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This is to certify that the
thesis entitled
PHYTOREMEDIATION OF POLYCYCLIC AROMATIC HYDROCARBON-
CONTAMINATED SOIL USING NATIVE MICHIGAN PLANT SPECIES

presented by
Cindy Shin Mai Wan

has been accepted towards fulfillment
of the requirements for

MS. degree in Crop & Soil Sciences

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PHYTOREMEDIATION OF POLYCYCLIC AROMATIC HYDROCARBON -

CONTAMINATED SOIL USING NATIVE MICHIGAN PLANT SPECIES

By

Cindy Shiu Mai Wan

A THESIS

Submitted to
Michigan State University
in partial fulfillment of the requirements
for the degree of

MASTER OF SCIENCE
Department of Crop and Soil Sciences

2002

ABSTRACT
PHYTOREMEDIATION or POLYCYCLIC AROMATIC HYDROCARBON -
CONTAMINATED SOIL USING NATIVE MICHIGAN PLANT SPECIES
By

Cindy Shiu Mai Wan

Phytoremediation is the use of plants to degrade, detoxify, or remove
environmental contaminants. The Rouge Manufacturing Complex (Dearborn, MI) Coke
Oven area is contaminated with polycyclic aromatic hydrocarbons (PAHS), which were
formed from 60 years of industrial coal processing. PAHs are carcinogenic, mutagenic,
and teratogenic organic contaminants with low water solubility. PAHs sorb strongly to
organic matter in soils and sediments and consequently, are not readily available for
biodegradation. In this study, 18 native Michigan plant species and an unplanted control
were evaluated for their abilities to reduce PAHs in amended Coke Oven area soil over
one growing season in a field demonstration plot. Four plant species treatments
significantly decreased soil total PAH concentration ([tPAH]) over time; two plant
species treatments had lower soil [tPAH] compared to the unplanted control in July, and
the soil [tPAH] for one plant species treatment was lower than that for the unplanted
control in September. By contrast, the unplanted control soil [tPAH] did not decrease
over time. This study identifies plant species with superior PAH-phytoremediation

abilities for further laboratory studies and field applications.

ACKNOWLEDGEMENTS

I would like to express my appreciation to my advisor, Dr. Clayton Rugh, for
giving me the opportunity to work in his lab, creating this exciting project, and for his
suggestions and advice. I would like to thank Dr. Stephen Boyd, Dr. Tom Fernandez, and
Dr. Phil Robertson for serving on my graduate committee and for theirv’a‘luable _
comments during the courseof my study at Michigan Sta‘t‘egUniversity (MSU).

I gratefully acknowledge my lab coworkers and former coworkers for their
contribution to transplanting, collecting soil samples, sieving, vial washing, discussions
and reviewing manuscripts: Christina Harzman, Dr. Pulla Kaothien, Sarah Kinder, Sarah
Marshall, Susan Redwine, Chris Saffron, Rachada Settavongsin, Endang Susilawati,
Sharon Stump, and Theresa Wood. A special thank you to Theresa Wood for teaching me
the extraction protocol and for analyzing the samples. I would like to thank Emily B.
Smith and Dr. Sasha Kravchenko for statistical discussions and assistance with the
analyses. I would like to express my gratitude to Dave Freville for discussions about soils
and soil mixtures and for allowing us to use the cement mixer. I would like to express my
thanks to Andy Fogiel for discussions about compost. I am grateful to Dr. Lee Jacobs and
Dr. Brian Teppen for‘prov‘iding technical support. I would like to thank North American
Prairies for their donation of seed.

I have been very fortunate to have had many inspiring and caring teachers in my
life. I would like to thank Dr. Nancy Dengler, Dr. Robert Jefferies, Dr. Tammy Sage, and
Dr. Rowan Sage at the University of Toronto for sharing their passions for their work
with me, being excellent undergraduate professors who taught me so much, giving me

thoughtful advice, and inspiring me to pursue further studies in plant biology and

iii

ecology. I would especially like to thank my previous mentor, Dr. Rowan Sage, forhis
advice, support, encouragement, and for always having my best interests at heart.

I would like to thank my new friends at MSU for chocolate, discussions and
support: Ping Ping Jiang, Sherill Baldwin, and Jihye Lim. I would also like to thank
Rachada Settavongsin for reminding me to eat when I forgot and for bringing medicine to
me when I was ill (even if I was allergic to it). To my lifelong friends, Cathy Chan, Chi
Phong Luong, Nela Veljkovic, Tahti Leesment, Ventura Wu, Diane Leal, Hang Huynh,
Elizabeth Tang, and Kristina Korogyi: “You were the rain when my spirits were dry”.
Last but not least, I would like to express my gratitude and appreciation to my parents,
sister, brothers, and brother-in-law for all they have taught me and for their love and

support.

iv

TABLE OF CONTENTS

PAGE
LIST OF TABLES ....................................................................... vi
LIST OF FIGURES ..................................................................... vii
LIST OF APPENDICES ................................................................ vii
INTRODUCTION ........................................................................ 1
CHAPTER 1. REVIEW OF LITERATURE ......................................... 3
References ....................................................................... 29
CHAPTER 2. FIELD STUDY OF MICHIGAN NATIVE PLANTS
FOR PHYTOREMEDIATION OF A PAH-
CONTAMINATED SOIL ............................................ 40
Abstract ........................................................................... 41
Introduction ...................................................................... 42
Materials and Methods ......................................................... 46
Results ........................................................................... 59
Discussion ........................................................................ 79
Summary and Conclusion ...................................................... 87
References ........................................................................ 89
APPENDICES ........................................................................... 92

LIST OF TABLES
PAGE

Table 1.1. PAH Chemical and Toxicological Properties ................................... 16

Table 2.1. Plant species at Phytoremediation Demonstration (Phyto Demo) Site. . ....47

Table 2.2. Soil properties of Phyto Demo site ................................................ 51
Table 2.3. Soil nutrients of Phyto Demo Site ............................................... 52
Table 2.4. PAH abbreviations and detection limits ......................................... 58
Table 2.5. Upland treatment means ........................................................... 61

Table 2.6.. Statistical Results using upland data from May, July, and September. . .....62

Table 2.7. Phyto Demo Upland [P1anted]/[Unplanted] treatment ratios ................. 67
Table 2.8. Phyto Demo plant [tPAH] in wetland and upland plots ....................... 77
Table 2.9. Phyto Demo plant [tPAH] in control plot ....................................... 78
Table A1. APGEN greenhouse study data .................................................... 93
Table A2. APGEN greenhouse [Planted]/[Unplanted] treatment ratios ................ 95
Table A3. Phyto Demo plant mortality ........................................................ 96
Table A4. Phyto Demo plant individual PAH concentrations ....... l ...................... 97
Table A5. 1. Greenhouse (GH) study treatment codes ..................................... 103
Table A5.2. Soil properties for GH study ................................................... 106
Table A5.3. Soil nutrients for GH study ..................................................... 108
Table A5.4. GH study soil [tPAH] ............................................................ 112

Table A5.5. GH study soil [tPAH] maximum pot ranges and treatment ranges. . . . . ...1 13

Table A5.6. GH study plant [tPAH] .......................................................... 115

vi

LIST OF FIGURES

PAGE
Figure 1.1. PAH structures ...................................................................... 15
Figure 2.1. Overhead view of Phytoremediation Demonstration site ...................... 48
Figure 2.2. Phyto Demo Treatment Plot cross-section schematic .......................... 49
Figure 2.3. Phyto Demo Treatment Plots Layout schematic ................................ 54

Figure 2.4 (a—c). Plot cell [tPAH] changes May — July — Sept indicated by colors. . ....63
Figure 2.5 (a —b). Cell percentage decrease from May — Sept indicated by colors. . ....64

Figure 2.6 (a-l). Percentage reduction in soil [tPAH] and individual PAH

concentrations [iPAH] for all treatments ............................................. 69
Figure A5.1 (a-c). Plant shoot dry weight ..................................................... 116
Figure A5.2 (a—c). Plant root dry weight ...................................................... 1 17

vii

LIST OF APPENDICES

Page
Appendix 1. APGEN Greenhouse soil [tPAH] data ...................................... 93
Appendix 2. APGEN planted soil [tPAH] /unplanted soil [tPAH] ratios .............. 95
Appendix 3. Phyto Demo plant mortality data ............................................. 96
Appendix 4. Plant individual PAH compounds ........................................... 97

Appendix 5. Greenhouse Study of Effects of Plant Species and Compost on PAH
Phytoremediation ..................................................................... 101 ,

viii

INTRODUCTION

Phytoremediation, the use of plants to degrade, detoxify, remove or contain
environmental contaminants, is an emerging field in environmental rehabilitation.
Phytoremediation is a sub-discipline of bioremediation, which more commonly describes
the use of microbes for treatment of contaminants. The use of plants to remediate a site
has advantages over traditional engineered cleanup or bioremediative techniques. Plants
can stabilize soil by intercepting the impact of raindrops, absorbing and taking up water
from the soil, thereby minimizing soil erosion. Vegetation increases organic matter in the
soil and prevents the loss of organic matter by wind erosion by decreasing the soil surface
area exposed to convection. Unlike microbes, plants are able to reduce leaching of water-
soluble contaminants because they utilize water fi'om soils. In addition, plants harvest and
utilize the sun’s energy, whereas most engineering remediation technologies require the
expensive input of energy for operating machinery to decontarninate soils.

Studies on phytoremediation of inorganic and organic contaminants have focused
on plant selection, emphasizing screening for superior species, selected plants, and
symbiotic interactions between plants and microorganisms. Recently, research has
investigated the influence of factors such as soil amendments. Most phytoremediation
research has been conducted under laboratory or greenhouse conditions. This thesis
describes laboratory-scale screening of a variety of Michigan native plant species
followed by field-scale application of selected species.

The Ford Rouge Manufacturing Complex in Dearborn, MI has been
manufacturing steel and automobiles for eighty years. By-products from these activities

have led to contamination of parts of the site. Areas of the site are contaminated with

polycyclic (polynuclear) aromatic hydrocarbons (PAHs or PNAs), which were formed
from the production of coke for the smelting of iron ores. PAHs are carcinogenic,
mutagenic, and teratogenic. They are environmentally persistent, due in part to their low
water solubility, which cause them to sorb strongly organic matter in soils and sediments.
Consequently, they are not readily bioavailable and are resistant to biological
degradation. As a part of the Rouge Heritage Initiative Renovation designed by Bill
McDonough & Partners (Charlottesville, VA), the Ford Motor Company Environmental
Quality Office, Ford Land Corporation, and Michigan State University’s
Phytoremediation Lab are collaborating to develop a phytoremediation strategy for the
Ford Rouge Facility.

The objectives of this thesis were to evaluate the efficacy of plant Species for the
phytoremediation of PAHs and to broaden our understanding of the phytoremediation of
organic contaminants with low water solubility in soils. This study was designed to
characterize potential plant species to be used for PAH phytoremediation. Plant species
that exhibit the greatest rate of PAH reduction could be applied in future laboratory
studies and large-scale environmental rehabilitation efforts such as at the Rouge
Manufacturing Complex. Furthermore, determining which species have the highest PAH-
soil decontaminating capacities is a primary step in identifying plant species for further
research, which could lead to greater understanding of the biochemical reactions and

mechanisms involved in PAH phytoremediation.

 

CHAPTER I

REVIEW OF LITERATURE

1. INTRODUCTION

Environmental contamination of land and water natural resources with hazardous
materials is a worldwide concern. The source of contamination on hazardous sites is
frequently anthropogenic, typically resulting from industrial and military activities. As of
August 2001 there were 1235 Superfiind National Priorities List sites in the United States
being cleaned up under the Superfund program (EPA, 2002b). A brownfield is an urban
site that is either abandoned or under-used because it has real or perceived environmental
contamination, though potential for redevelopment or reuse. The number of brownfield
sites has been estimated to be more than 2900 in Canada (National Round Table on the
Environment and the Economy, 1998). In United States there are over 5000 brownfields
(EPA, 2002a). In many cases, contaminated sites are abandoned, have low soil fertility
and are poorly vegetated. Such areas can further deteriorate via wind and water erosion
resulting in the loss of soil nutrients and organic matter. Environmental rehabilitation is
necessary to prevent further land degradation, and to protect humans and wildlife from
exposure to hazardous pollutants.

Persistent organic pollutants (POPS) are toxic chemicals that do not readily

undergo biogeochemical reactions, remain in soils for a long time, and are prone to

LI)

biomagnify through the food chain. Biomagnification refers to the increase in
contaminant concentration at sequentially higher levels of the food web. PAHS are
harmful to humans and wildlife because they are carcinogenic, mutagenic and
teratogenic. Examples of POPS include the pesticide l,l,1-trichloro-2,2-bis(p-
chlorophenyl)ethane (DDT), polychlorinated biphenyls (PCBS), and polycyclic aromatic
hydrocarbons (PAHS). Unlike synthetic pesticides and PCBS, PAHS are formed naturally
in the environment as a result of forest fires, volcanic eruptions, thermal geologic
reactions, and plant and bacterial reactions (Blumer, 1976). Since the 18005, the
beginning of the industrial revolution, anthrOpogenic activities have led to the production
of vast amounts of PAHS, which have exceeded the levels that are naturally degraded,
and created an imbalance between PAH formation and degradation (Hites, 1977; Suess,
1976). Anthropogenic sources of PAHS include the burning of fossil fuels, railroad
industries, manufactured gas plants, and coke production.

Current strategies for remediation of PAH-contaminated soil include physical,
chemical and or biological treatment, though each has its drawbacks. Isolation and
containment use physical, chemical or hydraulic barriers to inhibit the distribution of the
contaminant, but do not reduce the level of the contaminant. For example, capping
involves mixing the soil with clay to reduce hydraulic conductivity (Cunningham and
Berti, 1993). In Situ thermal desorption treatment of soil for petroleum hydrocarbons
involves the application of heat and vacuum via thermal wells to vaporize,
decontaminate, or transport contaminants to the surface for further treatment (Conley,
2000). Thermal desorption may be effective, but this treatment requires the high input of

energy and the installation of wells. Soil washing consists ofmixing the soil. and

separation of the pollutant portion (silt and clay) of the soil from the portion with less
pollutant (sand and gravel). Soil washing is a commonly used strategy that requires
intensive labor and money (EPA, 2001). Excavation of contaminated material is often
favored because it rapidly removes contamination from a site, but this method of
remediation only transfers the contamination from one location to another where the
pollutant will persist (Cunningham and Berti, 1993).

Biological treatments of contaminated soil include natural attenuation, microbial
bioremediation, and phytoremediation. Natural attenuation is the use of indigenous soil
processes without intervention. Natural attenuation has reduced organic contaminants via
microbial degradation (Hiebert, 2000). Natural biodegradation of hydrocarbons and
chlorinated hydrocarbons occurred at an oilfield service facility. The intermediates and
products of microbial degradation of hydrocarbons and chlorinated hydrocarbons
were observed to increase over time. Elevated levels of methane were detected indicating
microbial degradation of chlorinated hydrocarbons and ethene (by product of
biodegradation of tetrachlorethene (PCB) and trichloroethene (TCE)). Carbon dioxide,
the final product of biodegradation of hydrocarbons and some chlorinated hydrocarbons,
increased (Hiebert, 2000). Natural attenuation is low-cost, but may not be effective if
initial contaminant concentration is high or toxic to plants and microorganisms. Microbial
bioremediation uses microorganisms to metabolize complex organic molecules.
Microbial degradation of PAHS has been extensively demonstrated in research literature
(Cerniglia, 1992; Cerniglia, 1979; Bumpus, 1985; Field e1 (11., 1992). Microbial
remediation typically requires nutrient inputs and adjustment of soil properties, such as

pH or temperature, so that the degrading bacteria and or fungi can persist, a process

‘JI

known as biostimulation. The contaminated soil and microorganisms are sometimes
mixed in situ, which is a disruptive, though sometimes beneficial, procedure. In some
cases, reactors are used for bioremediation (Civilini and Sebastianutto, 1996; C ivilini er
al., 1996; Lilja e! (21., 1996). The use ofa bioreactor involves transport ofthe soil,
possibly destroying the Site and often incurring high operating costs. Landfarming of
contaminated materials has also been shown to be a feasible method of remediation and
involves routine soil tillage (8 -12” depth) and the addition of fertilizer to enhance
microbial degradation of organic contaminants in the absenCe of plants (Sayles et al.,
1999; Reilley et al., 1996). This technique is often used for petroleum hydrocarbons, but
dissipation Slows over time (Sims and Overcash, 1983).

Remedial programs can also combine biological and chemical methods. For
instance, soil contaminated with PAHS and pentachlorophenol was treated in a laboratory
experiment using chemical oxidation by adding Fenton’s reagent (ferrous iron and
hydrogen peroxide) to generate free hydroxyl radicals followed by indigenous microbial
biodegradation of the chemically oxidized compounds (Allen and Reardon, 2000). For
large cleanup operations, the cost of this method may be prohibitively high.

Phytoremediation, or vegetated treatments, is a method of environmental
rehabilitation that could potentially reduce the concentration of contaminants and
improve soil quality simultaneously. Phytoremediation is the use of plants to degrade.
detoxify or remove inorganic and organic contaminants (Cunningham and Berti, 1993).
Phytoremediation has been demonstrated to accelerate contaminant biodegradation
during natural attenuation, microbial bioremediation, and landfarming. Previous studies

showed vegetated soil leads to greater rates of PAH reduction compared with unplanted

6

soil (Aprill and Sims, 1990; Nedunuri et (11., 2000; Pradhan et al., 1998; Yateem e! (11.,
2000). Vegetated landfarming was 30-44% more effective for PAH reduction than soil
landfarming with no plants (Reilley et a]. , 1996). Few studies, however, have reported the
individual effects of a variety of plant species on phytoremediation of PAHS. Plants may
secrete different compounds that support soil microflora, and some plant species may
favor the PAH-degrading microorganisms via exudation of specific compounds.

