Ecology Of Ecto- And Ericoid Mycorrhizal Systems
In Petroleum Hydrocarbon-Contaminated Sub-Boreal Forest Soils

Susan J. Robertson
B.Sc, University of Calgary, 1990
P.B.D., Simon Fraser University, 1996
M.Sc., University of Northern British Columbia, 2003

Dissertation Submitted In Partial Fulfillment Of
The Requirements For The Degree Of
Doctor of Philosophy
in
Natural Resources and Environmental Studies

The University of Northern British Columbia
December 2008

© Susan J. Robertson, 2008

1*1

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Abstract
The impacts of petroleum hydrocarbon (PHC) contaminants in northern forest soils are not
well understood from either eco-toxicological or microbial ecological perspectives. The
purpose of this research was to examine interactions between PHCs and ecto- (ECM) and
ericoid (ERM) mycorrhizal communities at the rhizosphere scale, where microbial activities
underpin processes such as decomposition, carbon and nutrient cycling, and primary
production at landscape scales. Several methodological approaches were simultaneously
used to assess changes in physical, chemical and biological properties of plant-soil systems
treated with ecologically relevant concentrations (i.e. ~7-22 tonnes ha"1) of PHCs. From
microscopy and community fingerprinting (LH-PCR) studies, we found few differences in
community structure attributable to PHC contamination. PHC treatment also did not appear
to alter broad patterns of C metabolism for either bacterial (CLPP) or ECM fungal (laccase
assay) communities. Habitat changes, which generally included increased C:N ratios,
slightly more acidic pH, and hydrophobicity or water-logging in organic and mineral layers,
respectively, did not appear to inhibit microbial communities. Together, these findings point
to resilience within intact mycorrhizal systems, mainly due to sorption of PHCs within
organic soil layers and protective properties of the mycorrhizosphere habitat. Soil PHC
analysis (GC-FID) showed a general decrease in contaminant levels 16 weeks after PHC
treatment, indicating an intrinsic capacity for biodegradation within the soil communities.
ECMs appeared to play a vital role in this process through provision of habitat and cosubstrates for heterotrophic bacterial communities (i.e. mycorrhizosphere effect) and via
secretion of laccase, which opens aromatic ring structures for subsequent bacterial attack.
These results emphasize the importance of synergistic functions among microbial guilds with

ii

respect to ecological processes. Finally, we found that the spatial patterns of mycorrhizal
communities within the rhizosphere depended primarily on properties of the host plant and
soil environment. The extent that different properties influenced community structure varied
between the three groups of microorganisms. This systems approach addressed fundamental
questions in mycorrhizal ecology by considering PHC pollution as a form of environmental
disturbance. Conservation of the integrity of mycorrhizal systems in contaminated forest soils
may be key for sustainable management in terms of ecosystem resilience and remediation.

in

Table of Contents
Abstract
Table of Contents
List of Tables
List of Figures
Acknowledgements

ii
iv
vii
viii
xi

Introduction

1

Chapter 1: Petroleum hydrocarbon contamination in boreal forest soils: a
mycorrhizal ecosystems perspective
Abstract
Introduction
Mycorrhizas
Classification and structure
Diversity
Ecology of ecto- and ericoid mycorrhizal systems
Soil habitat
Community interactions
Mycorrhizosphere bacteria
Plant linkages
Ecosystem processes
Decomposition
Primary production
Summary
Petroleum hydrocarbon contamination of forest soils
Disturbance
Chemical toxicity
Soil properties and processes
Biodegradation
Bacterial pathways
Fungal cytochrome P450 and ligninolytic systems
Metabolic potential of ECM/ ERMfungi
Genetic controls
Implications for management
Conclusions
References

5
5
7
11
11
13
18
18
24
24
27
29
30
34
36
38
38
39
43
49
51
57
60
63
65
74
77

Chapter 2: Interactions between petroleum hydrocarbon contaminants and ectoand ericoid mycorrhizal communities in sub-boreal forest soils
Abstract
Introduction
Materials and methods
Field site
Bioassay and PHC treatment

94
94
96
100
100
103

iv

Experimental design and sampling
Morphological assessment of ECM communities
DNA extraction and length heterogeneity PCR
Laccase assays
Results
ECM morphotypes: development and response to PHCs
ECM and ERM community structure
Harvest time and PHC effects
Plant effects
Organic layer effects
Laccase activity
Discussion
PHC impacts on mycorrhizal communities
Potential for direct mycorrhizal role in biodegradation
Conclusions
References
Chapter 3: Enhanced biodegradation of petroleum hydrocarbons in
mycorrhizosphere of sub-boreal forest soils
Abstract
Introduction
Materials and methods
Field site
Bioassay and PHC treatment
Experimental design and sampling
PHC extraction and quantification using GC-FID
Soil nutrient analysis and pH
LH-PCR and fragment analysis
Community level physiological profiles
Results
PHC quantification from soils
Analysis of PHC fractions within plant - organic soil systems
Soil properties
Bacterial community structure
Bacterial community level physiological profiles
Discussion
PHC Biodegradation
Mycorrhizosphere effect
Conclusions
References

v

Chapter 4: Root-associated microbial communities differ with Pinus contorta var.
latifolia and Vaccinium vitis-idaea co-inhabiting sub-boreal forest soils
Abstract
Introduction
Materials and methods
Field site
Bioassay, PHC treatment and sampling
Soil Analysis
LH-PCR and fragment analysis
Results
Soil properties
ECM morphotypes
Fungal community structure
Bacterial community structure
Discussion
ECM and ERM root communities
Soil properties
FH and CWD systems
Vertical segregation
Conclusions
References

184
184
186
189
189
190
191
192
195
195
196
199
205
209
209
212
214
215
220
221

Chapter 5: Conclusions and future considerations for microbial ecology and
sustainability management in PHC-contaminated northern forest ecosystems
Rhizosphere model for mycorrhizal systems
Ecological integrity and resilience
PHC biodegradation and functional redundancy
Plants and soils
Management in PHC contaminated forests
References

225
226
227
229
232
235
237

Appendix A: ECM morphotype descriptions

238

Appendix B: Photographs of field site, soils, bioassay, and ECM morphotypes

240

VI

List of Tables
Table 2.1: Summary of plant, organic soil layer, and PHC treatment variables for 20042005 study (n=2)
105
Table 2.2: Summary of plant, organic soil layer, and PHC treatment variables for 20052006 (single-plant) and 2006-2007 (double-plant) studies (n=3)*
105
Table 2.3: One-way ANOVA plus Fisher's LSD (a = 0.05) of Simpson diversity (1/D)
between PHC treatments (within harvest groups): PHC-0 = control; PHC-1 = 73 mg
cm"2; PHC-2 = 146 mg cm"2; PHC-3 = 219 mg cm"2

117

Table 2.4: One-way ANOVA plus Fisher's LSD (a = 0.05) of Simpson diversity (1/D)
between harvests (within PHC treatment groups) at 1, 4, 7, 10, 13, and 16 weeks
following PHC treatment

118

Table 3.1: Summary of plant, organic soil layer, and PHC treatment variables for 20052006 (single-plant) and 2006-2007 (double-plant) studies (n=3)*
147
Table 3.2: Means (± standard errors) for soil properties (C, N, pH) in control and PHCtreated plant - organic soil systems at 1 and 16 weeks. Percent C and N are reported
on an air-dry basis
164
Table 4.1: Plant, organic soil layer, and PHC treatment variables (n=3 for each
combination)

vn

191

List of Figures
Figure 2.1: Maps showing the study site at the Kenneth Creek field site, located about
100 km east of Prince George in the sub-boreal spruce (SBSwkl) zone of the central
interior of British Columbia, Canada
102
Figure 2.2: Relative abundance of ECMs on pine roots in FH and CWD soil systems
sampled at 3-week intervals over 16 weeks with 0, 1, 2, or 3 mL of crude oil added to
the soil surface
115
Figure 2.3: Relative abundance of ECMs on birch roots in FH and CWD soil systems
sampled at 3-week intervals over 16 weeks with 0, 1, 2, or 3 mL of crude oil added to
the soil surface
116
Figure 2.4: NMS ordination of mycorrhizal fungal showing multivariate effects of
harvest time (1 and 16 weeks) and PHC treatment (stress = 19.66; instability = 0.12).. 121
Figure 2.5: Pairwise comparisons of fungal community structure (genotypes) in PHCtreated and control systems at 1 and 16 weeks. Significant differences (MRPP) are
represented by dark arrows and corresponding p-values; light (slashed) arrows
represent no differences between groups

121

Figure 2.6: NMS ordination of mycorrhizal fungal communities of single (closed) and
double (open) plant systems (stress = 19.66; instability = 0.12)

123

Figure 2.7: NMS ordination of organic soil layer (FH, CWD, and FHoil) effects in single
plant systems (stress = 17.64; instability = 0.024)
125
Figure 2.8: Table showing laccase activity of ECM morphotypes and ERM hair roots
assessed by intensity of colour development (-, none; +, pale green; ++, dark green) at
1,16 and 24 h incubation times with ABTS on the left. Symbols in brackets indicate
less typical reactions over numerous trials (n=5+); Photograph showing range of
colour development for ECMs (n=3, vertically) incubated in microplate wells with
ABTS for 24+ h on the right
127
Figure 3.1: Overlay of two chromatograms generated from GC-FID analysis showing a
reduction in PHC peak areas from 1 to 16 weeks. The vertical lines represent the
boundaries (based on retention times of standards) for analysis of the F2 (nClOnC16), F3a (nC16-nC23), F3b (nC23-nC34) and F4 (nC34-nC50) PHC fractions

152

Figure 3.2: Concentration of total PHCs (ppm) in three organic soil layers (FH, CWD
and FHoil) for PHC-treated and untreated controls at 1 and 16 weeks (data pooled for
plant treatment). Bars represent standard errors of the means. Significant losses of
PHCs within organic soil treatment groups are indicated by *
157
Figure 3.3: Concentrations of F2 (nC10-nC16), F3a (nC16-nC23), F3b (nC23-nC34),
and F4 (nC34-nC50) PHCs (ppm) in organic layers of plant - soil systems at 1 and 16
weeks after PHC treatment. Plant treatments include: pine (P), birch (B), lingonberry
(L), pine and lingonberry (P+L), and no plant (N). Bars represent standard errors of
the means (n=3); significant differences (1-way ANOVA) within treatment groups are
indicated by *
159

vin

Figure 3.4: Concentrations of F2 (nC10-nC16), F3a (nC16-nC23), F3b (nC23-nC34),
and F4 (nC34-nC50) PHCs (ppm) in Ae layers of plant - soil systems at 1 and 16
weeks after PHC treatment. Plant treatments include: pine (P), birch (B), lingonberry
(L), pine and lingonberry (P+L), and no plant (N). Bars represent standard errors of
the means (n=3); significant differences (1-way ANOVA) within treatment groups are
indicated by *
160
Figure 3.5: Concentrations of F2 (nC10-nC16), F3a (nC16-nC23), F3b (nC23-nC34),
and F4 (nC34-nC50) PHCs (ppm) in Bf layers of plant - soil systems at 1 and 16
weeks after PHC treatment. Plant treatments include: pine (P), birch (B), lingonberry
(L), pine and lingonberry (P+L), and no plant (N). Bars represent standard errors of
the means (n=3); significant differences (1-way ANOVA) within treatment groups are
indicated by *
161
Figure 3.6: Schematic diagram showing plant effects on bacterial genotype richness and
community structure. Arrows (solid, genotype richness and community structure;
dashed, richness only) between plant treatments indicate significant differences.
Treatments not connected by arrows are not significantly different
166
Figure 3.7: NMS of bacterial community structure associated with single- (pine, birch or
lingonberry) and double- (pine + lingonberry) plant systems (stress = 16.42;
instability = 0.06)
167
Figure 3.8: PCA ordination of areas under C substrate curves showing the relative
influence of PHC, organic soil and plant treatment variables. Plant effects are
indicated by different symbols

169

Figure 4.1: Vertical distribution of ECM morphotypes (Cen, Cenococcum; Amp,
Amphinema; Lac, Lactarius; Rus, Russulaceae; Rh-Sl, Rhizopogon-Suillus 1; Rh-S2,
Rhizopogon-Suillus 2; MRA; E-strain), ECM/ ERM gentotypes, and soil properties
(%C, %N, pH) in PHC-treated and untreated organic (FH or CWD), Ae and Bf soil
layers of a pine-lingonberry system
198
Figure 4.2: NMS ordination of fungal community structure by plant (pine or lingonberry)
and organic (FH and CWD) and mineral (Ae, Bf) soil layers (stress = 18.98;
instability = 0.08)
200
Figure 4.3: NMS ordination of fungal community structure by soil layer in the two soil
systems (FH-Ae-Bf and CWD-Ae-Bf) (stress = 18.98; instability = 0.08)

202

Figure 4.4: Pairwise comparisons of fungal community structure (genotypes) in pine
(ECM) and lingonberry (ERM) groups: a) within soil layers; b) within and between
layers of FH and CWD soil systems. Significant differences (MRPP) are represented
by dark arrows and corresponding p-values; light (slashed) arrows represent no
differences between groups
204
Figure 4.5: NMS of bacterial community structure associated with plant (pine or
lingonberry) in organic (FH or CWD) and mineral soil layers (stress = 15.96;
instability = 0.06)
Figure 4.6: Pairwise comparisons of bacterial community structure (genotypes) in all
plant systems (P+L), as well as in pine (ECM) and lingonberry (ERM) groups: a)
IX

206

within soil layers; b) within and between layers of FH and CWD soil systems.
Significant differences (MRPP) are represented by dark arrows and corresponding pvalues; light (slashed) arrows represent no differences between groups
208

x

Acknowledgements
I could not have finished this thesis without the support of many people. I especially thank
my co-supervisors, Hugues Massicotte and Mike Rutherford, for contributing so much of
their time to discussing ideas and data, reading and editing papers, and supporting my
emotional break-downs. I also thank my supervisory committee, Bill McGill, Paul Sanborn,
Keith Egger, and Shannon Berch, for their valuable contributions to the design, techniques,
analysis, and interpretations of this research, as well as insightful comments on earlier drafts
of this work. I am additionally indebted to Alex Hawley for helping me to connect the dots.
I thank fellow fungal researchers, Nabla Kennedy, Kei Fujimura, Monika Gorzelak, Brian
Pickles, and Jocelyn Campbell, for friendship and good advice in multivariate statistics,
science, and life. I am particularly grateful to Linda Tackaberry for her sympathetic ear,
regular coffee breaks, editorial suggestions, and guidance through exhausting administrative
mazes. My gratitude, also, to the people who provided materials or helped with technical
aspects of this work: Dawn Stubley (Tree Seed Center, Surrey), Barb Rayment (Birch Creek
Nursery, Prince George), Ralph Alms (Husky Inc., Prince George), Steve Storch and John
Orlowsky (Enhanced Forestry Lab), Quanji Wu (PHC analysis), Anna Scarpino, Allen Esler,
and Clive Dawson (soil analysis), and Dana O'Bryan and Mark Thompson (DNA analysis).
I am also very grateful for my extremely understanding family and friends for still talking to
me after several years of sporadic contact. Special thanks to Andy Kendrick, who has
vicariously endured eight years of grad school in eight years of marriage, and Misha, for
hanging in there.
Finally, this research would not have been possible without financial support provided by the
Natural Sciences and Engineering Research Council of Canada through grants to Hugues
Massicotte and Mike Rutherford and a PGS-B scholarship to me. I am also very grateful to
the University of Northern British Columbia for providing laboratory facilities and travel
grants for conferences.

XI

Introduction
The ubiquity and enormous biomass of mycorrhizal systems (i.e. plant-fungal symbioses and
bacterial communities associated with the mycorrhizosphere) in northern forest soils implies
a key role in forest ecosystem processes. Although recent studies have revealed enormous
taxonomic and genetic diversity of mycorrhizal communities associated with certain plants in
some ecosystems, the specific activities of individual or groups of taxa and their
contributions to processes across heterogeneous landscapes are not well understood. The
complexity of the soil environment and the multifunctional nature of many microorganisms
have made it difficult to predict how ecosystems may respond to environmental disturbances
such as soil contamination with crude oil. Very little is currently known regarding the fate
and impacts of petroleum hydrocarbon (PHC) contaminants in forest soil ecosystems.

The purpose of this research was to examine interactions between PHCs and mycorrhizal
communities in plant-soil systems, where different guilds of microorganisms interact within
the larger trophic group tightly linked to decomposition, carbon and nutrient cycling, and
primary production. The microbial groups assessed in the current study include ecto- (ECM)
and ericoid (ERM) mycorrhizal fungi and associated bacterial communities. A combination
of methodological approaches were simultaneously used to gain a better understanding of the
physical, chemical and biological changes that occurred in the rhizosphere following
contamination with ecologically relevant (i.e. equivalent to several tonnes per hectare) levels
of oil. The specific objectives of each study are summarized in the following paragraphs for
each of the five thesis chapters. All chapters are written in the first person plural in
recognition of contributions of others to this work.

1

Chapter 1 is a review of literature from several different fields, including botany, mycology,
microbial ecology, soil science, and environmental toxicology. This synthesis is presented
from the perspective of mycorrhizal systems, which represent the structural and functional
interface between decomposition and primary production processes in northern forest
ecosystems. It is an attempt to form linkages between the often disparate bits of knowledge
that contribute to the current understanding of the fate of PHCs in boreal forest soils in both
ecological and toxicological contexts. The literature review helped to inform the direction of
research presented in the subsequent chapters. This paper was published in Biological
Reviews (Robertson et ah, 2007).

A fundamental question is addressed in Chapter 2: does PHC contamination of forest soils
negatively impact established (or establishing) mycorrhizal systems? We hypothesized that
the structure (composition) and diversity (richness and relative abundance) of ECM/ ERM
fungal and associated bacterial communities would be altered in response to PHC treatment,
either via soil habitat changes, chemical toxicity, or altered C or nutrient regimes. We further
expected that systems with different host plant (e.g. root structure and exudation patterns) or
soil properties (e.g. previous PHC contamination history or high lignin content) would
contain mycorrhizal communities adapted to PHC contamination conditions to varying
extents. Using a bioassay (single- and double-plant systems established in reconstructed
forest soil layers in pots), morphological and molecular (LH-PCR) techniques were used to
assess changes in mycorrhizal community structure in response to different PHC
concentrations over 16 weeks. Laccase assays were also conducted to assess changes in

2

enzyme activity associated with PHC treatment. Results of these assessments are discussed
in terms of ecological integrity and remediation/ restoration.

Chapter 3 focuses on the indirect role of the mycorrhizosphere in the biodegradation process
(i.e. provision of habitat for consortia of bacteria involved in metabolizing PHCs) and
examines effects of PHCs on structure and metabolic function of bacterial communities. We
hypothesized that indigenous bacterial communities would have the capacity for PHC
biodegradation, at least for the smaller fraction PHCs, and that this capacity would be
enhanced in the presence of mycorrhizal root systems that provide C substrates (exudates) for
co-metabolism. In addition, we expected that the mycorrhizosphere would provide some
protection of bacterial communities from potentially toxic effects of some PHC chemicals.
Single-, double- and no-plant systems were treated with the highest PHC level (i.e. 219 mg
cm"2) tested previously. GC-FID analysis was used to determine concentration of PHCs (in 4
fractions) in soil layers at 1 and 16 weeks; soil layers were analyzed for C:N ratio and pH.
Genotypic profiles of root-associated bacterial communities were assessed using LH-PCR
analysis; metabolic profiles were based on C substrate use after 7 weeks. These results are
discussed in terms of the mycorrhizosphere effect, functional redundancy, and syntrophic
biodegradation of PHCs.

In Chapter 4, the relative contributions of plant and soil properties to spatial distribution
patterns of ecto- (ECM) and ericoid (ERM) mycorrhizal fungi as well as root-associated
bacterial communities inhabiting the shared rhizosphere of pine (ECM host) and lingonberry
(ERM host) were investigated in PHC-treated and control systems. Community profiles (i.e.

3

based on the relative abundance of all genotypes) were generated for all root systems using
LH-PCR and primers targeting the ITS (fungi) and 16S (bacteria) regions of ribosomal DNA.
ECM composition, relative abundance and variation between soil layers were also assessed
using light microscopy. Spatial distribution patterns of the three groups of microorganisms
(i.e. ECM and ERM fungi and associated bacteria) are compared and discussed with respect
to habitat and niche considerations in the plant-soil systems.

Final conclusions and considerations for future research are presented in Chapter 5. It is a
general discussion along several themes that emerged during analyses in the three data
chapters, including the rhizosphere as an ecological unit, system resilience to disturbance,
syntrophic biodegradation of PHCs, functional redundancy within and between trophic
groups, importance of scale, variation and uncertainty in ecological data, and ecological
foundations for sustainable forest management.

Robertson, S.J., McGill, W.B., Massicotte, H.B. and Rutherford, P.M. (2007) Petroleum
hydrocarbon contamination in boreal forest soils: a mycorrhizal ecosystems perspective.
Biological Reviews 82: 213-240.

4

Chapter 1: Petroleum hydrocarbon contamination in boreal forest soils: a mycorrhizal
ecosystems perspective

Abstract
The importance of developing multi-disciplinary approaches to solving problems relating to
anthropogenic pollution is now clearly appreciated by the scientific community, and this is
especially evident in boreal ecosystems exposed to escalating threats of petroleum
hydrocarbon (PHC) contamination through expanded natural resource extraction activities.
This review aims to synthesize information regarding the fate and behaviour of PHCs in
boreal forest soils in both ecological and sustainable management contexts. From this, we
hope to evaluate potential management strategies, identify gaps in knowledge and guide
future research. Our central premise is that mycorrhizal systems, the ubiquitous root
symbiotic fungi and associated food-web communities, occupy the structural and functional
interface between decomposition and primary production in northern forest ecosystems (i.e.
underpin survival and productivity of the ecosystem as a whole), and, as such, are an
appropriate focal point for such a synthesis. We provide pertinent basic information about
mycorrhizas, followed by insights into the ecology of ecto- and ericoid mycorrhizal systems.
Next, we review the fate and behaviour of PHCs in forest soils, with an emphasis on
interactions with mycorrhizal fungi and associated bacteria. Finally, we summarize
implications for ecosystem management. Although we have gained tremendous insights into
understanding linkages between ecosystem functions and the various aspects of mycorrhizal
diversity, very little is known regarding rhizosphere communities in PHC-contaminated soils.
This makes it difficult to translate ecological knowledge into environmental management
strategies. Further research is required to determine which fungal symbionts are likely to

5

survive and compete in various ecosystems, whether certain fungal - plant associations gain
in ecological importance following contamination events, and how PHC contamination may
interfere with processes of nutrient acquisition and exchange and metabolic processes.
Research is also needed to assess whether the metabolic capacity for intrinsic decomposition
exists in these ecosystems, taking into account ecological variables such as presence of other
organisms (and their involvement in syntrophic biodegradation), bioavailability and toxicity
of mixtures of PHCs, and physical changes to the soil environment.

Key words: ectomycorrhiza, ericoid mycorrhiza, mycorrhizal ecosystems, boreal forest soils,
ecosystem processes, petroleum hydrocarbons, soil pollution, biodegradation,
bioremediation.

6

Introduction
Boreal and sub-boreal forest ecosystems include arctic, sub-arctic and northern mid-latitude
forest regions that are dominated by a cold climate and are able to support only a few
coniferous and broadleaf tree genera (Burton et al., 2003). Petroleum hydrocarbons (PHCs)
are complex mixtures of aliphatic, alicyclic and aromatic compounds (Miller and Herman,
1997; Potter and Simmons, 1998) plus constituents that contain N, S or O in addition to H
and C. PHCs may find their way into terrestrial ecosystems by surface spills or leaks from
pipelines or storage tanks. The microbial ecology of boreal forest ecosystems, with or
without PHCs, is incompletely understood. It is known, however, that symbiotic fungi
colonize and extend beyond the roots of dominant plant species, thereby forming an
intimately interwoven belowground mycorrhizal system. Mycorrhizal fungi account for most
of the microbial biomass in organic soil horizons (Lundstrom et al., 2000; Dahlberg, 2001).
The traditional role of individual symbioses involves the exchange of soil nutrients for
carbohydrates fixed through plant photosynthesis (Smith and Read, 1997). Nutrients are
obtained from inorganic sources inaccessible to plants or accessible, but more readily
obtained, by the mycobiont. However, some mycorrhizal systems appear to possess welldeveloped saprotrophic capabilities (i.e. oxidative and hydrolytic enzyme systems) that
mobilize nutrients from organic sources. Such capabilities may have developed through
selection in ecosystems characterized by slow decomposition and retention of nutrients in
organic polymers (Hibbett et al., 2000; Burke and Cairney, 2002; Cairney and Meharg, 2002;
Read and Perez-Moreno, 2003). Mycorrhizal systems capable of metabolizing exogenous
organic compounds therefore may be candidates for use in remediation of soils contaminated
with PHCs.

7

Mycorrhizal fungal mycelia and surrounding soil (i.e. mycorrhizosphere) provide suitable
habitats for diverse communities of microorganisms due to increased availability of highenergy metabolic substrates and surfaces for colonization (Sarand et al., 2000; Sen, 2003;
Heinonsalo et al., 2004). This enhances bacterial decomposition of plant materials because
mycelia provide a path, together with associated water films, through which bacteria can
migrate to substrates in micropores. Metabolic synergism between fungal and bacterial
members of soil communities ensures that virtually all organic compounds are subject to
biotransformation (if available to decomposer organisms) and that nutrients and energy-rich
compounds are exchanged between plants and the soil environment via mycorrhizal fungal
networks (Simard et ah, 1997; Read and Perez-Moreno, 2003; Diaz, 2004; Heinonsalo et al.,
2004). Consequently, in addition to their direct transformation of organic compounds,
mycorrhizal systems may indirectly enhance degradation of PHCs in soil by modifying the
structure of associated bacterial communities (Cairney and Meharg, 2002).

Oil extraction, refinement and transportation activities in boreal regions have resulted in
surface and near-subsurface soil contamination with PHCs including crude (or synthetic
crude) oil, gasoline, diesel and creosote (Kanaly and Harayama, 2000). The current standard
against which environmental impacts are evaluated is sustainability (maintenance of
ecological integrity) using various ecological indicators as measures. Sustainability requires
management strategies for large areas and long periods of time that satisfy diverse
environmental, social and economic needs (Burton et al, 2003). The relationship between
soil microbial communities and ecosystem processes (e.g. decomposition and

8

biogeochemical cycling) provides insights into how communities and ecosystems respond to
environmental change. Microbial diversity (the variety of taxonomic, genetic and functional
characteristics of organisms) helps sustain terrestrial ecosystems by conferring ecosystem
stability (the ability to withstand change), resilience (the ability to recover from change) and
resistance (the inherent capacity to withstand disturbance) (Andren and Balandreau, 1999;
Tiedje et al, 1999; Nannipieri et al, 2003; Swaminathan, 2003; Fitter et al., 2005). Lower
diversity or higher specialization occurs in disturbed soil systems due to: (1) extinction of
populations that lack sufficient tolerance to the change imposed, and/or (2) selective
enrichment of populations that tolerate or thrive under the new conditions (Diaz, 2004;
Hofman et al., 2004). To understand the basis of community differences associated with
changes in environmental conditions, it is necessary to integrate the functional properties and
environmental requirements or tolerances of communities with processes at an ecosystem
level (Bengtsson, 1998; Cairney, 1999; Dahlberg, 2001; Read and Perez-Moreno, 2003).

