00E+01
1.00E+02
1.00E+03
1.00E+04
1.00E+05
1.00E+06
1.00E+07
1.00E+08
Soil 1 30C Soil 2 30C Soil 3 30C
Conc
entr
atio
n (D
NA
Copi
es/m
l)
Total Bacteria
Total Archaea
51
These results show that there are bacteria and archaea present in the supernatant of the majority
of the microcosms at 8 weeks showing higher concentrations in archaea populations than
bacteria populations. At this time, Soil 1 shows the highest concentrations of total bacteria and
archaea followed by Soil 2 and Soil 3. This suggests that at 8 weeks of incubation an increase in
microbial population is correlated with an increase in peak temperature experienced during
thermal remediation, however, this is not the case over an extended monitoring period.
The presence of archaea is an indicator of methanogenic activity occurring within the
microcosms. This is not surprising in an environment with petroleum hydrocarbon degradation,
as bacteria typically degrade the hydrocarbons to a methanogenic precursor such as formate,
acetate or H2, and the methanogens continue the degradation process, degrading these precursors
to end products of methane and carbon dioxide [65]. Methanol is also a precursor to
methanogenic activity [66] and was present in the initial microcosms as a cosubstrate in the
naphthalene spiking solution between 1-2% of the microcosms’ aqueous solution. With a lack of
naphthalene degradation occurring at this time (discussed in the previous section), the growth of
the archaea population is most likely due to the presence of methanol. Methanol degraders are
widely distributed in the environment [26] and, therefore, it is likely that these degraders were
present in this study. Methanol degradation has been found to utilize many different electron
acceptors in the degradation process and would not be limited by the re-dox conditions in the
microcosms [67]. Furthermore, methanol degraders isolated from natural sources typically thrive
in the neutral pH conditions as are present in this study [67]. In the case that the pH fluctuates,
there have been methanol degraders that have been found to degrade in acidic and basic
conditions [67]. Given the conditions in the microcosms, it is possible that methanol degradation
was responsible for this growth in archaea populations. This activity may have a positive
influence on the potential for naphthalene degradation.
Methanol has been found to enhance PAH degradation as a cosubstrate. It accomplishes this
through the promotion of microbial growth, as it is typically more readily degradable than the
PAH. As microbial populations increase, PAH degradation ensues as the cosubstrate source is
depleted [68]. Concentrations of 1000mg/L were found to enhance phenanthrene degradation
[68]. Limited information is available in the literature regarding the direct impact of methanol
on naphthalene degradation. Additionally, there is limited information on the potential negative
impacts of methanol on naphthalene degradation, however, it is understood that the methanol
52
concentration may reach inhibitory concentrations. Methanol can be toxic to microbial
populations at high concentrations, generally >100,000ppm [67]. The concentration in the
microcosms of this study is 13,000 ppm, and therefore falls below this inhibitory concentration.
qPCR at 21 weeks
One triplicate microcosm from each soil and temperature scenario was destructed at 149 days
following microcosm preparation to sample a soil slurry mixture for further microbial analysis.
5ml samples were taken for both qPCR analysis and pyrotag analysis to be conducted at a later
date. The pyrotag samples were extracted for DNA and frozen for future use at a temperature of -
20ºC or less. qPCR samples were extracted for DNA and analyzed immediately. The results are
shown in Figure 16.
Figure 16: qPCR results for total bacteria and total archaea at 149 days for each soil and
temperature scenario. S1, 2, 3-30, 45, 60 represent the soil number followed by the
temperature of that scenario. For example, S1-30 represents a triplicate microcosm for Soil
1 at 30ºC.
The conditions have changed quite significantly from 8 weeks to 21 weeks showing a significant
growth in archaea for Soil 3 at 30ºC with a decrease in archaea for Soil 1 and 2. This suggests
1.00E+00
1.00E+01
1.00E+02
1.00E+03
1.00E+04
1.00E+05
1.00E+06
1.00E+07
S1-30 S2-30 S3-30 S1-45 S2-45 S3-45 S1-60 S2-60
Conc
entr
atio
n (D
NA
Copi
es/m
l slu
rry)
Total Bacteria
Total Archaea
Bacteria MDL
Archaea MDL
53
that Soil 3 at 30ºC provides the most favourable conditions for archaea growth. This is the only
condition that had an archaea concentration above the detection limit and further exceeded the
bacteria population. Soil 1 and 2 microcosms do not contain archaea concentrations above the
detection limit. It will require further monitoring to determine if this growth is slower for these
soil conditions or if the soils were sterilized during thermal remediation.