The Ford Rouge Manufacturing Complex in Dearborn, MI, once the largest
integrated industrial facility in the world, has areas contaminated with byproducts from
eighty years of steel and automobile manufacturing. PAHS have accumulated in areas of
the facility used during coal processing for coke production for iron ore smelting. PAHS
are highly hydrophobic organic contaminants that tend to sorb strongly to the soil and
sediment organic matter fraction. As a consequence, PAHS are difficult to biodegrade and
remain in the soil for extended periods. PAHS pose health hazards to humans and wildlife
because they are carcinogenic, mutagenic, and teratogenic. The purpose of this thesis is to
evaluate the effectiveness of various native Michigan plant Species for phytoremediation
of PAHS in soil from the Rouge Manufacturing Complex. It is hoped that information
gained from this study will advance our understanding of processes involved in
phytoremediation of PAH pollutants.

2. LITERATURE REVIEW

2.1 Phytoremediation

Phytoremediation is the use of plants to degrade, detoxify or remove
environmental contaminants (Cunningham and Berti, 1993) and has been reviewed in

numerous papers (Alkorta and Garbisu. 2001; Cunningham and Berti, 1993; Macek er (1].,

2000; Salt et al., 1995). Bioremediation refers to the using biological means to remove
contamination. In the literature, “Bioremediation” typically refers to the use of bacteria
and fungi to remove pollutants from the environment. Phytoremediation is a subdiscipline
of bioremediation and consists of a variety of strategies based on different mechanisms of
contaminant removal. Phytoextraction is the use of plants to remove inorganic
contaminants, typically metals, from soil by concentrating them in harvestable plant
parts. Phytoextraction of useful or valuable metal pollutants (e. g. Zn, Cu) with
subsequent harvesting and recovery is referred to as biomining (Cunningham and Berti,
1993) or phytomining (Pletsch et (11., 1999). For plants to decontaminate a site within a
reasonable number of harvests, it has been proposed that plants must accumulate 1 to 3%
of a metal per dry weight aboveground biomass (Cunningham and Ow, 1996). Plants that
can accumulate a contaminant in high concentrations are known as hyperaccumulators.
Phytostimulation, or plant-assisted bioremediation, is the enhancement of microbial
biodegradation in the rhizosphere. Rhizofiltration is the use of plant roots to absorb
mineral or heavy metal contaminants from water and aqueous waste streams and
subsequent disposal of laden biomass. Phytostabilization is the use of plants to reduce
motility of pollutants in the environment by sequestration, lignification, or humification
in plant or soil matrices. Phytostabilization is usually used on metal—contaminated sites to
prevent erosive particles from increasing the area of contamination. Phytovolatilization is
the use of plants to uptake a contaminant and then convert it to a volatile form that is
released into the atmosphere. Phytodegradation (also known as phytotransformation) has
been defined as the absorption and conversion by catabolism or anabolism in the plant

root or shoot. Phytodegradation has also been defined as the use of plants and associated

microorganisms to degrade organic pollutants (Cunningham et al., 1995), however in this
thesis the previous definition of phytodegradation will be used because the terms
phytodegradation and phytostimulation distinguish between plant degradation of the
contaminant and plant-assisted microbial degradation, respectively.

Plants have been demonstrated to be an effective approach for remediation of
inorganic pollutants. Lead can be removed from soil by phytoextraction by
hyperaccumulators such as T hlaspi rotundifolium (Cunningham and Ow, 1996; Reeves
and Brooks, 1983). Lead uptake by Brassicajuncea (Indian mustard) was enhanced
when the synthetic chelator ethylene diamine tetraacetic acid (EDTA) was added to
hydroponic solution (Vassil et al., 1998) or soil (Blaylock et al., 1997). Selenium (Se)
can be phytoextracted by Brassica napus (canola) (Banuelos and Mayland, 2000). The
Se-enriched shoots of B. napus may then be harvested and used as forage for Se-deficient
livestock (Banuelos and Mayland, 2000). B. juncea has also been shown to
phytovolatilize selenium (de Souza et al. , 1998). Soils contaminated with arsenic can be
remediated by phytoextraction using Pteris viltata (brake fern) (Ma et al., 2001) or B.
juncea (Pickering et al., 2000). Arsenic uptake is enhanced by addition of
dimercaptosuccinate, a chelator of dithiol arsenic (Pickering et al., 2000). Phytoextraction
of nickel can be accomplished by T hlaspi goesigense (Kramer et al., 1997; Persans et al.,
1999) and several Brassicaceae species (Baker, 1989). Thlaspi caeru/escens
(Brassicaceae), a hyperaccumulator, can phytoextract zinc (Tolra (2101., 1996) and
cadmium (Whiting er al., 2000). Phytoremediation may take 2—20 years depending on
clean-up goals, the volume of contaminated soil, the distribution and concentrations of

contaminant, soil characteristics, depth of contamination. plant growth rate, and climate

(Naval Facilities Engineering Center, 2002). Phytoremediation of metals in soil costs
$25-$100 per ton of soil. Conventionally-used remediation techniques can cost
considerably more: soil washing ($50-150/ton), in situ soil flushing ($75-$210/ton), ex
situ solidification/stabilization ($75-$150/ton), in situ solidification/stabilization ($1 1 l-
205/ton), thermal desorption ($150-$500/ton), thermal treatment ($200-$450/ton), and
landfilling ($100-$500/ton)(Schnoor, 2002). The excavation of one acre of sandy loam
soil to a depth of 50 cm would cost $400 000 for excavation and storage using
conventional soil removal methods. By contrast, phytoextraction of the same soil would
cost $60 000 — $100 000 (Salt et al., 1995).

Phytoextraction can be also used to remediate soils contaminated with

radionuclides. Redroot pigweed (Amarant/ms retroflexus) has been shown to

. . . 137 .
hyperaccumulate radioactive ceSIum Cs, a byproduct of nuclear fiSSIOn (Lasat et al..
1998). A recent phytoextraction study Showed that A. retroflexus. B. juncea and

Phaseolus acutifolus A. Gray (tepary bean) removed 908r and 137Cs from soil in a field

study (F uhrmann et al. , 2002). Radionuclide concentration ratios (plant contaminant

concentration divided by that in soil) for that A. retroflexus, B. juncca and Phaseolus

. 137 . 9O
acunfolus were 2.58, 0.46, 0.17 for Cs, respectively and 6.5, 8.2, 15.2 for Sr,
respectively (F uhrmann et al. , 2002). A plant to soil concentration ratio greater than one

. . . . . . 134
Indicates that the plant 1S accumulating the contaminant. High levels of CS were taken

up by A groslis capillaris (bent grass) (Sanchez 6! al., 1999). Brassica narinosa (Chinese

mustard), Brassica chinensis (Chinese cabbage) and B. jzmcea have demonstrated

hyperaccumulation potential of uranium in the presence of citric acid (Huang et al.,
1998).

Phytoremediation has been used to treat soils containing organic contaminants
such as TCE (trichloroethene), BTEX compounds (benzene, toluene, ethylbenzene,
xylene), TNT (2,4,6-trinitrotoluene), RDX (Royal Demolition Explosives; 1,3,5-trinitro-
1,3,5-triazine). pesticides and PAHS principally by phytodegradation or phytostimulation.
Numerous field studies used hybrid poplars (Populus trichocarpa x Populus deltoia’es
and P. trichocarpa x P. maximowiczii) to metabolize TCE to metabolites such as chloral
hydrate, trichloroethanol, di- and trichloroacetic acid, and carbon dioxide(Newman et al.,
1997). Hybrid poplars were shown to degrade TCE due to plant dehalogenase enzyme
activity (Schnoor et al. , 1995). Phytoremediation has also been demonstrated for
nitroaromatic compounds, such as nitrobenzene (McFarlane et al., 1990) and hybrid
poplar (P. deltoides x P. nigra) metabolism of TNT (Thompson et al. , 1998). Plants have
potential for PAH phytodegradation Since they possess oxygenase, peroxidase, and
laccase enzymes, but this ability has not been clearly demonstrated (Criquet et al., 2000).
These studies indicate there is potential for phytodegradation to effectively remediate
organic contaminants.

In addition to phytodegradation, plants can also remediate organic contaminants
by phytostimulation. Poplars can phytostimulate microbial degradation of TCE (Walton
and Anderson, 1990). Microorganisms in the rhizosphere can degrade TCE to form

metabolites such as ciS-1,2-dichloroethylene and vinyl chloride, and degrade TCE

completely to carbon dioxide (Walton and Anderson, 1990). Greater mineralization of

14 . . . . . . . .
C—TCE and microbial respiration were observed in rhizosphere 8011 than in unvegetated

11

soil (Walton and Anderson, 1990). Plant roots have been Shown to increase the microbial
count and enhance mineralization in soils contaminated with the pesticides parathion and
diazinon (Hsu and Bartha, 1979). Plants such as Morus rubra (Mulberry), Rhus
aromatica (sumac), Malclura pomifera (osage orange), Helianthus maximillani
(perennial sunflower) can provide PCB-degrading bacteria with cometabolites such as the
phenolic compounds flavonoid and coumarin (Donnelly et al., 1994; Fletcher et al., 1995;
Fletcher and Hegde, 1995). Plants can provide cometabolites, e. g. phenolics or terpenes,
for PAH-degrading microbes (Hegde and Fletcher, 1996). Phytoremediation of PAHS
will be discussed in more detail later in this chapter.

The cost of phytoremediation of organic contaminants is lower compared with
other remediation strategies. Phytoremediation using fine-rooted grasses costs $10-$35
per ton of soil (Schnoor, 2002). By contrast, the costs of other approaches are: in situ
bioremediation $50—$150/ton, soil venting $20-$220/ton soil washing $80-$200/ton,
thermal treatment $120-$300/ton, solidification/stabilization $240-340/ton, and
incineration $200-1500/ton (Schnoor, 2002).

Biotechnological methods have been employed for improvement of plants for
environmental clean-up. Plant may be genetically altered to change plant morphology to
favor remediation processes. Plants can be genetically transformed by using
Agrobacterium rhizogenes to produce increased root biomass (Stomp et al., 1993; Stomp
et al., 1994; de Araujo et al., 2002; Shanks and Morgan, 1999). This transformation
would enhance the root surface area, possibly increase root exudation, which in turn
could increase microbial activity and contaminant biodegradation. Increased root biomass

may also lead to increased contaminant uptake (Nedelkoska and Doran, 2000b;

12

Nedelkoska and Doran, 2000a). Plants can also be genetically altered to produce enzymes
that can degrade or transform contaminants. For instance, Arabidopsis thaliana plants
and Nicotiana tabacum (tobacco) were transformed with the bacterial merA gene, which
encodes mercuric reductase, and merB gene, that encodes organomercurial lyase (Rugh et
al., 1996; Rugh et al., 1998; Bizily et al., 2000). MerB enzyme catalyzes the degradation
of organic mercury to Hg(II) and the MerA enzyme catalyzes the reduction of Hg(II) to
Hg(0), a much less toxic form of mercury that volatilizes to the atmosphere (Summers,
1986). The mer gene transformed plants evolved substantial amounts of elemental
mercury compared with the control and were able to tolerate 25-100 uM HgClz, levels
toxic to untransformed plants (Rugh et al., 1996). Transgenic plants have been developed
for the phytoremediation of organochlorides such as TCE. Plants engineered to express to
a mammalian cytochrome P450 gene were capable of 400 times greater degradation of
TCE than wildtype plants (Doty et al. , 2000). Transgenic poplar plants have also been
developed that can overexpress y-glutamylcysteine synthetase, the rate-limiting step in
glutathione synthesis (Rennenberg, 1997; Gullner et al., 2001), Glutathione binds
organochlorides, which makes them less toxic and tags them for vacuolar import
(Edwards et al. , 2000). Transgenic tobacco plants expressing the bacterial nitroreductase
gene from Enterobacter cloacae showed increased tolerance and detoxification of TNT
(2,4,6-tiinitrotoluene) compared to wildtype (Hannink et al., 2001). Field studies and
research experiments to evaluate the safety and cross-fertilization of transgenic plants
with wild populations need to be conducted before these biotechnological advances can

be practically used in phytoremediation.

13

Phytoremediation is a remediation strategy that has many advantages over other
clean-up technologies. Phytoremediation may be implemented with minimal disturbance
to a site, Simultaneously rehabilitating the soil, and with reduced ii 31: of contaminant
distribution. Phytoremediation can enhance bioremediation by providing carbon sources,
cometabolites, and improved soil properties, such as decreased pH, increased porosity,
decreased bulk density. Transgenic plants may be easier to control compared with
transgenic microorganisms. Phytoremediation requires relatively low maintenance, is
aesthetically pleasing, and is compatible with restoration ecology. 1n the United States,
the use of conventional technologies for cleanup of existing contaminated sites is
estimated to cost $10 billion and treatment of hazardous wastes to be at least $400 billion
(Salt et al. , 1995). The costs of phytoremediation are expected to be‘lower than standard
engineering-based approaches.

Despite the potential for ecological and economic advantages, phytoremediation
has its limitations. Vegetated treatments cannot access deep contaminants, may take
longer than most other methods, and are restricted to the growing season. Plant-based
remediation may not be effective at high levels or for all contaminants. In spite of these
shortcomings, phytoremediation is a relatively new field with potential to enhance and
complement other remediation strategies.

2.2 Polycyclic Aromatic Hydrocarbons (PAHS)

Polycyclic (polynuclear) aromatic hydrocarbons are widely distributed
environmental pollutants. PAHS consist of two or more fused benzene or furan rings
arranged linearly, angularly or in clusters (Blumer, 1976) (Fig. 1.1). Heterocyclic

aromatic compounds are formed when the carbon in the benzene is substituted with

14

Naphthalene

Acenaphthylene

Acenaphthene 0%

F luorene
Phenanthrene GOO
Anthracene 000
F luoranthene 0.0 1

Figure 1.1. Structures of some PAHS.

fi

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Benz(a)anthracene 000‘
Chrysene 00
Benzo(b)fluoranthene $0.0
Benzo(a)pyrene CO. I

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16

nitrogen, sulfur, oxygen, or other elements (Blumer, 1976). PAHS are formed as a result
of incomplete combustion of organic material such as when there is an insufficient supply
of oxygen (Heil, 1998). Temperature and combustion conditions, rather than fuel type,
influence the Specific PAHS formed (Giger, 1974; Jenkins et al., 1996; Blumer, 1975).
There are some general chemical and toxicological trends for PAH compounds. PAHS of

successively higher molecular weights have lower aqueous solubility, greater

hydrophobicity as indicated by higher log KOW values, and greater carcinogenicity (Table

1.1). The log KOW is a term that describes the hydrophobicity of a compound and is the

logarithm of the concentration of a compound dissolved in the octanol phase divided by
the concentration in the water phase in a partitioning assay.

PAHS are formed by both natural and anthropogenic processes. Prior to the
twentieth century, there was a natural balance between the production and degradation of
PAHS (Hites, 1977; Suess, 1976). PAHS occur naturally as a result of thermal geologic
reactions associated with fossil fuel and mineral production, during the burning of
vegetation in forest and brush fires, and also by some plant and bacterial reactions
(Blumer, 1976). Human activities such as combustion of wood and fossil fuels, petroleum
refining, coal and oil Shale conversion, and chemical manufacturing lead to the formation
of PAH compounds creating areas of high PAH deposition and accumulation (Hites,
1977; Suess, 1976).

Background levels of PAHS prior to the industrial age were substantially lower
than levels currently seen around the world. The main source of terrestrial PAHS is from
atmospheric deposition of PAHS from the combustion of fossil fuels. At the Rothamsted

Experimental Station in southeast England, PAH concentrations in soil core samples from

the mid-18003 to the present were analyzed (Jones, 1989). The total PAH concentration
found in soil samples from the mid 18003 was 350 ng/g dry weight and has since
increased at an accelerating rate to 1770 ng/g dry weight reflecting the rise in
anthropogenic activities, such as fossil fuel combustion and growing worldwide
industrialization (Jones, 1989).

PAHS are hazardous compounds and major environmental problems. Several
PAHS are carcinogenic, teratogenic, and mutagenic (Shabad, 1975; Sims and Overcash,
1983; Dipple, 1990). Fish from industrially contaminated water with benzanthracene and
other contaminants had four times more tumors than fish from unpolluted waters (Brown,
1973).Since PAHS can biomagnify, accumulation in organisms such as oysters (Lee,
1978) and fruit flies (Southworth, 1978), has consequences for animals higher in the food
web. PAHS are environmentally persistent due to their tendency to partition to organic
matter. This results in strong sorption to soils and sediments, hindering their degradation
and resulting in their persistence as stable soil complexes (Means, 1980). The
biogeochemical fate of PAHS in soil is controlled in part by surface adsorption (Reilley er
al. , 1996) and sorption to other soil components. Research has shown that even when
soil microbes were abundant, PAHS were not degraded due to their lack of bioavailability
(Carmichael and Pfaender, 1997).

Regulatory agencies have set enviromnental limits for PAHS that attempt to

balance economic and health interests. The US. Environmental Protection Agency states

. . . . 3
that particulate concentrations of PAHS in air above 150 rig/m are unhealthy and above

420 rig/m3 are hazardous (Heil, 1998). As indicated by the Michigan Department of

Environmental Quality (MDEQ) clean-up regulations. PAHS are toxic at different levels.

18

According to the MDEQ, the 4-ring PAHS - benz(a)anthracene, Chrysene, pyrene - and
the 5 and 6 ring PAHS are permitted in soils and waters at particularly low levels (Table
1.1). The MDEQ has designated acceptable levels for PAH industrial and commercial
long—terrn ingestion groundwater cleanup criteria, and dermal exposure to contaminated
soil (Table 1.1). These criteria provide targets for environmental rehabilitation efforts.
2.3 PAHS in soil

Microbial degradation

Biodegradation of PAHS is positively correlated with water solubility (Aronstein
et al. , 1991). PAHS with more rings and higher molecular weight have lower water
solubility and tend to sorb strongly to organic matter and soil (Bossert and Bartha, 1986;
Reilley et al., 1996; Carmichael and Pfaender, 1997). Subsequently, two- and three-ring
PAHS biodegrade more readily than those of four-, five- and six-ring PAHS (Bossert and
Bartha, 1986).