The potential toxicity of some PHCs to human, plant and animal receptors is used in
managing contaminated sites, but the physical, chemical and biological impacts on soil
microbial communities are less extensively studied and used (Miller and Herman, 1997;
Nicolotti and Egli, 1998). Controlled experiments have provided valuable information
regarding the toxicological impacts of chemicals on test organisms, which forms the
scientific basis for current remediation standards. In soils, toxicity of PHCs to soil organisms
including plants occurs concurrently with physical and chemical changes to the soil habitat
following PHC contamination (Tarradellas and Bitton, 1997; Blakely et al., 2002; Trofimov
and Rozanova, 2003). Is it possible to separate the effects of chemical toxicity from habitat

9

changes such as hydrophobicity, lowered redox potential or reduced nutrient supply in PHCcontaminated soils? Are methods available for assessing the fate and behaviour of PHCs in
forest soils that include bioavailability and indicators for ecological integrity that also
complement measures for plant productivity? In addition, many PHCs are structurally
analogous to organic compounds naturally found in the soil environment and appear to be
degraded by soil microbial communities using the same biochemical pathways (McGill et al.,
1981; Siciliano and Germida, 1998). Can functional aspects of microbial populations and
communities (e.g. exocellular enzymes) be manipulated for bioremediation of contaminated
soil?

Numerous reviews have addressed various aspects of mycorrhizal systems (e.g. Meharg and
Cairney, 2000; Dahlberg, 2001; Burke and Cairney, 2002; Allen et al, 2003; Read and
Perez-Moreno, 2003; Fitter et al., 2005) or of PHC behaviour and biodegradation in soil
(McGill et al, 1981; Riser-Roberts, 1998; Alexander, 1999, 2000; Prince and Drake, 1999;
Diaz, 2004; Stokes et al, 2005; Rombke et al, 2006). How might we advance the
understanding of the fate and behaviour of PHCs in boreal forest soils in both ecological and
sustainable management contexts? We aim to do so by synthesizing information regarding
the interactions between mycorrhizal communities and PHC contaminants in boreal soils.
From this, we hope to evaluate potential management strategies, identify gaps in knowledge
and guide future research. Our central premise is that mycorrhizal systems occupy the
structural and functional interface between decomposition and primary production in
northern forest ecosystems and as such are an appropriate focal point for such a synthesis.
Information in this synthesis should be useful to professionals ranging from ecologists to

10

engineers involved in the management and remediation of contaminated boreal forest soils.
Our approach is first to provide pertinent basic information about mycorrhizas, followed by
insights into the ecology of ecto- and ericoid mycorrhizal systems. Next we review the fate
and behaviour of petroleum hydrocarbons in forest soils, with an emphasis on interactions
with mycorrhizal fungi and associated bacteria. Finally, we summarize implications for
ecosystem management.

Mycorrhizas
Classification and structure
Mycorrhizas are symbioses between plant roots and an array of soil-inhabiting, filamentous
fungi. These associations are virtually ubiquitous and generally considered mutualisms (i.e.
reciprocally increase the fitness of both partners) as they are based on a bidirectional
exchange of nutrients that is essential to the growth and survival of both partners (Smith and
Read, 1997; Peterson and Massicotte, 2004; Sapp, 2004). The fungal partner acquires
nitrogen (N), phosphorus (P) and other nutrients from the soil environment and exchanges
them with the plant partner for photosynthetically derived carbon (C) compounds that fuel
fungal metabolism. The structural attributes of mycorrhizas are related to their primary
function of nutrient exchange and provide the basis for broad classification into seven
currently recognized groups: ectomycorrhizas, ericoid mycorrhizas, ectendomycorrhizas,
arbuscular mycorrhizas, arbutoid mycorrhizas, monotropoid mycorrhizas and orchid
mycorrhizas (Peterson et ah, 2004). In boreal forest ecosystems, most trees typically form
ectomycorrhizal (ECM) symbioses, whereas the major constituents of the understorey

11

vegetation often form arbuscular (AM), ericoid (ERM) or arbutoid (ARM) mycorrhizas. The
ECM and ERM groups will be considered herein in the greatest detail.

Ectomycorrhizas, the associations between ECM fungi and the roots of woody plants, are
characterized by three structural components: the mantle, the Hartig net and the extraradical
mycelium (Smith and Read, 1997). The mantle is a sheath of fungal tissue that covers the
highly active tips of the lateral roots of the plant and forms the boundary between the root
and the soil environment. Its compact, but also variable, morphological nature provides a
buffering capacity that helps to prevent root cell dehydration or penetration by pathogenic
organisms (Brundrett, 1991). Fungal cells (hyphae) emanate from the outer mantle as
extraradical mycelia and grow into the surrounding soil where they reach micropore areas
and absorb nutrients that may otherwise be inaccessible, both physically and biochemically
(i.e. enzymatic processing of organic compounds), to roots (Soderstrom, 1992; Perez-Moreno
and Read, 2000). Some ECM fungi also form rhizomorphs, which are thick linear aggregates
of hyphae that are specialized for long-distance translocation of nutrients and water (Agerer,
2001). Lipids, phenolic compounds, proteins and polyphosphates may accumulate in the
hyphae of the outer mantle, which may also bind heavy metals and thereby prevent their
uptake into roots (Peterson et ah, 2004). The inner mantle consists of repeatedly branched
hyphae, suggesting a role in nutrient exchange such as enabling absorption of glucose and
fructose from the root and conversion to fungal sugars (e.g. trehalose, mannitol or glycogen)
(Peterson et ah, 2004). At the interface of nutrient exchange is a highly branched structure
known as the Hartig net, which is formed by multidigitate growth of fungal hyphae between
epidermal and cortical cells of the root, and is the probable site for exchange of resources

12

between symbionts (Peterson et al., 2004). Subtle variations in morphological attributes
viewed using light microscopy are often used to distinguish between ECM fungal taxa;
development and differentiation of extraradical mycelia may provide predictive features
relevant to the ecological classification of ECMs (Agerer, 1987-2002, 2001).

The common feature of plants that form ericoid mycorrhizas is the formation of very fine
lateral roots that are composed of a vascular cylinder, one or two rows of cortical cells and an
epidermal layer of enlarged cells (Peterson et al., 2004). ERM fungi do not form mantles or
Hartig nets, but rather colonize the epidermal cells of these fine roots and develop
intracellular hyphal coils that are specialized for nutrient exchange (Peterson et al., 2004).
The intracellular fungal symbiont is separated from the plant cytoplasm by a plant-derived
membrane, which invaginates to follow fungal growth and coil formation (Perotto et al.,
2002). ERM fungal taxa cannot be distinguished by morphological characters using light
microscopy. From molecular studies, it appears that ERM roots are composite structures that
house multiple fungal symbionts, which implies that epidermal root cells may potentially
function as separate units colonized by a variety of fungi (Perotto et al., 2002).

Diversity
Mycorrhizal symbioses have been an important force in evolution (Pirozynski and Malloch,
1975; Blackwell, 2000; Cairney, 2000; Sapp, 2004). Based on reconstructions of
evolutionary lineages (phylogenies) from fungal DNA and the fossil record, it is currently
accepted that the first mycorrhizal associations were pivotal in allowing plants to colonize
the terrestrial environment about 600 million years ago and they form the evolutionary basis

13

of present plant communities (Pirozynski and Malloch, 1975; Blackwell, 2000). Redecker et
al. (2000) reported fossilized fungal hyphae and spores found from the Ordovician of
Wisconsin (about 460 million years old) that strongly resemble modern Glomerales-like AM
fungi. Modern AM fungal species persist in most extant plant species and form a single
monophyletic group descended from these first mycorrhizas (Cairney, 2000). The AM group
represents four orders (Archaeosporales, Paraglomerales, Diversisporales and Glomerales) of
fungi within the phylum Glomeromycota (Smith and Read, 1997).

ECM fungal diversity appears to have arisen about 200 million years ago, corresponding to
changes in climate that allowed for colonization of the land with trees and increased organic
matter content of some ancient soils (Cairney, 2000). Although ECM plant partners
(phytobionts) represent only about 8000 species (mostly in the families Pinaceae, Betulaceae,
Fagaceae, Dipterocarpaceae, Salicaceae and Myrtaceae), these species are of global
importance because of their disproportionate occupancy and domination of terrestrial
ecosystems in boreal, temperate and subtropical forests (Smith and Read, 1997). It has been
estimated that 5000-6000 species of fungi (of the sub-divisions Basidiomycotina,
Ascomycotina and Zygomycotina) form ECM symbioses (Molina et ah, 1992; Horton and
Bruns, 2001), but these numbers are expected to rise as more regions are progressively
explored in detail (Cairney, 2000). Phylogenetic analyses reveal that ECM fungi have
originated from several independent lineages and that symbiosis with plants has been
convergently derived (and perhaps lost) many times over millions of years (Hibbett et al.,
2000). Some ECM taxa are closely related to, or descended from, wood-rot fungi and some
are related to other saprotrophic fungal taxa (Tanesaka et ah, 1993; Hibbett et al., 2000).

14

This variation in the ability to degrade wood may have helped drive fungal speciation to
avoid competition between closely related species that would otherwise use the same
resources and occupy the same niche (Tanesaka et al, 1993; Bruns, 1995; Martin et ah,
2000). The ability to degrade the complex aromatic chemical structures of lignin in wood
may also confer an ability to transform similar structures in PHCs.

ERMs evolved about 100 million years ago, as sclerophyllous vegetation (i.e. plants with
small, tough foliage and tissues that are rich in lignin and cellulose, but deficient in N and P)
emerged in nutrient-poor soils (Cairney, 2000). Many plants of the family Ericaceae (e.g.
Vaccinium, Rhododendron, Gaultheria, Ledum species) are common components of the
understorey vegetation in northern forests and usually form typical ERMs (Vralstad et ah,
20026). In the Southern hemisphere, plant species of the family Epacridaceae form ERMs
(Cairney and Ashford, 2002). ERM fungi were thought to belong to the Ascomycotina, of
which fungal strains in the Rhizoscyphus ericae - Scytalidium vaccinii species complex
(Helotiaceae, Helotiales, Ascomycota) are most commonly studied and reported (Vralstad et
al., 2002a; Zhang and Zhuang, 2004). In addition, ERM fungi identified as Oidiodendron
(anamorphs of the ascomycete family Myxotricaceae) as well as a broad range of sterile
mycelia with divergent morphologies and unknown identifications have been described
(Vralstad et al, 2002<2). Recent morphological (clamped hyphae and dolipore septae
forming typical ERM coils on Vaccinium, Rhododendron and Gaultheria species) and
molecular (rDNA sequences) evidence indicates that some ERM fungi may belong to the
Basidiomycotina (Berch et ah, 2002; Perotto et ah, 2002). It has become increasingly

15

apparent that a wider spectrum of taxa is involved in the ERM symbiosis than had been
previously imagined.

ECM communities appear to consist of large numbers of fungal species (i.e. exhibit high
species richness), even within small areas with little heterogeneity in plant communities, soil
properties, climate and disturbance patterns (Bruns, 1995; Kranabetter et al, 1999; Taylor et
al, 2000; Mah et al., 2001; Robertson et al., 2006). ERM fungal communities also appear to
exhibit high richness. For example, Monreal et al. (1999) isolated 20 fungi (five of which
formed ERM in vitro) from sixty segments (each 3 mm long) of fine roots from an 8-cm-long
salal {Gaultheria shallon Pursh) rhizome. This ERM fungal richness is consistent with other
reports of species-rich communities of mycorrhizal and non-mycorrhizal endophytes in
individual root systems of other ericaceous [e.g. Calluna vulgaris (L.) Hull] and
epacridaceous [e.g. Woollsiapungens (Cav.) F. Muell.] plants. All groups of ericoid fungi
reported globally have been found associated with salal from a single site on Vancouver
Island (British Columbia, Canada) and all ERM groups reported on salal have been found
associated with other plant species elsewhere in the world (Berch et al., 2002). It is currently
hypothesized that sterile mycelia with ERM behaviour represent a heterogeneous group of
fungal taxa that are mostly unidentified and appear to include a variety of unculturable
mycobionts (Berch et al., 2002; Perotto et al., 2002). High species richness and abundance
may represent ecological adaptation to local environmental heterogeneity and is thought to
provide forests with a range of strategies to maintain efficient functioning under an array of
environmental conditions (Cairney, 1999; Nannipieri et al, 2003).

16

Establishing whether diversity is important for ecosystem processes has become a central
issue in ecology (Leake, 2001). In general, soil microbial communities appear to comprise
groups of organisms that fulfill broadly similar ecosystem functions (i.e. exhibit functional
redundancy) (Yin et al., 2000). Functional diversity represents the value and range of
capabilities that are possessed by organisms present in a given ecosystem and are relevant to
ecosystem processes (Allen et al., 2003; Sobek and Zak, 2003). There is a growing body of
evidence suggesting that the functional characteristics of component taxa are at least as
important as species richness for maintaining essential ecosystem processes (Naeem, 2002;
Nannipieri et al., 2003). Knowledge of the individual roles of mycorrhizal fungal species, or
of their distribution either in relation to each other or to the physical and chemical
environments of the soil, is limited (Goodman and Trofymow, 1998; Rosling et al., 2003)
and insufficient for determination of community needs and responses by building up from the
species level. Moreover, in mycorrhizal ecosystems, we hypothesize that the functional
significance of individual taxa is overshadowed by the integrated functional capability of the
community, which is likely not an additive function of the independent capabilities of
component species. The tendency to generalize ecological functions from a few fungal
isolates reveals little information about the intrinsic physiological potential of most taxa
(Cairney, 1999; Cairney and Meharg, 2003) or of the community. Current evidence suggests
that ongoing parallel evolution of plant and fungal partners in response to environmental
change on local and global scales may most readily explain extant patterns of mycorrhizal
diversity and specificity (Cairney, 2000). Although functional redundancy almost certainly
exists within mycorrhizal communities, high taxonomic and genetic diversity of ECM (and

17

probably ERM) fungi may indicate that they also exhibit a high level of functional
heterogeneity (Cairney, 1999).

Are all the pieces of an ecosystem essential for restoration? Following a disturbance, should
the management target be to maintain (or reintroduce) the original species richness at all
costs, or, alternatively, to nurture the survivors (stress-resistors) so that they can contribute to
the restoration of habitats in a future (altered) state? Is it likely that resistant organisms will
modify the environment in ways that favour only themselves (i.e. preserving a specialized
community), or do modifications lead eventually to succession by organisms that are
incapable of tolerating the initial conditions (as suggested by most concepts of ecological
succession)? Species richness may be a critical aspect of ecosystem resilience and
functioning, but within a restoration context, more emphasis should perhaps be devoted to
the resistant biota and their contribution in restoring pre-contamination conditions.
Community specialization may indicate environmental stress, but we hypothesize
specialization may also be a desirable response to stress, and a useful characteristic in
allowing stressed ecosystems to achieve long-term stability and diversity.

Ecology of ecto- and ericoid mycorrhizal systems
Soil habitat
Soils are living, open, dynamic systems. They contain structured and heterogeneous
matrices, generally store nutrients and energy, and support high microbial diversity and
biomass (Nannipieri et al., 2003). To thrive, soil microorganisms must mobilize energy and
nutrients stored in soil. Soil structure provides a complex and variable set of microbial

18

habitats ranging from energy-rich to barren, or aerobic to anaerobic, over micrometre
distances. Soil structure is determined by soil aggregation, which occurs when soil particles
within aggregates cohere more strongly to each other than to adjacent aggregates (Hartel,
1998). Aggregates are composed of sand, silt and clay particles that are held together by
organic matter, precipitated inorganic materials, microorganisms and the products of their
metabolic activities (Griffiths and Caldwell, 1992; Hartel, 1998). Aggregates are dynamic,
constantly forming and disintegrating. Organic substrates and plant residues are entrained
and protected during aggregate formation and released during aggregate disintegration
(Plante and McGill, 2002). The solid phase adsorbs important biological molecules (e.g.
DNA, enzymes, etc.) and many soil reactions are catalyzed at the surfaces of soil minerals
such as clays, Mn (III and IV) oxides and Fe (III) oxides (Nannipieri et ah, 2003). In
addition, the zeta potential of charged mineral and organic surfaces generates a steep pH
gradient around them. For example, McLaren and Skujins (1968) cite examples of the pH
optima of enzymes being several units higher in colloidal systems than in solution,
apparently due to the lower pH in the immediate environment of the enzyme, close to
colloidal surfaces. Water occupies the aggregate pore spaces and forms a meniscus around a
central pocket of air, which provides an aerobic and aqueous habitat suitable for supporting
bacterial communities (Wardle, 2002). Pore water also retards gas exchange, thereby
creating anaerobic microsites. Pore water also participates in hydrolysis and mediates other
soil reactions (Hartel, 1998).

Boreal forest soils are typically acidic with seasonal or intermittent availability of mineral
nutrients (N and P) and high C:N ratios due to the surface accumulation of recalcitrant

19

organic matter resulting from incomplete oxidation of plant material (Prescott et ah, 2000;
Allen et ah, 2003). This organic layer (mor humus) stores nutrients and also contributes to
moisture retention and soil structure (Prescott et ah, 2000). The forest floor is the most
metabolically active fraction of these soils and is heavily colonized by ECM and ERM root
systems of trees and understorey vegetation (Lundstrom et ah, 2000). Wallander et ah
(2001) estimated the extraradical mycelia biomass of ECMs to represent about 820 kg ha"1 in
boreal forest soils. Fungal metabolic activities produce organic acids that percolate with rain
water down through the soil profile and contribute to accelerated weathering of mineral soils
(Griffiths and Caldwell, 1992; Heinonsalo et ah, 2004). Soluble complexes are formed
between the organic acids and Fe and Al ions in the upper mineral soil, thereby fostering
leaching of Fe and Al ions and creating a weathered, eluvial horizon (Lundstrom et ah,
2000). These complexes percolate further downward and precipitate, creating a
characteristic rust-coloured illuvial B horizon overlying the parent material (Lundstrom et
ah, 2000). These changes with depth in soil chemical and mineralogical properties create
contrasting habitats for microorganisms. For example, Rosling et ah (2003) found that the
species composition of the ECM community varied between organic and mineral horizons of
boreal podzolic soils and that most taxa occurred in only one part of the soil profile.

Less than 5% of the soil volume is occupied by microorganisms, but these sites of increased
biological activity are where the majority of soil reactions are mediated (Diaz, 2004). The
availability and nutrient content of organic matter are key factors influencing microbial
biomass and community composition (Tiquia et ah, 2002). Other major factors controlling
the distribution and abundance of soil microbial communities include: (1) properties of the

20

soil environment (e.g. pH, O2 supply and availability of water and nutrients such as N, P, Fe);
(2) factors affecting dispersal (e.g. soil structure, micro-aggregate stability and routes of
dispersal); and, (3) the controls of population turnover (e.g. nematode or protozoan grazing,
controls on lytic enzymes, protective soil matrices) (Tiedje et ah, 1999). Introduction of
PHCs alters all three of these fundamental characteristics. For example, O2 supply is often
reduced, water movement is restricted and soil fauna including nematodes and protozoa are
temporarily lost from the contaminated ecosystem.

Microbial growth in soils is typically resource-limited (most often energy-limited) and
increases rapidly in response to addition of reduced C to provide energy for the large chemoorganotrophic biomass (Nannipieri et ah, 2003; Morgan et ah, 2005). Actively growing
roots leak or secrete (exude) soluble and insoluble organic compounds into the surrounding
soil (rhizosphere) that provide most of the low molecular weight C available to
microorganisms (Darrah, 1991; Garbaye, 1994). Rhizodeposition is concentrated at the root
tips and at sites of lateral branch formation, which correspond to sites of greater microbial
population density and community complexity compared to bulk soil (Linderman, 1988;
Chanway, 1997; Sarand et ah, 2000; Soderberg et ah, 2004). Soluble forms of C (e.g.
monosaccharides, amino acids and organic acids) are readily metabolized by microorganisms
to CO2 or converted to biomass; insoluble forms of C (e.g. mucilages, sloughed cortical cells
and dead root hairs) are less readily metabolized (Darrah, 1991), but they may form new
microbial habitats, which are eventually consumed. As with fungi, bacterial richness and
functional redundancy are both high, at least at coarse scales. Using fatty acid methyl ester
profiles (FAME analysis) and 16S rRNA gene sequences, Axelrood et ah (2002a, b)

21

described immense bacterial richness (isolates representing 42 known bacterial genera and
clones spanning nine divisions, respectively) in surface organic matter and mineral soil
samples from forests in the central interior of British Columbia. Culture collections were
well represented by Pseudomonas, Bacillus, Paenibacillus and Arthrobacter species
(Axelrood et al., 2002a), whereas molecular clones were represented by Bradyrhizobium,
Rhizobium, Pseudomonas and Burkholderia species (Axelrood et al., 20026). These genera
are considered common soil inhabitants and important components of rhizosphere
communities with respect to nutrient cycling and transformation of minerals and complex
organic substrates (Axelrood et al., 20026).

It has not yet been fully appreciated that the establishment of mycorrhizal symbioses
substantially alters the morphology and physiology of plant roots (e.g. alters permeability of
root membranes), which also changes root exudation patterns as well as the types of C
substrates exuded (Linderman, 1988; Ingham and Molina, 1991; Rygiewicz and Andersen,
1994). The extraradical mycelia generate increased volumes of mycorrhizosphere soil
compared to noncolonized roots and not only support microbial growth through exudation of
energy-rich substrates, but also provide surfaces for colonization and contribute to formation
of soil structure (Griffiths and Caldwell, 1992). The presence of ECM mycelia alters
bacterial community structure by stimulating proliferation of selected bacterial populations,
among other mechanisms (Frey et al., 1997; Heinonsalo et al., 2000). Fluorescent
pseudomonads isolated from the mycorrhizosphere of Douglas-fir (Pseudotsuga menziesii
(Mirbel) Franco) appear preferentially to use trehalose, a carbohydrate derived from fungal
metabolism (Frey et al, 1997). Fluorescent pseudomonads and actinomycetes have been

22

observed around ECM roots of birch, closely associated with the mantle and in proximity to
fungal exudates (Ingham and Molina, 1991). There is also some evidence that diverse
microbial communities may be selectively present in association with certain ECM mycelia
(Garbaye, 1994; Read and Perez-Moreno, 2003). For example, Olsson and Wallander (1998)
found that structure of soil bacterial communities, assessed using phospholipid fatty acid
(PLFA) analyses, depended both on ECM fungal species and soil type. Fungal mycelial
(mat) communities are unique soil habitats that contribute to maintenance of high richness of
bacterial and fungal taxa within ecosystems (Griffiths and Caldwell, 1992).

In summary, conditions within soil habitats vary by orders of magnitude over micrometre
distances, in response to physical (structure and aggregates), chemical (pH, 02, soluble
substances) and biological (microorganisms, soil fauna, plant roots) variables. Soil habitats
may also be substrates (e.g. plant residues). Perhaps because of this almost infinite variety of
habitats at the microbial-size scale, it is difficult to find any soil sample that is missing major
genera of the known microbiota of terrestrial ecosystems. Molecular techniques continue to
show increasingly large ranges of genetic material within soils (e.g. Axelrood et ah, 20026;
Prosser, 2002), with most (more than 99%) of the bacterial genotypes represented currently
not culturable (Pace, 2005). With greater sampling effort, the number of known bacterial
divisions has expanded substantially in recent years (Pace, 2005). From a management
perspective, the genetic potential to mediate virtually any biogeochemical reaction and the
habitat needed to support it appears to exist in most soils, with only specialized capabilities
potentially missing.

23

Community interactions
Mycorrhizosphere bacteria
As mycorrhizal fungi constitute the most significant rhizosphere communities, they have
immense potential for interactions with other soil organisms such as bacteria, fungi,
protozoa, nematodes, arthropods and mammals, as well as with each other (Fitter and
Garbaye, 1994; Read and Perez-Moreno, 2003; Cairney, 2005). The primary factors that
influence the composition of associated communities are the quality and quantity of C
compounds present, competitive interactions between mycorrhizal fungi and free-living
microorganisms for mineral nutrients, and the beneficial, detrimental or neutral impacts of
secondary metabolites produced by symbiotic or free-living organisms (Siciliano and
Germida, 1998; Cairney and Meharg, 2002). The outcomes of interactions between ECM,
ERM and saprotrophic fungal mycelia may include mutual interference of growth (deadlock)
or replacement of one taxon with another through competition (Cairney, 2005).

The interactions between ECM/ ERM fungi and the heterotrophic bacterial community are
important for accessing mineral nutrients (Burke and Cairney, 1998). Observations that
enhanced decomposition of organic compounds occurs in (mycor)rhizosphere soils have been
attributed to the greater metabolic activities associated with higher densities of
microorganisms (Heinonsalo et ah, 2000). Enriched bacterial communities, often arranged
as biofilms (organised systems consisting of layers of biologically active cells), have been
noted at the surfaces of the ECM fungal mantle and extraradical mycelia, which are the sites
of nutrient mobilization, uptake and translocation (Sen, 2003). The exposure of microbial
biofilms to organic polymers such as cellulose and proteins appears to drive degradative

24

secondary metabolism; this enables plant and microbial uptake of simple compounds (e.g.
sugars, amino acids and mineral nutrients) that are released during the decomposition process
(Sen, 2003).

From the germination of fungal propagules in soil to establishment of true symbiosis,
mycorrhizal fungi experience a free-living stage during which they interact with bacteria
(known as mycorrhizal helper bacteria, MHB) that appear to be beneficial to the colonization
process via one or more of several proposed mechanisms (Garbaye, 1994). In axenic culture
with nutrient limitation, MHB may act by direct trophic interactions (where bacteria provide
C substrates or growth factors to the free-living fungi) or by metabolic detoxification of
fungal metabolites (e.g. polyphenols, etc.) (Duponnois and Garbaye, 1990). Bacteria that are
active at the time of mycorrhizal formation may facilitate recognition between the plant and
mycorrhizal fungus, improve the receptivity of the root for fungal colonization, or stimulate
fungal growth, thereby increasing encounters between roots and mycelia (Frey-Klett et al.,
1997). MHB also appear to colonize fungal hyphae and stimulate initial mycorrhizal
formation through production of vitamins, amino acids, phytohormones and/ or cell wall
hydrolytic enzymes, which may influence germination and growth rates of fungal structures,
enhance root development and/or decrease susceptibility to infection (Martin et al., 2000).
Shishido et al. (1996) found that three strains of fluorescent pseudomonads enhanced spruce
seedling growth through mechanisms unrelated to increased mycorrhizal colonization, but
growth promotion of pine by two strains was facilitated by an interaction with mycorrhizas.
Mycorrhizal root tips tended to support slightly higher populations of Pseudomonas spp. than
non-mycorrhizal root tips and additional colonization sites or altered/ enhanced exudation in

25

the mycorrhizosphere were observed. Frey-Klett et ah (1997) found that high levels of
bacterial inoculum (MHB Pseudomonas fluorescens BBc6) in the rhizosphere are not
necessary for a helper effect to occur.