Soil 3 at 30ºC reached the lowest peak temperature during thermal remediation (37ºC) and was
maintained at the lowest temperature throughout the microcosm study (30ºC). This scenario also
had the largest concentrations of precontamination present in the soil prior to initiating the study.
It is possible that this particular soil had maintained a microbial species that was sterilized at the
higher peak temperatures reached for Soil 1 and 2 (70 and 101ºC). It is also interesting that Soil 3
maintained at 45°C does not show this same archaea presence. This suggests that this species
does not thrive at temperatures as high as 45ºC. Analyses of the supernatant chemistry were
conducted to further the understanding of the processes occurring in this particular scenario. This
is discussed in the subsequent sections.
Bacteria populations decreased for both Soil 1 and 2 at 30ºC, while bacteria populations for Soil
3 at 30ºC remained constant. This may have been a result of the change in method. At 8 weeks
only the supernatant samples were analyzed and at 21 weeks a slurry mixture was analyzed.
Additional qPCR analysis testing the supernatant is required to determine if this was the result of
a method change. If these results suggest that the method was not responsible for this decrease,
analysis of the supernatant chemistry is required to further understand what may have caused this
decrease in bacteria for these particular scenarios.
4.3.3 Qualitative Observations
Qualitative observations showed a transformation of the soil colour from a tan, brownish colour
to a dark greenish colour in all of the active microcosms and a vibrant orange colour in the
control microcosms. Figure 17 shows the colour transformations. The colour was consistent
throughout the soil and did not accumulate on the surface of the soil or in a preferential location.
Soils at 30°C, particularly Soil 3, showed darker green colours and more vibrant orange colours
in the active microcosms and control microcosms respectively and were therefore chosen for
additional analysis to investigate this observation. This is consistent with the qPCR results which
determined the highest archaea concentrations in this scenario.
54
Figure 17 Colour transformation of microcosms. A) Soil at time = 0 B) Active microcosm at
time = 5 months C) Control microcosm at time = 5 months.
The only difference between the active microcosms and control microcosms was the addition of
sodium azide and mercuric chloride to the control microcosms for sterilization at time zero. This,
along with the qPCR results, indicates that the change in colour is most likely a result of
microbial activity.
4.3.4 Microscopy
A microscopy analysis was conducted at 119 days after the microcosms were prepared to
investigate the colour change further. Soil samples for Soil 3 at 30°C were taken for both the
active microcosms and the control microcosms as this scenario resulted in the most intense
colour change. The results are shown in Figure 18. There is a greater presence of
microorganisms in the active microcosms showing both cylindrical shaped organisms (rods) and
spherical organisms (cocci) [26]. It is difficult to determine the properties of these organisms
strictly by observing the morphology as archaea can have the same appearance as bacteria [26].
These organisms do not appear in the sterilized control microcosm, as expected. If the
fluorescence was a result of particulate within the microcosms, similar patterns would have been
seen in the control image, again suggesting that these are in fact microorganisms.
A B C
55
Figure 18: Images from a Leica DMI 3000 Inverted Microscope, slides stained with
Acridine Orange and viewed under a red filter. A and B are fluorescence images of sample
from active microcosm of Soil 3 at 30C. C is fluorescence image of sample from control
microcosm of Soil 3 at 30C
There was a wide range in sizes measured for the microorganisms on each slide. Cocci ranged
from 1.5-6 µm in diameter. Rod shaped bacteria ranged from a length to width measurement of
6-2µm to 23-4µm with aspect ratios ranging from 2-5. Cocci were more prevalent than rod
shaped organisms. The measured lengths and widths are typical for archaea and bacteria.
These results compliment the observations of colour change in the microcosms, indicating that
that the colour change is most likely due to microbial action.
4.3.5 Ions Analysis
A cation and anion analysis was conducted at 119 days to further understand the microbial
processes occurring within the microcosms.
A B
C 20µm
56
4.3.5.1 Cation Analysis:
A cation analysis generally showed greater concentrations of metals in the active microcosms
than in the control microcosms, shown in Figure 19. Refer to Table 12 for the specific results of
cations that reached concentrations above the detection limit.