Microorganisms (bacteria, fungi, algae) can degrade PAHS under aerobic
conditions using oxygenases to incorporate oxygen into the carbon ring (Dagely, 1975).
Microbial degradation of PAHS has been well summarized in a variety of comprehensive
reviews (Sims and Overcash, 1983; Wilson and Jones, 1993; Cerniglia, 1993; Walton,
1994)

There are two classes of oxygenase enzymes: monooxygenase and dioxygenase.
Eukaryotes, including mammals and fungi, possess monoxygenases, which incorporate
one oxygen atom into the aromatic substrate to form arene oxides (epoxides) followed by
enzymatic addition of water to yield trans-dihyrodiols and phenols (Cerniglia, 1993;

Wilson and Jones, 1993). The trans-dihydrodiol is oxidized to a catechol, which is then

19

subjected to ring cleavage enzymes (Sims and Overcash, 1983). Further catabolic
activities lead to the production of tricarboxylic acid cycle (TCA) intermediates such as
succinic, fumaric, pyruvic, acetic acids and acetaldehyde (Heitkamp, 1988a; Heitkamp,
1988b). Since the monoxygenase PAH biodegradation pathway forms epoxides, this may
be a cause of PAH ecotoxicity. Epoxides have been shown to bind DNA and RNA
initiating the formation of tumors in carcinogenesis (Sims and Overcash, 1983).

The dioxygenase pathway does not lead to the formation of mutagenic and

carcinogenic epoxides. Dioxygenases are found in prokaryotes, including bacteria and

some blue-green algae, and incorporate both atoms of molecular oxygen (Oz) into an

aromatic substrate (Sims and Overcash, 1983). Dioxygenase attack on an aromatic ring
results in the production of cis—dihydrodiol (Cerniglia, 1993; Wilson and Jones, 1993).
The cis-dihydrodiol is oxidized to catechol followed by catabolism to TCA cycle
intermediates.

Most fungal metabolism mechanisms of PAHS are cometabolic, which means that
PAHS are not the primary substrate and that intermediate compounds are formed rather
than carbon dioxide and water (Wilson and Jones, 1993). The two main groups of fungi
involved in PAH degradation are those that use monooxygenases (e. g. cytochrome P-
450) and those that use lignin peroxidases to initiate attack on PAHS (Cerniglia, 1993).
Lignin peroxidases oxidize PAHS and initiate a free radical attack by a single electron
transfer forming quinones (Reddy, 1995). A third group of fungal enzymes involved in
the degradation of PAHS, called laccases, are considered to contribute less to the
degradation of PAHS because of their relaxed substrate specificity (Cerniglia, 1993;

Harayarna, 1997).

20

Bacterial biodegradation of PAHS is considered to be the dominant process for
PAH reduction in soils (Reilley et' al., 1996). Bacteria can degrade PAHS either as the
sole carbon source or by cometabolic processes (Wilson and Jones, 1993).
Bacterial degradation of PAHS is predominantly due to dioxygenase activity, which does
not lead to the formation of epoxides or mutagenic, carcinogenic, or teratogenic
intermediates (Sims and Overcash, 1983). This factor may favor soil bacteria over fungi
for bioremediation of PAHS.
Photodecompostion, oxidation, hydrolysis, leaching and volatilization of PA HS
Absorption of ultraviolet radiation leads to photolysis of PAHS (Sims and
Overcash, 1983). PAHS also react with ozone (Zeng etal., 2000), other oxidants, nitrogen
oxides, and sulfur oxides to cause decomposition (Sims and Overcash, 1983). The
formation of singlet oxygen and radicals generated by photolytic cleavage of trace

carbonyl compounds or from enzymatic reactions in aqueous systems, alkylperoxy

(R02 0) and hydroperoxy (H02 0 ), can result in the oxidation of PAHS (Sims and

Overcash, 1983). PAHS generally do not undergo hydrolysis. Photodecomposition,
oxidation and hydrolysis of PAHS are not considered significant pathways for PAH
reduction in soils (Sims and Overcash, 1983).

Leaching is not considered an important pathway leading to the reduction of
PAHS in soil due to their low aqueous solubility (Reilley et al., 1996). Leachate from pots
in a greenhouse experiment with contaminated soils was analyzed and did not contain
detectable PAHS (<10 g/L) (Schwab, 1994). Volatilization also is not considered an

important PAH reduction process from soils. PAHS with three or more rings have very

low vapor pressures (Reilley etal., 1996) and the high log KOW of most PAHS suggest

soil sorption of PAHS would be far greater than volatilization (Sims and Overcash, 1983)
(Table 1.1; Fig. 1.1). Park et al. (1990) observed that volatilization accounted for 30%
decrease of naphthalene 48h after PAH addition to soil, though did not significantly
reduce PAHS of higher molecular weight.
Plant uptake and accumulation of PAHS

PAHS detected in plants may be the result of atmospheric deposition, plant
biosynthesis, adsorption, or uptake. Atmospheric deposition of PAHS on plants could
account for background plant PAH levels (Lodovici et al., 1994). PAHS'have been
reported to form by plant biosynthetic processes (Borneff, 1968), though this
phenomenon has not been demonstrated by others (Sims and Overcash, 1983). Using

predictive mathematical models based on previous literature values for various plant

physiological and biochemical parameters, compounds with log KOW 0-1 may be taken up

by roots and translocated because they are water-soluble; compounds with log Kow

between 1 to 4 can be taken up by roots and transported in the xylem, and compounds

with log K0W greater than 4 would adsorb to roots (Trapp, 2002; Cunningham and Berti,
1993). The graphical relationship between translocation and log Kow is bell-shaped

indicating that there is an optimum lipophilic range of log KOW 1.5 to 2.5 for plant

translocation of organic compounds (Briggs et al., 1982).

Plants can accumulate PAHS via sorption onto plant roots, e. g. naphthalene
(Schwab er al., 1998; Schwab, 1994), or volatilization through the plant (Watkins, 1994).
The more lipophilic a compound, the greater the likelihood it is to concentrate in roots.

Mentha pulegium plants accumulate polymeric dyes, which are PAH analogs, into

Ix)
Ix)

lignifying tissues (Strycharz and Shetty, 2002). It was not determined if the aromatic
substrate was cross-linked to the cell wall or if it merely accumulated in plant tissue.

Some studies suggest that higher plants may not translocate PAHS. In one experiment

various plants grown hydroponically with 8.0 ug/kg benzo(a)pyrene (B(a)P) Showed no

(<3 ug/kg) B(a)P (log KOW = 6.04) was translocated in any of the plants or plant tissues

including leaf and stem tissues for green beans (Blum, 1977). In a three-year field study,

a mixture of 14 PAHS with log KOW ranging 3.37 to 7.66 did not accumulate to detection

limit levels in grass tissues (Qiu et al., 1997). Similarly, insignificant amounts of
anthracene and pyrene were taken up by alfalfa after 24 weeks in spiked field soil with
initial starting concentration of 100 mg/kg in a greenhouse study (Schwab, 1994).
Plant biostimulation of microbial PAH-degradation

Plants may Stimulate microbial degradation of PAHS and thereby enhance
biological remediation. Vegetation improves physical and chemical properties of soils,
promotes soil microbial activity, and increases contact between root-associated microbes
and soil contaminants. Roots benefit soil structure by enhancing soil porosity and
subsequent water and gas movement. Increased oxygen in soil may be important for
bioremediation of PAHS Since the initial step in the main PAH-degradation pathway
requires oxygen. Roots can grow into dense soil aggregates, and thereby increase the
volume of soil exploited by plants and microorganisms (Aprill and Sims, 1990). The
rhizosphere promotes microbial activity by enhancing transport of water, air and
providing carbon substrates via decaying organic matter and root exudation (Brady and
Weil, 1999). In addition to enhancement of water infiltration, plants can also remove

excess water, an important role for biodegradation of PAHS since aerobic conditions are

1x)
J J

required. Plants may provide carbon sources in the form of carbohydrates or organic
acids to increase microbial activity and or numbers. Plants may enhance the rate of
microbial PAH degradation by providing cometabolites, e. g. phenolics or terpenes, for
PAH—degrading microbes (Hegde and Fletcher, 1996). Plants can also secrete surfactants
(lipids and sterols) that lubricate the root, solubilize contaminants, and thereby increase
contaminant bioavailability (Siciliano and Gennida, 1998a). Root exudates have been
experimentally demonstrated to enhance bioremediation of PAHS. The addition of root
exudates of A vena barbata Pott ex Link (slender oat) resulted in lower phenanthrene
concentrations compared to unamended controls after 20 days in a growth chamber
experiment (Miya and Firestone, 2001). This decrease may have been because the plant
exudates provided nutrients, enhanced phenanthrene solubility (e. g. exudates were
biosurfactants), or served as primary substrates for cometabolic metabolism of
phenanthrene or served as cometabolites themselves. Root exudates increased
heterotrophic and microbial phenanthrene-degraders in the soil compared to soil amended
with only root debris, likely because root exudates provided higher carbon and nitrogen
than root debris (Miya and Firestone, 2001). Plants possess oxygenase, peroxidase, and
laccase enzymes which may allow phytodegradation of PAHS, though this has not been
clearly demonstrated (Criquet et al. , 2000). As a result, phytostimulation, rather than
phytodegradation alone, is likely the main phytoremediation mechanism for PAH
biodegradation.

2.4 Phytoremediation of PAHS in soil

Numerous studies have been performed to evaluate the effectiveness of plants for

PAH biodegradation. Planted treatments using F estuca arundinacea Schreb. (fescue),

Sorghum vulgare L. (sudan grass), and Panicum virgatum L. (switchgrass) resulted in
significantly lower soil concentrations of pyrene and anthracene and higher soil microbial
counts in a greenhouse study after 24 weeks compared to the unplanted treatment
(Schwab, 1994). In a follow-up to the preceding study, the same research group added
another species, Medicago sativa L. (alfalfa) to the previous three plant species mix
which was shown to reduce the soil PAH concentration more effectively than the
unvegetated treatments under greenhouse conditions(Reilley et al., 1996). This research
team conducted phytoremediation field trials for treatment of industrially contaminated
PAH soils and demonstrated that planted treatments Significantly enhanced dissipation of
the target pollutants (F iorenza et al. , 2000). The authors of this report suggested that the
successful planted trials may have been due to persistence of drought-tolerant species
during the drought conditions of the study period, rather than comparison of relative
abilities between the three Species tested. Another research team also demonstrated that
PAHS (benz(a)anthracene, benzo(a)pyrene, Chrysene and dibenz(ah)anthracene)
disappearance was greatly enhanced in vegetated soils (Aprill and Sims, 1990). A
phytoremediation study conducted on a crude oil Spill site showed that Lolium annual
(rye grass) and Stenotaphrum secundatum L. (St. Augustine grass) were superior to
Sorghum biocolor L. (sorghum) and an unvegetated control in reducing contaminant
concentration (Nedunuri et al., 2000). These studies support the use of phytoremediation
of PAH-contaminated soil as an effective environmental rehabilitation technology.
2.5 Potential Sources of Variation in PAH Phytoremediation

Plant species and soil amendments may affect the results of PAH

phytoremediation. When designing a phytoremediation strategy, a broad range of plant

species and soil amendments should be thoroughly researched for effectiveness prior to
full-scale implementation. Different plant species have varied toxicity tolerances to
PAHS. Not all plants can germinate and grow well in PAH-contaminated soils. Plant
Species were shown to differ in germination rates in PAH-contaminated soil derived from
petroleum in a screen of 22 plant species (grasses, herbs, legumes) (Adam and Duncan,
1999). Certain plants may possess physiological characteristics beneficial to
phytoremediation of PAHS. It has been proposed that flood-tolerant or wetland species
may enhance aerobic PAH microbial metabolism by more efficiently transporting air to
the rhizosphere and thereby improve the soil conditions for the initial oxidative step in
bacterial PAH metabolism (Shimp et al., 1993). Plants can provide varying amounts and
types of carbon sources that support microorganisms in general (Yoshitomi, 2001).
Different plant species produce different types and quantities of exudates, such as
cometabolites, that may potentially favor PAH-degrading microbes (Leigh et al. , 2002;
Miya and Firestone, 2001; Siciliano and Gennida, 1998b). In contrast, some root secreted
compounds may inhibit PAH-degrading bacteria by providing allelochemicals (Brady
and Weil, 1999; Leigh et al. , 2002). When evaluating phytoremediation as a potential
environmental rehabilitation strategy, it is essential to examine numerous plant species
due to the wide range of potential rhizosphere contributions and plant adaptations to
varied climate and edaphic conditions.

Different plant species have been demonstrated to possess varying abilities to
decrease PAH concentration in soils. Nine plant species were tested individually for
pyrene degradation on soil Spiked with 86.6 mg/kg pyrene. Plant species treatments had

pyrene concentration reductions of 55-74% in the soil compared to reductions of no more

26

than 40% from the implanted soil over a period of 8 weeks (Liste and Alexander, 2000).
Degradation rates of total petroleum hydrocarbon-contaminated soil by three different
plant Species treatments were compared and the leguminous plants resulted in the most
degradation (Yateem et al., 2000).

Since each contaminated site condition iS unique, it is essential to pre-test plant
and soil amendment combinations to achieve optimal phytoremediation. Soil additives
such as compost can have several beneficial effects on phytoremediation of contaminated
soil. Amendments can increase nutrient levels, organic matter content, cation exchange
capacity, and nutrient availability, and decrease bulk density and pH (Brady and Weil,
1999). Compost can increase the microbial activity and specifically the number of
pollutant degraders in soil. Addition of steer manure compost to contaminated soil was
shown to enhance degradation of 1,3-dichloropropene after 8 weeks of treatment
(Ibekwe, 2001). It has been demonstrated that the bacterial and fungal populations were
increased in numbers and diversity in pesticide-contaminated soil amended with yard
compost compared to unamended soil (Cole, 1994). Poultry litter and peat moss soil

amendments have been demonstrated to increase plant biomass and percentage

. . . . 137 90 .
accumulation of radioactive contaminants, Cs and Sr, compared to controls With no

amendments (Entry et al., 2001).
Addition of soil amendments, however, does not always lead to improved
remediation. Amending the soil with chemical cometabolites, inorganic/organic nutrients

and surfactants in an attempt to enhance bioremediation led to decreased mineralization

of 14C-phenanthrene and l4C-pyrene in 5 spiked soils (Carmichael, 1997). These

supplements increased the population of heterotrophic microorganisms. but not that of

PAH-degrading microorganisms (Carmichael, 1997). It was hypothesized that degrader-
microbes used the amendments as carbon sources rather than degrading the target
contaminant (Carmichael, 1997). Furthermore, organic contaminants can become less
available for microbial degradation if organic amendments are used because organic
contaminants sorb to organic matter. The use of amendments in environmental
rehabilitation Should be evaluated for effectiveness in greenhouse experiments and pilot-
field studies prior to large-scale application because amendments may not always
improve remediation.

SUMMARY & CONCLUSION

Persistent organic pollutants (POPS), such as polycyclic aromatic hydrocarbons
(PAHS) are hydrophobic chemicals that biomagnify through the food chain. As a result of
their low water solubility, PAHS sorb strongly to soil and sediment and become
unavailable for biodegradation.

Phytoremediation is a form of environmental remediation that uses plants to treat
organic and inorganic contaminants. Phytoremediation has several advantages over other
remediation strategies; it is minimally destructive, cost-effective, aesthetically pleasing,
and potentially self-sustaining. Soil PAHS are reduced predominantly by microbial
degradation, which has been observed to be enhanced in rhizosphere conditions. Abiotic
processes are considered relatively insignificant for natural reduction of soil PAHS.
Phytoremediation has been successfully demonstrated by numerous laboratory and field
studies as a promising technology for treatment of PAH-contaminated soils.

Selection of plant taxa and application of soil amendments Should be researched

carefully because not all treatments have been Shown to be beneficial for

28

phytoremediation. In this thesis study, a variety of native Michigan plant species were
evaluated for PAH phytoremediation in industrially-impacted soils collected from the
Rouge Manufacturing Complex. It is hoped that this study will enhance our knowledge of
PAH-phytoremediation processes and produce results that will be useful for large-scale

environmental rehabilitation efforts.

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CHAPTER II

FIELD STUDY OF MICHIGAN NATIVE PLANTS FOR PHYTOREMEDIATION

OF PAH-CONTAMINATED SOIL

40

ABSTRACT
Field Study of Michigan Native Plants for Phytoremediation of PAH-Contaminated Soil
Phytoremediation of polycyclic aromatic hydrocarbons (PAHS) was evaluated as

a strategy to rehabilitate soil contaminated from coking operations of the Ford Rouge
Manufacturing Complex (Dearborn, MI). A Phytoremediation Demonstration Site
consisting of 3 plots (wetland, upland, and control) was constructed in Sept 2000 at the
Allen Park Claymine (Allen Park, MI). The effects of 18 Michigan native plant species
on the concentrations of 14 PAHS in amended Rouge soil were evaluated in the upland
plot over the 2001 growing season. The soil total PAH (sum of 14 PAHS) concentration
(soil [tPAH]) decreased at a rate of 10 mg/kg per 2 months in the entire plot from May to
September 2001. The soil [tPAH] of the plot in May, July and September were
approximately 109 mg/kg, 100 mg/kg, and 91 mg/kg, respectively. Four of the 18 plant
Species significantly reduced soil [tPAH] in the upland plot from July to September,
unlike unplanted control cells which showed no significant reduction. Only one treatment
(Eupatorium purpureum) had significantly lower soil [tPAH] than the implanted control
at the end of the 2001 season. Two planted treatments (Andropogon scoparius and
Eupatorium perfoliatum) had significantly lower soil [tPAH] than the implanted
treatment in July, but this early-season effect did not persist for the September samples.
The soil [tPAH] results agree with those from preliminary greenhouse studies conducted
in collaboration with Applied Phytogenetics research labs. The study will run for 3 years.
during which time vegetated soils could continue or even accelerate PAH biodegradation.
Early observations from this field experiment indicate phytoremediation may be an

effective strategy for rehabilitation of PAH-contaminated soils.