Another group of naturally occurring, free-living soil bacteria that colonize roots and
enhance plant growth when added to seeds and roots are known as the plant growth
promoting rhizobacteria (PGPR) (Chanway and Holl, 1991). PGPR activity has been
reported in Azospirillum, Bacillus, Clostridium, Hydrogenophaga, Serratia, Staphylococcus,
Streptomyces, and Micro bacterium species (Chanway, 1997). Holl and Chanway (1992)
found that growth of mycorrhizal pine was stimulated by inoculating the rhizosphere with
Bacillus polymyxa strain L6, which appeared to be a function of the size of the bacterial
population. Plant growth promotion was not attributed to increased symbiosis by the ECM
fungus Wilcoxina, and was also unlikely to be due to N fixation as this Bacillus strain
contributed to only 4% of seedling foliar N. Rather, stimulation of pine growth may have
been a result of bacterial production of plant growth substances such as indoleacetic acid.
Other researchers have suggested that PGPR may, at least in the short term, improve the C
supply to mycorrhizas by providing an increased supply of N (fixed from the atmosphere) to
the plant (Ingham and Molina, 1991; Martin et ah, 2000). Microorganisms may directly
stimulate plant growth by providing nutrients (e.g. N, P, S) or growth factors (e.g. auxin,
cytokinin, gibberellin), increasing root permeability or inducing plant systemic resistance to
pathogens. Indirectly, microorganisms may influence other rhizosphere components that
influence plant growth, such as increasing legume or alder root nodule number and size,

26

increasing colonization frequency of mycorrhizal fungi, or suppressing deleterious
rhizobacteria (Chanway, 1997).

Plant linkages
Plant communities in northern forest ecosystems are linked below ground via the extensive
extraradical mycelial network of mycorrhizal fungi (Dahlberg, 2001; Simard and Durall,
2004). Host-specific fungi form intraspecific plant linkages, whereas fungi with more
general host requirements may form interspecific linkages that allow for nutrient and C
transfer between different tree species. In a microcosm experiment, radiolabeled C transfer
through the soil mycelial network has been demonstrated between Sitka spruce [Picea
sitchensis (Bong.) Carr.] and pine species (Pinus contorta Dougl. ex Loud, and P. sylvestris
L.) (Finlay and Read, 1986). In the field, Simard et al. (1997) demonstrated bidirectional C
transfer between Douglas-fir and paper birch {Betula papyrifera Marsh.) via a common
mycelial network, with a significant net gain by the shaded Douglas-fir. Mycorrhizal
networks appear to have the capacity to mediate significant N transfer among interconnected
plants (Casuarina and Eucalyptus pairs); N gradients (between N-rich donors and N-limited
receivers) may drive unidirectional N transfer (He et al., 2005). Similarities in the
composition of ECM communities associated with various host species in bioassays and field
surveys indicate the potential for linkages between varieties of plant species (Kranabetter et
al, 1999; Massicotte et al, 1999).

The coexistence of ECM and ERM plants in boreal forests provides many opportunities for
sharing ECM and ERM fungi that link plants and translocate nutrients, although little

27

research on this issue has been conducted (Perotto et al., 2002). Vralstad et al. (2000)
demonstrated that fungal strains derived from ECM morphotype Piceirhiza bicolorata
constituted assemblages of very close relatives to ERM type Rhizoscyphus ericae. Similarly,
Monreal et al. (1999) showed sequence similarity (ITS2 region) between the ECM fungus
Phialophora finlandia andi?. ericae. In a resynthesis experiment using 12 R. ericae strains
on ECM and ERM hosts, Vralstad et al. (20026) showed that genetically close relatives of
the ERM fungus R. ericae are true ECM partners with conifer (spruce and pine) and
angiosperm (birch) species, but no isolates tested formed both ECMs and ERMs. These
studies indicate that ECM and ERM plants may share mycobionts of this species complex
(known as the R. ericae aggregate) and, based on ITS phylogeny, the ability to form both
ECM and ERM symbioses may have evolved with the R. ericae aggregate. Villarreal-Ruiz et
al. (2004) recently reported the ability of a fungus from the R. ericae aggregate to form
simultaneously both ECMs and ERMs in culture with Pinus sylvestris and Vaccinium
myrtillus seedlings, respectively, based on rDNA sequencing and microscopy.

Due to the complexity of the molecular mechanisms involved in establishment of a tight
(host-specific) symbiosis, the type of fungal associations with different plant hosts may not
be of great physiological importance under non-contaminated conditions, but may gain
ecological importance under stressed environmental conditions (Perotto et al., 2002).
Mycelial linkages may influence fungal and plant ecology by providing a source of fungal
inoculum to newly growing roots, allowing the C demands of the mycelium to be met by
more than one plant and facilitating the transfer of C and mineral nutrients between
neighbouring trees (Jones et al, 2003). It has been proposed that ECM and ERM fungi may

28

contribute to development of plant communities if the net transfer of C and nutrients is
predominantly from a pioneer plant species to a late successional species, but a greater
awareness of these processes is important for understanding the interactions between trees
and understorey vegetation (Dahlberg, 2001). Kernaghan et ah (2003) demonstrated a
positive correlation between ECM fungal richness and overstory host tree richness that was
explained by resource heterogeneity in combination with the preference (specificity) of ECM
fungi for certain plant hosts. Recently, DeBellis et ah (2006) showed that the distributions of
ECM fungi in southern mixed-wood boreal forests are influenced by the relative proportions
of host tree species. Conservation of stand diversity should therefore support diverse fungal
communities. Whether such communities are essential for ecosystem recovery following
PHC contamination is still not known. Regardless, minimizing overstory disruption
increases the possibility of preserving the integrated below-ground mycelial network with its
associated communities, and maximizes its potential to hasten site recovery.

Ecosystem processes
Biogeochemical cycling of nutrients and energy through ecosystems is driven by ordering
(e.g. photosynthesis, growth, humus formation) and dissipative (e.g. respiration, senescence,
decomposition) processes (Addiscott, 1995). Mycorrhizal systems form the functional
interface between decomposition (release of carbon and nutrients from organic substrates)
and primary production (formation of biomass) for both above-ground and below-ground
communities. In the below-ground food web, chemo-organotrophic organisms (those that
obtain energy and carbon from organic substrates) appear to be ultimately responsible for
governing nutrient availability for plant productivity (Wardle, 2002; Wardle et ah, 2004). In

29

reconstructed soil profiles (mini-ecosystems), the microflora (bacteria and fungi) were found
to exert a greater influence on nutrient mobilization and tree growth than either the fungusfeeding mesofauna or predator trophic groups (Setala et al, 2000). Setala et al. (2000)
reported that although species composition of the trophic groups was important for system
functioning, species richness within functional groups had a negligible impact on primary
production. Soil fauna (nematodes, protozoa, enchytraeids, microarthropods, earthworms,
termites, etc.) that feed on the microflora are also important in stimulation of primary
production (Wardle, 2002) by preventing nutrient sequestration within inactive microbial
biomass.

Decomposition
Heterotrophic bacteria and fungi directly decompose complex carbohydrates (mainly
cellulose and lignin) in plant detritus (Wardle, 2002). Cellulose is readily biodegradable as it
consists of P(l-4) linkages of D-glucose that form flat, linear chains H-bonded together to
create microfibril sheets (Evans and Hedger, 2001). By contrast, lignin is a threedimensional aromatic polymer consisting of P(0-4) linkages of monomeric units of either
cinnamyl alcohol: coumaryl alcohol (grasses), coniferyl alcohol (gymnosperms) or sinaptyl
alcohol (angiosperms) that surround the microfibrils and provide rigidity to plant cell walls
(Evans and Hedger, 2001). Due to its complex and uniquely heterogeneous structure (i.e.
hydrophobicity and thermodynamic stability), lignin is highly resistant to degradation (i.e.
recalcitrant) and inhibits decomposition by up to a few years by limiting access of
microorganisms or enzymes to substrates (Prescott et ah, 2000; Steffen, 2003).

30

In the early stages of decomposition, soluble compounds and cellulose are rapidly
metabolized under conditions where C is available and N is usually limiting (Prescott et al.,
2000). Decomposition rates slow over time due to changes in substrate compounds
(increased lignin fraction) and succession of microorganisms able to compete for various
substrates (Berg, 2000). In the later stages of decomposition, there is a net loss of lignin and
N is mineralized from humus (Prescott et ah, 2000). Growth may become N-limited in
habitats with high C:N ratio substrates, whereas in habitats with low C:N ratio (<30:1)
substrates, decomposition of organic matter may result in C limitation (Tiquia et al., 2002).
With the loss of cellulose, relative lignin and N concentrations increase and the higher N
concentration can slow the decomposition rate. Such slowing may be due to low molecular
mass N-containing compounds reacting with lignin residues during humification (Prescott et
al, 2000), creating more recalcitrant aromatic compounds; or mineral N may repress
synthesis of lignin-degrading enzymes in a wide range of soil organisms (Gallo et al., 2004);
or a combination of these (Magill and Aber, 1998). Higher initial N concentration (lower
C:N ratio) leads to lower decomposition extent (i.e. lower mass loss) and more stabilized
organic matter in forest soils (Berg, 2000). The process of humus formation (i.e.
humification) is thought to involve microbial modifications of lignin and condensation of
proteins or amino acids into humus precursors, which polymerize into structurally intricate
humus molecules (Prescott et al, 2000). Compared to the original plant material, humus is
low in carbohydrates (e.g. cellulose and hemicellulose) and high in polyphenolics (e.g. lignin
constituents) and immobilized N, which is sparingly available to plants (Prescott et al.,
2000).

31

Although it is generally accepted that mycorrhizas play important roles in decomposition and
cycling of C, N and P in ecosystems, details of their functions in nutrient dynamics and
regulation of nutrient and energy flows are continuing to be developed and refined (Martin,
2001; Allen et al., 2003). The traditional view of the role of mycorrhizas in obtaining
limiting nutrients from forest soils involves fungal exploration for nutrients [e.g. amino acids,
ammonium ( N H / ) , nitrate (NO3") and inorganic P] that are released during decomposition of
plant organic matter by heterotrophic fungal and bacterial communities or that are bound to
the soil matrix (e.g. insoluble forms of Al and Ca phosphates) (Martin et al., 2000).
However, molecular studies have revealed that some fungal species, previously regarded as
decomposers of woody debris (saprotrophs), are both frequent and abundant components of
ECM communities (Hibbett et al., 2000; Koljalg et al., 2000). This, along with other
conceptual advances in biocomplexity theory, have led to re-evaluation of how mycorrhizas
function within ecosystems and how interactions between multiple species of plants,
mycorrhizal fungi and soil saprotrophs regulate community composition and ecosystem
processes (Allen et ah, 2003).

It has been hypothesized that the distribution of the different mycorrhizal categories is related
to specialization for nutrient acquisition in particular environments (Read and Perez-Moreno,
2003). In higher latitude and higher elevation forest ecosystems, where seasonally low
temperatures and dry conditions result in very slow rates of decomposition, natural selection
may have favoured ECM and ERM symbioses with the capacity to mobilize nutrients from
organic material and provide them to plants (Read, 1991; Perotto et al., 2002; Read and
Perez-Moreno, 2003). Fungal specialization for N acquisition and utilization may be

32

important for determining community structure, lending strong theoretical support to the idea
that ECM diversity increases the effectiveness of nutrient acquisition from different spatial
locations and different substrates in soil (Martin et al, 2000; Leake, 2001).

Some ECM fungi appear to be directly involved in nutrient mobilization from organic
compounds through production of a wide range of hydrolytic and oxidative enzymes such as
polyphenol oxidases (e.g. laccase, catechol oxidase and tyrosinase) and endochitinases
(Martin et al., 2000; Burke and Cairney, 2002; Lindahl and Taylor, 2004). Most ECM fungi
so far investigated have demonstrated limited phenol-degrading activities, but few have been
studied. By contrast, ERM fungi (e.g. R. ericae) appear to have well-developed saprotrophic
abilities and degrade most polymeric components (e.g. polysaccharides, lignin, protein, chitin
and pectin) of plant and fungal cell walls (Martin et al, 2000). It is widely accepted that
enzymatic degradation of organic polymers, through production of an array of hydrolytic
enzymes in the extraradical mycelia and translocation of nutrients to the root, is the major
benefit of ERM symbioses to plants (Perotto et ah, 2002). ERM plants may enhance their
exploitation of complex soil substrates by broadening their metabolic capabilities through an
association with several fungi endowed with different functional enzymes (Martin et al.,
2000). This implies that simultaneous associations with a variety of symbiotic fungi may be
an important strategy to broaden the range of functions in the colonization of different
substrates (Perotto et al., 2002). Whereas many ECM and ERM fungi appear to have the
ability to access N and P directly from organic compounds, the extent to which they
contribute directly {via enzymatic catabolism) or indirectly (by influencing soil microbial

33

community structure) to decomposition remains unclear (Cairney and Meharg, 2002;
Heinonsalo et ah, 2004).

Primary production
Regardless of the mechanisms involved, symbioses with mycorrhizal fungi improve plant
nutrient acquisition by facilitating access to organic sources, potentially increasing the uptake
of nutrients via extensive growth of the mycelia and circumventing nutrient depletion zones
in the soil (Buscot et ah, 2000). Nutrients are absorbed across fungal membranes and are
either retained by the fungi for biosynthesis and growth, or transported distances of
centimeters to meters to the plant roots, where translocation to the host enhances the
photosynthetic machinery of the plant (Allen et ah, 2003). Plant photosynthetic rates depend
on the concentrations of N (for enzymes), P (for ATP and ADP), Fe and Mg (for
chlorophyll), internal CO2 and water (to keep stomata open to fix CO2) (Buscot et ah, 2000).
The resulting nutrient sink (root cells) allows for nutrient absorption and translocation
through fungal cells that occurs more quickly than diffusion to the roots through soil (Martin
et ah, 2000). It is thought that, as long as N or P is limiting in the soil environment, plants
will support their fungal partners through continued allocation of C (Allen et ah, 2003).

By absorbing, assimilating and translocating nutrients (e.g. nitrate, ammonium and amino
acids such as glutamate, glutamine and alanine), fungi create a C sink in the mycorrhizal
roots (Allen et ah, 2003). Heinonsalo et ah (2004) recently reported equivalent 14C
allocations to roots and ECMs in organic and mineral A and B horizons of a podzol, using a
Pinus sylvestris mini-rhizotron system. The sink strength controls the rate of photosynthate

34

production and the sugar supply appears to regulate some fungal gene expression (Buscot et
al., 2000). Carbon metabolism provides fungal mycelia and plant cells with the energy,
reducing power and biomass required for synthesis of various metabolites (e.g. amino acids)
required for growth (Martin et al, 2000). Martin et al. (2000) have suggested that
differences in ECM morphological features (e.g. abundant emanating hyphae with increased
metabolic activity versus few emanating hyphae with decreased activity) may reflect
differences in the need for C between taxa. Mycorrhizal fungi acquire most or all C via host
photosynthesis and translocation (average 10-20% of net photosynthetic yield), but may also
obtain C through assimilation following biodegradation of organic polymers in the soil
(Martin et al., 2000; Allen et al., 2003).

The ecological significance of using organic polymers as C sources, which may decrease the
need for C from the host plant, is unknown (Martin et al, 2000). A substantial amount of
fungal C is allocated to the synthesis of recalcitrant compounds such as chitin (60% of the
fungal cell wall) that can persist in the environment for years and increase soil aggregation,
stability, C storage and water-holding capacity. It has been estimated that as much as 20% of
N in boreal podzolic forest floors may be retained in chitin (0-1,4 linked Nacetylglucosamine units) present in dead and alive fungal mycelia (Lindahl and Taylor,
2004). The remaining C is respired (43-60%), accumulated as fungal storage sugars (e.g.
mannitol and trehalose) or lipids, or deposited in the mycorrhizosphere as labile compounds
(sugars, amino acids) that support the growth of bacterial communities (Allen et al., 2003;
Heinonsalo et al., 2004). The release of microbial communities from C limitation provides
the potential for them to play a major role in decomposition and nutrient mobilization (Read

35

and Perez-Moreno, 2003). The C cost for maintaining the external mycelium is unknown,
but the extent to which C is allocated from roots to mycelial systems may be intrinsically
linked to growth and nutrient-foraging activities of ECM fungi. Using digital
autoradiographic techniques, Leake et al. (2001) showed that patterns of C allocation within
ECM mycelia are highly dynamic and responsive to changes in niche (caused by spatial
variability in resource quality or interactions with other organisms). It has been hypothesized
that coexistence among fungi may be explained by the differential partitioning of C resources
among fungal species (Allen et al., 2003). The key role of mycorrhizas in C cycling
(particularly in the positive feedback loop between plant growth, decomposition and leaf
litter quality) may have important consequences for the C gains and losses of ecosystems and
thus for the C budget at local, regional and global scales (Read, 1991; Cornelissen et al.,
2001; Allen e/ al, 2003).

Summary
The influences of mycorrhizal fungi on plant populations and communities are not merely the
sum of effects on the individuals within populations (Dahlberg, 2001; Koide and Dickie,
2002). Due to their long history and multiple evolutionary events, different plants and fungi
bring independent characteristics to the symbiosis, resulting in extensive physiological
variation among mycorrhizas (Allen et al., 2003). For example, some fungal species,
previously regarded as saprotrophs, are both frequent and abundant components of ECM
communities (Hibbett et al., 2000; Koljalg et al., 2000). The spatial scales within which
individual mycelia operate as physically or physiologically integrated entities in nature are
also not clear (Cairney, 2005). For example, all groups of ERM fungi reported globally have

36

been found associated with salal from a single site (Berch et ah, 2002). Whereas a distinct
suite of functions can be assigned to a single mycorrhiza, the many genomic combinations
(genetic diversity) of symbionts, environmental heterogeneity and the extensive
connectedness of mycorrhizal root systems result in a complex suite of ways that
mycorrhizas can function in ecosystems (Cairney, 1999; Allen et ah, 2003). For example,
certain species of mycorrhizal fungi, rhizosphere organisms and plants may interact such that
there is a net immobilization of nutrients, which results in slower rates of decomposition
(Allen et ah, 2003). Other combinations of organisms (guilds) may not influence the
equilibrium of either nutrient cycling or decomposition, or may interact to increase nutrient
quality of litter and decomposition rates.

High species richness and abundance may represent ecological adaptation to local
environmental heterogeneity and is thought to provide forests with a range of strategies to
maintain efficient functioning under an array of environmental conditions (Cairney, 1999;
Nannipieri et ah, 2003). From a management perspective, a key challenge is to discover if
modifications of the environment send mycorrhizal ecosystems in predictable directions. Or,
alternatively, is it the combination of fungal and plant species that directs the trajectory?
Current evidence suggests a vast range of genetic potential in most mycorrhizal ecosystems
ready to respond to changing environmental conditions. Consequently, the hypothesis that
environment is predominant in determining the outcome warrants testing.

37

Petroleum hydrocarbon contamination of forest soils
Disturbance
Analysis of changes to the structure and taxonomic diversity of soil communities following
disturbance can provide clues to genetic and functional diversity by revealing some features
of the surviving organisms. Although rarely considered in this way, discrete PHC
contamination events such as oil spills are disturbances that disrupt ecosystem, community or
population structure and alter the physical environment and resource or substrate availability
(White and Pickett, 1985). Forest soil contamination is usually from a point source (i.e.
involving discrete, localized and often readily measurable discharge of chemicals) and often
results in rapid surface contamination, sometimes with large quantities of PHCs. For
economic and toxicological reasons, the fate and behaviour of organic pollutants in soils has
been the subject of intense research, with special interest focused on those chemicals that can
be taken up or transformed by living organisms (Alexander, 1999; Semple et ah, 2003).
However, the capacity of soil microorganisms to biodegrade organic pollutants not only
depends on whether they have the necessary metabolic pathways, but also on whether the
chemicals inhibit the microorganisms or are biologically available. Short- and long-term
changes to soil microbial communities may result if some populations are susceptible to
chemical toxicity (i.e. inhibition of cellular metabolic processes) or to changes to their
physical (soil) and chemical (substrates/ inhibitors) habitat. Whether these changes alter
community functions depends on the degree of redundancy within soil communities.

38

Chemical

toxicity

Petroleum products are complex mixtures that can contain numerous aliphatic (linear and
branched chains), alicyclic (unsubstituted and alkyl substituted structures) and aromatic
(unsubstituted and alkyl substituted structures with at least one unsaturated ring) compounds
(Miller and Herman, 1997; Potter and Simmons, 1998). Numerous methods are available for
analysing petroleum hydrocarbon mixtures in environmental media (Weisman, 1998).
Natural gases are generally composed of methane, ethane and small amounts of higher
molecular mass hydrocarbons whereas most crude oils contain compounds such as paraffins,
aromatics, naphthenics and asphaltenes that are present in varying proportions (McGill et ah,
1981; Potter and Simmons, 1998). Gasoline typically contains compounds in the nC4 to
nC12 range, while diesel compounds are in the nC8 to nC21 range (Potter and Simmons,
1998). Light paraffinic crude oils are dominated by molecules with C numbers less than 16
and consisting mainly of alkanes (paraffins) and cycloalkanes (naphthenes). These chemicals
are generally first metabolized by microorganisms (Westlake et ah, 1973; Riser-Roberts,
1998). The heavier oils have greater proportions of aromatic hydrocarbons and heterocyclic
NSO compounds (i.e. containing N, S, or O) with C numbers usually above 20 (McGill et ah,
1981;Delilleefa/.,2004).

Using chromatographic techniques, crude oils can be resolved into four categories of
compounds: asphaltenes, saturates, aromatics and polars (eg. NSO compounds) (McGill et
ah, 1981; Pollard et ah, 1992; Weisman, 1998). Asphaltenes are a mixture of pentaneinsoluble, colloidal compounds including polyaromatic and alicyclic molecules with some
alkyl substitutes (usually methyl groups) that vary in molecular mass between 500 and

39

several thousand (McGill et ah, 1981). The structures of these compounds, particularly those
with higher mass, share characteristics with proposed structures of humic acids (Prescott et
ah, 2000). Saturated and aromatic hydrocarbons (mainly n-alkanes, branched alkanes, mono, bi-, and polycyclic alkanes (naphthalenes) and mono-, bi- and polyaromatics) usually
account for 75% of the mass of crude oils (McGill et ah, 1981; Potter and Simmons, 1998).
Up to 25% of the total mass may be n-alkanes, with cyclic hydrocarbons accounting for 3060% of the total mass. Monocyclic aromatic compounds (e.g. toluene, benzene and xylene)
and bicyclic types (e.g. naphthalene, biphenol) represent 1-2%; polycyclic aromatics (usually
methylated derivatives of fluorene, phenanthrene, anthracene, chrysene, benzofluorene and
pyrene) are present in lower amounts (McGill et ah, 1981). The NSO fraction contains polar
compounds such as naphthenic acids, mercaptans, thiophenes and pyridines. Most of the N
in crude oil is contained in the distillate residue as part of the asphalt and resin fraction and
usually accounts for less than 0.2% (rarely exceeds 1%) by mass. The S content varies
between 0.3 and 3% whereas the O content usually does not exceed 3% (McGill et ah, 1981).

Aliphatic compounds are generally less toxic than aromatics, and toxicity has been found to
vary with compound size (McGill et ah, 1981; Edwards et ah, 1998). The quantity and
composition of polycyclic aromatic hydrocarbons (PAHs) are major considerations in the
evaluation of toxicity of PHC mixtures (Miller and Herman, 1997). PAHs with four or more
benzene rings are known to be genotoxic to humans and other ecological receptors; in
general, as relative molecular mass and polarity (i.e. degree of oxidation) of PAHs increase,
carcinogenicity also increases and acute toxicity decreases. This is due to the metabolic
production of highly reactive electrophilic intermediates that can access biological molecules

40

such as DNA, RNA and proteins and react to form adducts or lesions (Landis and Yu, 1995).
Little is known about PHC toxicity to plant and microbial communities in forest soils as the
majority of studies have excluded the complex interactions between combinations of
chemicals, interacting communities and the soil environment that may exert synergistic,
potentiative or antagonistic effects (Landis and Yu, 1995; Evans and Hedger, 2001; Koivula
et ah, 2004). It is likely that toxicity varies with the type of pollutant, the extent of pollution
and the general condition (i.e. extent of obvious signs of stress or disease) of the ecosystem
prior to chemical disturbance (Seghers et al., 2003).

In agricultural clay soils, the maximum toxicity of crude oil (indicated by worm survival,
seed germination, bacterial bioluminescence and photosynthesis inhibition) was highest
immediately after introduction of oil (Chaineau et ah, 2003). An initial decrease in microbial
density has often been observed immediately following the addition of PHCs to soil. For
example, the addition of 10% (volume/ mass) toluene to soil resulted in survival of only
about 1% of the indigenous bacteria, which eventually recolonized the soil to reach a high
cell density (Huertas et al, 2000). Biotransformations of PHC substrates may also lead to
the release of an array of potentially toxic metabolites into the surrounding environment
(McGill et al., 1981; Riser-Roberts, 1998). For bacteria, toxicity resulting from PHC
contamination has been inferred from decreases in enzyme (hydrogenase and invertase)
activity (Suleimanov et al., 2005), although reduced enzyme activities may also result from
competition for limiting nutrients following PHC contamination.

41

The toxicity of non-ionized organic contaminants for microorganisms is primarily due to a
nonspecific mode of action that involves partitioning of organic chemicals into the
hydrophobic (lipophilic) layer of the cell membrane and disruption of membrane integrity
(i.e. increased membrane permeability) (Miller and Herman, 1997). Kirk et al. (2005)
suggested that interference with fungal membranes may explain the reduced extraradical
hyphal growth of AM fungi {Glomus species) in PHC medium with soil. In general, fungi
are considered to be more tolerant of high concentrations of polluting chemicals than
bacteria, possibly due to differences in cell wall structure (Blakely et ah, 2002). Some yeasts
(e.g. Saccharomyces cerevisiae) have been found to alter their membranes (i.e. increase
hydrophilicity) to exclude hydrophobic contaminants (Park et al., 1988). It is possible that
similar compensation mechanisms occur in other fungi as well. Long-term resistance to
PHCs could also be due to the ability of ECM fungi to produce spores that resist
environmental stress factors and germinate when the concentration of toxicants associated
with PHC contamination has decreased sufficiently over time (Nicolotti and Egli, 1998).

PHCs may directly kill plants on contact, slow their growth, inhibit seed germination, create
nutrient-deficient conditions or, at lower concentrations, stimulate plant growth (McGill et
al., 1981). Nicolotti and Egli (1998) have suggested that crude oil has a caustic or lethal
effect on plants only when it comes into direct contact with tissues and that reduced growth
and biomass may be manifestations of changes to soil communities. In general, the taller the
trees and the deeper their roots, the greater their tolerance to increased PHC concentrations in
soil (Trofimov and Rozanova, 2003).

42

Soil properties and processes
Blakely et al. (2002) found that creosote impacted soil food webs and decomposition
processes more by altering the habitat of microinvertebrates and their prey (i.e. fungi and
bacteria) than via direct chemical toxicity. Kirk et al. (2005) suggested that PHCs may
interfere with plant-fungus communication by altering root exudation patterns or changing
the soil environment such that migration of diffusible chemical signals (e.g. flavonoids,
auxins, etc.) is prevented. Many of the major impacts of PHCs on soil biota and plants in
forest ecosystems appear to be associated with disturbances to water, nutrient and oxygen
supplies related to the hydrophobicity and fluidity of oily products (Tarradellas and Bitton,
1997; Trofimov and Rozanova, 2003).