Figure 19 Cation analysis at 119 days of microcosm monitoring
Table 12 Concentration of Metals in µg/L at 5 months of microcosms running, where *BDL
is below the detection limit
Sample Concentration (µg/ml)
Element (Limit of
Quantitation µg/L)
Soil 3 (Active)
Soil 3 (Active)
Soil 3 (Control)
Soil 2 (Control)
B (0.01) 1.1 0.9 0.3 0.7
Ba (0.001) 0.8 1.0 0.7 0.4
0
20
40
60
80
100
120
140
160
180
B Ba Ca Co Fe K Mg Mn Na Ni Si
Conc
entr
atio
n (µ
g/L)
Cation
Soil 3 (Active 1) Soil 3 (Active 2) Soil 3 (Control 1) Soil 2 (Control 2)
57
Ca (0.002) 159.2 166.5 52.0 45.0
Co (0.02) 0.4 0.4 0.1 BDL
Fe (0.01) 148.4 143.6 BDL BDL
K (0.005) 4.7 4.7 4.7 2.1
Mg (0.01) 84.2 87.5 30.3 37.9
Mn (0.01) 48.3 53.9 9.4 5.7
Na (0.005) 4.1 4.0 37.4 33.3
Ni (0.01) 0.2 0.2 0.1 BDL
Si (0.05) 17.2 17.4 6.6 9.9
This suggests that the metals, most notably iron, were reduced in the active microcosms and
oxidized in the control microcosms. The increase in concentrations is a result of the ferric iron,
Fe3+, reduced to the more soluble ferrous iron, Fe2+. This is indicative of anaerobic iron
bioreduction. The ferric iron in the control microcosms was oxidized to form iron oxides. This
finding compliments the observations of the colour transformation. Ferric oxides typically have a
yellow to orange colour which is exactly what is observed in the control microcosms. The green
colour of the active microcosms may be attributed to the formation of iron sulfides which have a
characteristic green colour [69]. Reducing environments with high ferrous iron in the aqueous
phase have been found at a methanol contaminated site in South Carolina, suggesting that these
are characteristic conditions for methanol degradation, which might be occurring at this stage in
the microcosms [70].
The decrease in manganese and magnesium is not surprising in this scenario as they would co-
precipitate with the iron, therefore reducing the concentrations in the control microcosms as
shown in the results. The high concentrations of sodium, Na, in the control microcosms may be
attributed to the sodium azide that was added as a sterilizing agent.
58
4.3.5.2 Anion Analysis:
The anions were also investigated to further understand the water chemistry throughout the
microcosm study. The concentrations of anions initially added to the microcosms in the synthetic
groundwater and the concentrations reported at 155 days are shown in Table 13.
Table 13 Concentration of anions in synthetic groundwater and Soil 3 supernatant at 155
days. Concentrations in mg/L
There are significantly higher concentrations of anions in the control microcosms than the
original synthetic groundwater. This suggests that additional anions leached out of the soil once
combined with the synthetic groundwater in the microcosm. The active microcosms show a
significantly lower concentration of each anion than the control microcosms with nitrite and
phosphate completely undetected in the active microcosms. As nitrite is reduced to nitrate in a
denitrification process nitrite would be the first anion to be depleted. This is represented by the
results in the active microcosm and suggests that denitrification may have taken place. The
results are also indicative of sulfate reduction as the sulfate concentrations are much lower in the
active microcosm than in the control microcosm. Phosphate concentrations are also depleted in
the active microcosm as they are utilized for growth by microbial populations [71].
The sulfate reduction along with denitrification in the active microcosms is indicative of the
methanol degradation. Similar conditions to those found in this study were found at the methanol
Anion Synthetic
Groundwater Time = 0
Microcosm Supernatant,
Soil 3 Active
Microcosm Supernatant,
Soil 3, Control
Sulfate 4.0 6.1 55.1
Nitrate 0.7 0.3 3.1
Chloride 132.5 149.4 253.4
Nitrite - - 2.1
Phosphate - - 26.8
59
contaminated South Carolina site, discussed in the previous section [70]. This is further
confirmation that methanol degradation might be the target of biodegradation in the microcosms
at the time of analysis. Once the methanol is depleted, it is expected that the more resistant
naphthalene degradation will commence. This will require further monitoring of the microcosms.
4.3.6 pH, Specific Conductivity and ORP
The pH, specific conductivity and ORP were measured at 149 days following microcosm setup.
Each temperature and soil scenario was tested during a destructive sampling event. The samples
were taken in an anaerobic glovebag and measured immediately to prevent significant changes
that may result from exposure to an aerobic environment. The measurements were taken in the
order of ORP, specific conductivity and finally pH. The results are found in Table 14.