41

INTRODUCTION

Former industrial sites are often impacted with hazardous materials, are poorly
vegetated, and possess infertile soils and sediments. If such land remains abandoned,
wind and water erosion can accelerate soil degradation to the point at which natural soil
rehabilitation is not possible. The Ford Rouge Manufacturing Complex (Dearborn, MI,
USA.) was once the world’s largest integrated industrial site and is a historical icon of
the American industrial age. Eighty years of automobile and steel manufacturing,
however, have led to contamination of the facility with byproducts of these activities.
Phytoremediation, the use of plants to degrade, detoxify, or remove environmental
. contaminants, is one method of environmental rehabilitation that is being considered as a
potential remedial treatment for PAH-contaminated soils in unused areas of the Complex.

The primary contaminants found at the Coke Oven area of the Rouge
Manufacturing Complex are polycyclic (polynuclear) aromatic hydrocarbons (PAHS or
PNAs), which were formed by the processing of coal during coke production for the
smelting of iron ores. PAHS consist of two or more fused benzene and furan rings
arranged linearly, angularly or in clusters (Fig. 1.1). PAHS are formed as a result of
incomplete, oxygen-deficient combustion of organic materials (Heil, 1998). Temperature
and combustion processes rather than fire] type determine which PAHS are formed
(Giger, 1974; Jenkins et al., 1996; Blumer, 1975). PAHS are derived fiom both natural
and anthropogenic sources (Hites, 1977). PAHS are formed naturally as a result of
thermal geologic reactions, forest and brush fires, and plant and bacterial reactions
(Blumer, 1976). Anthropogenic activities over the last century, such as the combustion of

fossil fuels, wood burning, industrial coke production, petroleiun refining, coal and oil

42

shale conversion and chemical manufacturing, have led to high levels of PAH discharges
disrupting the natural balance between PAH production and degradation (Hites, 1977;
Suess, 1976). PAHS are highly hydrophobic and have low solubility in water (Table 1.1)
causing them to be highly recalcitrant to decay in soils and sediments (Harayarna, 1997).
PAHS pose environmental hazards because they are carcinogenic, teratogenic, and
mutagenic (Shabad, 1975; Sims and Overcash, 1983; Dipple, 1990). It is therefore
necessary to remediate land contaminated with PAHS to prevent harmful exposure to
humans and wildlife.

Phytoremediation, vegetation-based environmental detoxification, could be an
effective method of reducing soil PAH concentration. Microbial bioremediation, the use
of fungi and bacteria to degrade contaminants, has been used to treat PAH-contaminated
soils. Plants have been shown to be beneficial for PAH-bioremediation of contaminated
soils. Soils planted with F estuca arundinacea Schreb. (fescue), Sorghum vulgare L.
(sudan grass), and Panicum virgatum L. (switchgrass) resulted in significantly lower soil
concentrations of pyrene and anthracene compared to unplanted soils after 24 weeks in a
greenhouse study (Schwab, 1994). Plants were also shown to accelerate the
disappearance of several high molecular weight PAHS compared to unvegetated soils
(Aprill and Sims, 1990). Vegetated soil had Significantly lower soil PAH concentrations
compared with the unvegetated control (Reilley et al., 1996). This study showed the
presence of plant roots and added organic substrates, such as organic acids, were essential
for sustained PAH degradation. This research suggests that effective PAH biodegradation

may depend on consistent supplies of root exudates and other readily metabolizable

43

carbon resources (Reilley et al., 1996). Collectively, these studies indicate that vegetated

soils may be superior to unplanted soils for treatment of PAH-contaminated soils.

Plants may assist soil bioremediation by several mechanisms, including induction
of microbial degradation gene activity and enhancement of microbial biomass and
community diversity. Several studies have demonstrated that plant-produced compounds
may stimulate microbial PAH degradation. Plants secrete carbon compounds such as
organic acids and carbohydrates for utilization by microorganisms (Yoshitomi, 2001).
Root exudates may promote microbial cometabolic degradation of organic pollutants
(Leigh et al., 2002). High molecular weight PAHS or PCBs may be degraded by
microorganisms, but cannot function as the sole carbon source. Some root secreted
compounds may serve as both metabolizable carbon sources and inducers for expression
of PAH-degradation pathway genes. Plants have been shown to release exudates similar
in structure to organic contaminants and lead to increased microbial activity and
contaminant degradation (Siciliano and Germida, 1998). Roots of Morus rubra (red
mulberry) to produce phenolic compounds which were demonstrated promote the growth
and activity of PCB degrading microbes (Hegde and Fletcher, 1996). PCB-degrading
bacteria, Alcaligenes eutrophus H850, Corynebacterium Sp. MBl, and Pseudomonas
putida LB400, were observed to be increased in numbers and degrading activity by plant
root compounds (Donnelly et al., 1994). Plant phenolic compounds may also
cometabolically induce PAH-degrading bacteria as well as PCB-degrading bacteria
(Donnelly et al., 1994; Hegde and Fletcher, 1996). In addition, plants possess the
enzymes oxidase, laccase, and peroxidase, which may contribute to PAH oxidation and

metabolism (Criquet et al., 2000). Plant root 2,7-diaminofluorene-peroxidases were

44

demonstrated to increase in abundance by the presence of anthracene and mycorrhizal
fungi (Criquet et al., 2000). These studies have shown that plants themselves may have
enzymes capable of PAH degradation and also produce root exudates that may act as
cometabolic carbon sources for induction of microbial PAH-degrading genes.
Rhizosphere soils have been Shown to possess higher microbial counts compared to

unplanted soils resulting in more effective PAH bioremediation. Vegetated soils were

observed to possess 1400 x 105 bacterial colony forming units (CFU) per gram of soil,

while soil in the absence of plants had 6 x 105 CFUs (Schwab, 1994). Similarly,

researchers showed that vegetated contaminated soil had 4-8 log CFU/ g compared with
non-vegetated contaminated soils, which had a log CFU/ g soil range from 3-6, a ten- to
hundred-fold increase (Y ateem et al., 2000). For both studies, the observed increase in.
bacterial numbers for vegetated soils was proposed to be responsible for enhanced PAH
degradation.

PAH phytoremediation is not currently a well-developed environmental
rehabilitation technology. A limited number of plants species have been demonstrated
and reported to be effective for PAH degradation in field conditions. Different plant taxa
are known to possess widely different capabilities for PAH phytoremediation with little
understanding of the range of potential mechanisms. Previous PAH phytoremediation
studies have made either very general, empirical observations or focused on limited
numbers of plant species. Most research publications report PAH phytoremediation
experiments using only a Single species (Qiu et al., 1997), few individual species
(Nedunuri et al., 2000; Pradhan et al., 1998; Reilley et al., 1996), or a mixture of small

numbers of species (Aprill and Sims, 1990; Fiorenza et al., 2000).

45

Phytoremediation remains an attractive method of soil rehabilitation, though few
studies have tested a comprehensive range of different plant species under field
conditions. The project reported in this thesis attempts to use a broad spectrum of
Michigan native plants to evaluate each species’ relative effectiveness for soil PAH
phytoremediation. Eighteen selected plant species were evaluated for phytoremediation
of 14 PAH compounds in a pilot-scale field study. It is expected that this experiment will
provide additional resources to enhance our understanding of PAH phytoremediation and

its utility for rehabilitation of PAH-contaminated soils.

MATERIALS AND METHODS

Plant species selection

A preliminary 14-week greenhouse study was conducted to identify plant taxa
with potential to reduce PAHS. The experiment tested ~40 native Michigan native plant
species for soil PAH reduction in collaboration with the Applied PhytoGenetics
laboratory (APGEN; Athens, GA). Soil was obtained from the Rouge Coke Oven area,
amended with 30 % perlite (v/v), placed in 400 cc plastic pots and planted with plant
plugs. Planted and implanted control pots were watered and fertilized weekly with 6% of
a stock solution of Peter’s N-P-K (20-20-20). Untreated control pots were not amended
and were maintained in the greenhouse without fertilization or watering. Native Michigan
plants were obtained from Wildtype Native Plant Nursery (Mason, MI). Soil and plant
tissue sampling was performed after treatment periods of 4, 6, 10 and 14 weeks. Single,
whole-pot samples were analyzed at each interval for each species.

Twenty species were selected from preliminary data after 6 weeks of the APGEN

study (Table 2.1). For ease of presentation and discussion, 6 letter abbreviations were

46

assigned for plant species based on the first 3 letters of the both the first and second

names of the plant scientific name (Table 2.1).

Table 2.1. Species planted in Phytoremediation Demonstration Facility. For Plot: W =

Wetland, U = Upland, C = Control.

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

Scientific Name Common Name Abbreviations Plot
Amorpha canescens Leadplant AMOCAN U, C
Andropogon gerardii Big Bluestem ANDGER U, C
Andropogon scoparius Little Bluestem ANDSCO U, C
Aster novae—angliae New England Aster ASTNOV W, U, C
Carex sprengelii Sprengel Sedge CARSPR W, U, C
Ceanothus americanus New Jersey Tea CEAAME U, C
Cirsium discolor Pasture Thistle CIRDIS U, C
Eupatorium perfoliatum Boneset EUPPER W, U, C
Eupatorium purpureum J oe-Pye Weed EUPPUR U, C
Geum triflorum Prairie Smoke GEUTRI U, C
Hystrix patula Bottlebrush Grass HYSPAT U, C
Lobelia cardinalis Cardinal Flower LOBCAR U, C
Mimulus ringens Monkey-Flower MIMRIN W, C
Physocarpus opulifolius Common Ninebark PHYOPU U, C
Scirpus atrovirens Bulrush SCIATR W, U, C
Silphium teribinthinaceum Prairie-dock SILTER U, C
Solidago patula Swamp goldenrod SOLPAT W, C
Spartina pectinata Prairie Cordgrass SPAPEC W, U, C
Spirea alba Meadowsweet SPIALB U, C
Viburnum dentatum Arrowwood Viburnum VIBDEN U, C

 

 

 

 

 

Field Trial of Selected Native Michigan Plant Species

The Rouge Manufacturing Complex is still in operation for steel production and is
not suitable for field experimentation. Therefore, a Phytoremediation Demonstration
(Phyto Demo) Site was constructed at the Allen Park Claymine (Allen Park, MI) in
conjunction with URS Corporation under the direction of Michael Coia, Project Engineer
(Willow Grove, PA) in the summer of 2000 (Fig. 2.1). Pits were excavated for the

construction and installation of the treatment plots measuring 20’ x 50’ each for the

47

 

 

Walkway

 

OO
01:]

Leachate
Reservoir

   

Wetland Upland Control

Figure 2.1. Overhead view of Phytoremediation Demonstration Site. Distances not drawn
to scale.

48

Discharge sump

—h/

 

 

Geotech

24in. “Her

 

 

 

 

 

 

 

 

 

20 feet

Figure 2.2. Phyto Demo Treatment Plot cross-section schematic. I-IDPE refers to high
density polyethylene.

49

wetland and control plots and 20’ x 100’ for the upland plot. The upland plot was twice
as large as the wetland because most of the terrain at the Rouge Manufacturing Complex
is upland and a potential remediation installation at this site would be for this habitat.
Each plot was bordered by ~1’ berm of uncontaminated soil and overlaid with 60 mil
high-density polyethylene (HDPE) liner to prevent escape of contaminated soil or
leachates from the treatment plots. Approximately 4” of pea gravel was laid on the
HDPE-lined plot in the graded slope at the bottom of each plot (3% incline from long
axis side to center) to facilitate drainage and overlaid with geotech fabric to contain the
soils (Fig. 2.2). Sump tanks and float-ball actuated pumps were installed to remove
excess leachate to nearby holding tanks for testing and eventual discharge into a sanitary
sewer.

PAH-contaminated soil was obtained from the area adjacent to the Coke Oven
system of the Ford Rouge Manufacturing Complex (Dearborn, MI). Uncontaminated
control soil was topsoil from the Claymine landfill. PAH-contaminated and
uncontaminated control soils were amended with (5% v/v) poultry manure (Herbruck’s
Poultry Ranch, Saranac, MI) and (10% v/v) yard compost (Charter Township of
Ypsilanti, MI). All soil and compost amendment components for the Phyto Demo site
were sieved then mixed using a mechanized soil shaker-screen (2” debris exclusion) and
a front-end loader. Compost-arnended, Coke Oven soil was placed in the wetland and
upland plots and amended, uncontaminated soil was placed in the control plot. Compost-
amended soil from the Phyto Demo site and native Coke Oven soil were tested for
agronomic and physical properties (Tables 2.2 & 2.3) by MDS Harris Laboratories

(Lincoln, Nebraska). Both the native Coke Oven soil and the Phyto Demo soil were

50

Table 2.2. Properties [mean, standard deviation (SD)] of Rouge Manufacturing Complex

Coke Oven soil (unamended—no yard compost or poultry manure) and Phytoremediation

Demonstration site soil [amended with 10% yard compost and 5% poultry manure (v/v)],

N = 2. Amended soils were sieved (4.75 mm). NA means standard deviation was not

available because N = 1. Soil pH and soluble salt determined using 1:1 water to soil ratio.

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

CEC

% % Bulk Soluble salts Sodium % (meq /

Sand Silt % Clay density pH (mmhos/cm) (mg/kg) OM 100g)
Unamended Rouge soil

Mean 72 24 4 1.1 8.3 0.3 140 7.85 17.05

SD 2.8 2.8 0 0.07 O 0.02 5.67 0.07 0.92

Amended Rouge soil
Mean 70 24 6 1.2 8.1 1.2 124 2.8 18.7
SD NA NA NA 0 0 0.14 2.83 0.28 1.56

 

51

 

Table 2.3. Concentrations of nutrients [mean, standard deviation (SD)] of soil Rouge
Manufacturing Complex Coke Oven soil (unamended—no yard compost or poultry
manure) and Phytoremediation Demonstration site soil [amended with 10% yard compost
and 5% poultry manure (v/v)], N = 2. Amended soils were sieved (4.75 mm). Note, N is
nitrate determined by cadmium reduction method, P is determined by Bray I or Olsen
method, cations (Ca, Mg, K, Na) extracted by modified ammonium acetate method, trace
elements (Zn, Mn, Cu, Fe) extracted by modified diethylenetiiaminepentaacetic acid
(DTPA) method;gorganic matter determined by loss-on-ignition, sulfur (S) and boron (B)

were determined by inductively coupled plasma (ICP) emission spectroscopy.

 

Nutrient (mg/kg)

 

N P K Mg Ca S Zn

 

 

 

 

 

 

 

 

 

 

 

 

Unamended Rouge soil

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

mean 2.5 3.5 92 240.5 2877 29.5 16.6 22.2 5 27.7 1.1
SD 0.7 0.7 9.9 33.2 79.2 3.5 2.0 0.9 0.9 5.0 0.1

Amended Rouge soil

mean 72 82 570 348.5 2756 89.5 17.35 18.5 3.9 50.4 2.0
SD 5.7 0 59.4 38. 9 213.6 7. 8 2.1 2.3 0.4 5.9 0.2

 

 

52

 

classified as sandy loam (MDS Harris Laboratories, Lincoln, NE). Amended soils were
placed into each of the plots at a loose-filled depth of 24” (Fig. 2.2). The plots were
overlaid with wood planks on the soil surface to demarcate individual cells (~3.5’ x 4’ =
~20 sq. ft). The wetland and control plots consisted of 30 cells and the upland plot
consisted of 60 cells (Fig. 2.3). Individual cells were assigned plant treatments designated
alphanumerically as shown (Fig. 2.3).

Plants were planted at the Rouge Phyto Demo site in September 2000. Plug- or
quart-sized plants were taken out of pots, roots cleaned of potting soil, and transplanted
into the individual cells on ~9” centers (12 plants per cell). Plants were planted in 3 cells
per species in either or both upland and wetland plots according to their ecological habitat
range (Table 2.1). The control plot had one cell per plant species and was used to assess
herbivory or pathogen problems and background soil PAH concentrations. The Phyto
Demo field plots were fertilized with an N-P-K solution (20-20-20) at a concentration of
~475 ppm weekly. A substantial number of plants (about 50%) were replaced in May
2001 due to overwinter mortality.

Sample Handling

For the summer 2001 growing season, soil samples were collected three separate
times at 9-week intervals in mid May, July and September. At each sampling time, three
2” diameter-cores of 8” depth were removed from each cell. Each core sample excluded
the first 2 inches of the surface to avoid soil PAH content altered by photolysis or
evaporation. Rocks were separated from core samples in plastic tubs and the soil placed
in 150 mL amber jars with teflon caps. The amber jars containing the soil samples were

transported from the field in coolers containing ice-packs and stored in the 4°C walk-in

53

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54

refi'igerator at the MSU Phyto laboratory. Soil samples were sieved using a stainless steel
8-inch diameter 2.36 mm sieve (Gilson Co.) to remove rocks, mulch or other debris
before analysis. May soil samples were sieved immediately before each samples were
extracted. July and September soil samples were sieved approximately one week after
sampling and stored at 4°C until extraction. Plant leaf samples were obtained from 3
plants from each cell in July. Fresh soil samples, ~5.0 :1: 0.1 g, were placed in a drying
oven for 48 hours at 105 °C for soil moisture and dry weight determination. Plant dry
weight was determined after drying plant tissue samples in paper bags at 80 °C for 48
hours.
Extraction Protocol

Plant and soils were analyzed for PAH content by dichloromethane extraction.
Plant tissue extractions were performed on ~1.5 i 0.1 g fresh weight unless there was
insufficient tissue, in which case lesser amounts were used for PAH extraction and dry
weight determination. Soil cores were analyzed for PAH concentration by
dichloromethane extraction of 3.0 i 0.1 g F W subsamples (1 subsarnple for May, 3
subsamples for July and Sept). Plant and soil PAHS were analyzed by phase extraction in
3 mL saturated potassium chloride solution and 10 mL of the organic extraction solvent,
dichloromethane, in a 20 mL amber vials. The extract mixtures were vortexed for 20
seconds, sonicated for 10 minutes, and placed on a rotating shaker (~125 rpm) overnight.
Sample extracts were filtered using 3 mL polypropylene sterile disposable (B&D, Fisher
Scientific) syringes and 13 mm 0.45 pm PTFE teflon syringe filters (SGE, DC Scientific)

during transfer to 2 mL gas chromatography (GC) vials. Extraction vials were re-used

55

after being washed in soapy water, rinsed sequentially with acetone and pure water, and
oven-dried.