The disturbance caused by PHC contamination leads to considerable changes in physical and
chemical properties that are not typical of unpolluted soils. PHC constituents may be found
in mobile form, fixed in the soil pores and fissures, adsorbed on the surface of organic and
mineral soil constituents or forming a continuous cover on the soil surface (Trofimov and
Rozanova, 2003). Morphological changes in PHC-contaminated podzolic soils exhibit
fragmentary patterns resulting from the unevenness of chemical distribution in the soil mass,
increased amounts of iron in the upper horizons, and increased amounts of cemented soil
aggregates (Trofimov and Rozanova, 2003). The extent of physical movement in the soil
profile depends on temperature, PHC viscosity, moisture content, soil structure and soil
texture (McGill et al, 1981). Greater lateral spread of PHCs occurs in cold conditions; in hot
and dry soil conditions, vertical movement into the water-unsaturated zone may occur more
frequently (McGill et al, 1981). In sandy soils, frontal migration of PHCs down the soil

43

profile along the paths of roots and fissures altered the soil profile to a depth of greater than 1
m (Trofimov and Rozanova, 2003). In gray forest soils, heavy fractions of PHCs were
retained in the upper plow layer and filled the largest infiltration, aeration and drainage
pores; lighter fractions were largely retained in illuvial horizons and filled fine water
retention pores (Suleimanov et al., 2005). This can lead to waterlogging and reducing
conditions in the soil profile, both of which inhibit decomposition processes (Trofimov and
Rozanova, 2003). Large amounts of oily material in soils may also indirectly increase soil
temperature (by 1-10°C) if there is loss of surface vegetation. In some cases, more damage to
the soil may occur due to the high osmotic potential of associated brine water (salinity 40,000
-45,000 ug mL"1) than to the presence of PHCs alone (McGill et al, 1981).

Water insolubility, hydrophobicity and soil sorptive properties increase with increasing size
(number of aromatic rings) and complexity (molecule topology or pattern of ring linkages) of
chemicals; PAHs with three or more rings tend to be strongly sorbed to the soil (Reilley et
al, 1996; Alexander, 2000; Kanaly and Harayama, 2000; Cerniglia and Sutherland, 2001;
Chaineau et al, 2003). Chemical persistence in soil also depends on several environmental
factors, including the type and quality of clay particles (as well as cation exchange capacity),
the type and concentration of solutes in surrounding solution, soil organic matter (SOM)
content and composition, pH and temperature (Alexander, 1999; Semple et al., 2003).
Organic chemicals may be sorbed to and retained by soil particles by adsorption or
partioning. Adsorption entails chemical processes (ion exchange) or, more often, physical
forces (H bonding or van der Waals forces) to surfaces of organic polymers or the external
surfaces of 1:1 clay minerals and the external and internal surfaces of 2:1 (expanding) clays

44

(Alexander, 1999; Miller and Herman, 1997; Ellerbrock et al., 2005). Sorption to minerals
must compete with water and may be very low for nonionized organics in hydrated systems.
Sorption to organic solids may occur via physical binding, which concentrates chemicals on
outer surfaces or within the pores of a solid. Partitioning of organic chemicals into the SOM
is a process of transfer from the bulk state of one phase to the bulk state of another by
mechanisms analogous to dissolution and leads to a distribution of molecules within a
portion or the entire volume of the organic matter (Alexander, 1999; Chaineau et al., 2003).
The extent of chemical retention in the SOM fraction is directly correlated with the octanolwater partition coefficient of the substance (K0Vi, measure of chemical hydrophobicity), the
amount of SOM in the solid phase and its degree of oxidation or polarity (Xing et al., 1994;
Alexander, 1999; Wang et al., 2005). In forest soils, the sequestration of organic pollutants
in SOM (i.e. sorbed inside soil aggregates or at inactive particle surfaces) may decrease
toxicity of chemicals through physical separation from biological receptors, which also
decreases substrate bioavailability for enzymatic degradation (Alexander, 2000; Ellerbrock et
al., 2005). Toxicity of hydrophobic organic contaminants has been found to be less severe
for organisms in soils with high humus content (Salminen and Haimi, 1997).

PHC-polluted soils are characterized by lower values of hygroscopic moisture, hydraulic
conductivity and water retention capacity (i.e. wettability) compared to unpolluted soils
(Trofimov and Rozanova, 2003; Suleimanov et al., 2005). This is related to the spatial
arrangement of hydrophobic components within SOM (Roy and McGill, 2000). Higher
molecular mass components and their degradation products remain near the soil surface and
form crusts that decrease water availability and limit water and gas exchanges between the

45

soil and the atmosphere. The creation of discrete and continuous water-repellent fronts
parallel to the soil surface is also recorded in post-fire forest soils (Certini, 2005).
Hydrophobic films on the exterior surfaces of soil aggregates reduce the wettability of the
soil and increase structure stability (McGill et al., 1981; Certini, 2005). Many PHCcontaminated soils eventually take up water and remain wet; however, long-term (years)
hydrophobicity of crude oil contaminated agricultural soils has been documented in western
Canada (Roy et al, 1999).

The longer some chemicals remain in soil, the more they appear to resist desorption and
biodegradation. Weathered (aged) chemical residues have considerable time to interact with
the physical and chemical components of soil. Interactions may entail: (1) sorption, most
likely via partitioning; or (2) irreversible incorporation into soil organic matter {via
humification) by the catalytic activity of a variety of oxidative enzymes present in the soil
matrix (Miller and Herman, 1997; Alexander, 1999). PHC pollution has been found to
substantially increase the organic C (humic acid) content of soils (Trofimov and Rozanova,
2003). Humification of PHC constituents is explicit to transformation processes. Covalent
bonding between organic chemicals and humic polymers (humin, fulvic acid and humic acid)
in soil can form stable linkages to dialkylphthalates, alkanes and fatty acids that are resistant
to microbial degradation and are not readily extractable with many organic solvents (McGill
et al, 1981; Alexander, 1999). Petroleum residues (as indicated by dichloromethane
extraction) are associated with soil organic matter (Roy et al., 2003). Sorption of volatile
PHCs from adjacent soil has generated hydrophobicity in soils not directly contaminated with
PHCs (Roy and McGill, 2000). They may be either directly incorporated through H bonding

46

of phenolic and benzene carboxylic acids into the molecular structure of soil humic materials
or adsorbed to the surface of the molecule (McGill et ah, 1981). They are not entirely
associated with the humic component, however, because exhaustive extraction with NaOH
did not eliminate hydrophobicity of soils (Roy and McGill, 2000). It is not known whether
complexes between hazardous organic chemicals and soil humic materials are cleaved in
nature to give detectable levels of the original compound, whether these complexes are
assimilated by animals and plants, or whether they pose problems of present or future
toxicological significance (Alexander, 1999).

In PHC-contaminated soils, the C (energy) supply increases, which promotes metabolic
activity on the part of all the microorganisms not directly inhibited by the PHCs and
concurrently the C:N ratio tends to increase. The carrying capacity of a soil is the maximum
level of microbial activity that can be supported under existing environmental conditions,
which depends on the size of the population, availability of O2 or nutrients, temperature and
water availability. The carrying capacity of soils may be exceeded as a result of large inputs
of C from PHCs (Miller and Herman, 1997). The intensive growth of PHC-oxidizing
microorganisms in response to increased C availability is accompanied by consumption of
soil nutrients, resulting in decreased nutrient availability for plants (Xu and Johnson, 1997;
Tiquia et ah, 2002; Trofimov and Rozanova, 2003). A decrease in available N may also be
partly due to the inhibition of nitrification and ammonification processes or from loss of
nitrates (McGill et al, 1981; Suleimanov et al., 2005). Some studies have shown that
nitrifying bacteria were not found in freshly contaminated soils, but that nitrification
processes were eventually regained; stable organic matter and total N content have increased

47

significantly following PHC contamination (McGill et al., 1981). An increase in total N has
been attributed to increased atmospheric N2 fixation during PHC biodegradation (McGill et
al., 1981). Thus, as the immobilization of mineral N present in the soil increases, the amount
of available N decreases such that organisms benefiting from N2 fixation, or consortia
capable of recycling N from microbial biomass, are the only organisms that can thrive under
these conditions (i.e. selection of PHC-tolerant species) (Nicolotti and Egli, 1998). Oxidative
degradation may also alter the composition of soil bacterial communities, so that aerobic
cellulolytic and proteolytic species decrease and anaerobic N- fixing species increase
(Nicolotti and Egli, 1998). Under disturbed water and extreme O2 limitation, P may be
reduced and escape to the atmosphere as hydrogen phosphide (Suleimanov et al., 2005),
although this is a very small loss mechanism.

It does not appear that a single addition of PHCs limits microbial communities in the long
term. However, several studies indicate that although total microbial numbers tend to
increase over time, species richness often decreases, which may or may not have deleterious
impacts on ecosystem functions (McGill et al., 1981; Hofman et ah, 2004). In general, PHC
contamination is expected to lead to an initial loss of richness, followed by rapid proliferation
of metabolically competent members of communities inhabiting the new environmental
conditions imposed by the chemical contaminants (Gramss et al., 1998; Seghers et ah, 2003;
Diaz, 2004). Some studies have reported drastic reductions in overall ECM biomass and
colonization potential in soil following a spill, whereas some fungi appeared resistant to the
PHCs and may have benefited from their presence (Nicolotti and Egli, 1998). In greenhouse
experiments, spruce seedlings grown in crude-oil-contaminated soils exhibited shifts in ECM

48

community structure in response to increased contaminant concentrations (Nicolotti and Egli,
1998). Nitrogen availability may be a major factor structuring ECM fungal communities;
mineral N and foliar nutrient ratios (N:P, P:A1) were found to be excellent predictors of
fungal taxonomic richness in organic horizons and organic nitrate availability was a good
predictor of their relative abundance (Lilleskov et ah, 2002). From studies of impacts of acid
rain on soil communities, changes to soil chemical status and functions of the decomposer
community have been suggested to lead to imbalances in nutrient cycling and ecosystem
productivity (Pennanen et al., 1998).

Biodegradation
The ability of heterotrophic bacteria and fungi to degrade organic pollutants appears to be
inherent in most natural microbial communities and it is generally accepted that biological
processes eventually degrade or transform most bioavailable (i.e. accessible by organisms or
their enzyme systems) organic compounds (McGill et al., 1981; Sarand et ah, 2000;
Nannipieri et ah, 2003; Delille et ah, 2004; Diaz, 2004). Many xenobiotic chemical
constituents (e.g. PHCs) are structurally analogous to compounds naturally found in the soil
environment (e.g. plant material, fungal and root exudates and allelopathic chemicals) and
appear to be biodegraded through the same biochemical pathways (Miller and Herman, 1997;
Siciliano and Germida, 1998). In addition to accidental releases, low levels of PHCs may
also enter soils from natural seepages or via atmospheric deposition after burning of fossil
fuels (Knox et al., 1999; Kanaly and Harayama, 2000; Trofimov and Rozanova, 2003;
Certini, 2005). Biodegradative potential does not appear to be a distinguishing taxonomic
character as metabolic ability (via different genes and biochemical pathways) is widespread

49

among many species of ubiquitous genera of bacteria and fungi (Siciliano et ah, 2003;
Chaillan et ah, 2004).

The complete catabolic conversion (mineralization) of organic substrates to inorganic
products (H 2 0 and CO2) and use of nutrient constituents (C, N, P, S and other elements) for
synthesis of cellular components is known as growth-linked biodegradation (Alexander,
1999). Biodegradation of some organic pollutants appears to result from transformations by
microbial populations that are unable to use the substrate as a source of energy (even if the
conversion is an energy-releasing oxidation reaction) or as an essential nutrient (i.e. not
significant sources for biosynthesis) and is known as cometabolism (Miller and Herman,
1997; Alexander, 1999; Sarand et ah, 2000). Cometabolism is usually attributed to the
activity of enzymes with relaxed specificities that act on structurally related substrates
(Hickey, 1998). Organic substrates may be transformed to products that are not typical
intermediates of central metabolism and the organism may not possess enzymes to convert
further the compound into metabolic intermediates for biosynthesis or energy production.
Alternatively, products may inhibit enzymes for subsequent metabolic conversions, suppress
the growth of the organism, or the organism may require a second substrate (cofactor) to
bring about a particular reaction (Alexander, 1999). Cometabolic reactions also have impacts
in nature that are different from growth-linked biodegradation. Whereas the rate of growthlinked biodegradation characteristically increases with time as populations that are able to
use the substrate as a source of energy and nutrients multiply, the environmental
consequences of a population's inability to grow at the expense of the substrate are slow rates
of biotransformation (due to small microbial biomass) and accumulation of organic products

50

that tend to persist in the environment (Alexander, 1999). The potential for substantial
biodegradation of PHCs in soil (via growth-linked and cometabolic pathways) results in
release of a nearly limitless array of metabolites into the soil environment. Some metabolites
may be toxic to the soil biota; some may react with soil constituents or may be quickly
degraded by other microorganisms present (McGill et al., 1981; Riser-Roberts, 1998).

In a community context, biodegradation involves synergism. Syntrophic biodegradation
occurs when two or more populations carry out transformations that one population alone
cannot perform or performs slowly (Alexander, 1999). Thus, even if a particular population
can metabolize only a small number of the chemical substrates available, other populations
occupying the same habitat may possess complementary degradative enzyme capabilities that
may ultimately result in complete chemical mineralization. Studies of mixed populations
(i.e. communities) of bacteria have revealed more complex and rapid biodegradation than
was previously believed possible based on studies of pure cultures (McGill et al, 1981).
Little is known concerning syntrophic biodegradation by specific guilds of organisms. If
functional redundancy is the norm in most ecosystems, is the degradation of a substrate in
soil dictated by specific guilds of organisms, or are the properties of the substrate and soil
environment more important?

Bacterial pathways
Because of their metabolic versatility, bacteria are able to obtain energy from virtually every
organic compound (Romantschuk et al., 2000; Diaz, 2004). The most common electron
acceptor for microbial respiration is O2 and aerobic processes provide the highest amount of

51

energy to cells. Oxygen is not only the electron acceptor, but also participates in activation
of the substrate via oxygenation reactions (Diaz, 2004). Anaerobic (anoxic) conditions are
prevalent in aquifers, aquatic sediments and waterlogged soils. Here, biodegradation is
carried out by strict anaerobes or facultative organisms using alternative electron acceptors
such as nitrate (e.g. denitrifying organisms such as Pseudomonas, Alcaligenes and
Flavobacterium), sulphate (e.g. sulphate reducers such as Desulfobacterium), Fe(III) (e.g.
ferric iron reducers such as Geobacter), CO2 (e.g. methanogens such as Methanospirillum) or
others such as chlorate, Mn or Cr (McGill et ah, 1981; Diaz, 2004). Use of alternative
acceptors depends on their availability as well as competition between different
microorganisms for electron donors. The energy obtained using Fe(III) or nitrate is almost as
efficient as using O2, but less energy is generated by sulphate reducers and methanogens.
Fermentative strains may be energy-limited and restricted to syntrophic existence, requiring
other populations to consume the potentially toxic endproducts of fermentation.
Photosynthetic organisms use energy from the sun to degrade aromatics anaerobically to
acetyl-CoA, which is subsequently used in biosynthetic reactions (Diaz, 2004). In both
aerobic and anaerobic degradation, structurally diverse compounds are degraded through
many different peripheral pathways to a few intermediates that are further channelled via
biochemical pathways (i.e. reactions leading to the formation of Krebs cycle intermediates)
to the cell's central metabolism (Diaz, 2004).

Many microorganisms can use the aliphatic compounds present in PHCs as C sources. The
mid-size straight-chain n-alkanes (nCIO to nC18) appear to be metabolized more readily than
n-alkanes with shorter or longer chains, and saturated (single C bonds) are degraded more

52

readily than unsaturated (double C bonds) compounds (Miller and Herman, 1997; Delille et
al., 2004). The extent and location of hydrocarbon sidechain branching or halogen
substitution slows the biodegradation of the compound (Miller and Herman, 1997). The
most common aerobic biochemical pathway involves direct incorporation of one atom of O2
into the alkane by a mixed function oxidase or monooxygenase enzyme, but both O2 atoms
can also be incorporated, hi either case, a primary fatty acid is formed that is subjected to
consecutive removal of two-carbon fragments (P-oxidation), which are converted to acetylCoA. This intermediate enters the Krebs cycle where complete mineralization to CO2 and
H 2 0 occurs (Miller and Herman, 1997). For alkenes, the first step is attack at the terminal or
subterminal methyl group or at the double bond to yield an alcohol or epoxide that can be
further oxidized to a primary fatty acid and enter P-oxidation. Degradation of alicyclic
hydrocarbons (which are major components of crude oil, constituting 20-67% by volume) is
thought to occur primarily via cometabolic reactions to open rings and subsequently cleave
linearized products for entry into the Krebs cycle (Miller and Herman, 1997).

Under anaerobic conditions, however, oxygenation of hydrocarbons using O2 is not possible.
Aromatic ring structures may be activated under anaerobic conditions using a reductive
rather than oxidative process. There is growing evidence that under anaerobic conditions
some microbial communities are able to use O from H2O or CO2 for ring cleavage of
aromatics or to prepare for ring cleavage of aromatics. For example, Schink et al. (1992)
propose H2O as the O source for ring cleavage during benzoate metabolism by fermenting
and denitrifying bacteria, and CO2 as the O source for carboxylation in preparation for ring
cleavage of aniline by a sulphate-reducing bacterium.

53

Westlake et ah (1973) found that the ability of mixed populations of bacteria to use crude oil
as a sole C source depended on the composition and amount of n-saturates, asphaltenes and
NSO compounds and also that the aromatic fraction of crude oil was capable of supporting
bacterial growth. It seems that PHC biodegradation is more closely related to the intrinsic
biodegradability of chemicals (and their bioavailability) than to the particular enzymatic
capacities of the microorganisms involved (Chaillan et ah, 2004).

Bacteria such as Pseudomonas, Mycobacterium, Rhodococcus,

Flavobacterium,

Acinetobacter, Arthrobacter, Bacillus and Nocardia are considered the primary degraders of
polycyclic aromatic hydrocarbons (PAHs) in soil (Kanaly and Harayama, 2000; Chaillan et
ah, 2004). Pseudomonas has been the most extensively studied, owing to its ability to
degrade so many different contaminants and its ubiquity in soils containing PHCs (McGill et
ah, 1981; Axelrood et ah, 2002a; Delille et ah, 2004; Diaz, 2004). Most aerobic peripheral
pathways involve oxygenation reactions carried out by monooxygenases and/or
hydroxylating dioxygenases that incorporate one or two atom(s) of O2 into the aromatic ring
structure to generate dihydroxy aromatic intermediates (e.g. catechol, protocatechuate,
gentisate, hydroxyquinol and hydroquinone) (Miller and Herman, 1997; Siciliano and
Germida, 1998; Cerniglia and Sutherland, 2001; Watanabe, 2002; Diaz, 2004). These
compounds are the substrates of ring-cleavage enzymes that use molecular oxygen to open
the aromatic ring between the two hydroxyl groups (ortho cleavage, catalyzed by intradiol
dioxygenases) or proximal to one of the two hydroxyl groups (meta cleavage, catalyzed by
extradiol dioxygenases). After several subsequent steps, the linearized products are

54

incorporated into the Krebs cycle. Anaerobic peripheral pathways usually converge to
benzoyl-CoA, which is dearomatized by specific multicomponent reductases and consumes
ATP (Diaz, 2004).

The initial hydroxylation step is considered to be rate limiting and the enzymes involved
generally determine the substrate range of microorganisms, although other factors (e.g.
substrate specificity of transcriptional regulators and membrane transporters) may also
contribute (Watanabe, 2002). In a recent study of biodegradation by indigenous microbial
communities in sub-antarctic soils, PAHs with greater than three rings were generally not
degraded (Delille et ah, 2004). There is little evidence that microbial growth can be
sustained with PAHs with four or more rings as a sole substrate (although they may be
degraded by syntrophic cometabolism) and there is very limited information regarding
bacterial degradation of PAHs with five or more rings (Reilley et ah, 1996; Kanaly and
Harayama, 2000). However, bacterial degradation of pyrene (a pericondensed four-ring
PAH) has been reported from sediment near a hydrocarbon source, and a Mycobacterium
species isolated from that sediment has been shown to degrade pyrene using an inducible
enzyme system (Kanaly and Harayama, 2000).

The metabolic flexibility of bacteria is related to their genetic adaptability. For example,
Siciliano et ah (2003) found that a substantial decrease of aged PHCs in soil was related to a
greater presence of catabolic genes (i.e. alkB, alkane monooxygenase; ndoB,
naphthalenedioxygenase; xylE, catechol-2,3-dioxygenase) in bulk and rhizosphere soil.
However, it was unclear whether the number of organisms containing these genes increased,

55

or if the number of genetic elements present in the community increased. The genes
responsible for aromatic biodegradation pathways are usually arranged in clusters (operons)
in mobile genetic elements (e.g. plasmids or transposons). Gene clusters contain catabolic
genes (encode enzymes for catabolic pathways), transport genes (responsible for active
uptake of the compound) and regulatory genes (adjust expression of the catabolic and
transport genes to the presence of the compound to be degraded) (Diaz, 2004). For example,
the catabolic genes of the ortho and meta pathways are organized as operons with flanking
transposon elements on the TOL (toluene) plasmid (Sarand et al., 1998). This facilitates
horizontal transfer of the respective genes and rapid adaptation of microorganisms to the
presence of new substrates (Diaz, 2004). Conjugation (transfer of genetic material from one
microorganism to another) appears to be important in the dissemination of catabolic genes in
the indigenous environment (Sarand et al, 2000; Siciliano et al., 2003).

Depending on chemical structure, contaminant concentration and environmental conditions,
the onset of PHC biodegradation generally follows a period of acclimation in which no
chemical degradation is evident (Alexander, 1999). Adaptation most commonly occurs by
induction of the enzymes necessary for biodegradation, followed by increases in populations
of biodegrading organisms (Miller and Herman, 1997). Chronic exposure to PHC substrates
(e.g. near natural seepages or in areas where frequent spills occur) results in shorter
acclimation periods (due to maintenance of biodegradation pathways within adapted
communities) and subsequently increased transformation rates (Miller and Herman, 1997;
Alexander, 1999). This pollution-induced community tolerance appears to increase
proportionally with increased exposures (Seghers et al., 2003). The end of this period is

56

marked with a rise in respiration and increase in density (varying from slight to several
orders of magnitude) that reflects growth of hydrocarbon-degrading populations as well as
increased growth of organisms such as protozoa that graze microflora or decompose necrotic
tissue (McGill et al., 1981). Subsequent declines in microbial respiration may occur due to
complete degradation of labile fractions or to limiting availability of N and P (McGill et al.,
1981). High abundance, rapid growth and the ability to transfer genes horizontally allow for
rapid microbial adaptation to changes in environmental conditions (Romantschuk et ah,
2000; Diaz, 2004).

Genetic changes such as mutations (i.e. appearance of new genotypes) may occur when
communities are faced with chemicals that do not have natural chemical analogues (Miller
and Herman, 1997). Such events occur at low frequency; however, if new genotypes possess
physiological characteristics that provide a selective advantage (e.g. new metabolic
capacities), they may multiply {via horizontal gene transfer) within the surviving community
(Alexander, 1999). The length of time required for a genetic change or for selection and
development of an adapted community is not yet predictable (Miller and Herman, 1997).
However, given enough time and favourable environmental conditions, the capacity to
degrade almost any organic compound is likely to evolve in or immigrate to a contaminated
site (Romantschuk et al., 2000).

Fungal cytochrome P450 and ligninolytic systems
Many fungi (e.g. Aspergillus, Penicillium, Fusarium) isolated from PHC-contaminated soils
and cultured on PHC-containing medium have been found to use crude oil as a sole C and

57

energy source (Chaillan et ah, 2004). In general, eukaryotic organisms oxidize aromatic
compounds to water-soluble products via a cytochrome P450 monooxygenase reaction,
incorporating one atom of molecular O2 into the aromatic ring to form a transient arene oxide
and reducing the second atom of O2 to H2O. The arene oxide is immediately hydrated by an
epoxide hydrolase to yield a trans-dihydrodiol or, alternatively, is non-enzymatically
isomerized to form phenols that can conjugate with sulphate, glucuronic acid or glutathione
(Miller and Herman, 1997; Cerniglia and Sutherland, 2001). These reactions increase both
the water solubility and bioavailability of chemical substrates; soil conditions that favour
fungal activity may initially increase the toxicity of the parent chemicals (Reilley et ah,
1996). Whereas complete mineralization results in innocuous endproducts (CO2 and H2O),
partial biodegradation can produce intermediate metabolites with unchanged, reduced or
increased chemical toxicity. Toxic chemical intermediates with increased water solubility
are of particular concern as this can result in the transport and spread of contaminants
through the environment (Miller and Herman, 1997).

The ligninolytic enzyme system of white rot fungi (WRF) has been extensively studied due
to structural analogies between lignin and PAHs as metabolic substrates (Scheel et ah, 2000).
Although lignin is a much larger and more heterogeneous polymer than the fused benzene
ring structures of PAHs, it is also hydrophobic and insoluble, thereby posing similar
problems for enzyme catalysis (Harvey and Thurston, 2001). WRF degrade lignin using a
complex nonspecific enzyme system, often while simultaneously obtaining C from cellulose
and hemicellulose (i.e. cometabolism) (Scheel et al., 2000; Steffen, 2003). As with the
cytochrome P450 system, the oxidizing enzymes of the ligninolytic system increase the

58

bioavailability, solubility and redox status of the chemical substrates for subsequent
metabolism (Harvey and Thurston, 2001). Different fungi appear to possess different
combinations of oxidizing enzymes (Harvey and Thurston, 2001).

The initial hydroxylation step of the pathway is accomplished with small, diffusible
oxidizing agents (highly reactive radicals) generated by three groups of extracellular
enzymes: lignin peroxidases (LiP), manganese peroxidases (MnP) and laccases (Collins and
Dobson, 1997; Harvey and Thurston, 2001). LiP (EC 1.11.1.14) and MnP (EC 1.11.1.13) are
heme-containing enzymes that function at low pH and catalyze the oxidation of lignin, humic
substances and many organopollutants (Schlosser and Hofer, 2002). Both enzymes require
H2O2, which is generated through fungal glucose oxidase, glyoxal oxidase and arylalcohol
oxidase reactions (LiP and MnP) or oxidation of organic acids (MnP only) (Evans and
Hedger, 2001). In the white rot basidiomycete Phanerochaete chrysosporium, PAHs with
ionization potentials at or below about 7.55eV are substrates for direct one-electron oxidation
by LiP, whereas those with ionization potentials above this threshold appear to be acted upon
by radical species formed during MnP-dependent lipid peroxidation reactions (Bogan,
Schoenike and Lamar, 1996). For LiP, radical cations are produced from one-electron
oxidations of non-phenolic compounds, which act as non-specific redox mediators and
extend the substrate range and redox capacity of LiP (Harvey and Thurston, 2001). During
the catalytic cycle of MnP, the active centre is oxidized by H2O2. Reduction of the resting
enzyme is achieved by two successive one-electron transfers that oxidize Mn 2+ to Mn3+,
which is facilitated by fungal organic acids (e.g. oxalate or malonate) upon chelation of the
highly reactive Mn 3+ state. Schlosser and Hofer (2002) found evidence supporting a

59

physiological role of laccase-catalyzed Mn

oxidation in providing H2O2 for extracellular

oxidation reactions and demonstrated a novel type of laccase-MnP cooperation relevant to
biodegradation of lignin and organic pollutants.