Table 14 Supernatant parameters at 149 days
Temperature (ºC) Soil Type pH ORP
(mV)
Specific Conductivity
(µS)
30
Soil 1 4.5 36 1417
Soil 2 5.8 95 643
Soil 3 5.36 -53 1.97
45
Soil 1 4.97 89 973
Soil 2 5.62 73 600
Soil 3 5.59 50 630
60 Soil 1 5.22 53 680
Soil 2 5.45 107 595
The pH for each scenario is slightly acidic, ranging from 4.5-5.8. These results are lower than the
synthetic groundwater initially used to prepare the microcosms, however, the in-situ conditions
were also recorded to be slightly acidic, between pH 5 and 7 [46]. It is not unexpected that the
60
microcosms would reach equilibrium closer to the pH of the site as the synthetic groundwater
was not buffered. Furthermore, varying forms of the reaction for the anaerobic degradation of
methanol have been found to produce hydrogen ions which may contribute to a lower pH [72].
Many PAH degraders and methanol degraders degrade most efficiently at neutral pH, however,
degradation is possible at more extreme values [67].
Soil 3 at 30ºC is a notable scenario, showing a significantly lower ORP value of -53mV and a
lower specific conductivity of 1.97µS. This further compliments the findings in the previous
analyses suggesting this scenario is the most microbially active microcosm in a reducing
environment.
4.3.7 Methane Production
Headspace sampling was conducted at 149 days following microcosm setup to measure the
methane production within the microcosms. Methane was not detected within the majority of the
microcosms with the exception of the microcosm containing Soil 3 at 30ºC. The methane
concentration in this scenario far exceeded the calibration of the analytical method for methane
analysis previously developed. Concentrations were greater than 45 mg/L in the gaseous phase.
This finding once again compliments the previous analyses suggesting that Soil 3 at 30º has
significant methanogenic activity occurring within the microcosm, most likely due to methanol
degradation. High methane production is often associated with methanol degradation and has
been found at methanol contaminated sites [67] [70]. This is also confirmed by the high
concentration of archaea found in this scenario through qPCR analysis at 149 days. Archaea are
often associated with methanogenic activity.
Soil 1 and 2 at 30ºC did not show methane production, suggesting that the microbial community
responsible for producing methane was not present in either of these soils at this time. This may
be indicative that the higher temperatures reached for Soil 1 and 2, 101 and 70º C respectively,
inactivated this microbial community in the soil. It is also possible that these microbial
communities may take longer to acclimate post thermal remediation. This was also
complimented by the qPCR results showing little archaea presence at 149 days. Further
monitoring and analysis is required to determine if these microbial communities are present in
Soils 1 and 2.
61
Soil 3 was also analyzed for methane at 45ºC to determine if the microbial community could
survive at higher temperatures. Methane production was not found under this condition. This
suggests that this increased temperature was not a suitable condition to host this microbial
community. These results are consistent with the qPCR results.
62
Chapter 5 Conclusions and Recommendations
5.1 Conclusions In this study, microcosm batch studies were conducted to determine the impacts of thermal
remediation on the biodegradation of naphthalene by indigenous anaerobic bacteria at a
previously hydrocarbon contaminated site. Soil samples from a former bulk fuel terminal site
heavily contaminated with leaded and unleaded gasoline, diesel fuel and kerosene that underwent
thermal remediation in 2011-2012 were used in this study to simulate in-situ conditions. Three
peak temperatures reached during thermal remediation were tested (37, 70 and 101ºC) to
determine the impacts of peak temperature on the presence of microbial communities in the soil.
Microcosm batch studies were conducted to test each of these conditions at temperatures
experienced along the respective cooling profiles of each soil (30, 45 and 60ºC).
Naphthalene degradation was not detected in any of the scenarios tested within 150 days of
monitoring. This may be a result of long acclimation periods required for anaerobic
environments as acclimation periods of 18 months have been found for anaerobic hydrocarbon
degraders [60]. To further investigate this, the microcosms will continue to be monitored for a
longer period of time while maintained at their respective temperatures.
Although naphthalene degradation had not yet initialized at 150 days, there was evidence of
microbial activity occurring within the microcosms. Visual observations were the first indication
of microbial activity occurring as the active microcosms turned green, indicating a reduced
environment and the control microcosms turned orange, indicating an oxidative environment.