The May soil samples and a portion of the July soil samples were shaken with the
vials in the upright position. A test was performed to compare recovery rates when vials
were placed in the upright versus the horizontal position. Results from this test indicated
samples from vials placed horizontally on the shaker had 30-70% higher concentrations
than samples from vials placed in an upright position (data not shown), therefore
invalidating the results from the upright extracted samples. Compromised samples were
re-extracted with the vials placed horizontally on the shaker using remaining soil from
the jars stored at 4°C. For May samples, only one subsample from each jar was taken for
[tPAH] determination due to the insufficient volume of the remaining soil for 3
subsamples.

Gas Chromatography-F lame Ionization Detector (GC—FID) Analyses

PAH analyses were performed on an Agilent 6890 Gas Chromatograph equipped
with an Agilent 3396B/C Integrator and Agilent 7683ALS auto injector. An Alltech AT-5
capillary column with inside diameter of 0.53 mm, purchased length 30.0 rn and film
thickness 1.20 pm (Alltech: Deerfield, IL) was used for PAH compound separation. The
column was cut to minimum length 15.0 m to exclude residue build up at the front of the

column as it aged. The carrier gas was helium delivered at a rate of 5.4 mL/minute and

fuel source for the FID was H2 delivered at 40.0 mL/minute. The make-up gas consisted

of N2 (flow rate of 45 mL/minute) and 0.1 grade air (flow rate of 450 mL/minute). The

capillary column oven was set with an initial isothermal period of 100° C for 1 minute

followed by elevation at 100 °C/minute until 3100 C was reached. The volume of the

56

injected sample was 5 11L and the temperatures of the injection port and the detector were
270° C and 330° C, respectively.

The standards were made from EPA 610 PAH standard mix (Supelco, Bellefonte,
PA), which includes the PAH compounds in Table 2.4. Calibration curves consisted of 3
to 8 points using 1% to 50% dilutions made of EPA 610 mix stock reagent. We assumed
a standard curve though the origin. The determined concentrations of the first four
compounds that eluted, naphthalene, acenaphthylene, acenaphthene, fluorene, were not
reliable because the concentrations found in the samples were often lower than the lowest
standard read by the GC-F ID. However, these concentrations are included in the
determination of total PAH concentration. Vials containing only 3mL KCl and 10mL
dichloromethane or 10mL dichloromethane alone were analyzed as “blank” controls.
Calculations

1. GC-FID output (20.2 ug/mL) sample extract concentration (mg/kg = ug/ml) corrected
by extraction volume and dilutions:

e.g. 20.2 ug/mL X 10 ml = 202 ug in that sample

2. ug tPAI-I/g FW X g FW/g DW = ug tPAH/g DW tissue or soil

e.g. 202 ug PAH/g FW X 1.5 g FW/0.3 g DW = 1010 mg/kg
(or 1010 ug/g DW)

The total PAH concentration [tPAH] was determined by taking the sum of the calculated
concentrations of the compounds listed in Table 2.4. For ease of presentation and
discussion, the full names of the PAH compounds analyzed were abbreviated (Table 2.4).
Lowest calibration standard (1%) of EPA 610 standard mix for these compounds are

displayed in (Table 2.4).

57

Table 2.4. Abbreviations and lowest calibration standard in 2001 for PAH compounds

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

analyzed.

Compound Abbreviation Lowest standard (pg/mL)
Naphthalene Naph 1 0
Acenaphthylene Acny 20
Acenaphthene Acne 1 0
Fluorene Flre 2
Phenanthrene Phen 1
Anthracene Anth 1
Fluoranthene Flra 2
Pyrene Pyre 1
Benz(a)anthracene Baan 1
Chrysene Chry 1
Benzo(b)fluoranthene Bbfl 2
Benzo(a)pyrene Bapy 1
Dibenz(a,h)anthracene Daha 2
Benzo(ghi)perylene B ghp 2
Sum of concentrations for the
above 14 PAH compounds [tPAH] 56

 

 

 

58

 

Statistical Analyses

The upland soil [tPAH] data were tested for normality by analyzing stem-leaf
plots, normal probability plots and residual plots. The upland May soil total PAH
concentration ([tPAH]) data were analyzed using ANOVA and Students’ t-test for
significant differences between each pair of treatment means. The ANOVA test requires
an equal number of subsamples per core. For May there was only one subsample per core
and for July and September there were 3 subsamples per core. In order to compare May
data to July and September data, two subsamples were arbitrarily excluded fi'om the
upland July and September soil [tPAH] data, and a two-way AN OVA statistical test was
performed using May, July and September soil [tPAH] data. July and September soil
[tPAH] data were analyzed using two-way ANOVA with all 3 subsamples. The Student’s
t-test was used to test for significant differences between pairs of treatment means at each
sampling time and for significant differences between treatment means at different
sampling times. A less conservative statistically significant level, a = 0.1, as opposed to
the conventional level a=0.05, was used to enhance the ability to detect differences
between treatments and between sampling times. All statistical analyses were done using
SAS version 8.01.

RESULTS

Plant species selection

The APGEN greenhouse screen of 40 species showed general trends in soil total
PAH concentration ([tPAH]) reduction for most treatments (Appendix 1). Eighteen plant

species treatments out of 36 achieved greater reduction in soil [tPAH] than the unplanted

59

pots. This is represented by calculation of treatment index, i.e. average planted soil
[tPAH] divided by unplanted soil [tPAH] (Appendix 2).
Plant Mortality
Substantial over-wintering plant mortality was observed in May 2001 at the

Phyto Demo site after the initial planting in September 2000 (Appendix 3). A. scoparius.
A. novae-angliae, C. americanus, C. discolor, P. opulifolius, S. teribinthinaceum, and S.
patula had mortality rates greater than 50% after over-wintering for the first year
(Appendix 3). Plant mortality was greatly reduced in ensuing seasons (Appendix 3).
Wetland

Wetland plot mean (including all planted and implanted control treatments) soil
[tPAH] appeared to fluctuate over time. The mean total PAH concentration [tPAH] and
standard error of the wetland plot for May, July, and September were: 98.9 a: 4.4 mg/kg,
122.1 d: 5.3 mg/kg, and 76.1 d: 2.3 mg/kg. Standard errors represent variability among all
cells for the plot.
Upland

The upland plot (including all planted and implanted treatments) soil [tPAH]
means i standard error of mean (SEM) for May, July and September were 108.80 :t
3.2mg/kg, 100.3 i 3.2 mg/kg, and 90.9 i 3.2 mg/kg, respectively. Soil [tPAH] was
observed to decrease over time for most upland treatments (Table 2.5). The overall
decrease in soil [tPAH] of the upland plot is illustrated by color assignment to average
cell soil [tPAH] ranges (Fig. 2.4 a — c). Cell color codes are easily observed to shift from
an abundance of red-orange-yellow cells (i.e. high [tPAH]) to an increase in blue-green

cells (i.e. low [tPAH]) from May to September. Most cells were observed to decrease in

60

Table 2.5. Soil total PAH concentration [tPAH] (mean i SEMY) for treatments in the

upland plot from sampling times in May, July, and September 2001, N = 3 cells for each

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

treatment.

Soil Concentrations (mg/kg)

Treatment May July September
Amorpha canescens 101.3 i 15.0 90.4 :1: 0.5 81.9 3: 6.3
Andropogon gerardii 106.7 i 19.7 106.4 i 4.8 84.4 :t 5.31"
Andropogon scoparius 105.8 i 7.4 79.2 :t 8.9* 80.3 :t 10.9
Aster novae-angliae 117.9 i 16.6 104.021: 4.2 84.1 i 1.0T
Carex sprengelii 106.1 :1: 4.2 92.5 :t 14.9 91.8 :1: 2.6
Ceanothus americanus 106.5 i 4.5 103.9 :1: 12.7 96.6 i 4.3
Cirsium discolor 108.5 :t 6.4 92.3 :1: 6.9 87.5 :t 5.8
Eupatorium perfoliatum 97.1 i 7.6 79.1 i 4.3* 84.3 i 8.4
Eupatorium puipureum 110.8 i 13.5 91.9 d: 7.5 72.1 :t 10.5*i'
Geum triflorum 111.5 :1: 15.2 93.9 d: 6.7 114.3 d: 2.3*‘l'
Hystrixpatula 101.1 :t 12.7 105.3 :1.- 8.7 105.4 :t 8.4
Lobelia cardinalis 109.5 :t 14.4 107.8 :1: 7.4 93.2 :t 4.7
Physocarpus opulifolius 1183 i 6.9 84.5 d: 5.8 101.0 i 8.7
Scitpus atrovirens 124.7 :t 14.1 95.7 d: 7.9 104.8 :1: 19.4
Silphium teribinthinaceum 110.9 :1: 20.0 118.6 i 15.7 92.9 :1: 3.7T
Spartina pectinata 100.2 :t 4.5 90.6 i 5.4 82.7 i 3.4
Spirea alba 127.3 :1: 25.0 85.9 i 2.3 95.65 3: 4.9
Viburnum dentatum 90.2 i 6.8 113.4 :t 6.1 107.8 :E 14.4
Unplanted 112.9 :1: 7.3 100.8 i 5.1 93.7 i 5.4
SEM (ANOVA mixed

13.07 8.09 8.09
model)

 

 

 

 

 

7 Standard errors represent variability between the averages of soil [tPAH] from each cell

for a given treatment.

6 Standard error of mean based on ANOVA mixed model.

* Significantly different from unplanted at that sampling time (t -test, a = 0.1).

T Significant difference between July and September means (t-test, a = 0.1).

61

 

Table 2.6. Statistical results from two-way ANOVA analyses for the upland plot.

 

 

 

 

Source Df MS F P-value
May, July, Sept (1 subsample per core)

Treatment" 18 1153.8 0.7 0.78
Error (MST) 28.9 1644.5

Time 2 13677.3 8.0 <0.01
Treatment x Time 38 1123.5 0.7 0.89
July & Sept (3 subsamples per core)

Treatment 18 472.2 2.0 0.04
Error (MST) 38 242.1

Time 1 526.7 3.5 0.07
Treatment x Time 18 248.0 1.6 0.10
Error (MSE) 38 150.9

 

*Note: treatment includes plant species’ and implanted treatments.

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64

soil [tPAH] by 0-20 % (Fig. 2.5 a). For comparison of planted treatments to color codes,
treatment ID codes are included in the adjacent figure (Fig. 2.5 b).

The analysis using one subsample showed the effect of sampling time on soil
[tPAH] was significant (Table 2.6), the plant species’ treatment effect (including
unplanted and untreated treatments) was not significant, and the treatment x time
interaction effect was not significant (Table 2.6). The one subsample analyses revealed
that there were significant differences in mean soil [tPAH] between the May and July (t-
test, P = 0.064) sampling times, July and September (t-test, P = 0.045) sampling times,
and May and September (t-test, P < 0.001) sampling times.

Diflerences in soil [tPAH] over time for Upland Plot

For July and September sampling times the treatment effects were significant, the
time effect was significant, and the treatment x time interaction effect was significant
(Table 2.6). F our plant species (A. gerardii, A. novae-angliae, E. purpureum, S.
teribinthinaceum) treatments showed significant decreases in soil [tPAH] from July to
September (3 subsample analysis) (Table 2.5). Soil [tPAH] increased significantly in the
G. triflorum treatment. The most effective phytoremediation treatment, E. purpureum,
decreased soil [tPAH] 91.9 i 7.5 to 72.1 d: 10.5 fi'om July to September. By contrast, the
soil [tPAH] in unplanted cells showed no significant difference between July and
September samples (Table 2.5). Significant differences for specific treatments between
May and July and between May and September could not be determined using 3

subsamples per core because May only had one subsample per core taken.

65

Differences in soil [tPAH] between treatments

There were no statistical differences among the 19 treatments (plant species and
unplanted) for the May sampling time (P = 0.95; Table 2.5). Treatments A. scoparius and
E. perfolz’atum had soil [tPAH] significantly lower than the unplanted soil [tPAH] in July
(Table 2.5). The September mean soil [tPAH] in cells planted with E. purpureum was
significantly lower than that in the unplanted treatment (Table 2.5). By contrast, cells
planted with G. triflorum had a significantly greater mean soil [tPAH] than the unplanted
at the September sampling time (Table 2.5).

To compare APGEN data and Phyto Demo data, soil indices were calculated by
dividing the soil [tPAH] for a planted treatment by the soil [tPAH] in the unplanted
treatment for each sampling time, i.e. Planted soil [tPAH]/ Unplanted soil [tPAH]. An
index of less than 1 indicates the planted treatment has reduced the soil [tPAH] to a
greater extent than the unplanted treatment. An average soil index for each of the
sampling times (APGEN, 4 sample times; Phyto Demo, 3 sample times) was calculated
for each species common to both studies. In the Phyto Demo study, ten of the 18 plant
species treatments had average soil indices less than one (Table 2.7). In addition, seven
plant species treatments had soil indices less than one for all three sampling times. In
both the APGEN and Phyto Demo studies E. perfoliatum, A. scoparius, A. canescens, C.
discolor, P. opulifolius and A. novae—angliae treatments had average soil indices less than
one (Table 2.7 and Appendix 2). Plant species treatments S. alba, H. patula, and S.
atrovirens had average soil indices greater than one in both studies. Eleven out of the 17
plant treatments (71%) that were common to both studies have similar soil index values

relative to the unplanted.

66

Table 2.7. Upland Phytoremediation Demonstration site 2001 soil indices ranked by

"Avg" index. "Avg" is the average of May, July, and September soil indices.

 

[PIanted]/[Unplanted] Ratios

 

Each" indicates
ratio < 1 at one

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

Treatment May July Sept Avg sampling time
E. perfoliatum 0.86 0.78 0.90 0.85 ***
A. scoparius 0.94 0.79 0.86 0.86 ***
E. purpureum 0.98 0.91 0.77 0.89 ***
M. canescens 0.90 0.90 0.87 0.89 ***
S. pectinata 0.89 0.90 0.88 0.89 ***
C. discolor 0.96 0.92 0.93 0.94 ***
C. sprengelii 0.94 0.92 0.98 0.95 ***
A. gerardii 0.94 1.06 0.90 0.97 **
P. opulifolius 1.05 0.84 1.08 0.99 *
A. novae-angliae 1.04 1.03 0.90 0.99 *
nplanted 1.00 1.00 1.00 1.00

S. alba 1.13 0.85 1.02 1.00 *
C. americanus 0.94 1.03 1.03 1.00 *
L. cardinalis 0.97 1.07 0.99 1.01 **
H. patula 0.89 1.04 1.12 1.02 *
V. dentatum 0.80 1.12 1.15 1.02 *
G. triflorum 0.99 0.93 1.22 1.05 **
S. teribinthinaceum 0.98 1.18 0.99 1.05 **
S. atrovirens 1.10 0.95 1.12 1.06 *

 

67

 

 

Soil [tPAH] percentage reduction

Planted treatments reduced soil [tPAH] 0—35 % from May to September 2001,
while the unplanted control reduced soil [tPAH] by 17 % (Fig. 2.6 a). Treatments had
different effects on a given PAH contaminant. For example, E. purpureum substantially
decreased naphthalene, while the soil naphthalene concentration in the unplanted control
did not change. The planted treatments that led to the greatest reduction in soil [tPAH]
were E. pwpureum, A. novae-angliae, S. alba, A. scoparius, and A. gerardii and these
planted treatments had percentage reductions from 20—35 % from May to September
2001.
Individual PAH compounds percentage reduction

The percentage reductions of 11 individual PAH compounds and total PAH
concentrations were not uniform among all treatments (Fig. 2.6 b-l). When each of the
treatments are presented along the x-axes of graphs for individual PAH compounds in the
ranked order of % [tPAH] reduction, apparent differences for % reduction for each given
individual PAH compound are observed between the planted treatments (Fig. 2.6 b-l). In
general, higher molecular weight PAHS (Chrysene, benzo(b)fluoranthene,
benzo(a)pyrene, dibenzo(ah)anthracene, and benzo(ghi)perylene) seemed to be reduced

to a lesser extent (Fig. 2.6 g-l) than lower molecular weight PAHS (Fig. 2.6 b-f).

68

Figure 2.6 (a-l). Percentage reduction (mean 5: SEM) in soil [tPAH] and individual PAH
compounds for all treatments (May — September 2001) in Phyto Demo upland plot. Note
naphthalene data may not be accurate because the concentrations in the samples were
occasionally below the lowest standard of the calibration curve. Data for acenaphthene,
acenaphthylene, and fluorene are not presented because the concentrations of samples
were below the lowest calibration curve standard. The treatments are presented in order

of decreasing % reduction in soil [tPAH] along the horizontal axes.