Laccases (benzendiokoxygen oxidoreductase, EC 1.10.3.2) are (blue) multicopper enzymes
(glycosylated polyphenoloxidases) that are an essential component of a complex nonspecific
enzyme system secreted by different kinds of fungi that have been shown to oxidize lignin
and various organic contaminants (Schlosser and Hofer, 2002; Gonzalez et al., 2003;
Hoegger et al., 2004). In addition to lignin depolymerization and polyphenol degradation,
laccases are thought to be involved in the release of N from insoluble protein-tannin
complexes, mycelial pigmentation, humus formation, fruiting body formation and
detoxification of phenolic compounds, which protects fungi against soil pollutants and host
defense compounds (Kanunfre and Zancan, 1998; Burke and Cairney, 2002; Hoegger et al.,
2004). Most reports refer to laccase activity as extracellular, but some WRF may also have
intracellular laccases (Burke and Cairney, 2002). Laccases catalyze the reduction of O2 to
H2O (4 electron reduction without formation of free reduced oxygen species) using a range of
phenolic compounds as hydrogen donors (Burke and Cairney, 2002; Schlosser and Hofer,
2002). An electron is removed from the phenolic hydroxyl groups of lignin to form free
phenoxy radicals, which are further oxidized to quinines (Hoegger et al., 2004).

Metabolic potential ofECM/ ERMfungi
As some ECM and ERM fungi are closely related to WRF, some researchers have suggested
that they may have retained some ability to degrade organic substrates, including PHCs

60

(Hibbett et al, 2000; Meharg and Caimey, 2000; Meharg, 2001). Braun-Lullemann,
Huttermann and Majcherczyk (1999) reported on the ability of 16 species (27 strains) of
ECM fungi that were isolated from middle European forests to degrade PAHs (1500 ppm of
phenanthrene, pyrene, chrysene and benzo[#]pyrene) in pure, liquid culture. The slow but
efficient metabolism of benzo[a]pyrene by ECM fungi was comparable to results of
experiments with WRF (Braun-Lullemann et al, 1999). A study in which 58 fungal isolates
from different physio-ecological groups were exposed to a range of PAHs showed that all
fungal groups could degrade PAHs, but that ECM fungi were 19% as efficient as WRF
(Gramss et al, 1999). Similarly, Meharg and Cairney (2000) tested 42 ECM fungal species
with several types of persistent organic pollutants and found that 33 species were able to
degrade one or more classes of chemicals. Only one of 21 ECM fungal species could not
degrade at least one PAH; degrading species seemed to prefer chemicals with four to five
rings. In each case, the direct oxidative activities correlated with production of extracellular
enzymes that appeared to metabolize aromatic rings (Braun-Lullemann et al, 1999; Gramss
et al, 1999). Green et al. (1999) reported that the ECM fungus Tylospora fibrillosa
degraded 4-fiuorobiphenyl to significant extents via sequential hydroxylation reactions.

A variety of ECM and ERM fungi are known to produce polyphenol oxidases (e.g. laccase,
catechol oxidase and tyrosinase) in culture conditions, but there is little evidence for
production of extracellular peroxidases (e.g. LiP and MnP) (Cairney and Burke, 1998; Burke
and Cairney, 2002). Timonen and Sen (1998) assayed macerated fungal mycelia from
several regions of a microcosm system and found that levels of enzyme activity in the
environment were lower for ECM fungi than for saprotrophic fungi, possibly due to

61

avoidance of competition or preferential exploitation of substances of a particular quality.
Gramss et al. (1998) reported that most cultured ECM fungi (from sporocarps collected from
previously contaminated forest plots and industrial sites) exhibited oxidase activity: high
extracellular enzyme activities were found for Lactarius and Russula species and high
intracellular enzyme activities were found for Suillus, Hebeloma, Leccinum and Tricholoma
species. Donnelly and Entry (1999) found extracellular enzyme activity at the advancing
hyphal front of ECM fungi and suggested the possibility of a lack of complete dependence
for C on the plant partners.

Few studies have considered mycorrhizal fungi in symbiosis with a plant for the degradation
of organic pollutants (Koivula et al., 2004). Meharg et al. (1997) found that the
mineralization rate of 14C-labelled 2,4-dichlorophenol by ECM fungi (Suillus and Paxillus
species) in symbiotic culture with P. sylvestris seedlings was increased by 50% and 250%
compared to respective rates of those ECM fungi in axenic culture. As mineralization was
extremely slow in vermiculite (i.e. no bacteria present), these data were interpreted to suggest
that fungal patch differentiation led to greater enrichment and stability of bacterial
communities at the fungal-soil interface. It is unknown how the C contributions of the
phytobiont influenced fungal responses or how synergistic or antagonistic interactions
between mycorrhizas and other microorganisms altered their ability to mineralize or degrade
organic pollutants.

62

Genetic controls
Ligninolytic enzymes are typically produced by WRF as multiple isoenzymes. This
biochemical diversity had been attributed to post-transcriptional modifications of a single
gene product, but characterization of several laccase gene families suggests that at least part
of this diversity could be due to the multiplicity of laccase genes in fungal genomes
(Gonzalez et ah, 2003). For example, Hoegger et al. (2004) found that Coprinopsis cinerea
has at least eight different laccase genes within the haploid genome, which is the largest
laccase gene family reported so far from a single haploid fungus. Bogan et al. (1996) found
that the genome of Phanerochaete chrysosporium contains at least 10 structurally related
genes {lipA through lipJ) encoding LiP proteins and at least three MnP genes (mnpl, mnp2,
mnp3).

Screening genomes for genes that encode laccases and peroxidases may represent a reliable
means of identifying potential enzymatic activities in ECM and ERM fungi (Burke and
Cairney, 2002). As DNA sequences for laccase and peroxidase genes are now available for
several saprotrophic fungi, opportunities exist to design molecular probes or primers for
identification of similar genes and/ or mRNA transcripts in mycorrhizal fungi (D'Souza et
al., 1996; Burke and Cairney, 2002). For example, Chen et al. (2003) used laccase gene
primers to screen ECM basidiomycetes for laccase-like genes, which were amplified from
Lactarius, Russula, Piloderma and Tylospora species. Timonen and Sen (1998) examined
gene expression in identified functional components of P. sylvestris mycorrhizal systems and
found expression of isozymes (i.e. polyphenol oxidase and acid phosphatase) was increased
in hyphal fronts of Paxillus involutus and Suillus bovinus ECM systems as they advanced in

63

the humus. Through gene amplification and sequencing, Chambers et ah (1999) reported
evidence for MnP genes and peroxidase activity in cultured Tylosporafibrillosa.

Chen et ah

(2001) extracted DNA from dried basidiomes or fungal cultures and amplified and sequenced
genes for LiP and MnP using primers based on Phanerochaete chrysosporium genes.
Although they reported the presence of LiP genes in a broad range of ECM fungal taxa and
MnP genes in some ECM fungal taxa (three Atheliaceae taxa), Cairney, Taylor and Burke
(2003) recently attempted to repeat these experiments and reported a lack of evidence to
support the presence of peroxidase genes in ECM fungi.

Extracellular laccase is constitutively produced in small amounts by several fungi, but
enzyme expression is considerably enhanced by a wide range of substances, including a
variety of different aromatic compounds (Burke and Cairney, 2002; Gonzalez et ah, 2003).
Regulation of laccase production appears to be complex and vary between taxa. For
example, in the WRF Pycnoporus cinnabarinus, laccase activity is increased with an increase
in C:N ratio whereas in Phanerochaete chrysosporium, laccase activity is repressed by
glucose regardless of N content and increased in the presence of cellulose when N is also
increased (Burke and Cairney, 2002). From assays of liquid cultures of the ECM fungus
Thelephora terrestris, Kanunfre and Zancan (1998) found increased secretion of extracellular
laccase with a decreased C:N ratio. At the molecular level, Collins and Dobson (1997)
demonstrated that laccase gene {Ice) transcription was activated by copper and nutrient N and
that induction occurred at the level of gene transcription in the presence of two aromatic
compounds. Chen et ah (2003) reported that some laccase-like genes amplified from

64

Lactarius, Russula, Piloderma, and Tylospora species appeared to be regulated at the
transcriptional level, with transcription enhanced by higher N content.

Implications for management
Two broad objectives dominate management goals for contaminated forest sites: (1)
reduction of risk; and, (2) long-term forest sustainability. In Canada, as in many
jurisdictions, contaminated site remediation is required if there is risk to human or
environmental health; this is not well defined for forest ecosystems and has led to situations
of either over- or under-management. Here we wish to reflect on both these objectives and
show how they converge.

Risk arises from the conjunction of three conditions: a contaminant (toxicant), a pathway and
a receptor. Mycorrhizal ecosystems in contaminated sites may attend to the toxicant by
metabolizing and removing or immobilizing it, or by transforming and altering its mobility
and toxicity. Moreover, mycorrhizal ecosystems may also be among the critical receptors.
Consequently, the concept of risk entails both remediation and ecotoxicology.

Bioremediation can be defined as the use of organisms to detoxify contaminants through
immobilization, chemical transformation and mineralization processes (Diaz, 2004).
Bioremediation efficacy is influenced by a variety of substrate and soil conditions, including:
PHC composition; soil temperature, texture and structure; length of time the PHCs have been
in the soil; and associated bioavailability, together with associated toxicants (Pollard et ah,
1994). Use of microorganisms as management tools requires knowledge of which organisms

65

(or functional guilds of organisms) are likely to be present in a particular ecosystem, how
they respond to different types of physical and chemical disturbances, and methods for
ascertaining whether organisms are actually healthy and not just surviving (Blakely et ah,
2002). From previous sections of this review, we can ask: (1) does the genetic potential exist
(or is it likely to exist) to metabolize the array of substrates expected in the PHCs at a given
site; (2) if so, what environmental conditions are likely to foster its expression and can these
conditions be achieved; (3) is there need for added genetic potential through genetic
engineering technologies; (4) if so, what constraints might limit its expression and what
precautions might be needed; (5) is bioaugmentation needed to increase genetic potential;
and, (6) if so, what precautions might be necessary, and how might its potential be best
exploited? Although much remains unknown, considerable insights have been gained from
recent and ongoing research regarding these questions.

Observations that virtually all organic substrates appear to be transformed by soil
microorganisms if they are accessible (e.g. Simard et al., 1997; Read and Perez-Moreno,
2003; Diaz, 2004; Heinonsalo et al, 2004), combined with the continually increasing
diversity of soil microbial communities revealed by molecular techniques (e.g. Axelrood et
al., 2002Z>; Berch et al, 2002), the observation that catabolism of PHCs and plant residues
share many common elements, and the absence of reports of soils that lack the ability to
metabolize PHCs, all point to the ubiquitous genetic potential by soil communities to
transform and perhaps completely catabolise PHCs. Consequently, bioremediation appears
to be a sensible and potentially feasible intervention and has been extensively used. The
focus of bioremediation strategies tends to be on contaminant disappearance, but is based on

66

a limited understanding of links between degradation and the basic nutritional needs of the
responsible soil microbial community (Mills et al., 2003). In addition, the optimum
environmental conditions to sustain communities that degrade PHCs are understood only in
broad terms. Continued progress may be expected by careful attention to, and documentation
of, the connection between the environmental conditions imposed on a site by PHC
contamination, as well as catabolic response and environmental preference by persistent
communities. In essence, PHCs impose their own environment, including varying
concentrations of toxicants. Consequently, PHCs control the community or guild that
survives, which in turn dictates the optimum conditions for its functioning. Based on first
principles, and observations on a wide range of chemo-organotrophic microorganisms, it is
reasonable to expect that a slightly acidic pH, well oxygenated and nutrient sufficient
environment would favour functional guilds that would metabolize PHCs. Attaining such
conditions, however, can be challenging in forest ecosystems without disrupting them.

Is there a need to use recombinant organisms? In the early development of bioremediation
technology, investigators recognized the scope of environmental pollution and the diversity
of chemical pollutants, and invested significant effort into metabolic engineering to
manipulate specific catabolic pathways or particular host cells (Alexander, 1999). Metabolic
engineering has created recombinant organisms with novel hybrid pathways of
biodegradation and increased substrate ranges; it has completed incomplete pathways,
created multiple pathways, and provided mechanisms that enhance chemical bioavailability
(Diaz, 2004). Because of issues (e.g. biosafety or inability to compete for resources)
associated with introducing recombinant bacteria to contaminated ecosystems,

67

bioremediation (using recombinant bacteria) is often conducted ex situ, under relatively
controlled conditions (Diaz, 2004). Ex situ bioremediation (with or without recombinant
bacteria) is also used in situations where a high degree of control of environmental conditions
is wanted (Riser-Roberts, 1998) and where added energy inputs such as in rotating
bioreactors are desired. Recombinant organisms face public resistance due to the fear of
their escape from the site, or transfer of genetic material to indigenous populations. Further,
they may not always compete well with indigenous populations, or may require specialized
environments. Consequently, they may be of limited potential for use on a large scale in
forested ecosystems.

There are fewer biosafety issues associated with bioaugmentation, the introduction of exotic
microorganisms isolated from unrelated sites, but adapted to contaminant biodegradation
(Ward et ah, 2003) or to extreme soil conditions (Cunningham et ah, 2004; Stallwood et ah,
2005). Such additions can be made in a variety of ways, including industrial production and
subsequent slurry applications. Soil samples from adjacent contaminated and remediated
sites used as an inoculum would seem to be reasonable candidates as well. Although it can
be readily used in the field, bioaugmentation lends itself better to highly engineered systems
(e.g. slurry bio-reactors), to recalcitrant or novel contaminants for which the indigenous
population may be ill-equipped, or for extreme environments (e.g. Cunningham et ah, 2004;
Stallwood et ah, 2005). Cost is also a factor from a management perspective. Leavitt and
Brown (1994) reported on three case studies comparing bioaugmentation using a commercial
supplement with stimulation of indigenous soil organisms for removal of PHCs in a
bioreactor and a land-treatment facility, and acetone or Ws-2-chloroethyl ether in a waste-

68

water facility. Based on cost and efficacy, they concluded that bioaugmentation was not
warranted in their situations and that biostimulation of indigenous organisms was the best
choice. In situ bioremediation through enhancing contaminant biodegradation by indigenous
soil populations and communities (i.e. syntrophic bioremediation) is considered less
destructive and more cost-effective for remediating contaminated soils on large scales
(Delille et ah, 2004; Doelman and Breedveld, 1999) and has proven successful when
properly implemented (Nelson et ah, 1994). In situ bioremediation is more likely to maintain
the desired integrity of below ground mycelial networks.

Phytoremediation refers to all plant-induced biological, chemical and physical processes that
aid in the remediation of contaminants (Cunningham et ah, 1996). Traditionally, research in
this area has focused on use of agricultural plant species for the remediation of agricultural or
industrial soils. Reilley et ah (1996) found that the presence of vegetation significantly
enhanced the dissipation (and likely biodegradation) of anthracene and pyrene in the soil
environment. Others have reported that various grasses, legumes and woody plants facilitate
the degradation of PHCs in soil (Aprill and Sims, 1990; Chaineau et ah, 2000; Liste and
Alexander, 2000; Palmroth et ah, 2002; Merkl et ah, 2005). Plants and associated
mycorrhizospheres may increase the activity of PHC-degrading organisms, either via general
enhancement [i.e. (mycor)rhizosphere effect] or due to proliferation of specific microbial
groups (i.e. altered functional component of the microbial community) (Siciliano et ah,
2003). They may also mediate desorption of contaminants bound to soil constituents by
altering pH and redox potential, as well as concentration and types of organic compounds in
the (mycor)rhizosphere. Research has shown that sites containing plants and expected

69

mycorrhizal associations experience more rapid reduction in toxicity of PHCs (Parrish et ah,
2005).

An increased awareness of the abundance and diversity of mycorrhizal systems in vegetated
soils has led to their consideration for in situ bioremediation (Meharg, 2001). Where there is
little risk to human or ecological health, the purposeful planting of trees inoculated with
specific mycorrhizal fungi is expected to establish these mycelial systems in soil and allow
gradual decontamination over a period of several years (Braun-Lullemann et al., 1999;
Meharg and Cairney, 2000). A related approach is to transplant plugs or sprigs of vegetation
from non-contaminated soil, as is done in various restoration ecology projects (e.g. Fraser
and Kindscher, 2005), into contaminated areas for the final stages of clean up. This approach
allows for simultaneous remediation and revegetation of sites without further disruption to
physical and chemical properties of the soil and provides an inoculum of a soil community
adapted to the site. Although it may require several months or years for tree root systems and
associated mycorrhizal biomass to establish, mycelial systems would be expected to remain
in a vital state for several decades, whereas other organisms (e.g. WRF) may complete their
life cycles in a few days or weeks and then rest as spores (Gramss et al, 1999). A more
thorough knowledge of which fungal symbionts are likely to survive and compete in various
ecosystems, as well as which fungi contribute directly (exhibit biodegradative capabilities) or
indirectly (provide suitable habitat for other microorganisms that exhibit biodegradative
capabilities) to bioremediation of contaminated sites is required as part of management
strategies that adopt this approach. Other phenomena, such as fungal specificity to plant
hosts, may also require consideration (Molina et al., 1992). The spectrum of possible plant

70

hosts that can be selected by a particular mycorrhizal fungus can vary from a few to many.
At the same time, the host receptivity (the number of different fungi accepted by a particular
plant) can also differ. Both may impact the ecological contribution made by the symbiosis.
For example, alder, compared to Douglas-fir, is very selective, typically initiating symbioses
with a very restricted number of fungal species. Some fungi may form symbioses with one
plant genus, whereas other fungi are less selective, initiating symbioses with potentially
hundreds of plant hosts. How this selective nature between fungi and plants impacts
ecological functions in general, and bioremediation in particular, remains unknown.

Intrinsic bioremediation (contaminant biodegradation by adapted indigenous communities)
may be an acceptable management strategy where risks to human or ecological health are
low (Alexander, 1999). Nicolotti and Egli (1998) showed that some ECM fungi surviving in
contaminated forest soil may metabolize chemicals in crude oil and suggested that crude oil
spills in mixed agricultural and forest areas do not cause long-term environmental damage of
the kind associated with coastal ecosystems, possibly due to intrinsic bioremediation by the
soil community. Intrinsic bioremediation of PHC-polluted soils differs substantially between
ecosystems and depends on the particular combination of soil-forming factors, soil
properties, microbial communities and the content and composition of PHCs and their
products (Trofimov and Rozanova, 2003). Studies of indigenous microbiota along with key
physical and chemical parameters provide hope for the eventual ability to predict how natural
attenuation will proceed at contaminated sites. Intrinsic bioremediation may be an
appropriate management strategy in boreal forest ecosystems in some situations.

71

If external action to improve the bioremediation of a soil seems essential, environmental and
ecological knowledge is required to choose the least destructive methods (Romantschuk et
al., 2000). Ideally, bioremediation strategies should be based on knowledge of the
microorganisms present in polluted environments, their metabolic abilities and how they
respond to changes in environmental conditions in an ecological context (Blakely et al.,
2002; Diaz, 2004). The current state of knowledge, however, does not permit predictions or
management strategies to be built up from the species level. Studying the physiology,
biochemistry and genetics of soil microorganisms is important for contaminated site
bioremediation, as well as for biomonitoring the impacts of chemicals as disturbance agents.
Unfortunately, this is mostly unknown and current management is largely based on empirical
rather than theory-based deductions. As demonstrated by the research described in this
review, factors that alter the survival or activity of soil biota are important considerations for
ecosystem management (Setala et al., 2000). However, the key question in terms of
sustainability is how contamination events impact ecosystem functions in the near and longterm future.

Ecosystem health is not well defined, but has been described in terms of vigour
(productivity), organization (diversity and mutual dependence) and resilience (maintenance
of structures and patterns in the presence of environmental change) and interpreted by
correlating biological indicators for processes that are considered critical for ecosystem
function (Blakely et al, 2002; Lu and Li, 2003). Traditional indicators for contaminated
sites include impacts of chemical contaminants on plant condition and biomass, with more
recent interest in specific organisms capable of biodegradation. Fewer studies have

72

examined changes in soil properties that define its fertility and agro(eco)logical properties
(Suleimanov et al, 2005). It has been suggested that criteria for remediation of PHCcontaminated soils should be amended to include suppression or significant modification of
the plant community, reduction in plant biomass, disturbance in functioning of soil biota,
simplification of the soil community, decreases in the biological activity of the soil, and
movement of PHCs into surface water or ground water (Trofimov and Rozanova, 2003). Soil
quality guidelines in Canada have been developed using most of these criteria for
agricultural, residential/ urban parkland, commercial and industrial lands, but not for forest
ecosystems (Ouellet et al., 2002).

Many biological and chemical-physical approaches have been proposed to predict or measure
the bioavailability of organic compounds for biodegradation or to ecological receptors (i.e.
toxicity). Biological measures of bioavailable toxicants include seedling emergence and
growth tests, along with various soil invertebrate tests of acute toxicity, chronic toxicity,
behaviour and reproduction (Stephenson et al, 2002). Rombke et al. (2006) recently
identified potential invertebrate species and testing methodologies for assessing ecotoxicity
of contaminants in boreal forest soils. Other biological measures include contaminant uptake
and impact on organism biomolecules, and impacts on organism-mediated processes (e.g.
nitrification) (Svendsen et al., 2002). Chemical-physical approaches include: kinetics of
PHC desorption, mild solvent extraction, solid phase extraction (SPE), supercritical fluid
extraction and cyclodextrin extraction. SPE correlates well with biological measures of
ecotoxicity (and bioavailability) and does not have the potential to disrupt the structure of
soil organic matter phases as do chemical methods (Ehlers and Loibner, 2006).

73

Conclusions
(1) The importance of developing multi-disciplinary approaches to solving problems relating
to anthropogenic pollution is now clearly appreciated by the scientific community, and this is
especially evident in boreal ecosystems exposed to escalating threats to PHC contamination
through expanded natural resource extraction activities, hi this review, we have presented a
mycorrhizal ecosystems perspective on PHC contamination in boreal forest soils in order to
identify gaps in knowledge and to guide future research in both ecological and sustainable
management contexts so that scientists, land and facilities managers, industrialists and
government officials will be better prepared to manage the inevitable accidents that will
occur.

(2) We know that the taxonomic, genetic and functional diversity of mycorrhizal ecosystems
in boreal soils is immense and continues to expand with increased sampling effort. We also
know that the functioning of these communities underpins survival and productivity of the
ecosystem as a whole. It appears that redundancy in broad-scale biodegradative functions is
essential for ecosystem recovery following PHC contamination, to account for the loss of
community components that are unable to tolerate the altered physical and chemical
conditions imposed by the PHCs. What remains to be determined are the details and the
translation of this information into effective ecosystem management.

(3) The ubiquity and enormous biomass of extraradical mycelia of mycorrhizal fungi in
forest soils implies a key role in forest ecosystem processes. Recent studies have highlighted

74

the high taxonomic and genetic diversity of ECM or ERJVI fungal communities associated
with certain plants in some ecosystems, though many potential hosts and types of ecosystems
have not yet been surveyed.

(4) Although there is thought to be some relationship between high diversity (species
richness) and ecosystem health due to some degree of redundancy, the functional basis of
ECM and ERM fungal diversity is virtually unknown. The physiological mechanisms of
nutrient exchange between fungal and plant partners are also not well understood,
particularly with respect to nutrient acquisition from the soil environment and especially for
PHC-contaminated soils. A more thorough knowledge of which fungal symbionts are likely
to survive and compete in various ecosystems is required, as well as a better understanding of
whether certain types of fungal associations with different plant hosts gain in ecological
importance following disturbance events. Whereas community responses (e.g. shifts in
community structure) to some types of disturbances (e.g. fire, forestry practices, etc.) have
been described in the recent literature (although it is unknown if these are related to shifts in
function), responses to PHC pollution are not well understood. In fact, very little is known
regarding rhizosphere communities in forest soils subjected to PHC contamination.

(5) Studies of PHC contamination in forest soils are rare, as are the impacts on soil
organisms and the intrinsic decomposition in these systems. The scientific basis for current
remediation standards is based on information from experiments examining the toxicological
impacts of PHC chemicals on test organisms. However, sequestration of organic pollutants
in forest SOM may decrease the chemical toxicity of chemicals through physical separation

75

from biological receptors, which also decreases substrate bioavailability for enzymatic
degradation. More research in this area is needed. Also necessary are improved methods for
assessing the fate and behaviour of PHCs in forest soils, including determinations of
bioavailability and development of a wider variety of indicators for ecological integrity than
the traditional measures of plant productivity. Future research is needed to determine how
toxicity varies with type of pollutant, mixtures of pollutants, extent of pollution, and the
general condition of the ecosystem prior to chemical disturbance.

(6) Few studies have examined whether the coexistence of ECM and ERM plants in boreal
forests provides opportunities for sharing ECM and ERM fungi that link plants and
translocate nutrients, and virtually nothing is known of how PHC contamination may
interfere with processes of nutrient acquisition and exchange.

(7) Recent studies have shown that some ECM and ERM fungi appear to play a direct role
(via enzymatic catabolism) in biodegradation of complex organic substrates (including
PHCs). However, few studies have examined various fungi in detail, or have examined
mycorrhizal fungi in symbiosis with a plant. It is unknown as to how PHC contamination
might interfere with fungal metabolic processes.

(8) Incomplete biodegradation can produce potentially toxic intermediate metabolites; toxic
intermediates with increased water solubility are of particular concern as this can result in the
transport and spread of contaminants through the environment. More research is required in
this area.

76

(9) Few studies have considered the indirect role of ECM and ERM systems in
biodegradation through their interactions with the mycorrhizosphere-associated bacterial
communities and little is known regarding syntrophic biodegradation by different functional
guilds of organisms. Most fungi have been examined in isolation from an ecosystem context,
thereby excluding interactions of individual ECMs and ERMs with each other, their soil
environment and other members of the plant and microbial communities. Thus, information
gained from these studies may have little ecological relevance for understanding how forest
ecosystems function or for informing bioremediation management strategies for
contaminated soils.

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Chapter 2: Interactions between petroleum hydrocarbon contaminants and ecto- and
ericoid mycorrhizal communities in sub-boreal forest soils

Abstract
The impacts of petroleum hydrocarbon (PHC) contamination on mycorrhizal communities in
boreal forest soils are not well understood. In this study, we used a bioassay approach to
determine whether ecologically relevant concentrations of PHCs altered ecto- (ECM) and
ericoid (ERM) mycorrhizal fungal communities and whether mycorrhizal communities
played a direct role in PHC biodegradation through secretion of laccase. Surface-sterilized
seeds (Pinus contorta, lodgepole pine; Betulapapyrifera, paper birch) or seedlings
(Vaccinium vitis-idaea, lingonberry) were planted into Cone-tainer™ pots containing
reconstructed soils: an organic layer (mor humus, coarse woody debris, or previously
contaminated humus) overlying sandy mineral horizons (Ae and Bf) of field-collected forest
soils obtained from central BC, Canada. After 4 months, BC light crude oil (0, 73, 146, or
219 mg cm"2) was applied to the soil surface around the seedling stem; systems were
destructively sampled over 16 weeks following treatment. ECM communities (composition,
relative abundance and spatial distribution) were assessed on pine and birch roots using light
microscopy and fungal community profiles were generated for all root systems using length
heterogeneity PCR and primers targeted at the ITS region of rDNA. In addition, selected
mycorrhizal root tips (ECM and ERM) were tested for laccase activity in assays with 2,2'azinobis-3-ethylbenzthiazoline-6-sulfonate (ABTS). There were no significant changes in
ECM or ERM fungal community structure associated with PHC treatment; however, PHC
treated systems appeared to exhibit some community differences over time. Plant and
organic soil properties appeared to have a greater influence on mycorrhizal community

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structure than PHC contamination. In addition, laccase activity was observed for many
dominant ECMs, indicating potential for biodegradation of aromatic PHCs. Thus ecological
integrity of the plant - soil system may confer resilience against effects of PHC
contamination and also contribute to PHC biodegradation.