This observation was most prominent in the scenario of Soil 3 at 30ºC. This scenario reached the
lowest peak temperature during thermal remediation (37°C) and was maintained in the lowest
temperature water bath (30°C). This scenario was tested further to investigate the microbial
activity resulting in this change in colour. qPCR confirmed that there existed a higher
concentration of archaea in Soil 3 at 30°C. In addition, cation analysis confirmed the observation
showing higher concentrations of the more soluble reduced form of the ions in the active
microcosms than in the control microcosms. Microscopy analysis additionally complimented
these results as the active microcosms contained both rod and cocci shaped microorganisms
63
whereas the control microcosms did not. Significant methane production in Soil 3 at 30ºC further
confirmed these results indicating anaerobic microbial activity.
It is believed that the microbial activity occurring up to this point is due to the degradation of
methanol. Methanol is more readily degradable than PAHs and is typically used as an
amendment to soils to increase the microbial activity to reach populations large enough to
degrade the more resilient hydrocarbons, such as PAHs. It is expected that once the methanol is
further depleted, naphthalene degradation will be initiated. Further monitoring of the microcosms
will be required to determine these results.
5.2 Further Work In order to determine the impact of thermal remediation on the degradation of naphthalene by
indigenous microorganisms further, monitoring of the microcosms, along with additional
analyses are required. This will be continued by another member of the University of Toronto,
Civil Engineering Groundwater Research Group in the upcoming months. The work to be
continued is as follows:
• Microcosms will continue to be monitored for naphthalene concentrations over a longer
period of time to determine if acclimation of microbes to naphthalene occurs.
• Anions, cations, and microscopy analyses should be conducted for each scenario (each
soil type at each temperature tested). This will determine if the reducing environment
found in Soil 3 at 30°C is also occurring in the other scenarios. These less favourable
scenarios may require longer acclimation times and therefore should be monitored for a
longer period of time.
• Pyrotag sequencing should be conducted on the DNA samples that were frozen
throughout the microcosm study. This will give an indication of what species of microbes
are present.
64
5.3 Recommendations In addition to the work conducted in this thesis there are some analyses that are lacking in the
literature that would be useful to fulfill the objective of combining thermal remediation
technologies and bioremediation.
Hydrocarbon contaminated sites often occur in anaerobic conditions which are not as efficient
for biodegradation as aerobic conditions. The addition of oxygen to the subsurface, using air
sparging, during thermal remediation may be a feasible option to enhance biodegradation at these
high temperatures. Aerobic thermophiles capable of degrading PAHs have been isolated
suggesting that it is possible to enhance aerobic biodegradation at thermally remediated sites.
During an air sparging feasibility pilot study at the North Carolina Site, aerobic biodegradation
was found to occur. This study did not, however, determine if increased temperatures in
combination with air sparging would promote thermally enhanced biodegradation. It has been
found that oxygen transfer rates are increased with increasing temperatures and would be more
readily available for aerobic microbes at increased temperatures [12]. It is possible to test these
impacts on the currently monitored microcosms by introducing oxygen.
It would also be interesting to test the impacts on indigenous microbial communities throughout
the course of a thermal remediation project. As stated earlier, this has been conducted at PCE and
TCE contaminated sites, however literature is lacking for PAH and other hydrocarbon
contaminated sites. Unfortunately, for this study, soil samples were only available post thermal
remediation and so there was no baseline of microbial populations prior to or during thermal
remediation. It is recommended that a site be characterized for microbial populations throughout
the thermal remediation process to gain a better understanding to how the heating and cooling of
a site impacts the microbial communities. This type of field scale investigation will be more
useful in the optimization of a combined thermal remediation and bioremediation technology.
A field scale analysis also accounts for an open system where as the microcosm study performed
above simulates a closed system. An open system allows for the recharging of the thermally
treated area, including nutrients required for biodegradation along with microbial communities
present in the surrounding subsurface. This type of study may reflect a greater potential for
biodegradation post thermal remediation and should be considered in the future.