69

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Plant PAH concentration

Plant leaf tissue tPAH concentrations were determined to be unusually high and
variable, ranging in mean i SEM from 260.3 2t 33.0 mg/kg for C. americanus to 7979.5 3:
2787.9 mg/kg for E. purpureum in the upland plot (Table 2.8). For the wetland plot,
plant leaves had [tPAH] concentrations from 260.6 d: 30.0 mg/kg for C. sprengelii to
5292.9 :1: 1771.4 mg/kg for S. patula (Table 2.8). Plant leaf tissue in the control plot had
a concentration range from 72.6 mg/kg for G. triflorum to 9191.7 mg/kg for S. patula
(Table 2.9). Since there was only one cell per plant species treatment, there were no
standard errors of means in the control plot. The most abundant PAH compounds
detected in plants and their respective range of concentration values were acenaphthylene
(30-12000 mg/kg), benzo(a)anthracene (30-1000 mg/kg), Chrysene (20- 7000 mg/kg),
benzo(a)pyrene (30-4000 mg/kg), and benzo(ghi)perylene (40- 960 mg/kg). Individual

PAH compound concentrations for plants are presented in Appendix 4.

76

Table 2.8. Plant total PAH concentrations in plant leaves (mean :t SEM) in mg/kg dry
weight. Leaf tissues were collected from three plants per cell and pooled for [tPAH]
analysis. Each mean represents the average concentration for three cells per plant
treatment, with the exception of C. americanus treatment with only two cells sampled.

Not all Species in the Phyto Demo study were sampled due to low available biomass for

 

 

some plants.
Treatment [tPAH] (mg/kg)
Upland
Amorpha canescens 1229.9 :1: 70.6
Andropogon gerardii 834.6 :1: 86.7
Andropogon scoparius 548.2 i 123.4
Aster novae-angliae 901.0 i 311.2
Carex sprengelii 310.2 d: 24.8
Ceanothus americanus 260.3 :1: 33.0
Cirsium discolor 2649.4 :1: 2294.2
Eupatorium perfoliatum 2111.0 :1: 198.5
Eupatorium purpureum 7979.5 :1: 2787.9
Geum triflorum 132.5 :t 19.9
Hystrixpatula 1011.5 :t 119.0
Lobelia cardinalis 2579.2 i 1120.5
Physocarpus opulifolius 549.6 i 224.0
Scimus atrovirens 304.2 i 15.5
Silphium teribt'nthinaceum 1508.3 :1: 681.5
Spartina pectinata 797.7 :t 38.9
Spirea alba 1139.7 i 336.2
Viburnum dentatum 723.5 i 172.9
Wetland
Aster novae-angliae 775.6 i 112.8
Carex sprengelii 260.6 :t 30.0
Eupatorium perfoliatum 1376.3 :t 156.5
Mimulus ringens“ 452.0 i 62.4
Scirpus atrovirens 274.6 i 37.4
Solidago patula* 5292.9 :t 1771.4
Spartina pectinata 496.8 i 78.8

 

Note: * means species was not in upland plot.

77

Table 2.9. Plant total PAH concentrations (mg/kg dry weight) in plant leaves from the
control plot (uncontaminated soil). Samples were taken from three plants in each cell and
pooled. The control plot contained only 1 cell per plant treatment. Not all species were

sampled because of limited plant growth for some plants.

 

 

Cell Treatment [tPAH]

(mg/kg)
CO6C Amorpha canescens 1256.8
C09A Andropogon gerardii 638.0
C 10A Andropogon scoparius 803.5
CO3A Aster novae-anglz'ae 1722.9
C 1 OB Carex sprengelii 340.2
C04B E upatorium perfoliatum 2 125.9
C1 0C E upatorz'um purpureum 8563 .9
COBB Geum trzflorum 72.6
COZC Hystrix patula 779.5
C03C Lobelia cardinalis 4933.1
CO7C Mimulus ringens 490.7
C07B Physocarpus opulifolius 353.1
COSC Scimus atrovirens 1 5 1 .9
C09B Silphium teribinthinaceum 1283.1
COIC Solidago patula 9191.7
C09C Spartina pectinata 792.0
C08C Spirea alba 1429.9
C04C Viburnum dentatum 51 8.9

 

78

DISCUSSION

Plant species selection

The APGEN greenhouse study results (Appendix 1) indicate varying abilities
among the tested plants for soil [tPAH] reduction. Anomalous points in the APGEN
greenhouse study, such as for Elymus virginicus, week 6, may be the result of a small
quantity of highly PAH-contaminated material in that particular sample. The variability
in PAH concentration in the soil samples may be reflective of the heterogeneous
distribution of organic matter since PAHS sorb strongly to organic matter. The goal of
the APGEN screening experiment was to achieve rapid identification of candidate species
for more extensive analysis under both field and greenhouse conditions. Consideration of
both apparent reductions and consistent trends for soil [tPAH] reduction over time
allowed selection of suitable species for future research.
Wetland Data

The wetland plot soil data is of concern due to extreme and unlikely fluctuations
over the course of the season. The wetland soil [tPAH] values were perceived as initially
low (May = 98.9 i 4.4 mg/kg), then much higher (July = 122.1 i 5 .3 mg/kg), and then
greatly reduced (September = 76.1 :t 2.3 mg/kg). The May data was obtained from
samples stored under suboptimal conditions. It is suspected that there was a substantial
loss of PAHS from the May soil samples due to a prolonged storage time in mostly empty
jars having excessive unfilled headspace. September wetland data apparently
misrepresent soil concentrations as unusually low, since a small number of grab samples
taken in November 2001 and May 2002 had much higher soil [tPAH] of 110-120 mg/kg

(Dr. Rugh, personal communication). Due to these inconsistencies and the likelihood

79

that the Wetland samples were inadvertently compromised, the wetland soil [tPAH] data
is not considered further in this thesis.
Upland data

Phytoremediation and microbial degradation resulted in plot decreases in soil
[tPAH] of ~10 mg/kg over 2 months. This is in accordance with previous studies, which
have shown soil [tPAH] was reduced in planted treatments and the unplanted control
(Pradhan et al., 1998; Liste and Alexander, 2000; Aprill and Sims, 1990). As in previous
studies, planted treatments reduced soil [tPAH] more than unplanted treatments (Qiu et
al., 1997; Schwab, 1994; Aprill and Sims, 1990; Nedunuri et al., 2000; Pradhan et al.,
1998; Yateem et al., 2000) Plant-assisted, aerobic bacterial degradation of PAHS was
likely the primary mechanism for soil [tPAH] reduction in the upland plot over the May
to September treatment period. Previous literature indicates bacterial aerobic degradation
of PAHS contributes most to PAH reduction in soil compared to volatilization, leaching,
or photolysis (Park et al., 1990; Reilley et al. , 1996). PAH degradation by plant processes
alone has not been clearly demonstrated as a means of effectively reducing soil [tPAH].

A. gerardii, A. novae-angliae, E. purpureum, and S. teribinthinaceum had
significant declines in soil [tPAH] fi‘om July to September, indicating that these plant
species are capable of phytostimulation of PAHS in soils. A. scoparius and E. perfoliatum
(July) and E. purpureum (September) treatments had lower soil [tPAH] compared with
unplanted, indicating that these taxa may also enhance soil [tPAH] reduction. Soils grown
with G. triflorum had significantly higher soil [tPAH] compared to the unplanted control
in September and the soil [tPAH] increased significantly from July to September. The

low July concentration, 93.9 :t 6.7 mg/kg, was not significantly different from the May

80

concentration 111.5 3: 15.2 mg/kg. G. triflorum soil data may be indicative of sample
variability than actual plant-soil phenomena. In this study, most plants were generally
observed to enhance biodegradation of PAHS, which is consistent with previous studies
(Aprill and Sims, 1990; Liste and Alexander, 2000; Pradhan et al., 1998; Reilley et al.,
1996; Yateem et al., 2000).

Variable plant effects on soil PAH concentrations are possibly the result of
differences in plant exudation. Plant taxa secrete different amounts and types of exudates
(Fletcher and Hegde, 1995; Siciliano and Germida, 1998). Some of these chemical
exudates have been shown to promote or inhibit microbial PAH-degraders (Leigh et al.,
2002; Schwab, 1994). The composition and amount of root exudate can change at
different growth stages of a plant. During senescence, the amount of phenols that support
PCB-degrading bacteria and possibly PAH-degrading bacteria released were observed to
increase in mulberry plants (Hegde and Fletcher, 1996). In addition, greater root volume
or biomass can result in more exudation, enhance the stress tolerance of a plant, or
increase oxygen in the soil and thereby facilitate the first step in PAH metabolism, all of
which promote PAH phytoremediation. Thus, in this study, the variable plant effects on
the concentrations of PAHS may have resulted from the combined effects of taxa-specific
root exudation, plant age, or root biomass.

Other published research that used the same plant species as this study had
different results with regard to PAH reduction. Specifically, the A. scoparius treatment
was not significantly different from the unplanted control in September. Yet in another
study, A. scoparius enhanced PAH reduction by 8% to 50%, in contrast to unplanted

controls (0% and 26%) in two soils over 6 months (Pradhan et al., 1998). Likewise, the

81

A. gerardii treatment was not significantly different from unplanted control in September,
yet it has been shown to decrease PAH concentration (Aprill and Sims, 1990).
Benz(a)anthracene, Chrysene, benzo(a)pyrene, and dibenz(ah)anthracene biodegradation
was reportedly enhanced by a mixture of plant species that included A. scoparius, A.
gerardii and other prairie grasses after 151 days (Aprill and Sims, 1990). The
discrepancies between these plant species effects on soil PAH reduction presented here
and previous literature may be due to insufficient treatment time and high variation
among soil analytes, rather than ineffectiveness of the Andropogon species.

The use of single subsamples for May soil core analysis restricted the ability to
statistically analyze differences within the May data and compared to the other sampling
times. The lack of significance of treatment and the treatment x time interaction effects in
the AN OVA test using one subsample indicated that the use of one subsample was not
sufficient to distinguish significant differences in soil [tPAH] between treatments. Pair-
wise least significant differences between treatments are not considered significant unless
the AN OVA F-test for treatment is significant (Carmer and Swanson, 1973;
Montgomery, 1997). In contrast, the two-way AN OVA analysis using 3 subsamples
showed that treatment effect was significant. The treatment x time interaction term was
also significant for July and September, which means there was a treatment effect that
varied by sampling date.

The limited statistical power of single soil sample analysis was exacerbated by
wide sample variation in observed soil [tPAH] levels among subsamples. It has been

suggested that better soil homogenization be achieved by pulverization, use of finer

82

sieves, or use of greater soil volumes for extraction in future studies (Dr. G. Phil
Robertson, MSU; personal communication).
Comparison of field results to greenhouse results

The results from the Phyto Demo field plots are consistent with those obtained
from the pilot greenhouse study conducted by APGEN (Carreira and Rugh, unpublished).
The species with the greatest potential to decrease soil [tPAH] in the Phyto Demo upland
plot as indicated by significant differences in soil [tPAH] compared to the unplanted
control or over time were A. gerardii, A. scoparius, A. novae-angliae, E. perfoliatum, E.
purpureum, and S. teribinthinaceum (Table 2.5). These 6 plant species had soil [tPAH]
lower than the unplanted control either 4 out of the 4 sampling times or 3 out of the 4
sampling times in the APGEN study (Appendix 2). Out of 38 species in the APGEN
study, A. novae-angliae, E. perfoliatum and S. teribinthinaceum were 3 species
treatments that had lower soil [tPAH] than the unplanted control 4 out of the 4 sampling
times (Appendices 1 and 2). Similarly, A. gerardii, A. scoparius, and E. puipureum were
3 treatments of 15 plant species treatments that had lower soil [tPAH] compared with the
unplanted control three out of the four sampling times in the APGEN study. Eleven out of
the 17 treatments common to both studies have soil indices (average of planted soil
[tPAH] divided by unplanted soil [tPAH]) that rank the same relative to the unplanted
soil (Table 2.7 and Appendix 2). The agreement between these two studies indicates
greenhouse studies can provide useful information for environmental field studies.
Individual PAH compounds in soil

Different individual PAH compounds vary in biodegradability and respond

differently to a given treatment. Although soil concentrations decreased for

83

benzo(a)pyrene, dibenz(ah)anthracene, and benzo(ghi)perylene in most treatments, these
high molecular weight PAH compounds (Fig. 2.6 g — k) showed smaller decreases
compared with lower molecular weight PAHs (Fig. 2.6 b - t). This trend is likely a
reflection of the physiochemical properties of PAHS (Park et al. , 1990). PAHS with
higher molecular weight are more recalcitrant to soil desorption and biodegradation
compared to lower molecular weight PAHS (Trapp, 2000; Cerniglia, 1992). In this study,
we observed that phytoremediation was more effective for reduction of lower molecular
weight PAHS compared with those of higher molecular weight.

Plant [tPAH]

The concentrations of PAHS observed in leaf tissue from plants in the Phyto
Demo study (Appendix 4) were higher than values reported in published literature. We
are currently uncertain of the reason(s) for the high PAH concentrations detected in plant
leaf tissues in this study and suspect that there may be analytical problems. Data from
previous literature indicate that the plantconcentration values obtained in this study may
be erroneous since leaf PAH concentrations are typically detected in only parts per billion
(ppb) and in this study the levels are in hundreds and thousands of parts per million
(ppm)-

PAHs in plant tissue may occur by plant uptake, plant biosynthesis,
biomagnification, atmospheric deposition or wind-blown dust, but the literature has not
reported values in the hundreds to thousands of parts per million in plants from these
sources. Natural background levels of PAHS can occur in plants with concentrations from
10-90 ug/kg dry weight for each PAH compound (Sims and Overcash, 1983), which are

1000 times lower than levels detected in plants in this study. Previous studies indicated

84

plants accumulate PAHS at levels ranging from undetectable to low natural background
concentrations (Sims and Overcash, 1983). Very low to undetectable PAH levels have
been observed in grasses (Qiu et al., 1997) alfalfa (Schwab, 1994) and carrot foliage
(Wild and Jones, 1992). These studies indicate these plants do not readily uptake and
accumulate PAHS, and that background PAH concentrations are typically undetectable to
low.

Biomagnification, which can be defined as plant to soil concentration ratios
greater than one, of PAHS is one possible explanation for the high concentrations
detected in plant shoot tissues in this study, though has not been shown to result in the
high PAH concentration values detected here. In this study the plant-to-soil (July)
benzo(b)fluoranthene concentration ratios for grasses were: A. gerardii 1.8 (20.0 mg/kg
dry weight in plant; 10.9 mg/kg dry weight in soil), A. scoparius 2.8 (21.3 mg/kg dry
weight in plant; 7.4 mg/kg dry weight in soil), H. patula 7.7 (83.0 mg/kg dry weight in
plant; 10.8 mg/kg dry weight in soil), and S. pectinata 11.1 (103.0 mg/kg dry weight in
plant; 9.3 mg/kg dry weight in soil). These values are not the highest plant to soil
concentration values reported in the literature. Crop to soil concentration ratios for
benzo(b)fluoranthene in wheat can be as high as 59.4 (119 ug/kg dry weight in plant; 2
ng/kg dry weight in soil; yield = 44.9 g dry weight) (Sims and Overcash, 1983). Previous
literature, however, may not provide an equivalent comparison because previous reports
of plant PAH concentrations did not have soil PAH concentrations as high as in this
study. In addition, high plant PAH concentrations were seen in leaf tissues obtained from
plants grown in uncontaminated soils, which would be indicative of extremely high and

implausible biomagnification from low soil [PAH] (Table 2.9).

85

Alternatively, PAHS detected in plant tissue in this study may have resulted from
atmospheric deposition (Lodovici et al., 1994; Sims and Overcash, 1983), but again this
source of PAHS has not been shown to result in hundreds or thousands of parts per
million in plants. Lodovici (1994) detected PAH concentrations on the level of hundreds
of ppb in leaves of evergreen tree Laurus nobilis.

It is also possible that PAH-enriched dust contributed to the high levels of PAH
concentration in our study, though high concentrations of PAHS were also detected in
greenhouse grown plants prior to exposure to PAH-contaminated soils (Appendix 5).
These findings indicate that dust was not a source of high leaf PAH concentrations in this
study. To minimize the possibility of wind-blown PAH-enriched dust contribution to
detected concentrations in future studies, it is recommended to rinse settled dust from
plant tissues prior to extraction.

It is possible that our analytical methods are the source of the elevated leaf PAH
concentrations. The GC-F ID may have detected plant compounds with similar
chromatographic properties to PAHS, however, previous literature has not reported such
problems. The compounds we identified as PAHS should be verified using other
analytical detection methods such as gas chromatography—mass spectrometry (GC-MS).
In addition, our plant extraction protocol should be compared to other published methods
or modified to use larger quantities of tissue to avoid magnification of “noise” from

residual PAHS in extraction glassware or the GC column.

86

SUMMARY AND CONCLUSION

Phytoremediation using native plants has potential to become an effective
remediation strategy for PAH-contaminated soils. Soil [tPAH] in the upland plot was
reduced 10 mg/kg over each 2 month period between May to September of the 2001
growing season. This study has also shown that the selection of plant species is an
important consideration for implementation of phytoremediation. Certain plant species
treatments showed enhanced soil [tPAH] reduction compared to the unplanted control.
Unplanted control soils decreased soil [tPAH] by 17%, while other plant species
treatments, notably E. puipureum, resulted in accelerated soil [tPAH] reduction of up to
35%. The species best able to achieve soil remediation over the first growing season were
A. gerardii, A. novae- angliae, E. purpureum, and S. teribinthinaceum, which decreased
soil [tPAH] significantly from July to September sampling times. By contrast, the
unplanted control soil [tPAH] did not significantly decrease. These results indicate that
these plant species enhanced soil [tPAH] reduction, possibly by stimulation of microbial
metabolic processes for PAH degradation.

The effect of a plant species on a given contaminant was not uniform across all
individual PAH compounds for all treatments. Different PAHS varied in biodegradability;
PAHS with 5 or 6-rings did not change in soil concentration or increased in relative
percentage from May to September. The results of this study are in accordance with
previous reports which demonstrated higher molecular weight PAHS are generally more
environmentally persistent than those of lower molecular weight.