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Introduction
Oil spills that occur in northern forest regions are localized and discrete disturbance events
that often result in rapid surface contamination, sometimes with large quantities of petroleum
hydrocarbons (PHCs). Very little is known regarding the impacts of PHC contamination on
soil communities, particularly on mycorrhizal systems that represent the dominant microbial
biomass in forests soils and underpin ecological processes (e.g. decomposition, nutrient
cycling, primary production, C sequestration, etc.) both below- and above-ground (Read and
Perez-Moreno, 2003). The major impacts of PHC contamination appear to be associated
with disturbances to soil water, nutrient and oxygen regimes through changes to physical and
chemical properties of soil (Blakely et ah, 2002). Changes to soil chemical status and
decomposer community functions could lead to imbalances in nutrient cycling and ecosystem
productivity (Pennanen et ah, 1998). Enhanced hydrophobicity of soils after fire has been
shown to alter soil water-holding capacity and lead to changes in both quantity and
composition of microbial and soil-dwelling invertebrate communities, potentially reducing
plant productivity (Certini, 2005). Thus, factors that alter the survival or activity of
mycorrhizal communities are important considerations for sustainable management of forest
ecosystems (Setala et ah, 2000). However, this requires that environmental and functional
properties assessed at population and community levels be integrated with ecological
processes at the ecosystem or landscape scale; this represents a primary challenge of
microbial ecology (Bengtsson, 1998; Standing et ah, 2007).

In contaminated forests, oil recovery and soil remediation strategies tend to be based on
socio-economic considerations and environmental concentrations of some PHCs (e.g.

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polycyclic aromatic hydrocarbons, PAHs) with potential toxicity/ carcinogenicity to human
and other ecological receptors (Tarradellas and Bitton, 1997; Nicolotti and Egli, 1998;
Blakely et ah, 2002; Trofimov and Rozanova, 2003). Despite considerable environmental
interest in PAHs, a general absence of documented adverse effects among terrestrial
receptors (i.e. plants, invertebrates, birds, herpetofauna, or mammals) exposed to PAHs in the
environment currently exists in the literature, although this is partly due to lack of research
(Kapustka, 2004). Nearly all information regarding PHC toxicity on soil microorganisms has
been compiled from laboratory tests that have focused on a few (e.g. PAHs) of the hundreds
of chemical compounds that comprise crude oil. Results have generally shown that fungi are
more tolerant than bacteria, possibly due to features of cell wall structure (Blakely et al.,
2002). However, in addition to reducing the variety and complexity of the biological
systems affected by the release of crude oil into the forest environment, these studies nearly
always disregard the symbiotic relationship between fungi and plant hosts, which may be
very important in terms of buffering negative impacts. Soil-based studies have found that
ectomycorrhizal communities exhibit high resilience to environmental stress when organic
layers (e.g. humus, woody debris, etc.) of forest soils have not been severely disrupted
(Setala et al., 2000; Jones et al., 2003). Greater environmental disturbance may occur when
extreme management measures (e.g. tree or contaminated soil removal) are taken as
compared to the original PHC contamination event. The lack of understanding of PHC
impacts on mycorrhizal communities in forest soils places remediation strategies in potential
conflict with other current forest management objectives.

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In situ bioremediation (i.e. enhanced contaminant biodegradation by indigenous soil
communities) is considered less destructive and more cost-effective for remediation of
contaminated soils on large scales and is also more likely to maintain the desired integrity of
mycelial networks (Doelman and Breedveld, 1999). PHC biodegradation is likely due to
species-specific fungal attributes that favor survival and degradation activities, extension of
the root area in contact with soil-adsorbed compounds, and influence over a complex
consortium of bacteria expressing a range of enzyme activities in response to carbon
availability in the mycorrhizosphere (Heinonsalo et al., 2000; Meharg and Cairney, 2000).
For their part, many ecto- (ECM) and ericoid (ERM) mycorrhizal fungi appear to secrete
various hydrolytic and oxidative enzymes that enable direct involvement in the degradation
and/ or detoxification of various organic contaminants (Timonen and Sen, 1998; Burke and
Cairney, 2002; Perotto et ah, 2002). For example, laccase (benzendiol:oxygen
oxidoreductase, EC 1.10.3.2), part of a complex, nonspecific enzyme system usually
associated with ligninolytic fungi, has been of particular interest with respect to
bioremediation (Gramss et ah, 1998; Donnelly and Entry, 1999). This enzyme catalyzes the
reduction of O2 to H2O (4 electron reduction without formation of free reduced oxygen
species) to open aromatic ring structures of a range of organic substrates, thereby overcoming
the thermodynamically rate-limiting step in microbial metabolic pathways (Burke and
Cairney, 2002). In addition to lignin and PAH depolymerization, laccase is thought to be
involved in the release of N from insoluble protein-tannin complexes, humus formation,
mycelial pigmentation, and fruiting body formation (Kanunfre and Zancan, 1998; Burke and
Cairney, 2002). It should be of no surprise that extracellular laccase appears to be
constitutively produced in small amounts by several mycorrhizal fungi; it is unknown

98

whether enzyme expression is enhanced in the presence of different aromatic compounds as
appears to be the case for some saprotrophic fungi (Burke and Cairney, 2002).

This study addresses a fundamental question: does PHC contamination of forest soils
negatively impact established (or establishing) mycorrhizal systems? Using a bioassay
(single- and double-plant systems established in reconstructed forest soil layers in Conetainer™ pots), morphological and molecular (LH-PCR) techniques were used to assess
changes in mycorrhizal community structure in response to different PHC concentrations
over 16 weeks. In these systems, organic soil layers (i.e. forest floor [FH], coarse woody
debris [CWD], and forest floor previously contaminated with PHCs [FHoil]) provided the
initial fungal inoculum for the developing rhizospheres; the composition of fungal
communities was expected to be influenced by the relative abundance of lignin (i.e. in FH
and CWD) and/ or previous exposure to PHCs (i.e. in FHoil). ECM (Pinus contorta var.
latifolia and Betulapapyrifera) and ERM (Vaccinium vitis-idaea) systems were expected to
represent a sub-set of the mycorrhizal fungal community in situ. Laccase assays were also
conducted to assess potential for direct role of ECM and ERM fungi in PHC biodegradation
and assess changes in enzyme activity associated with PHC treatment.

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Materials and methods
Field site
The Kenneth Creek field site is located in the wet, cool subzone of the sub-boreal spruce
(SBSwkl) biogeoclimatic zone of central British Columbia, Canada, about 100 km east of
Prince George (53°34'N, 122°47'W) (Figure 2.1). In 1982, the forest was logged and
burned, then subsequently planted with lodgepole pine (Pinus contorta Dougl. Ex Loud. var.
latifolia Engelm.); currently, the site is a mature, even-aged pine stand with small hybrid
white spruce {Picea glauca x engelmannii Parry ex Engelm.) and lesser numbers of western
hemlock (Tsuga heterophylla (Raf.) Sarg.). Young subalpine fir {Abies lasiocarpa (Hook.)
Nutt.) and sitka alder (Alnus crispa var. sinuata (Reg.) Rydb.) are present at the edge of the
forest, along the access road; western redcedar {Thujaplicata Donn) and trembling aspen
{Populus tremuloides Michx.) are also present in an unlogged stand across the main road.
The site has a thick understory of oval-leaved blueberry {Vaccinium ovalifolium Sm.);
mosses and lichens (e.g. Peltigera) cover the forest floor, with some Lycopodium species.

The soils on this site were described by Arocena and Sanborn (1999). Soils are classified as
Eluviated Dystric Brunisols (Soil Classification Working Group, 1998), and consist of sandy
parent material with low clay content and few coarse fragments. The forest floor is mor
humus, from 2-5 cm thick with copious fungal mycelia present. The C:N ratio of the forest
floor is approximately 50 and the pH (water) is ~ 4.2. Gray Ae horizons are generally 1-2 cm
thick, with thicker pockets in some areas. Red Bf horizons extend to almost 30 cm, beneath
which are Bm (27-60 cm), BC (60-100) and C (> 1 m) horizons. The C:N ratios of the Ae
and Bf horizons are about 20 and 13, respectively. The pH (water) of the Ae horizon is 4.2

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while the pH of the Bf horizon is about 4.8, increasing with depth to about 6.0 at the
transition to the C horizon. Fine roots are found at depths greater than 1 m. Large coarse
woody debris (i.e. downed trees), the legacy of past forest management, is abundant all over
the site.

Organic layers and the top 20-30 cm of mineral soils (Ae and Bf horizons) were collected
from the forest site in September of 2004, 2005 and 2006. The organic layers included the
mor humus forest floor (FH) that had been undisturbed for approximately 20 years, coarse
woody debris (CWD) in an advanced state of decay (i.e. decay class 5), and previously PHCcontaminated mor humus (FHoil) that had weathered in situ for four months throughout the
summer. The FHoil soils were collected in 2005 (year 2) only. In early May of that year, 2.0
L of BC light crude oil (Husky Refinery, Prince George, BC) was applied to each of three 1m2 plots with the moss layer (but not seedlings and other plants) removed. The oil had been
previously bubbled with N2 gas to remove the light and volatile compounds; a watering can
was used for even application in the field. All soils were stored at 4°C prior to use in
bioassay experiments, which commenced within 10 days of soil collection.

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Figure 2.1: Maps showing the study site at the Kenneth Creek field site, located about 100
km east of Prince George in the sub-boreal spruce (SBSwkl) zone of the central interior of
British Columbia, Canada.

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Bioassay and PHC treatment
Forest soils (i.e. Ae [~1 cm] and Bf [~15 cm] mineral soil layers beneath organic [FH, CWD,
or FHoil] soil layers [~2 cm]) were reconstructed in Cone-tainer™ pots (3.8 x 21 cm, Stuewe
and Sons, Corvallis, Oregon) with two clay pellets in the bottom to prevent soil loss (Setala
et ah, 2000). Mineral soils (Ae and Bf layers) were homogenized and sieved through (1 cm2)
screens prior to potting. In 2004 and 2005 (years 1 and 2), surface-sterilized seeds of
lodgepole pine (Pinus contorta var. latifolia) and paper birch (Betulapapyri/era Marsh.),
collected from the SBS and obtained from the Ministry Tree Seed Center, Surrey, BC (Seed
lots DWD20050009A (location 079-B-008) and DWD20050009B (location 094-E-015),
respectively), were planted into each pot. In 2005 (year 2), lingonberry (Vaccinium vitisidaea L.) seedlings (rooted cuttings), obtained from Birch Creek Nursery (Prince George,
BC), were planted into 10x10x10 cm pots (i.e. Ae [~1 cm] and Bf [~7 cm] mineral soil layers
beneath organic [~2 cm] soil layers). In 2006 (year 3), pine seeds were planted into Conetainer™ soil systems (FH and CWD organic layers) that were already planted with
lingonberry cuttings. All pots were placed in the greenhouse (22°C day temperature, 15°C
night temperature, and 16 h photoperiod) and fertilized once a month (5 mL of NPK
fertilizer; providing 100 ppm each of NPK) for the first four months following seeding/
planting. The plants were watered two or three times per week for the duration of the
experiment.

After four months of growth, seedlings and mycorrhizas were well established. At this time,
BC light crude oil (with volatiles removed as described for the field application) was pipetted
onto the organic soil surface of each pot, around, but not touching, the seedling stem(s). To

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determine effects of different levels of PHC contamination on plant - soil systems, 1, 2, or 3
mL (corresponding to 73, 146, and 219 mg cm"2, respectively) crude oil was tested on pine
and birch seedlings in 2004-05 (year 1). In the two subsequent years of study (i.e. years 2
and 3), pine, birch, and lingonberry seedlings were treated with the highest (219 mg cm"2)
PHC level tested, which corresponded to a field application rate of nearly 22 tonnes ha"1.
There was no PHC loss from the bottom of the pot (i.e. no sheen on the wet surface below)
observed after watering for the duration of the experiment and the smell of crude oil
dissipated in the greenhouse within a week of PHC treatment. These observations provided
some confidence that most of the oil was retained within the plant - soil systems (i.e. not lost
through volatilization or washing out).

Experimental design and sampling
Experiments followed a randomized block design and tested different combinations of plant,
organic soil layer and PHC treatments (Tables 2.1 and 2.2) over three years of studies. In
2004-05 (year 1), treatment combinations included plant (P or B) and organic layer (FH or
CWD) systems in PHC contaminated and untreated (control) soils (Table 2.1). Plant - soil
systems (n=2 per treatment group) were destructively sampled at 1, 4, 7, 10, 13 and 16 weeks
following PHC treatment in order to assess ECM morphotype community changes over time
in response to different levels (0, 73, 146, and 219 mg cm"2) of contamination. Single-plant
(pine, birch, and lingonberry) and double-plant (pine and lingonberry) systems (2005-06 and
2006-07, respectively) were harvested at 1 and 16 weeks to further assess relationships
between ECM and ERM fungal communities and PHC biodegradation (Table 2.2).

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Table 2.1: Summary of plant, organic soil layer, and PHC treatment variables for 2004-2005
study (n=2).
Plant
Pine [P]
Birch [B]

Organic Soil Layer
Forest floor [FH]
Coarse woody debris [CWD]

PHC
No PHC (control)
PHC (73 mg cm"2)
PHC (146 mg cm"2)
PHC (219 mg cm"2)

Table 2.2: Summary of plant, organic soil layer, and PHC treatment variables for 2005-2006
(single-plant) and 2006-2007 (double-plant) studies (n=3)*.
Plant
Organic Soil Layer
PHC
Pine [P]
Forest floor [FH]
No PHC (control)
Birch [B]
Coarse woody debris [CWD]
PHC (219 mg cm"2)
Lingonberry [L]
Contaminated forest floor [FHoil]
Pine + Lingonberry [P+L]
* n<3 in B-CWD and B-FHoil groups due to lack of seed germination

Root systems (intact, but with the shoot excised) were shaken free from the soil (samples for
PHC and nutrient analysis, pH and wet weights were collected concurrently, as described in
Chapter 3) and then washed in sterile dH20. A detailed morphological analysis of pine and
birch ECMs was conducted in 2004-05 (year 1). For this, root systems were stored in glass
plates with sterile water at 4°C for up to a week following each harvest. Molecular analysis
of ECM/ ERM fungi was conducted in 2005-06 (year 2) for the single-plant systems and in
2006-07 (year 3) for the double-plant systems. Within a few days of seedling harvest, pine
and birch root systems were quickly assessed under a dissecting microscope to note ECM
presence, abundance and distribution within the soil system and to ensure that they were free
of other roots or hyphae and soil particles. Samples of lingonberry hair roots were assessed
for cortical cell colonization by root endophytes (i.e. cells appeared cloudy, not clear), some
of which were expected to be ERM fungi. Except for a few root tips removed for laccase

105

assays, which were also performed at the 1 and 16-week harvest times, entire root systems
were then collected in 2 mL tubes and stored at -20°C until DNA extraction.

Morphological assessment ofECM communities
Standard light microscopy techniques of Agerer (1987-2002), Ingleby et al. (1990) and
Goodman et al. (1996) were used to group, quantify and spatially describe ECM
morphotypes of PHC treated and untreated (control) pine and birch seedlings at 3-week
intervals over the 16-week duration of the experiment. ECMs were initially described and
grouped according to colour, texture, lustre, dimensions, tip shape, branching pattern, and
presence or absence of rhizomorphs (mycelial strands). Root squash mounts were examined
at 400-1000X magnification and descriptions of mantle features, emanating hyphae,
rhizomorphs, and other distinguishing features were used to further group different ECM
morphotypes. Root tips that appeared uncolonized or partially colonized (due to the lack of a
well-developed mantle) were categorized as non-mycorrhizal. When possible, preliminary
identifications to fungal families or genera were assigned; otherwise, a descriptive name was
assigned.

ECM community diversity was represented by the Simpson diversity index, a non-parametric
measure of relative abundance of each ECM morphotype weighted more towards the
abundant types (Magurran, 2004). Simpson diversity (D) was calculated for each root
system from D = S pj2, where pi represents the proportion of individuals in the ith species, and
reported as 1/D. Diversity values were compared using one-way ANOVA to determine
significant differences (a=0.05) with level of PHC treatment, over time, and between the

106

plant - soil systems. The post-hoc Fisher's LSD test (a=0.05) was performed when a
significant result was obtained.

DNA extraction and length heterogeneity PCR
Frozen root systems were crushed in liquid nitrogen and DNA was extracted using a CTAB
(hexadecyl trimethyl ammonium bromide) protocol with an extra phenol/chloroform-isoamyl
alcohol (1:1) purification step (Fujimura et al, 2008). DNA extracts were further cleaned
using the Wizard® PCR Preps DNA Purification System kit (Promega) to remove phenolics
and other oily contaminants. The cleaned DNA extracts were resuspended in TE buffer.

Fungal communities were characterized by amplicon length heterogeneity PCR (LH-PCR),
which provides an estimate of community structure based on relative abundance of genotypes
(Martin and Rygiewicz, 2005). Community fingerprinting methods such as LH-PCR and
terminal restriction fragment length polymorphism (TRFLP) provide little taxonomic
resolution of microbial communities compared to methods such as sequencing, but are
expected to provide sufficient resolution to separate communities based on broad variables
(Kuyper and Landeweert, 2002). The ITS2 region of ribosomal DNA was amplified using
the forward primer ITS3 (5'GCATCGATGAAGAACGCAGC) (White et al, 1990) and the
D3 fluorescent dye-labeled reverse primer NL4B (5'GGATTCTCACCCTCTATGAC)
(Martin and Rygiewicz, 2005). ITS3 is a universal primer that binds in a conserved domain
128 bp from the 3' end of the 5.8S rDNA while NL4B binds in the large subunit (28S) at
basidiomycete- (and ascomycete-) specific sites (Martin and Rygiewicz, 2005); PCR
products are expected to vary from approximately 400 to >600 bp in length. PCR reactions

107

consisted of 10X PCR buffer, 2 mM dNTPs, 50 uM MgCl2, 10 uM forward and reverse
primers (Proligo, CO), 0.7 U Platinum Taq DNA polymerase (Invitrogen Life Technologies),
and nuclease-free water (Integrated DNA Technologies, Inc.) to a final volume of 27 uL, to
which 3 uL DNA (diluted 1:10) was added. The DNA Engine DYAD™ thermocycler (MJ
Research, Inc., Watertown, MA) conditions were as follows: initial denaturation for 4 min at
94°C, annealing for 1 min at 48°C, and extension for 2 min, followed by 35 cycles of
denaturation (94°C for 30 s), annealing (48°C for 30 s) and extension (72°C for 1 min 30 s)
and final extension at 72°C for 6 min 30 s. All PCR products were run on 1.2% agarose gels
to confirm amplification.

PCR products (2 uL) were loaded into a CEQ™ 8000 sequencer (Beckman-Coulter Inc.)
along with CEQ 600 bp size standard mixture. Run conditions were 60°C separation
temperature, 4 kV voltage, and 120 min separation time. Analysis was performed using the
amplicon fragment length polymorphism (AFLP) program of the CEQ™ 8000 sequencer and
the quartic model for size standard with a bin width of 1.5 bp. Peaks less than 11% of the
total sample peak height were not included. Profiles from separate DNA extractions and
PCR reactions were compared to assess reproducibility and suitability for analysis.

The relative abundance of genotypes comprising each sample (i.e. fungal community) was
calculated after relativizing the fluorescent signal strength of each fragment peak to the total
peak area within each sample. This step in the analysis set the minimum peak threshold by
removing the small peaks resulting from noise or reflecting the amount of DNA analyzed that
may impact the conclusions drawn (Osborne et ah, 2006).

108

Ordination techniques are used to arrange entities along single or multiple axes that
summarize the continuous trend within data; samples with similar community composition or
environmental characteristics are grouped more closely in the ordination space (McCune and
Grace, 2002). Nonmetric Multidimensional Scaling (NMS) is a preferred method for
analysis of community data because it does not carry assumptions of linearity among
variables (required for parametric approaches) (Ramette, 2007). NMS preserves similarity
distances in ranked order (i.e. nonparametric) and tends to linearize distances in species and
environmental space; it is not constrained to any specific distance measure. In this study,
NMS was calculated on the basis of a Sorensen distance measure with 50 runs with real and
randomized data and a maximum of 500 iterations to assess stability (instability criterion was
0.00001) using PC-ORD 5.0 software (McCune and Mefford, 1999). A stepwise reduction in
dimensionality (6D-1D) was used to minimize stress along with a random starting
configuration (user-provided seeds). Two measures were used to evaluate the structure of the
ordination results. Stress (i.e. goodness of fit measure) is the deviation from monotonicity
when distance is compared between the original species space and distance in the reduced
ordination space (McCune and Grace, 2002). For community data, it is typically in the 10-20
range, but should be interpreted cautiously at the upper end of this scale. Instability is a
measure of change in stress at each iteration. Stable, low stress solutions indicate strong data
structure. The final solution for NMS was accepted after comparing 50 runs with real to
randomized data using Monte Carlo simulations (McCune and Grace, 2002).

109

Univariate comparisons between treatment groups were tested statistically with MultiResponse Permutation Procedures (MRPP), a non-parametric method that tests the
hypothesis that there is no difference among groups (McCune and Grace, 2002). It provides
a statistic of the magnitude of differences between groups (analogous to effect size), given as
the chance-corrected, within group agreement (A) and a p-value. A is equal to 1 when all
items are identical within groups and 0 when heterogeneity within groups equals expectation
by chance. As most comparisons were made between unequal group sizes (i.e. unbalanced
analyses), multivariate differences were not tested statistically with nested permutational
multivariate ANOVA, and thus direct interaction effects could not be detected.

Laccase assays
Laccase assays were conducted in 2006 and 2007 (years 2 and 3) with pine and birch (ECM)
root tips and lingonberry (ERM) hair roots. Laccase activity was determined using a plate
assay with 2,2'-azinobis-3-ethylbenzthiazoline-6-sulfonate (ABTS; Sigma Chemical Co., St.
Louis, Mo.) as the substrate (Eggert et al. 1996). Freshly harvested plant roots were viewed
under a dissecting microscope to select ECM morphotypes and to confirm the presence of
colonized epidermal cells in ERM hair roots. These were removed from the root systems and
placed into individual wells of a microplate, each well containing 100 uL of 50 mM glycineHC1 buffer (pH 3). Following addition of 50 uL ABTS (2 mM) to each well, the plates were
incubated at ambient temperature (~22°C) for 24 h. The intensity of the bluish green color
development was rated as nil (-), light (+) or dark (++) after 1,16 and 24 h of incubation. In
an attempt to exclude reactions mediated by other potentially-present oxidative enzymes,
assays were conducted at pH 3, the optimum pH for laccase activity in the white rot fungus,

110

P. chrysosporium with ABTS as the substrate; peroxidase reactions with ABTS generally
occurred at pH 4-7. A small piece of field-collected Trametes fungus (collected behind the
EFL at UNBC) was used as a positive control (dark green colour developed within an hour of
incubation) for this assay; boiled fungus was used as the negative control (no colour
development after several days).

Results
ECM morphotypes: development and response to PHCs
All seedlings and root systems survived for 16 weeks following PHC treatment. A greater
proportion of seedlings generally appeared less healthy (i.e. yellowed or red needles/ leaves)
with the highest PHC treatment compared to controls, but untreated plants also occasionally
appeared chlorotic. There was no difference in seedling height or root system development
between control and PHC treated groups (data not shown).

Virtually all pine and birch root tips were visibly ectomycorrhizal at the first harvest time
(i.e. approximately 4 months post-seed germination and 1 week post-PHC treatment). The
richness of ECMs associated with each root system ranged from two to seven morphotypes,
with fewer distinct morphotypes observed on birch compared to pine roots. Morphotype
richness tended to increase over time (i.e. 1-16 weeks) for both plant species and organic soil
types. In general, ECMs, extraradical mycelia, and rhizomorphs were more extensively
developed by the 16-week sampling time, which may have contributed to their greater
visibility. Differences between pine and birch root system structures influenced the
distribution of ECMs through the soil system, as well as ECM density within the pot and

111

proximity to other root tips. Pine root tip density was greater in the organic layer of the soil
profile, but more root tips (i.e. up to 85%) occurred in the greater volume of the Bf layer.
Pine generally had a lower density of root tips compared to birch.

Morphotype descriptions for pine and birch ECMs are provided in Appendix A. ECM
morphotypes that were most frequently identified on root systems included Cenococcum,
MRA, E-strain, Amphinema, three Russulaceae (including Lactarius), two RhizopogonSuillus, and two Thelephoraceae types. Three unidentified types (one black and two white
types) were observed less commonly. The relative abundance of ECMs in FH and CWD soil
systems are shown for pine and birch in Figures 2.2 and 2.3, respectively.

Clusters of Cenococcum ECMs dominated the upper roots of pine and birch in the FH layer
(i.e. often representing 10-20% of the total root tip community), but were rarely observed in
the CWD layer. These tips were black and woolly, with characteristic stellate pattern of cells
of the outer mantle. Although relative abundance did not decrease in systems with higher
PHC concentrations, root tips appeared dry and less robust over time with PHC treatment.
The other ascomycetes, MRA and E-strain, were commonly found on root tips throughout the
soil profile, but with low relative abundance. All three ascomycetes were well-developed at
the 1-week sampling times; at later sampling times, evidence of secondary colonization was
observed, in which new ECM growth occurred on root tips previously colonized by MRA.

Amphinema ECMs and external mycelia were often associated with Cenococcum tips near
the organic-Ae layer interface, but were also occasionally observed lower in the soil profile.

112

Amphinema was characterized by copious yellow, curvy, clamped hyphae, but no
rhizomorphs were observed in this study. Piloderma hyphae, characterized by ornamentation
with needle-like crystals, were found associated with Cenococcum tips, but were never
observed as ECMs in this study, even though abundant Piloderma mycelia were observed in
the FH, CWD and FHoil organic layers in the field.

Russulaceae types dominated most root systems (i.e. ECMs observed on 80-90% of root tips)
of both pine and birch in FH and CWD soil systems. These ECMs were developed by the 1week harvest time, although sometimes with a thin mantle. Two Russulaceae types were
associated with pine. Root tips varied in color from yellow to orange-brown and were
generally unbranched and smooth or appeared velvety due to the few short undamped
hyphae that resembled cystidia early in their development. The cell arrangement of outer
mantles ranged from net to irregular synenchyma. The Russulaceae 2 ECM was identified as
a species of Lactarius based on the presence of laticifers, which contained latex material. On
birch, the Russulaceae 3 ECM, a cream-coloured smooth type, dominated all 96 root systems
assessed in the first year of the study. Although mainly found in the mineral soil layers,
Russulaceae ECMs were also common on the upper lateral roots, particularly in the CWD
organic layer.