65
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Appendix A
Groundwater Naphthalene Concentrations
Sample Zone
76
Appendix B
External Analytical Results
Table 15 Groundwater Chemistry at North Carolina Bulk Fuel Terminal [46]
Field Parameters AMW-6S SBMW-4R pH 5.63 6.73 Specific Conductivity (umhos/cm) 88 570 Temperature © 15.34 16.39 Dissolved Oxygen (mg/L) NA 0.05 Re-Dox Potential (mV) 386 -3.3 Turbidity (NTU) >1000 >1000 HACH DR /890 (mg/L) Ferrous Iron NA 0.59 Sulfide NA 0.03 Volatile Organics (µg/L) USEPA Method 8260B Acetone <50 146 Benzene <1 638 Chloroethane <2 <40 Chloroform 1.7 <40 1,2 - Dichloroethane <2 <40 Di-Isopropyl ether <2 <40 Ethylbenzene <2 1710 2-Hexanone <10 <200 4-Methyl-2-pentanone <10 <200 Methyl Tert Butyl Ether <2 <40 Tert Butyl Alcohol <20 <400 Toluene <2 2000 Xylene (total) <6 3690 Semi Volatile Organics (µg/L) USEPA Method 8270C 2,4-Dimethylphenol NA NA 2-Methylphenol NA NA 3&4-Methylphenol NA NA
77
Phenol NA NA Dibenzofuran NA NA Bis (2-ethylhexl)phthalate (DEHP) NA NA 2-Methylnaphthalene NA NA Naphthalene NA NA Metals (µg/L) USEPA Method 6010B Calcium 24000 71700 Iron, total 103000 95600 Lead, total 20.3 57.5 Magnesium 32100 59600 Manganese 2670 25700 Sodium <5000 <5000 General Chemistry (mg/L) Alkalinity as CaCO3 27.5 353 Chemical Oxygen Demand <20 23.4 Chloride 7.7 6.9 Nitrogen, Ammonia <.2 0.25 Nitrate as N 0.7 <.06 Nitrite as N <.01 <.01 Phosphorus, total 0.57 0.31 Solids, Total Dissolved 260 348 Solids, Total Suspended 10200 5300 Sulfate 4 2 Total Organic Carbon <1 8.4 Biological Oxygen Demand <2 >25.3 Total Inorganic Carbon 210 1000 Dissolved Gases (µg/L) Methane -CH4 2.12 786 Carbon Dioxide- CO2 5460 8290 NA - Not analyzed
78
Appendix C
Figure 21 Temperature profile for Soil 2 during thermal remediation
Figure 20 Temperature profile for Soil 1 during thermal remediation
79
Figure 22 Temperature profile of Soil 3 during thermal remediation
80
Appendix D
MDL Calculations
A standard 1-5 times the estimated detection limit was analyzed 7 times. The standard deviation
of these values was taken.
𝑀𝐷𝐿 = 𝑆𝑡𝑢𝑑𝑒𝑛𝑡 𝑡𝑣𝑎𝑙𝑢𝑒 𝑥 𝑠𝑡𝑎𝑛𝑑𝑎𝑟𝑑 𝑑𝑒𝑣𝑖𝑎𝑡𝑖𝑜𝑛
Method Detection Limit for SPME GC-FID Naphthalene Analysis
Values were determined as follows:
Standard 566.2µg/L Peak Area
1 651.7
2 536.8
3 464.1
4 588.6
5 489
6 588.7
7 604.5
Standard Deviation 75.5
T, 99%, df=6 3.143
Area Count MDL 210.1
Concentration MDL (µg/L) 99.9
81
The lowest standard above the highest blank is used to determine the method detection limit as
follows.
𝐶𝑞 𝐷𝑒𝑡𝑒𝑐𝑡𝑖𝑜𝑛 𝐿𝑖𝑚𝑖𝑡 = 𝐶𝑞𝑙𝑜𝑤𝑒𝑠𝑡 𝑠𝑡𝑎𝑛𝑑𝑎𝑟𝑑 𝑎𝑏𝑜𝑣𝑒 𝑡ℎ𝑒 ℎ𝑖𝑔ℎ𝑒𝑠𝑡 𝑏𝑙𝑎𝑛𝑘 − 1.6
𝑆𝑞 𝐷𝑒𝑡𝑒𝑐𝑡𝑖𝑜𝑛 𝐿𝑖𝑚𝑖𝑡 =(𝐶𝑞 𝑑𝑒𝑡𝑒𝑐𝑡𝑖𝑜𝑛 𝑙𝑖𝑚𝑖𝑡 − 𝑦𝑖𝑛𝑡𝑒𝑟𝑐𝑒𝑝𝑡 𝑜𝑓 𝑐𝑎𝑙𝑖𝑏𝑟𝑎𝑡𝑖𝑜𝑛 𝑐𝑢𝑟𝑣𝑒)
𝑠𝑙𝑜𝑝𝑒 𝑜𝑓 𝑐𝑎𝑙𝑖𝑏𝑟𝑎𝑡𝑖𝑜𝑛 𝑐𝑢𝑟𝑣𝑒
Method Detection Limit for qPCR Results
Where, Cq is the X and Sq is the Starting quantity.
L