Phytoremediation has advantages over engineered remediation approaches. Other

remediation methods that are commonly used, such as solidification, excavation and

87

landfilling, do not lead to decreases in contaminant concentration, are expensive, and
highly disruptive to a site. Phytoremediation is substantially less expensive, is minimally
disruptive during site rehabilitation, and can be used as an alternative or complement to
other remediation strategies. Identifying plant taxa and the cultivation conditions that
promote PAH-biodegradation will lead to improved PAH-phytoremediation technology.
Persistent organic pollutants (POPS), which include PAHS, are toxic chemicals
that are widespread in the environment. They do not readily undergo biogeochemical
reactions, remain in soils for a long time, and are prone to biomagnify through the food
chain. The rise in contaminated areas with POPS associated with anthropogenic activities
such as fossil fuel burning, and pesticide use Since the industrial revolution has presented
the world with major environmental crises. Environmental clean-up of these
contaminants is needed globally. Phytoremediation is an emerging field in environmental
rehabilitation. The major barrier for phytoremediation of organic pollutants is their lack
of bioavailability as a result of their strong sorption to organic material. In this study, we
evaluated the effectiveness of 18 native plant species to phytoremediate soil
contaminated with PAHS in field demonstration plots. PAH phytoremediation capabilities
are variable among plants and preliminary pilot greenhouse studies can provide good
estimates of plant effectiveness in the field. We have identified a group of plant Species
that accelerate biodegradation of PAHS after one growing season that can be used for

future field and laboratory phytoremediation applications.

88

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91

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soil index value <1.

 

 

Planted [tPAH]/ Unplanted [tPAH]

Appendix 2. APGEN greenhouse study results - soil [tPAH] ratios. Each * indicates a

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

Plant species 6 weeks 10 weeks 14 weeks AVG Trend
Mimulus ringens 0.45 0.36 0.24 0.35 ***
Eupatorium purpureum 0.19 0.76 0.37 0.44 ***
Solidago patula 0.54 0.47 0.45 0.49 ***
Amorpha canescens 0.40 0.59 0.71 0.57 ***
Aster novae-angliae 0.62 0.62 0.67 0.64 ***
Cirsium discolor 0.07 1.15 0.70 0.64 **
Silphium teribinthinaceum 0.59 0.84 0.63 0.69 ***
Eupatorium perfoliatum 0.82 0.61 0.67 0.70 ***
Viburnum dentatum 0.61 0.62 0.95 0.73 ***
Andropogon scoparius 1.37 0.43 0.51 0.77 **
Ceanothus americanus 0.89 0.74 0.82 **
Aristida purpurescens 0.92 0.93 0.63 0.82 ***
Quercus nigra 0.75 1.19 0.58 0.84 **
Liquidambar styraczflua 0.59 1 .26 0.73 0.86 **
Physocarpus opulifolius 0.52 1.48 0.86 0.95 **
Malus coronaria 0.76 0.81 1.36 0.98 **
unplanted control 1.00 1.00 1.00 1.00
Koeleria macrantha 1.32 1.12 0.59 1.01 *
Coreopsis tripteris 0.25 1.62 1.17 1.01 *
Aesculus glabra 1.02 0.59 1.52 1.04 *
Rudbeckia laccinata 0.92 1.69 0.64 1.08 *
Asclepias tuberosa 1.33 0.40 1.56 1.10 *
Hystrix patula 1.30 0.74 1.32 1.12 *
Verbena hastata 0.50 1.27 1.67 1.15 *
Nyssa sylvatica 0.74 1.74 0.97 1.15 **
Scirpus atrovirens 1.98 1.03 0.60 1.20 *
Panicum sp. 0.88 2.22 0.51 1.21 **
F raxinus pennsylvam’ca 0.93 2.94 0.04 1.30 **
Asclepias incarnata 0.79 2.51 1 .50 1.60 *
Andropogon gerardii 0.22 4.26 0.36 1.61 **
Spartina pectinata 0.74 3 .45 0.68 1.63 **
Corylus americana 1.23 3.74 0.77 1.91 *
Liam's aspera 2.66 1.48 2.07
Spirea alba 6.66 0.69 0.00 2.45 **
Carex sprengelii 3.1 1 3.97 1.56 2.88
Elymus virginicus 13.34 1.23 1.05 5.21

 

95

 

 

mortality rate.

Appendix 3. Phytoremediation Demonstration site plant mortality inventory. "M.R." is

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

Overwinter Season 1 Season 2

Scientific Name M. R.% (May01) M.R.°/o (JulyOl) M.R.% (Junedij
Amorpha canescens 37.5 27.3 2.8
Andropogon gerardii 31.3 23 .8 0.0
Andropogon scoparius 77.1 10.6 11.1
Aster novae-angliae 91.7 47.8 1.4
Carex sprengelii 33.3 17.0 0.0
Ceanothus americonus 100.0 62.5 50.0
Cirsium discolor 100.0 51.0 2.8
Eupatorium perfoliatum 44.0 38.8 9.7
Eupatorium purpureum not planted 0.0 8.3
Geum trzflorum not planted 6.3 16.7
Hystrix patula not planted 0.0 2.8
Lobelia cardinalis not planted 4.2 13.9
Mimulus ringens not planted 0.0 0.0
Physocarpus opulifolius 95.8 48.9 0.0
Scirpus atrovirens not planted 0.0 0.0
Silphium teribinthinaceum 85.4 48.3 0.0
Solidago patula 100.0 50.0 11.1
Spartina pectinata 4.2 2.1 0.0
Spirea alba not planted 0.0 0.0
Viburnum dentatum 4.2 2.1 0.0

Average MR. (‘70) = 61.9 22.0 6.5

 

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100

Appendix 5

Greenhouse Study of Effects of Plant Species and Compost on PAH
Phytoremediation

INTRODUCTION

Phytoremediation and microbial bioremediation are both biological methods of
environmental clean-up. Bioremediation often involves the construction of injection wells
for adding microorganisms, food sources, or chemicals to the site of contamination in
order to enhance remediation. Some bioremediation efforts include removing the soil and
placing it in bioreactors. Biological reactors (bioreactors) are chambers that are used to
mix contaminated soil, microorganisms and sometimes amendments for the purpose of
bioremediation. The use of reactors involves disruptive excavation and is expensive.
Phytoremediation uses plants to degrade, or detoxify environmental contaminants.
Phytostimulation is a phytoremediation strategy, which involves the stimulation of
microbial degradation of a contaminant by plants and exudates. Plants and
microorganisms rely on soil nutrients and water for survival, and for degradation
environmental contaminants. The availability of soil nutrients and water are dependent
upon the soil structure and properties. Addition of compost to contaminated soils can
improve soil structure, enhance soil nutrients, increase organic matter, increase porosity,
and increase soil cation exchange capacity. These changes could be favorable for plants
and microorganisms and enhance biological methods of remediation. This experiment
was conducted to evaluate the use of plants in combination with compost amendments for
biological degradation of PAHS in contaminated soils.

Composts have been shown to be beneficial to remediation of organic and

inorganic contaminants. Compost can increase the microbial activity and perhaps the

101

number of microbial degraders in contaminated soils. Using compost for bioremediation
of PAHS has been demonstrated to be an effective method of reducing PAH
concentration in bioreactors (Lilja et al., 1996; Civilini and Sebastianutto, 1996).
Addition of steer manure compost was shown to enhance degradation of 1, 3-
dichloropropene after 8 weeks (Ibekwe, 2001). Bacterial and fungal cell densities were
greater in yard compost—amended pesticide-contaminated soil when compared to
unamended contaminated soil (Cole, 1994). Poultry litter and peat moss amendments

have been demonstrated to increase plant biomass and percentage accumulation of

. . . 137 . 9O . .
radioactive contaminants Cesrum and Strontium compared to controls With no

amendments (Entry et al., 2001). Compost, therefore, has previously been beneficial to
environmental rehabilitation processes. In this study, the effects of plants in soils
amended with composts on the phytoremediation of PAHS were evaluated using three
plant species and an unplanted control.
MATERIALS AND METHODS

The experiment involved 3 different plant species treatments, 6 soil mixtures, and
3 sampling times. The 3 plant species were: AndrOpogon gerardii (Big bluestem),
Eupatorium perfoliatum (Boneset), Lobelia cardinalis (Cardinal flower). For ease of
presentation and communication, abbreviations were developed for each treatment. The
plant species was abbreviated as the first letter of its genus (Table A5.l). In addition, the
experiment had an unplanted soil control (coded “U”), and an untreated soil control. The
implanted soil control was watered and fertilized in the greenhouse along with the other
planted species’ treatments and was used as a control to assess the effects of plant

species. The untreated control treatment soil was stored in amber jars with teflon caps at

102

 

4 °C and was used as a control to assess the effects of abiotic (volatilization, leaching and

photolysis) and microbial processes on soil total PAH concentration [tPAH] that could

have occurred in the pots. This untreated control was also used to estimate an acceptable

storage time for soil samples at 4 °C.

The 6 soil mixtures were designated as treatments A, B, C, D, E, and F (Table

A5.l). Treatments B and B were soils amended with 15% yard compost (Charter

Township of Ypsilanti, MI) by volume, and soil treatments C and F were amended with

10% yard compost (sieved 5 2.36 mm) and 5% poultry manure (sieved _< 2.36 mm)

(Herbruck’s Poultry Ranch, Saranac, MI ) by volume.

Table A5.1. Treatment Codes.

 

 

 

 

 

 

Plant treatment code (Treatment
A Andropogon gerardii
E Eupatorium perfoliatum
L Lobelia cardinalis
U unplanted

 

Soil treatment code

 

A

contaminated soil

 

contaminated soil + 15% yard compost (v/v)

 

contaminated soil + 10% yard compost (v/v) + 5% poultry
manure (v/v)

 

uncontaminated soil

 

uncontaminated soil + 15% yard compost (v/v)

 

mmoow

uncontaminated soil + 10% yard compost (v/v) + 5%
poultry manure (v/v)

 

 

 

Samples were taken at weeks zero, four and eight. The total number of pots in the
experimental set up were calculated as follows: 4 plant species treatments (including
implanted) x 6 soil treatments x 6 replications x 2 sampling times, which amounted to
288 pots. The actual replication number sampled at each sampling time in the experiment
was four, allowing for two extra replications per sampling time in case of plant mortality
or accidents.

Soil

Uncontaminated soils

 

The uncontaminated soil consisted of a mixture of sandy loam and 2-NS sand
(sieved 5 4.75 mm). 2-NS sand as classified by the Michigan Depaitment of
Transportation typically possesses a high carbonate concentration (Dr. Delbert Mokrna,
MSU, personal communication). The uncontaminated soil was made by mixing 1/3
sieved 2-NS sand (40 L) and 2/3 sieved sandy loam (80 L) by volume using a cement
mixer.

Contaminated soils

 

Contaminated field soil was obtained from the coking oven area of the Rouge
Manufacturing Complex in Dearborn, MI and stored at 4° C until use in the greenhouse
experiment. The PAH-contaminated soil was mixed in a 1:1 (v/v) ratio with the
uncontaminated soil mix (1/3 of sandy loam + 2/3 2-NS sand), all sieved (f 4.75 mm)
prior to mixing.

Soil characteristics

 

Nutrient and structural characteristics for each soil treatment were determined by

MDS Harris (Lincoln, NE) at weeks zero and eight (Tables A52, A53). The Rouge

104

PAH-contaminated coke oven soil had a pH range from 8.0 - 8.1 and was classified as
loamy sand or sand (MDS Harris, Lincoln, NE). The pH of the contaminated soil was
maintained to favor indigenous microbial PAH degraders. The soil pH was measured

weekly from pot leachate with pH paper strips.

In general, contaminated soils had higher soluble salts, nutrients, sodium, and
cation exchange capacities than uncontaminated soils (Tables A5.2, A5.3). Amended
soils had apparently higher concentrations of nutrients compared with unamended soils
(Tables A5.3). Poultry manure treatments seemed to have higher phosphorus and
potassium than the other treatments. Yard compost treatments had the lowest pH values
(Table A5 .2).

Plants

Plant species included in this study, A. gerardii (Big bluestem), E. perfoliatum
(Boneset), and L. cardinalis (Cardinal flower), were chosen based on Chapter 2 field
results and seed availability. The seed gennination rates for A. gerardii, E. perfoliatum,
and L. cardinalis were roughly 44.9 %, 68.1 % and 12.2 %, respectively. Seeds for A.
gerardii were donated by North American Prairies (Annandale, MN) and the seeds for E.
perfoliatum and L. cardinalis were obtained from Wildtype Nurseries (Mason, MI).

Seeds were germinated in potting soil in plastic germination trays.

105

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

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107

Table A5.3. Soil nutrient concentrations at week 0 and week 8. Each value represents
the mean of two samples. Soil and plant codes defined in Table A5.1.

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

Nutrients (ppm)
Trmt

Soil Wks Plant N P K Mg Ca S Zn Mn Cu Fe B
A 0 none 77 15 129 479 5571 999 10 10 3 20 3
A 8 A 14 30 100 384 4435 879 12 11 3 20 2
A 8 E 6 65 38 303 2960 84 13 11 3 22 1
A 8 L 15 39 116 397 4481 930 12 11 3 21 2
A 8 U 3 63 96 316 3640 408 13 28 3 22 2
B 0 none 119 25 253 483 5129 999 11 9 2 70 3
B 8 A 15 51 123 348 3887 478 14 14 3 49 2
B 8 E 5 53 49 321 3134 80 15 13 3 40 1
B 8 L 20 52 177 398 4173 656 14 12 2 44 2
B 8 U 6 67 106 327 3395 204 15 37 3 48 2
C 0 none 1 14 88 878 608 5034 999 16 25 3 36 3
C 8 A 24 147 394 409 3888 711 19 28 3 52 3
C 8 E 15 174 82 296 3202 139 20 17 2 41 2
C 8 L 27 179 369 405 3713 595 20 17 3 45 3
C 8 U 4 201 228 376 3316 290 21 77 4 45 2
D 0 none 14 4 34 119 2659 97 1 4 1 31 0
D 8 A 7 21 9 144 2124 19 2 4 1 29 O
D 8 E 3 29 11 179 2136 17 2 4 1 3O 1
D 8 L 6 23 18 143 2156 18 2 4 1 30 O
D 8 U 4 26 14 173 2204 24 3 5 1 32 1
E 0 none 52 11 116 168 2682 82 3 4 1 44 1
E 8 A 7 28 25 196 2273 18 5 5 1 46 1
E 8 E 3 43 13 235 2357 14 5 5 1 44 1
E 8 L 7 33 36 203 2416 23 5 5 1 52 1
E 8 U 6 47 31 210 2198 22 4 5 1 46 1
F 0 none 25 83 603 220 2263 140 7 15 1 89 1
F 8 A 10 128 82 284 2200 31 10 19 2 52 1
F 8 E 9 117 28 269 2282 27 10 8 1 43 1
F 8 L 22 117 281 308 2318 40 13 9 1 111 1
F 8 U 4 124 70 275 2158 25 9 29 2 46 1

 

 

 

 

 

 

 

 

 

 

 

 

 

 

108

 

Experimental Set-up

A coffee filter was placed in each labeled plastic pot (10 cm x 10 cm x 4”) prior to
filling the pot with soil to minimize loss of soil. Contaminated soils were put in pots in
chemical safety fume hood. E. perfoliatum, L. cardinalis plantlets (each 6 weeks old),
and A. gerardii plantlets (3 weeks old) were transplanted into the pots containing the six
soil mixtures. E. perfolz'atum was severely wilted the second day after transplanting and a
total of 18 plants were replaced. To ease transplanting stress, the plants were covered
with Ziploc bags for 1 day. Plant viability was recorded weekly and photos of the planted
treatments were taken every two weeks. Each pot had been assigned a number from a
random number table (Moore and McCabe 1999) for placement in staggered randomized
rectangular grids (6 pots x 16 pots) spaced one pot width apart on greenhouse benches.
Greenhouse conditions

The experiment was conducted from February to April 2002, during which time
daylight hours increased from 9 hours to 11 hours. A LI-189 photometer (LI-COR,
Lincoln, NE) was used to measure photosynthetically active radiation (PAR) at pot
height. At noon on a relatively sunny day PAR levels were 350 — 700 uE-s’lm'z; on partly
cloudy days, PAR levels were 300 — 600 uE-s’lm'z. Although daylight hours were 9 toll
hours per day, artificial lights (400 Watts, high pressure sodium, intensity 970 candles,
General Electric) were on 16 hours per day. PAR levels at night when artificial lights
were illuminated were 40 - 75 uE-s'lm'z. Greenhouse temperature was typically between
20 °C to 30 °C except for a few days near the end of March when the greenhouse

temperature was ~38 °C due to temperature control malfiinction.

109

 

Watering & Fertilization

Plants were watered as required and unplanted treatments were watered daily to
field capacity. When the weather was cloudy, plants typically required watering once a
day and sometimes every two days. When the weather was sunny, plants required
watering at least three times a day to up to seven times a day for E. perfoliatum between
weeks four and eight. All treatments were watered with N-P-K fertilizer solution (20—20-
20, ~475 ppm) once a week.

Sample handling

Time zero samples were extracted and stored in amber jars with teflon lids at 4 °C
and used as untreated control samples for weeks 4 and 8. A representative 4 samples from
each plant species were used for extraction and a representative four plants were used for
determination of dry weights. Plant tissue dry weights were determined by oven drying at
~80 °C for at least 48 hours.

Plants and soil were destructively sampled at weeks 4 and 8. For each pot, the top
~1 cm of soil was discarded using a metal spatula. The root portion of the plant was
removed from the pot by gently prying into the soil around the roots with a metal spatula
and collecting the soil off the roots. Soil was rinsed off roots in a bucket of water and the
roots were blotted dry and weighed using an electronic balance. After the fresh weights
were recorded, the plants were dried in an oven for at least 48 hours at ~ 80 °C for
determination of dry weight.