Two Rhizopogon-Suillus types were found associated with root tips exclusively in the Bf
layer in FH and CWD soil systems associated with pine. The Rhizopogon-Suillus 1 ECM
was a dark brown, coralloid morphotype. The outer mantle had a net synenchyma cell
arrangement with amorphous globules at the surface; the copious hyphae (2-4 um)

113

characteristically bulged at the undamped septa and exhibited no other ornamentation. The
Rhizopogon-Suillus 2 ECM was white with a slightly tuberculate form, copious hyphae and
an outer mantle with prosenchymous (to net synenchymous) cell arrangement. By the 16week sampling time, both ECMs had developed extensive rhizomorphs and hyphal fans
(stained purple and orange, respectively, with KOH) that extended away from root tips into
the mineral soil. Although ECM/ mycelial biomass was not measured, it appeared that
rhizomorphic development may have been enhanced in PHC contaminated soils.

Two Thelephoraceae -like types were found associated with a few seedlings of pine and birch
in both FH and CWD soil systems. These ECMs were never observed prior to the 7-week
harvest and were present at all concentrations of PHCs. ECMs were smooth, ranged in
colour from orange to dark brown or black and had thin mantles of net or regular
synenchyma (respectively). Both had few long emanating hyphae, sometimes with clamps.

Three ECMs, one black and two white, were not identified in this study, mainly because they
were encountered so rarely. The black type resembled MRA, except that the outer mantle
was net to irregular synenchyma rather than the felt prosenchyma typically associated with
MRA. One white type was similar to Piloderma and was found exclusively with birch.

114

FH

1D0%

D Russulaceae 2
El Russulaceae 1

ao%

CWD

100%

Q Rrizcpogon-Suillus2

• E-strain

100%

60%

• Cenococcum

40%

• Rhizopogon-Suillus 1

100%

• black

80%

• Rhizopogon-Suillus 1

BlvFA

0%

D Russulaceae 2

80%

• Russulaceae 2

Hh/RA

I

o%

B Russulaceae 1
D Arrphinerra

60%

• Arrphinerra

40%

• E-strain

40%

• E-strain

ll

BIwRA

III

• Cenococcum

D Russulaceae 2
Q Russulaceae 1

80%

100%

•

• E- strain
alvRA

10%

7 wk
OThelephora-like 1
• RHzopogon-Sjillus 1

80%

D Russulaceae 2

• Arrphinerra
60%

• tVRA

t!

20%
0%

100%

4 wk

• Russulaceae 2

60%

0%

1 wk

• Rrizopogon-Suillus 1

80%

60%

HtvRA

40%

• Cenococcum
20%

3 0 % -H

B• iM

0%
100%

.1

80%

• Rrizopogon-Suillus2
• Rhizopogon-Suillus 1

D Russulaceae 1

I

40%
30%
0%

80%

r,

n be .,.
10 wk
0 Rhizopogon-Sjillus 3
• Rhizopogon-Suillus 1

80%-f

• Russulaceae 3

• Russulaceae 2

60%

100%

0%
1G0%

Pffl

:%
40% \

• Arrphinerra

60%

• B strain
Hr/RA

40%

• Cenococcum

B E-strain
Q tvRA
• Cenococcum

m Thelephors-like 1
• Rhizopogon-Suillus 2

20%
0%

13 wk

100%

• • l l
8 0 % -f

• Rhizopogon-Suillus 1

Q Rfizopogon-Suillus 2
• RhzopogorvSuillus 1
a Russulaceae3

• Russulaceae 2

60%

a Russulaceae 1

40%-}-

• Hloderrra
30%
0%

I

100%

• E strain
DtvRA

60%

0%

• Cenococcum

m

80%

20%

0 Thalephora-like 2

100%

• Rhizopogon-SUillus 2

30%

16 wk
• black
Q Rhizopogon-Suilus 3
• Rhizopcgon-Suilus 1

• Rrizopogon-SUillusI
• Russulaceae 2
D Russulaceae 1

40%

60%

D Russ daceae 3
B RussUaceae1

40%

• Bstrah

D Arrphinerra
30%

• E- strain
HWRA

0%

0

1 2

[PHCJ

3

• Cenococcum

30%

alvRA

0%
0

1 3

3

[PHC]

Figure 2.2: Relative abundance of ECMs on pine roots in FH and CWD soil systems
sampled at 3-week intervals over 16 weeks with 0, 1, 2, or 3 mL of crude oil added to the soil
surface.
115

FH

100%

• Russulaceae 3

CWD

• Cenococcum

80%

100%

D Rjssulaceae 3

1 wk

• Eistrain

80% U

DMA
60%

60%

40% +

4 0 % 4-

ao%

20% J-

o%

0%

100%

4wk

100%

• Rjssulaceae 3
• B strain

8 0 % 4-

w
D Russulaceae 3
• E- strain

eo%

a IwFA

60% I

60%

• Cenococcum

40%

4 0 % -H

23%

20%

0%

0%

100%

100%

• Russulaceae3
• E strain

60%

I

80%

60%

fc

60%

40%

I 40.

7 wk
B "Rielephora-like 1
D Rjssulaceae 3
• ^strain

• Cenococcum

i:

23%
0%

• -•-•-•.

100%

• Russulaceae 3
• Cenococcum

80%

g

D M

•7 r? 7

1

103%

10 wk
D Russulaceae 3
BlwRA

80% +

• Cenococcum
60%

60%

40%

40%

20%

23%

0%

0%

103%

100%

D white
D Russulaceae 3

80%

13 wk
D Russulaceae 3

80%

0IVRA
• Cenococcum

60%

• Cenococcirn

60% +
40%

40%

23%

23%
0% . M U U

•

100%

0%

16 wk
D white
B TVielephora-like 2

80%

100%

O"melephora-lite2
• RhizcpogorvSullus 1

80%

D Russdaceae 3

D Rjssulaceae 3

60% +

D Rloderrra

83%

QWRA

• Cenococcum

40% • •

40%

23% |

20%
0%

0%
0

1 2

[PHC]

3

7
0

-r. 7

TT,

1 2

3

[PHC]

Figure 2.3: Relative abundance of ECMs on birch roots in FH and CWD soil systems
sampled at 3-week intervals over 16 weeks with 0, 1, 2, or 3 mL of crude oil added to the soil
surface.
116

In general, Simpson diversity (1/D) values were lower in systems treated with the highest
PHC concentration (219 mg cm"2) compared to untreated controls, with intermediate
diversity values for the intermediate (i.e. 73 mg cm"2 and 146 mg cm"2) PHC concentrations
(Table 2.3). Differences between PHC treatments within harvest groups (n=2) were
significant for P-FH systems at 10 weeks (p=0.027) and for B-FH systems at 13 and 16
weeks (p = 0.015)

Table 2.3: One-way ANOVA plus Fisher's LSD (a = 0.05) of Simpson diversity (1/D)
between PHC treatments (within harvest groups): PHC-0 = control; PHC-1 = 73 mg cm"2;
PHC-2 = 146 mg cm"2; PHC-3 = 219 mg cm"2.
Harvest

PHC-0
mean (SE)

PHC-1
mean (SE)

PHC-2
mean (SE)

PHC-3
mean (SE)

1.23 (0.39)
1.94(0.39)
1.19(0.39)
1.36 (0.39) a
3.07 (0.39)
2.80 (0.39)

1.65 (0.37)
2.33 (0.37)
1.11(0.37)
2.11 (0.37) aft
1.77(0.37)
1.84(0.37)

1.29(0.27)
2.40 (0.27)
1.35 (0.27)
1.52(0.27)oi
1.99(0.27)
2.16 (0.27)

1.58(0.36)
1.13(0.36)
1.39(0.36)
2.44 (0.36) b
1.49(0.36)
1.54(0.36)

1.23 (0.33)
1.52 (0.33)
1.11(0.33)
2.01 (0.33)
1.12(0.33)
1.80(0.33)

1.11(0.21)
2.46(0.21)
1.23(0.21)
1.3(0.21)
1.20(0.21)
1.19(0.21)

1.24(0.27)
2.07 (0.27)
1.10(0.27)
1.24(0.27)
1.33(0.27)
1.41 (0.27)

1.11(0.22)
2.35 (0.22)
1.17(0.22)
1.17(0.22)
1.36(0.22)
1.49(0.22)

1.24(0.10)
1.23 (0.10)
1.22(0.10)
1.16(0.10)
1.21 (0.10) a
1.36 (0.10) a

1.35(0.11)
1.11(0.11)
1.13(0.11)
1.12(0.11)
1.17 (0.11) aft
1.12(0.11)6

1.10(0.02)
1.22(0.02)
1.22(0.02)
1.12(0.02)
1.18 (0.02) ab
1.08 (0.02) c

1.10(0.00)
1.10(0.00)
1.10(0.00)
1.10(0.00)
1.12(0.00)6
1.15 (0.00) d

1.12(0.07)
1.17(0.07)
1.12(0.07)
1.10(0.07)
1.16(0.07)
1.17(0.07)

1.11 (0.08)
1.16(0.08)
1.11(0.08)
1.1 (0.08)
1.07(0.08)
1.09(0.08)

1.53(0.11)
1.22(0.11)
1.53(0.11)
1.04(0.11)
1.13(0.11)
1.31 (0.11)

1.34(0.02)
1.22(0.02)
1.36(0.02)
1.10(0.02)
1.08(0.02)
1.07(0.02)

P-FH
1 wk
4wk
7 wk
10 wk
13 wk
16 wk
P-CWD
1 wk
4 wk
7 wk
10 wk
13 wk
16 wk
B-FH
1 wk
4 wk
7 wk
10 wk
13 wk
16 wk
B-CWD
1 wk
4 wk
7 wk
10 wk
13 wk
16 wk

117

Analysis of Simpson diversity values within PHC groups generally showed an increase over
the 16-week duration of the experiment (Table 2.4). These differences were significant
within the two higher PHC concentration groups (i.e. 146 mg cm"2 and 219 mg cm"2) for PFH (p=0.004), B-FH (p=0.002), and B-CWD (p=0.013) systems, as well as within the
untreated control group for P-FH (p=0.004).

Table 2.4: One-way ANOVA plus Fisher's LSD (a = 0.05) of Simpson diversity (1/D)
between harvests (within PHC treatment groups) at 1, 4, 7, 10, 13, and 16 weeks following
PHC treatment.
Harvest

PHC-0
mean (SE)

PHC-1
mean (SE)

PHC-2
mean (SE)

PHC-3
mean (SE)

1.23 (0.39) a
1.94 (0.39) aft
1.19(0.39)o
1.36 (0.39) a
3.07 (0.39) ft
2.80 (0.39) b

1.65 (0.37)
2.33 (0.37)
1.11(0.37)
2.11(0.37)
1.77(0.37)
1.84(0.37)

1.29 (0.27) a
2.40 (0.27) ft
1.35 (0.27) a
1.52 (0.27) aft
1.99 (0.27) aft
2.16 (0.27) aft

1.58 (0.36) aft
1.13 (0.36) a
1.39 (0.36) aft
2.44 (0.36) ft
1.49 (0.36) oft
1.54 (0.36) oft

1.23 (0.33)
1.52(0.33)
1.11(0.33)
2.01 (0.33)
1.12(0.33)
1.80(0.33)

1.11(0.21)
2.46(0.21)
1.23(0.21)
1.3(0.21)
1.20(0.21)
1.19(0.21)

1.24(0.27)
2.07 (0.27)
1.10(0.27)
1.24(0.27)
1.33 (0.27)
1.41 (0.27)

1.11(0.22)
2.35 (0.22)
1.17(0.22)
1.17(0.22)
1.36(0.22)
1.49(0.22)

1.24(0.10)
1.23 (0.10)
1.22(0.10)
1.1 6(0.10)
1.21 (0.10)
1.36(0.10)

1.35(0.11)
1.11(0.11)
1.13(0.11)
1.12(0.11)
1.17(0.11)
1.12(0.11)

1.10(0.02)00"
1.22 (0.02) ft
1.22 (0.02) oft
1.12 (0.02) c
1.18 (0.02) d
1.08 (0.02) ae

1.10(0.00)o
1.10 (0.00) a
1.10 (0.00) a
1.10 (0.00) a
1.12(0.00)o
1.15 (0.00) ft

1.12(0.07)
1.17(0.07)
1.12(0.07)
1.10(0.07)
1.16(0.07)
1.17(0.07)

1.11(0.08)
1.16(0.08)
1.11(0.08)
1.1 (0.08)
1.07(0.08)
1.09(0.08)

1.53(0.11)0
1.22 (0.11) oft
1.53(0.11)o
1.04 (0.11) ft
1.13 (0.11) ft
1.31 (0.11) aft

1.34 (0.02) a
1.22 (0.02) ft
1.36 (0.02) a
1.10 (0.02) c
1.08 (0.02) c
1.07 (0.02) c

P-FH
1 wk
4 wk
7 wk
10 wk
13 wk
16 wk
P-CWD
1 wk
4wk
7 wk
10 wk
13 wk
16 wk
B-FH
1 wk
4 wk
7 wk
10 wk
13 wk
16 wk
B-CWD
1 wk
4 wk
7 wk
10 wk
13 wk
16 wk

118

ECM and ERM community structure
Multivariate analysis using nonmetric multidimensional scaling (NMS) gave a threedimensional solution with a final stress of 19.66 and instability of 0.12. The NMS ordination
in Figures 2.4 and 2.6 show relative fungal community structure (genotypes) for all 109
plant-soil systems for which successful fragment analysis occurred. These included 24 pine,
12 birch, and 25 lingonberry single-plant systems, as well as 24 pine and lingonberry doubleplant systems in 48 FH, 43 CWD and 18 FHoil (used in single-plant systems only) soils.
Nearly half of all systems were treated with PHCs (53 treated and 56 untreated systems); 54
and 55 systems, respectively, were harvested from 1 and 16 weeks. Except for birch -CWD
and FHoil groups, all other (plant-soil-PHC-harvest) treatment groups were represented by
n=2-3. The total fungal community was represented by 91 DNA fragment lengths
(genotypes): 74 were associated with pine and 60 with lingonberry (3 genotypes were
exclusive to birch). Thirty genotypes were associated with double-plant systems only, 30
were associated with only single-plant systems, and 31 genotypes were common to both
single- and double-plant systems.

Harvest time andPHC effects
NMS ordinations of the harvest time and PHC treatment variables are shown along the first
and second axes of Figure 2.4. Overall, fungal community structure varied significantly
(p=0.005) between the 1 and 16-week harvest times. PHC treatment was not found to lead to
any significant differences in fungal community structure when considered as a single
variable. However, comparisons within PHC treated and control groups revealed that the
PHC treated group at the 16-week harvest significantly varied from both PHC treated

119

(p=0.005) and control (p=0.031) groups at the 1-week harvest, while fungal communities in
the control group did not vary after 16 weeks (Figure 2.5). In Figure 2.4, a great deal of
overlap is observed in the distribution of fungal communities at the two harvest times
(represented by diamond and inverted triangle symbols, respectively), but the 16-week group
appears to be more concentrated towards the upper left quadrant of the plot. The separation
of PHC treated communities (closed symbols) should appear more pronounced than
untreated control communities (open symbols).

ECM fungal communities of pine exhibited the same trends observed for the overall fungal
community; namely, that PHC impacts on fungal community structure were observed only
over time (p=0.031). ERM communities of lingonberry communities showed no differences
between PHC treatment and harvest time groups.

120

^

H1-PHC

<^> H1 - no PHC
,
•

o

H16-PHC
• H16-noPHC

•5-

•

<
•

•

o

16.8%)

*.-,
•

0*

•

,•
0

II
•

A,

0

Axis 2

J

•

•

•

•

•

•

•

o
•

o

•

•

Axis 1 (i2 = 18.8%)

Figure 2.4: NMS ordination of mycorrhizal fungal showing multivariate effects of harvest
time (1 and 16 weeks) and PHC treatment (stress = 19.66; instability = 0.12).

PHC ¥5 no PHC (control)

PHC-1wk

•* PHC-18wk

p>0.05

t p=§.§05
i p=0,Q3i

controM wk - "H—-*• control-16wk

Figure 2.5: Pairwise comparisons of fungal community structure (genotypes) in PHC-treated
and control systems at 1 and 16 weeks. Significant differences (MRPP) are represented by
dark arrows and corresponding p-values; light (slashed) arrows represent no differences
between groups.
121

Plant effects
Fungal community structure for different plant systems is represented along the first and
third axes of the same NMS ordination (Figure 2.6). The clearest distinction is between
single- and double-plant systems (represented by closed and open symbols, respectively).
The single lingonberry (closed triangles) systems appear grouped towards the left side of the
plot whereas single pine systems (closed squares) are more broadly distributed. Birch
communities (closed circles) are distributed within the single pine and lingonberry clouds.
Pairwise analysis using MRPP revealed that the pine and lingonberry communities from the
double-plant systems differed significantly from communities from each of the single-plant
systems, as well as from each other (p<0.001 in all cases). Single pine system communities
varied significantly from both birch (p=0.032) and lingonberry (p=0.028) communities,
which did not vary significantly from each other (note incomplete birch dataset).

122

I

Pine (single)

4 1 Birch (single)
£i

Lingonberry (single)

•

Pine (double)
\

Lingonberry (double)

Axis 1 (i^ = 18.8%)

Figure 2.6: NMS ordination of mycorrhizal fungal communities of single (closed) and
double (open) plant systems (stress = 19.66; instability = 0.12).

123

Separate analyses of pine and lingonberry communities reiterated differences between the
single- and double-plant systems. In both cases, genotype richness was greater in the doubleplant systems. For pine, single-plant systems had an average of 5.54 genotypes per root
system (n=24) compared to an average of 6.88 genotypes per root system (n=24) in doubleplant systems. For lingonberry, single-plant systems averaged 6.20 genotypes (n=25) and
double-plant systems averaged 7.63 genotypes (n=24). No differences in genotype richness
were associated with other treatment variables. For both plants, differences between fungal
communities in single- compared to double-plant systems had p-values of <0.001.

Organic layer effects
To eliminate the double-plant effect on the analysis of soil layers, 24 FH, 19 CWD, and 18
FHoil fungal communities were compared between single-plant systems only. Of a total of
61 genotypes, 31 were found only in FH soils, while CWD and FHoil soils each had six
unique genotypes. NMS analysis again gave a three-dimensional solution with a final stress
value of 17.64 and instability of 0.024. The ordination shown in Figure 2.7 shows the broad
distribution of genotypes in FH soils (dark squares), which reflects the heterogeneity of
fungal communities found in these systems. Along the second and third axes, the FHoil soils
(inverted triangles) are less broadly distributed than the FH soils; the CWD soils (open
diamonds) form the tightest cluster in the center of the ordination. Fungal community
structure varied significantly between all three types of soil systems: FH and CWD
(p=0.016), FH and FHoil (p=0.048), and CWD and FHoil (p=0.027).

124

Axis 2 (r8 = 21.4%)

Figure 2.7: NMS ordination of organic soil layer (FH, CWD, and FHoil) effects in single
plant systems (stress = 17.64; instability = 0.024)

125

MRPP comparisons between PHC treated and untreated (control) systems within soil groups
showed that most differences were related to soils, but that some may have been related to
PHC treatment. No PHC-treated soil systems (i.e. FH, CWD, or FHoil systems) varied
significantly from each other; all differences were related to the control soils. Fungal
communities in FH control soils varied significantly from both CWD (p=0.024) and FHoil
(p=0.041) control soils, but CWD and FHoil communities did not vary significantly from
each other. Fungal communities in FHoil control soils varied significantly (p=0.016) from
PHC treated FH soils, but not from either PHC treated CWD or FHoil soil communities.

Differences between fungal communities in the different soil systems were only found within
control groups (i.e. FH, CWD, and FHoil all varied significantly from one another), as well
as between the control FHoil and treated FH and CWD (but not between treated and control
FHoil soils). Lingonberry communities showed the same soil effects.

Laccase activity
Most ECM fungal root tips showed some level of laccase activity (i.e. development of a
green color over 24 h), as shown in Figure 2.8. The greatest level of activity was associated
with Rhizopogon-Suillus 1 and the two Russulaceae species that dominated pine roots (the
Russulaceae ECM associated with birch was not tested). In the numerous trials conducted,
Russulaceae (root tips) and Rhizopogon-Suillus 1 (root tips and/ or extraradical mycelia)
harvested from pine seedlings from different soils and PHC treatments consistently produced
a light green colour within the first hour of the assay, and a deep green colour by the 16-h
mark. The Rhizopogon-Suillus 2 ECM did not often give positive results for laccase activity,

126

but sometimes produced a light green colour by 16 or 24 hours. Amphinema ECMs usually
oxidized a low level of ABTS within the first hour of the assay; this light green colour did
not generally deepen over the subsequent 23 hours, possibly indicating a low level of laccase
secretion by this ECM. With respect to the ascomycetes, MRA usually showed low laccase
activity within the first hour and substantial oxidation of ABTS by 16 and 24 hours. Laccase
activity was seldom observed with Cenococcum ECMs; no enzyme activity was ever
observed for E-strain. Neither soil type (i.e. organic layer) nor PHC treatment had any
visible effect on laccase activity associated with these ECM root tips.

For ERM hair roots, little laccase activity was ever observed; however, in one trial (n=5),
hair roots from PHC-treated soils showed a strong positive reaction whereas roots from
untreated soils showed no activity.

Colour Devdopmoit
lh
lfih
24h
ECM
Russulaceae 1
Russulaceae 2 (Lactarius)
Fhi&pogpn-iMlhs 1
Rhizopogpn-SuilliG 2
AmpMnsma
MRA
Genococcum
E-slrain
ERM
ERM-PHC
ERM-no PHC

+
-(+)
++
-(+)
-

++
++
++
-(+)
+
+ (++)
-(+)

+ (-)

+ (-)

+ C-)

++
++
++
-(+)
+
+ (++)

-(+)
+ (-)

Figure 2.8: Table showing laccase activity of ECM morphotypes and ERM hair roots
assessed by intensity of colour development (-, none; +, pale green; ++, dark green) at 1, 16
and 24 h incubation times with ABTS on the left. Symbols in brackets indicate less typical
reactions over numerous trials (n=5+); Photograph showing range of colour development for
ECMs (n=3, vertically) incubated in microplate wells with ABTS for 24+ h on the right.

127

Discussion
We found that PHCs, when applied to plant-soil systems at rates comparable to a large
terrestrial oil spill event (i.e. equivalent to several tonnes of crude oil per hectare), had little
impact on established or developing mycorrhizal fungal communities over 16 weeks of
study.

PHC impacts on mycorrhizal communities
Detailed morphological analysis showed that the ECM morphotypes described here represent
a sub-set of ECMs commonly reported from northern forest soils (Rosling et al., 2003).
These included Cenococcum, MRA, E-strain, Amphinema, three Russulaceae (including
Lactarius), two Rhizopogon-Suillus, and two Thelephoraceae types, as well as three
unidentified ECMs. Notably absent from our bioassay were Piloderma ECMs, even though
their distinctive yellow mycelia were prevalent in FH, CWD, and particularly FHoil organic
layers of the sub-boreal forest field site. Piloderma has been associated with greater N
content that may be more typical of mature forest floors, which may explain why it was not
associated with the seedling roots in the current study (Lilleskov et al., 2002). It is thought
that Piloderma may specialize in efficient N uptake or increase nutrient availability through
enzymatic degradation of organic substrates, potentially including PHCs. Thus, preservation
of organic layers (i.e. Piloderma habitat) following PHC contamination may be an important
management strategy for sustaining this ecological function. Cortinarius, which also tends to
be associated with a high C:N ratio in mature forest soils (Lindahl et al., 2007) and may
possess biodegradation potential, was also not observed in this study.

128

We found that most ECMs appeared unaffected by PHC treatment at any of the levels tested
(i.e. ranging from approximately 7 to 22 tonnes ha"1). In one of the few comparable studies
of PHC-contaminated agricultural and forest soils (i.e. initiated following an oil well blowout
in northern Italy that contaminated an area of 1,500 ha with 18,000 m3 of crude oil), ECM
fungal responses ranged from no negative effects to reduced biomass and colonization
potential (Nicolotti and Egli, 1998). In subsequent experiments, in which different levels of
PHCs were added to culture media, Nicolotti and Egli (1998) found that some fungal species
were inhibited by oil whereas others (e.g. Laccaria sp.) appeared to grow better with PHCs,
even at very high concentrations. Although the current study encompasses more of the
complexity of the forest soil environment (particularly with respect to source of C via plant
photosynthesis) and is not directly comparable to culture-based studies, a similar spectrum of
ECM responses to PHC contaminants was observed. For example, Cenococcum ECMs may
have been inhibited by PHCs. The dry, flaccid appearance of Cenococcum on some root
systems may have partially resulted from toxicity of PHC chemicals, but was also likely due
to physical changes in the soil habitat (i.e. change in water holding capacity of the FH layer)
that occurred concurrently with PHC contamination. Soil drying may have been exacerbated
in our model plant - soil systems that lacked a protective moss layer and forest canopy, and
were thus potentially exposed to an even greater moisture loss due to evaporation compared
to field conditions. The consequences of Cenococcum reduction as a functional component
of the community are currently not known. On the other hand, rhizomorphic development of
Rhizopogon-Suillus types may have been enhanced by PHCs, as hyphal fans proliferated into
the contaminated mineral soil (biomass was not measured). The resulting expansion of
mycorrhizosphere space would be expected to support diverse microbial communities

129

involved in the biodegradation of PHC substrates. No differences in distribution or
abundance were observed for the Russulaceae types, which accounted for the majority of
ECMs on all pine and birch root systems, regardless of PHC treatment.

The diversity (i.e. Simpson diversity) of ECM communities on pine and birch root systems
generally decreased with greater PHC concentrations, but this trend was neither consistent
nor usually significant. At any one of the six sampling times, there were no substantial
changes of dominant ECMs in PHC treated systems compared to controls. ECM and ERM
fungal community structure (based on genotypes) also did not vary between PHC treated and
untreated (control) systems. As the plant - soil systems were treated with fairly high and
ecologically relevant concentrations of crude oil, these results indicate that the intact organic
soil layer, ECM mantle, and ERM hair root provided some level of protection against the
potential toxicity of PHC constituents (Blakely et ah, 2002). As also reported elsewhere
(Setala et ah, 2000), our results suggest that maintenance of the ecological integrity of plant soil systems provides resilience to environmental stress such as PHC contamination.

The temporal aspect of assessing mycorrhizal response to PHC treatment in dynamic systems
was emphasized in this analysis. The morphological studies revealed a general increase in
ECM richness and abundance over time, which was generally not altered by PHC treatment.
As seedling root systems and mycorrhizal associations were still growing and developing at
the time of PHC treatment, changes in diversity may have been due to increased ability to
identify ECMs, or as a result of fungal succession on root tips, as has been reported from
other studies (Massicotte et ah, 1999). The molecular studies also indicated shifts in ECM

130

fungal community structure over time and these time-related differences were only
significant in the PHC treated systems. Thus, important questions with respect to ecosystem
sustainability are whether PHC contamination alters the trajectory of mycorrhizal community
structure in the long-term, and whether ecosystem health may be negatively impacted in the
future as a result. Long term studies are required to address these questions.

ECM and ERM fungal community structure was largely determined by properties associated
with host plants than by PHC contamination. Differences in community structure between
single-plant systems reflect the many potential plant-fungal combinations that occur in
repeatable units across landscapes (Allen et al, 2003). More information on the individual
physiologies of these symbioses is required to understand their roles in ecological processes
and to predict how communities may adapt to future disturbances. Although fungal
community structure varied between all host plant systems, differences were most striking
between the single- and double-plant systems (i.e. more complex systems). These
differences appeared to be due to greater genotype richness per root system, as well as altered
genotype abundance patterns in the double-plant systems. Even though pine and lingonberry
mycorrhizas shared the same rhizosphere in the double-plant systems, the ECM and ERM
fungal communities remained distinct. The mycorrhizal community patterns observed in
double-plant systems may represent the non-linear, emergent properties associated with
complex systems whereby the influences of mycorrhizal fungi on plant populations and
communities are not merely the sum of effects on the individuals within populations
(Dahlberg, 2001; Allen et al, 2003).