At weeks 4 and 8, the soil from the center of the pot was collected and stored in
150 mL amber jars with a teflon lid at 4°C and extracted the day after harvesting. For

uncontaminated pots, one subsample was taken from each plant x soil treatment. Fewer

110

samples were taken from the uncontaminated pots because it was hypothesized soil
[tPAH] would be minimal, therefore, the data would not be useful and devoting more
resources to this would have been wasteful. Uncontaminated soil and plant treatment
samples were used for determination of background soil [tPAH] levels.

Analyses of samples

Extractions of PAHS from samples, and analyses for PAH concentration were
done as stated previously in Chapter 2 Materials and Methods section.

The total PAH concentration [tPAH] was determined by taking the sum of the
concentrations of the following compounds: naphthalene, acenaphthylene, acenaphthene,
fluorene, phenanthrene, anthracene, fluoranthene, pyrene, benzo(a)anthracene, chrysene,
benzo(b)fluoranthene, benzo(a)pyrene, dibenzo(a,h)anthracene, and benzo(g,h,i)perylene.
Refer to Table 2.4 for abbreviations and lowest calibration standards.

RESULTS
Soil total PAH concentration ([tPAH])

There were no obvious differences in soil [tPAH] among plant treatments
(including implanted and untreated treatments (4°C). In addition, there were no
differences among soil treatments at each sampling time (Table A5.4) during this 8-week
greenhouse study and no trends in soil [tPAH] over time. Soil [tPAH] was highly variable

in all planted, unplanted, and untreated treatments at each sampling time (Table A55).

111

Table A5.4. Soil total PAH concentration (mean :t SEM) in pots containing PAH-

contaminated soil over time (N = 4 pots, N = 1 as noted by “*” due to mortality).

Subsamples from each pot were averaged and standard errors represent the variability

of the average [tPAH] among pots for a given treatment.

 

Soil Concentrations (mg/kg)

 

 

 

 

 

 

 

 

Treatment Treifililient Week 0 Week 4 Week 8
A. gerardii A 226.3 d: 20.7 364.3 :1: 16.3 311.3 d: 34.8
A. gerardii B 233.9 i 27.5 251.6 :1: 24.8 331.2 :1: 63.9
A. gerardii C 256.9 :1: 34.4 370.9 :1: 783* 250.3 :1: 12.7
E. perfoliatum A 226.3 :1: 20.7 283.5 :t 26.6 384.7 :1: 62.9
E. perfoliatum B 233.9 :1: 27.5 222.0 d: 19.8 269.7 :1: 25.1
E. perfoliatum C 256.9 :1: 34.4 193.1 N 11.6 345.5 :1: 70.5
L. cardinalis A 226.3 :1: 20.7 321.9 d: 49.3 308.6 :1: 40.8
L. cardinalis B 233.9 :1: 27.5 418.8 :1: 100.1 296.1 i 56.2
L. cardinalis C 256.9 :1: 34.4 346.9 :L- 39.0 268.5 :t 32.1'
Unplanted A 226.3 :1: 20.7 420.0 :1; 34.0 247.3 :1: 22.7
Unplanted B 233.9 :t 27.5 223.6 :t 19.3 210.2 :1: 11.8
Unplanted C 256.9 3: 34.4 334.3 :t 63.4 261.7 at 40.3
Untreated A 226.3 i 20.7 322.1 :1: 61.7 337.9 :1: 105.1
Untreated B 233.9 :t 27.5 196.7 :1: 6.3 281.8 a: 29.6
Untreated C 256.9 :1: 34.4 283.5 :1: 38.6 447.5 :1: 88.9

 

112

 

 

 

Table A5.5. Soil [tPAH] maximum pot ranges determined from 3 subsamples per pot per

treatment and treatment soil [tPAH] ranges determined from means of subsamples from 4

individual pots at week 8.

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

Soil [tPAH] (mg/kg)
Treatment Soil T reatrnent Pot Range Treatment Range
A. gerardii A 220 - 522 237 — 376
A. gerardii B 194 - 986 222 — 484
A. gerardii C 140 - 335 218 — 275
E. perfoliatum A 258 - 692 270 — 542
E. perfoliatum B 180 - 430 239 — 320
E. perfoliatum C 195 - 1031 166 — 485
L. cardinalis A 207 - 526 230 — 416
L. cardinalis B 146 - 1081 223 — 464
L. cardinalis C 141 - 433 199 — 354
Unplanted A 292 - 401 191 — 301
Unplanted B 104 - 287 191 - 243
Unplanted C 178 - 608 150 — 331
Untreated A 157 - 1597 231 — 653
Untreated B 220 - 544 206 — 347
Untreated C 396 - 938 313 — 709

 

 

 

 

 

113

 

Plant [tPAH]
The [tPAH] in plant tissues were high (Table A56). Individual PAH compounds

detected in plants were predominantly benzo(a)anthracene, chrysene,
benzo(b)fluoranthene, benzo(a)pyrene, and benzo(ghi)perylene, though Eupatorium
leaves were also observed to contain elevated levels of acenaphthylene and acenaphthene
(data not shown).
Plant Dry Biomass

Plants had varied initial fresh weight at time 0 and were randomly assigned to
treatments. Further analyses of these data should utilize Analysis of Covariance with
initial plant fresh weight as a covariate. Contrasts should be utilized for treatment

comparisons.

Shoot Dry Weight

 

For each species, shoot dry weight increased over time (Fig A5.l). E. perfoliatum
plants had greater shoot dry weight than A. gerardii or L. cardinalis. For A. gerardii and
E. perfoliatum, plants grown in soil F (uncontaminated soil + yard compost + poultry
manure) showed substantially greater shoot dry weight than plants grown in soils A
through E at week 8. For A. gerardii there was a trend of increasing shoot dry weight in
soils D < E < P at week 8.

Root Dry Weight

Root dry weight increased over time from week 0 to week 8 in all soil treatments
(Fig. A52). In most cases, root dry weight was lower in contaminated soil treatments
than uncontaminated treatments.

It should be noted that that certain treatments showed higher mortality than

others. Treatments amended with poultry manure (treatments C and F) had plant

114

mortality. Shoot and root dry weights were low for treatments C and F at week 4 for all
species.

Table A56. PAH concentration (mean :2 SEM) mg/kg dry weight in plant shoot tissue
from plants grown in pots in the greenhouse. Standard errors represent the variability

among plants for a given treatment. N = 1, N = 3 as noted (*) due to mortality, and N = l

 

 

 

 

 

 

as indicated by T.
Plant [PAH] (mg/kg)

Plant Species Soil Week 0* Week 4 Week 8

A. gerardii A 1200.6 i 234.5 1593.8 2t 412.7 490.6 1 78.1
A. gerardii B 1200.6 i 234.5 562.1 :1: 81.0 605.3 :t 86.7
A. gerardii C 1200.6 :1: 234.5 2263.9 :1: N/AT 1666.1 i 8929*
A. gerardii D 1200.6 :1- 234.5 425.7 :1: 37.6 546.0 3: 24.3
A. gerardii B 1200.6 :1: 234.5 703.5 1 81.2 581.6 :1: 124.5
A. gerardii F 1200.6 3: 234.5 589.8 :1: 54.3 563.1 d: 90.4
E. perfoliatum A 1246.0 3: 272.7 1369.4 i 230.7 569.7 :t 88.3
E. perfoliatum B 1246.0 :1: 272.7 1024.5 i 210.2 597.2 i 79.8
E. perfoliatum C 1246.0 :1: 272.7 1683.9 i 320.3 448.5 i 28.5
E. perfoliatum D 1246.0 :t 272.7 1181.1 :t 300.8 444.3 i 92.6
E. perfoliatum E 1246.0 :1: 272.7 800.0 :t 132.6 543.5 3: 178.8
E. perfoliatum F 1246.0 :1: 272.7 2148.5 :t 1336.5 445.6 :h 63.3
L. cardinalis A 1198.1 i 330.0 322.] i 57.1 3422.3 :t 1802.3
L. cardinalis B 1198.] 3: 330.0 316.4 i 63.0 402.9 1 64.2
L. cardinalis C 1198.1 :1: 330.0 345.8 :1: 1051* 228.1 :1: N/AT
L. cardinalis D 1198.1 at 330.0 272.4 :1: 76.3 413.4 5: 171.2
L. cardinalis E 1198.1 i 330.0 299.5 i 64.0 278.8 :1: 59.2
L. cardinalis F 1198.1 i 330.0 267.2 :t 252* 239.5 N N/AT

 

 

* At time zero, a representative 4 plants were sampled per species.

115

 

2-0 - (a) A. gerardii

 

 

 

 

39
go 1.5 -
é’
i’.‘ 1.0 -
Q
‘5 0.5 -
o
":8
0.0 -
A B C D E F
Soil Treatment
6-0 ' (b) E. perfoliatum
3° 5.0 .
in 4.0 -
D
3 3.0 q
E 2.0 -
§ 1.0 -
.c:
m 0.0 4
A B C D E F
Soil Treatment

:63) 1'5 ' (c) L. cardinalis
é.
~35 1.0 -
3
a.
O 0.5 -
‘5
0
E

0.0 4

 

 

Soil Treatment

Figure A51 (a-c). Plant shoot dry weights (mean + SE) for plants in various soils at
week 0 (empty bar), 4 (grey bar), and 8 (black bar). N = 4, except for L. cardinalis x
Soil C (week 4, N = 3), L. cardinalis x soil F (week 8, N = 1), A. gerardii x soil C
(week 8, N = 3), L. cardinalis x soil C (week 8, N = 1), L. cardinalis x Soil F (week 8,
N = 1). In these instances sample sizes were reduced due to mortality.

116

 

 

1.4 -
1.2 -
1.0 -
0.8 -
0.6 -
0.4 -

O 2 d
.
':'.-\-‘- - '.
'.£ ' i :-, --
'u. 11 .2.- » -r In“

0.0 J

(a) A. gerardii

Root Dry Weight (g)

 

 

A B C D E F
Soil Treatment

7.0 -
6.0 -
5.0 -
4.0 -
3.0 -
2.0 4
1.0 -
0.0 -

(b) E. perfoliatum

Root Dry Weight (g)

 

 

Soil Treatment

3-0 ' (c) L. cardinalis
2.5 -

2.0-
1.5 -
1.0-

Root Dry Weight (g)

 

 

Soil Treatment

Figure A52 (a-c). Plant root dry weight (mean + SE) for plants in various soils at
week 0 (empty bar), 4 (grey bar), and 8 (black bar). N = 4, except for L. cardinalis x
Soil C (week 4, N = 3), L. cardinalis x soil F (week 8, N = 1), A. gerardii x soil C
(week 8, N = 3), L. cardinalis x soil C (week 8, N = l), L. cardinalis x Soil F (week 8,
N = 1). In these instances sample sizes were reduced due to mortality.

117

DISCUSSION

Soil Properties & Nutrients

Soil characteristics were varied among the different treatment soil mixtures. The
contaminated soil had higher nutrient concentration, which may partially reflect its higher
cation exchange capacity (Table A52). The pH increased from week 0 to week 8, which
is likely because the tap water in the MSU greenhouse has a high pH ~8 (Dave Freville,
personal communication), though the starting pH of the Rouge soil was observed to be ~
8.0 — 8.5 (Table A52). In this greenhouse study, soil compost amendments, specifically
the yard compost, decreased soil pH and increased soil organic matter, thereby improving
soil conditions for plant growth.

Soil [tPAH]

The soil [tPAH] values were highly variable and this variability masked any
effects of plant species and soil treatments. The soil from the Rouge Manufacturing
Complex Coke Oven area was highly variable in soil [tPAH], ranging from 500 - 900
mg/kg prior to mixing with uncontaminated soil. The experimental results may have been
improved with better homogenization of the contaminated soils prior to distribution
among the plant treatments. To reduce variability of soil [tPAH] for the samples in a
given treatment, more soil could be used for a single extraction, or soil samples could be
pulverized or finely sieved (Dr. G. Phil Robertson, personal communication). In addition,
the sample variation was very high and potentially not enough samples were taken to
accurately describe the variation, in particularly at time 0. Future studies may also

include more subsamples. Alternatively, the failure to observe reduction of soil [tPAH]

118

 

may indicate that 8 weeks was insufficient to demonstrate phytoremediation of such
highly PAH-contaminated soils.

The results in this study are not in accordance with the majority of previous
studies. There are several potential reasons that soil [tPAH] concentrations were not
observed to decrease in this study as has been shown in other studies (Aprill and Sims,
1990; Pradhan et al., 1998; Yateem et al., 2000) including the APGEN study and Chapter
2 of this thesis. The starting soil [tPAH] concentration for this experiment is higher and
more heterogeneous than in the Phyto Demo field study. It is possible that
phytoremediation may not be as effective at high concentrations. This explanation does
not hold for the comparison of the data in this study with the APGEN data, however,
since the APGEN soil [tPAH] data were just as high and in some cases higher (Appendix
1 vs. Table A55). The APGEN study was conducted differently using only Perlite as a
soil amendment and analysis of the entire pot contents, rather than only for rhizosphere
soil exclusive of roots as performed in this experiment. Concentration of PAHs have been
found to be 4- 5 times higher around plant roots as a result of increased mobility of PAHS
(Liste and Martin, 2000). A hypothesis explaining higher PAH concentration near roots is
that roots increase PAH mobility and exude organic compounds, resulting in sorption of
PAHS to these exudates and to root surfaces (Liste and Martin, 2000). Sampling
differences, therefore, may account for some discrepancy between this study and the
APGEN study. By week 8, plants roots filled the entire pot for most treatments and
separation of roots from soil was difficult. Small plant roots may have been included in

the extraction sample and this too may have contributed to the variability. Multiple

119

factors, alone or synergistically, may have led to discrepancies between this and previous
studies.
Plant [tPAH]

The exceedingly high concentrations of PAHS detected in plants in this study
cannot be explained by previous literature. Previous literature reports various results for
PAH concentration in plant tissue. Some previous studies have shown biomagnification
of PAHS occurs in plant tissues, though others indicate background levels at parts per
billion or lower. Data presented by (Sims and Overcash, 1983) indicate natural
background levels of PAHS such as anthracene, fluoranthene, benz(a)anthracene, pyrene
and benz(a)pyrene in plants at concentrations from 10-90 ug/kg dry weight for each PAH
compound. Thus, the plant PAH concentration values in this thesis are substantially
higher than values previously reported in the literature and the reasons for these high
concentrations in unknown at this time.

Plant Dry Biomass

There were differences in plant growth among plant species for the various
treatments. Transplanting stress and nutrient deficiency stress may have led to slow
grth rates of plants during the first weeks of the experiment. E. perfoliatum had greater
shoot dry weight than A. gerardii or L. cardinalis. Greater plant growth was observed in
uncontaminated soils than the contaminated soils. The higher plant biomass observed in
uncontaminated soils may indicate that plants were stressed in the contaminated soil
perhaps by the lack of nutrient availability or by the PAH contaminants.

The poultry manure amendment may have caused some toxic effects to the plants

in this study. The poultry manure had not been fully composted (Andy Fogiel, personal

120

communication) and the odor of ammonia was evident. At week 4, plants in the yard
compost + poultry manure—amended soil had lower shoot and root dry weights and
higher plant mortality than other treatments (data not shown). By contrast, at week 8, A.
gerardii and E. perfoliatum showed greater growth in uncontaminated soil amended with
yard compost + poultry manure than all other treatments. As plants grew bigger, they
may have become less susceptible to ammonium or salt toxicity and as a result slow
growth rates were not observed between weeks 4 and 8. It is therefore important to
consider the levels and quality of soil amendments prior to large-scale application in
remediation efforts.
SUMMARY & CONCLUSION

This study raised some issues and considerations for firture greenhouse studies.
The experimental results in these trials were confounded by high variability in soil
contaminant concentrations and high plant contaminant concentrations. The source of the
elevated plant tissue [tPAH] levels is unknown and remains to be resolved by reanalysis
of the apparent leaf PAH compounds by additional analytical procedures, such as GC-
MS. In this 8-week greenhouse study, no effect of plant treatment or soil amendments on
soil [tPAH] could be seen because high soil [tPAH] variation may have masked these
effects. Future phytoremediation greenhouse experiments could be improved by more
thorough homogenization of soils before treatment and upon sample analysis.
Additionally, larger sample volumes could be used to buffer the influence of
heterogeneous soil “hot spots” on soil [tPAH] determination. This study has raised

questions about high [tPAH] concentrations in plant tissues, provided important lessons

121

 

in sampling and analyses of PAH-contaminated soil samples, and has shed light on how
analytical protocols may be improved for future phytoremediation experiments.
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polycyclic aromatic hydrocarbon treatment in soil. Chemosphere 20, 253-265.

Civilini, M. and Sebastianutto, N. (1996). Degradation of naphthalene by microorganisms
isolated from compost. In The Science of Composting, M. de Bertoldi, P. Sequi,
B. Lemmes and T. Papi, eds. (Glasgow: Blackie Academic & Professional), pp.
870-883.

Cole, M.A., Liu, X., and Zhang, L. (1994). Plant and microbial establishment in
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Entry, J .A., Watrud, LS. and Reeves, M. (2001). Influence of organic amendments on
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Ibekwe, A.M., Papiemik, S.K., Gan, J ., Yates, S.R., Crowley, DE, and Yang, C.-H.
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Liste, H.-H. and Martin, A. (2000). Accumulation of phenanthrene and pyrene in
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Pradhan, S.P., Conrad, J .R., Paterek, J .R. and Srivastava, V.J. (1998). Potential of
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Sims, RC. and Overcash, MR. (1983). Fate of polynuclear aromatic compounds (PNAS)
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Yateem, A., Balba, M.T., El-Nawawy, AS. and Al-Awadhi, N. (2000). Plants-associated
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