131

In addition to assessing the integrity of established mycorrhizal fungal systems, we also
tested the ability of seedlings to germinate and grow in contaminated soils (i.e. FHoil
systems), which is an important consideration in the context of environmental restoration. In
these chemically complex systems, sensitivity to the initial conditions may create limits to
mycorrhizal diversity (Allen et ah, 2003), although this was not evident in either of our
morphological or molecular studies. Nicolotti and Egli (1998) found that crude oil (50 ppm
hand-mixed into homogenized forest soil) did not inhibit Norway spruce or poplar seed
germination or kill seedlings. In our study, using a much greater PHC concentration, longer
weathering period, and reconstructed soil layers, we also found that germination of pine
seeds did not appear to be inhibited by the presence of weathered crude oil. Furthermore,
seedling growth/ mycorrhizal colonization was sometimes more vigorous in the previously
contaminated soils compared to pristine (FH and CWD) layers. Birch seeds appeared to have
had lower rates of germination, possibly due to interference of PHC chemicals with initial
interactions between germinating roots, fungal propagules, and mycorrhizal helper bacteria
(Garbaye, 1994). However, the seedlings that grew in FHoil layer showed no apparent
differences to seedlings germinated in the other organic soil layers (i.e. FH and CWD layers).
The survival of lingonberry seedlings also did not differ between FH, CWD, and FHoil soil
systems.

Potential for direct mycorrhizal role in biodegradation
Nicolotti and Egli (1998) reported that some ECM fungi surviving in contaminated forest soil
may metabolize chemicals in crude oil. The distribution of laccase-secreting fungi represents
a part of the oxidative potential in soils (Luis et ah, 2005). We found that the metabolic

132

potential for laccase-mediated PHC biodegradation appears to exist among the dominant
members of ECM fungal communities including two Russulaceae (one Lactarius),
Rhizopogon-Suillus 1, and Amphinema types. High extracellular enzyme activities have been
reported for Lactarius and Russula species grown in culture (Gramss et ah, 1998) and
laccase-like genes have been amplified from Lactarius, Russula, Piloderma and Tylospora
fungi and soils (Chen et al. 2003; Luis et al, 2005). Increased extracellular enzyme activity
and expression of genes coding these enzymes have been found at the hyphal fronts of ECM
systems advancing in the humus of microcosms (Timonen and Sen, 1998; Donnelly and
Entry, 1999). The function of fungal laccase in the carbon cycle in soils, especially in the
formation, stabilization and degradation of the organic matter, is also of ecological interest
(Luis et al. 2004).

From our study, we know that the Russulaceae types dominated pine root systems in all soil
systems, both PHC treated and untreated, and could potentially influence large volumes of
forest soils (if distribution patterns are consistent with our bioassay). The RhizopogonSuillus 1 ECM, also influenced large volumes of mineral soil via rhizomorphic proliferation,
which may have even have been enhanced by high levels of PHC contamination.
Rhizopogon-Suillus ECMs are not well studied, partly because they are often found deeper in
the soil profile than most researchers sample (Rosling et al, 2003). Piloderma ECMs were
not described from our study, but it is likely that laccase-secreting Piloderma ECMs are
abundant in mature forest soils. These results indicate that soils with intact ECM systems
may possess high biodegradative potential for PHC contaminants via the combined activities
(syntrophic metabolism) of microorganisms in the mycorrhizosphere.

133

We did not find evidence of enhanced laccase activity (i.e. intensity of laccase reactions)
resulting from exposure of pine ECMs to PHC substrates, as has been reported from studies
in which a variety of different aromatic compounds were tested (Burke and Cairney, 2002).
It could be that the ECMs in this study were directly exposed to lower PAH concentrations in
the crude oil-contaminated soil compared to other studies. Alternatively, ECM fungi
inhabiting soils influenced by organic layers rich in lignin and humus may exist in a state of
enhanced laccase expression and activity. The warm, wet greenhouse conditions may have
further enhanced laccase activity through provision of favourable decomposition/
biodegradation conditions.

We generally did not observe much evidence of laccase activity when hair roots of
lingonberry were tested, even though ERM fungi (e.g. R. ericae) have been reported to have
well-developed saprotrophic abilities (Burke and Cairney, 2002; Perotto et al. 2002). In one
set of trials, laccase activity was consistently observed with hair roots that were recently
exposed to PHCs, but not with control roots. This result was not repeated, so no conclusions
can be drawn from this. As very little is known regarding biodegradation capacity in ERM
systems, further investigation is warranted.

Conclusions
In this study, we found that established ECM and ERM systems appear resilient to toxic
effects of crude oil, which was related to the integrity of the plant - soil systems. In addition,
ECM systems may directly enhance PHC biodegradation at the fungal - soil interface

134

through secretion of laccase, as well as indirectly aiding biodegradation through provision of
colonization surfaces and co-metabolic substrates for associated bacterial communities.
Thus, mycorrhizal systems may also be part of the remediation solution for PHC
contaminated forest sites within a sustainable ecosystem management context.

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Chapter 3: Enhanced biodegradation of petroleum hydrocarbons in the
mycorrhizosphere of sub-boreal forest soils

Abstract
The dynamics of petroleum hydrocarbon (PHC) biodegradation in boreal forest soils are not
well understood. We used a bioassay approach to determine whether differences in
mycorrhizosphere-associated bacterial communities corresponded to differences in PHC
biodegradation patterns. Surface-sterilized seeds {Pinus, Betula, Alnus sp.) or seedlings
(Vaccinium sp.) were planted into Conetainer™ pots containing reconstructed soils: an
organic layer (mor humus, coarse woody debris, or previously oil-contaminated humus)
overlying sandy mineral horizons (Ae and Bf) of field-collected forest soils obtained from
central BC, Canada. After 4 months, BC light crude oil (219 mg cm"2) was applied to the soil
surface around the seedling stem; systems were destructively sampled at 1 and 16 weeks
following treatment. Concentrations of PHCs in 4 fractions (based on equivalent normal
straight-chain boiling point ranges) were determined using acetone-hexane extraction
followed by GC-FID analysis. Genotypic profiles of root-associated bacterial communities
were generated using length heterogeneity-PCR analysis of 16S rDNA; metabolic profiles
were based on C substrate use after 7 weeks. Nearly all plant-organic soil layer combinations
showed significant loss of nC10-nC16 fraction PHCs from 1 to 16 weeks, indicating an
inherent capacity for biodegradation within these soils. Total PHC (nC10-nC50)
concentrations declined significantly in only planted (pine-woody debris and birch-humus)
systems, reinforcing the importance of the (mycor)rhizosphere for supporting degradative
microbial communities. Multivariate analyses showed that (mycor)rhizosphere type and
complexity influenced bacterial community structure, but that this was not related to PHC

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biodegradation. The level of PHC contamination used in this study appeared to have
minimal impact on soil bacterial community structure or broad metabolic functions.

Introduction
Boreal ecosystems are increasingly exposed to petroleum hydrocarbon (PHC) contamination
due to expanding natural resource extraction activities (Kanaly and Harayama, 2000). Longterm studies of oil-contaminated forest soils have reported reductions in PHC levels due to
biodegradation (or biotransformation) by indigenous microbial communities (Braddock et
ah, 2003; Prince et ah, 2003). From culture and microcosm experiments, many ubiquitous
soil fungi and bacteria have been demonstrated to biodegrade numerous PHC compounds
(Heinonsalo et ah, 2000; Sarand et ah, 2000; Genney et ah, 2004; Corgie et ah, 2003) and
some genetic and biochemical pathways have been elucidated (Meharg and Cairney, 2000;
Burke and Cairney, 2002; Watanabe, 2002; Diaz, 2004). It is generally accepted that the
capacity to biodegrade PHCs is intrinsic in most soils (Meharg and Cairney, 2000; Chaillan
et ah, 2004; Delille et ah, 2004) and requires metabolic synergy among different functional
guilds of organisms, including mycorrhizal fungi and the bacterial communities closely
associated with the mycorrhizosphere (Burke and Cairney, 2002; Diaz, 2004; Chaudhry et
ah, 2005). Very little is currently known of the dynamics of PHC biodegradation as it occurs
within the mycorrhizosphere of forest soil systems (Robertson et ah, 2007).

Soil microbial communities may shift in response to changes in environmental conditions
(Watanabe, 2002), but exhibit high resilience to environmental stresses when soil organic
layers (e.g. humus, woody debris, etc.) are not severely disrupted (Setala et ah, 2000). Major
impacts of PHCs on forest soil microbial communities appear to be associated with
disturbances to water, nutrient and oxygen regimes related to the hydrophobicity and fluidity
of oily products (Tarradellas and Bitton, 1997). Spilled PHCs initially spread laterally within

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the lignin-rich humus of the forest floor; eventually, lighter fractions move down the soil
profile, along the paths of roots and fissures (Trofimov and Rozanova, 2003; Suleimanov et
al, 2005), where changes with depth in soil chemical and mineralogical properties create
contrasting habitats for microorganisms (Dickie et al, 2002; Rosling et al, 2003). PHC
contamination is expected to lead to an initial loss of bacterial diversity, followed by rapid
proliferation of metabolically competent populations capable of inhabiting the new
environmental conditions imposed by the chemical contaminants (Gramss et al, 1998; Diaz,
2004). Part of this rapid adaptation within microbial communities results from lateral
transfer of mobile genetic elements carrying genes for PHC biodegradation (Sarand et al,
2000; Diaz, 2004) and is controlled by the composition and functional redundancy of the
community originally present (Setala et al, 2000; Delille et al, 2003). Changes in bacterial
community structure attributed to PHC contamination have been observed for several years
after the initial spill event occurred (Lindstrom et al, 1999).

Northern forests are dominated by plants forming ecto- (ECM) and ericoid (ERM)
mycorrhizal symbioses with fungi expected to have well-developed saprotrophic activities
(Read and Perez-Moreno, 2003). Mycorrhizal establishment alters the quality and quantity
of root exudates that supply energy to the large chemo-organotrophic biomass associated
with the mycorrhizosphere (Rygiewicz and Anderson, 1994; Nannipieri et al, 2003; Morgan
et al, 2005). Interactions between mycorrhizal fungal mycelia and associated bacterial
communities are important mechanisms for accessing mineral nutrients from organic
substrates (Burke and Cairney, 2002; Sen, 2003) and also seem crucial for cometabolic
biodegradation of PHCs in contaminated soils (Sarand et al, 1998; 2000). Mycorrhizosphere

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development and function play central roles in controlling bacterial communities and their
biodegradation activities in lignin-rich humus and PHC-contaminated soils (Heinonsalo et
al, 2000). Enhanced PHC biodegradation in mycorrhizosphere soils (i.e. mycorrhizosphere
effect) is attributed to the greater metabolic activities of higher densities of microorganisms
(Heinonsalo et al, 2000; Corgie et al, 2003; Siciliano and Germida., 1998).

Various combinations of host plants and mycorrhizal fungi may create specific
mycorrhizosphere characteristics that are important with respect to biodegradation capacity
(Rygiewicz and Anderson, 1994; Perez-Moreno and Read, 2000; Selmants et al, 2005). For
example, plants that also form symbioses with N-fixing organisms (e.g. Alnus sp. with
Frankia) have increased availability of N as well as C in the mycorrhizosphere (Selmants et
al, 2005; Roy et al, 2007). Interactions between overstory (predominantly ECM plants) and
understory (plants forming ERM or arbuscular mycorrhizas) vegetation may also alter
capacity for biodegradation (Read and Perez-Moreno, 2003). Thus, host-fungal effects on C
and nutrient availability should not be ignored in biodegradation capacity studies (Corgie et
al, 2003).

In the current study, we examined relationships between PHC biodegradation and indigenous
microbial communities in reconstructed sub-boreal forest soils. We compared
biodegradation patterns in different plant-soil systems that varied with respect to organic soil
layer (i.e. forest floor, coarse woody debris, or forest floor previously contaminated with
PHCs) and plant {Pinus, Betula, Alnus, or Vaccinium sp.) characteristics. In these systems,
organic soil layers provided the initial inoculum for the developing rhizospheres; the

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functional composition of these communities was expected to be influenced by the relative
abundance of lignin and/ or previous exposure to PHCs. Microbial community structure and
function were expected to be further influenced by individual (mycor)rhizosphere properties.
For example, although pine and birch both support ECM communities, they differ with
respect to density and depth of fine root tips. Alder forms symbioses with both ECM and Infixing communities, while lingonberry forms ERMs. Differences between single- and
double-plant (i.e. pine and lingonberry) systems were also assessed, as well as effects on
bacterial communities by the PHC treatment itself. Ultimately, the goal of this study was to
determine whether differences in bacterial community (genotypic and metabolic) profiles
corresponded to changes in biodegradation patterns in an ecologically relevant context.

Materials and methods
Field site
The Kenneth Creek field site is located in the wet, cool subzone of the sub-boreal spruce
(SBSwkl) biogeoclimatic zone of central British Columbia, Canada, about 100 km east of
Prince George (53°34'N, 122°47'W). In 1982, the forest was logged and burned, then
subsequently planted with lodgepole pine {Pinus contorta Dougl. Ex Loud. var. latifolia
Engelm.); currently, the site is a mature, even-aged pine stand with small hybrid white spruce
{Picea glauca x engelmannii Parry ex Engelm.) and lesser numbers of western hemlock
(Tsuga heterophylla (Raf.) Sarg.). Young subalpine fir {Abies lasiocarpa (Hook.) Nutt.) and
sitka alder (Alnus crispa var. sinuata (Reg.) Rydb.) are present at the edge of the forest,
along the access road; western redcedar {Thuja plicata Donn) and trembling aspen {Populus
tremuloides Michx.) are also present in an unlogged stand across the main road. The site has

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a thick understory of oval-leaved blueberry (Vaccinium ovalifolium Sm.); mosses and lichens
(e.g. Peltigera) cover the forest floor, with some Lycopodium species.

The soils on this site were described by Arocena and Sanborn (1999). Soils are classified as
Eluviated Dystric Brunisols (Soil Classification Working Group, 1998), and consist of sandy
parent material with low clay content and few coarse fragments. The forest floor is mor
humus, from 2-5 cm thick with copious fungal mycelia present. The C:N ratio of the forest
floor is approximately 50 and the pH (water) is ~ 4.2. Gray Ae horizons are generally 1-2 cm
thick, with thicker pockets in some areas. Red Bf horizons extend to almost 30 cm, beneath
which are Bm (27-60 cm), BC (60-100) and C (> 1 m) horizons. The C:N ratios of the Ae
and Bf horizons are about 20 and 13, respectively. The pH (water) of the Ae horizon is 4.2
while the pH of the Bf horizon is about 4.8, increasing with depth to about 6.0 at the
transition to the C horizon. Fine roots are found at depths greater than 1 m. Large coarse
woody debris (i.e. downed trees), the legacy of past forest management, is abundant all over
the site.

Organic layers and the top 20-30 cm of mineral soils (Ae and Bf horizons) were collected
from the forest site in September of 2005 and 2006. The organic layers included the mor
humus forest floor (FH) that had been undisturbed for approximately 20 years, coarse woody
debris (CWD) in an advanced state of decay (i.e. decay class 5), and previously PHCcontaminated mor humus (FHoil) that had weathered in situ for four months throughout the
summer. The FHoil soils were collected in the first year only. In May of 2005, 2.0 L of BC
light crude oil (Husky Refinery, Prince George, BC) was applied to each of three 1-m2 plots

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with the moss layer (but not seedlings and other plants) removed. The oil had been
previously bubbled with N2 gas to remove the light and volatile compounds; a watering can
was used for even application in the field. All soils were stored at 4°C prior to use in
bioassay experiments, which commenced within 10 days of soil collection.

Bioassay andPHC treatment
Forest soils (i.e. Ae [~1 cm] and Bf [-15 cm] mineral soil layers beneath organic [FH, CWD,
or FHoil] soil layers [~2 cm]) were reconstructed in Cone-tainer™ pots (3.8 x 21 cm, Stuewe
and Sons, Corvallis, Oregon) with two clay pellets in the bottom to prevent soil loss (Setala
et al, 2000). Mineral soils (Ae and Bf layers) were homogenized and sieved through (1 cm2)
screens prior to potting. Surface-sterilized seeds of lodgepole pine {Pinus contorta var.
latifolia), white birch (Betula papyrifera Marsh.), and sitka alder {Alnus crispa var. sinuata),
collected from the SBS and obtained from the Ministry Tree Seed Center, (Surrey, BC (Seed
lots DWD20050009A (location 079-B-008), DWD20050009B (location 094-E-015), and
DWD20050047A (location 002-E-001), respectively), were planted into each pot.
Lingonberry {Vaccinium vitis-idaea L.) seedlings were obtained from Birch Creek Nursery
(Prince George, BC) and planted into 10x10x10 cm pots (i.e. Ae [~1 cm] and Bf [~7 cm]
mineral soil layers beneath organic [~2 cm] soil layers). All pots were placed in the
greenhouse (22°C day temperature, 15°C night temperature, and 16 h photoperiod) and
fertilized once a month (5 mL of NPK fertilizer; providing 100 ppm each of NPK) for the
first four months following seeding/ planting. The plants were watered two or three times
per week for the duration of the experiment. Single-plant (pine, birch, alder, and

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lingonberry) systems were established in 2005-06; double-plant (pine and lingonberry)
systems were established in 2006-07.

Seedlings and mycorrhizas were well established after four months of growth. At this time, 3
mL BC light crude oil (with volatiles removed as for the field application) was pipetted onto
the organic soil surface of each pot, around, but not touching, the seedling stem. PHC
concentration (219 mg cm' 2 ) corresponded to an application rate of 21,900 kg ha"1 (i.e. ~22
tonnes ha"1). The smell of crude oil dissipated in the greenhouse pod within the first week
following PHC treatment and no PHC loss was observed from the bottom of the pot (i.e. no
sheen on the wet surface below) after watering for the duration of the experiment. These
observations provided some confidence that observed PHC losses between 1 and 16 weeks
were primarily due to biodegradation.

Experimental design and sampling
Experiments followed a completely randomized block design with a planned full factorial
structure for plant, organic soil layer and PHC treatments (Table 3.1).

Table 3.1: Summary of plant, organic soil layer, and PHC treatment variables for 2005-2006
(single-plant) and 2006-2007 (double-plant) studies (n=3)*.
Plant
Pine [P]
Birch [B]
Alder [A]
Lingonberry [L]
Pine + Lingonberry [P+L]

Organic Soil Layer
Forest floor [FH]
Coarse woody debris [CWD]
Contaminated forest floor [FHoil]
No organic soil [NO] (control)

PHC
No PHC (control)
PHC (219 mg cm"2)

n<3 (due to poor germination) for birch -CWD, birch -FHoil, and all alder treatments;
these treatments were removed from PHC and DNA analyses.

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For soil, PHC, and DNA analyses, all three soil layers (organic, Ae, Bf) of PHC-treated and
untreated control soil systems (n=3) were destructively sampled at 1 and 16 weeks following
PHC treatment. Treatments included single-plant (pine [P], birch [B], lingonberry [L], and a
control [N] with no plant (i.e. no mycorrhizosphere)) and organic soil layer (mor humus
[FH], coarse woody debris [CWD], PHC-contaminated mor humus [FHoil], and a control
[NO] with no organic layer) combinations; double-plant (P+L) systems and unplanted
controls (N) were grown in FH and CWD organic soil layers. Individual pots were emptied
into trays; plants were gently removed (for future mycorrhizal community analysis) from the
soil with as little disturbance as possible to the reconstructed soil horizons. Approximately 1
g (wet weight) of each soil layer was collected in duplicate in 20 mL pre-weighed glass vials
(Fisher Scientific, Ottawa, Ontario). One set of samples was dried overnight in a 105°C oven
to obtain soil dry weights (i.e. soil moisture contents); the other set was stored at 4°C until
PHC extraction. The remaining soils were combined within treatment groups and then
collected and air-dried for soil nutrient analysis and pH. Root systems (intact, but with the
shoot excised) were shaken free from the soil and then washed in 35 mL sterile H2O. Roots
were then divided into 2 mL tubes and stored at -20°C until DNA extraction.

For bacterial community level physiological profile (CLPP) analysis, 60 plant - organic soil
systems (n=2) were harvested seven weeks after PHC treatment in 2006. Treatments
included PHC-treated and control pots for all single plant (pine [P], birch [B], alder [A],
lingonberry [L], and no plant [N]) and organic soil layer (FH, CWD, FHoil) combinations.
After initially shaking the roots to remove excess soil, the intact root systems were placed in
50 mL tubes containing 35 mL of sterile dH 2 0. Deionized water was used instead of buffer

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as it does not contain nutrients that could potentially interfere with extracellular enzyme
analysis and because buffering may alter enzyme activity or bioavailability of contaminants/
substrates (Palmroth et al, 2005). The tubes were gently shaken for 20 s to remove
mycorrhizosphere soil clinging to the roots. This solution was used to inoculate Biolog
EcoPlates™ (Biolog, Hayward, CA).

PHC extraction and quantification using GC-FID
A sequential shake method was used for PHC extraction (Schwab et al, 1999; Siddique et
al, 2006). Ten millilitres of acetone:hexane (1:1, vol/vol) were added to approximately 1 g
of soil (wet weight) in 20 mL glass vials. The vials were shaken on a reciprocating platform
shaker at 120 cycles per minute for 30 min. Soil particles were allowed to settle before the
extract was removed. This process was repeated three times and extracts were then
combined.

The extracts were cleaned using a silica gel column procedure to remove polar organic
compounds (CCME, 2001). Glass columns (inside diameter of 16 mm) were plugged with
glass wool and filled with approximately 60 mm of 70-230 mesh Grade 60 A activated
(heated to 110°C for 12 h) silica gel followed by approximately 25 mm of ASC anhydrous
sodium sulfate (dried at 400°C for 4 h). Approximately 10 mL of acetone:hexane (1:1,
vol/vol) was used to condition the column prior to adding the 30 mL of PHC extract solution,
which was accomplished by pipetting the solution into the top of the column and letting it run
down the glass to the top of the sodium sulfate layer. After the extract had passed through
the column, the column was flushed with 10 mL cyclohexane to ensure all compounds of

149

interest were collected. The collection vessels were 40 mL glass vials with Teflon-lined
caps. Two milliliters of toluene were added to each vial and the caps were left open to
evaporate the lighter organic solvents. Once most of the solvents had evaporated, the
concentrated PHC extracts were transferred to 2 mL GC tubes for the final evaporation, and
topped up to exactly 2 mL with toluene if necessary. The PHC extracts were stored at 4°C
until GC-FID analysis. Acetone, hexane (GC grade) and other chemicals (AR grade) were
purchased from Fisher Scientific, Ottawa, Ontario.

The PHC extracts were analyzed on a Varian Model CP 3800 Gas Chromatograph (GC)
equipped with a flame ionization detector (FID). A 15 m x 0.25 mm ID with 0.25 urn film
thickness ZB-5 capillary column (Phenomenex Torrance, CA) was used for the separation of
the PHC extracts. Typically 1 uL of PHC extract was injected into the GC system using a
Varian CP 8400 auto-sampler. Splitless injection mode was performed on the 1079 PTV
injector and after 0.7 min, the split mode was activated at split ratio 10:1. Both the injector
and the detector (FID) temperatures were kept at 320°C during the analysis. The capillary
column temperature was initially held at 50°C for 1 min, then increased at 15.0°C min"1 to
110°C and further increased at 10.0°C min"1 to 300°C and held at 300°C for 10 min. The
total run time was 34 min for each sample. The carrier gas (helium) was maintained at a
constant flow rate of 1.5 mL min"1 for the whole analysis and no pressure pulse was used for
the injection.

PHCs from crude oil were quantified using the CCME reference method for determining
PHC fractions (each fraction based on equivalent normal straight-chain hydrocarbon (nC)

150

boiling point ranges) in soil (CCME, 2001). The PHC fractions F2 (nCIO- nC16), F3 (nC16nC34) and F4 (nC34-nC50) were determined according to the CCME Canada-Wide Standard
for Petroleum Hydrocarbons in Soil - Tier 1 Method (CCME, 2001). Earlier work showed
that the F2 fraction contained mainly aliphatics such as alkanes. The F3 fraction includes
aliphatic hydrocarbons as well as PAHs such as fluorene (C12H10), phenanthrene (C14H10)
and pyrene (Ci 6 Hi 0 ); here, we divided the F3 fraction into F3a (nC16-nC23) and F3b (nC23nC34) fractions. Previous studies have shown the Fl fraction (nC6-nC10) to be negligible
and therefore it was ignored in this study. Peak retention times (i.e. peak maximums) of the
external standards decane (nCIO), hexadecane (nC16), nonadecane (nC19), eicosane (nC20),
tricosane (nC23), dotriacontane (nC32), and tetratriacontane (nC34) (Sigma Aldrich,
Oakville, Ontario) dissolved in toluene were used to determine F2, F3a, F3b and F4 regions
on the GC-FID chromatograms (Figure 3.1); external standards were run concurrently with
samples at concentrations of 10, 25, 50, 125 and 250 ppm.

PHC concentrations (\ig mL"1) were calculated by dividing the areas under the GC-FID
curves by the response factor for the F2, F3a, F3b, F4 and total PHC (tPHCs). These values
were then multiplied by the volume of the GC vial (2.0 mL) divided by the equivalent dry
weight (g) of each soil sample to obtain concentrations in u.g g"1 (ppm). The final
concentrations of PHCs were compared at 1 and 16 weeks by 1 -way ANOVA (a = 0.05)
using Statistica 6.1 (StatSoft Inc., USA).

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F2(nC10-nC16)
4

F3a (nC16-nC23)
fc_

F3b (nC23-nC34)

F4 (nC34+)

Figure 3.1: Overlay of two chromatograms generated from GC-FID analysis showing a
reduction in PHC peak areas from 1 to 16 weeks. The vertical lines represent the boundaries
(based on retention times of standards) for analysis of the F2 (nC10-nC16), F3a (nC16nC23), F3b (nC23-nC34) and F4 (nC34-nC50) PHC fractions.

Soil nutrient analysis andpH
Organic (FH, CWD and FHoil) and mineral soil (Ae and Bf) samples from single-plant
systems were analyzed for total C and N content using <100-mesh samples (air-dried, then
ground in a Model MM200 ball mill; Retsch, Haan, Germany) by dry combustion using a
Model 1500 NC Elemental Analyzer (Fisons, Milan, Italy). The pH of organic soil layers
was measured in a 1:4 soil to deionized H2O suspension while 1:2 suspensions (in deionized
water) were used for mineral soils (Kalra and Maynard, 1991).

LH-PCR and fragment analysis
Mycorrhizosphere-associated bacterial communities (genotype richness and abundance) were
characterized by amplicon length heterogeneity PCR (LH-PCR) using the D4 fluorescent

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dye-labeled forward primer 27F (5'AGAGTTTGATCMTGGCTCAG) and unlabelled reverse
primer 355R (5'GTCGCCTCCCGTAGGAGT) as described by Mills et al (2003). Root
systems were crushed in liquid nitrogen and DNA was extracted using a CTAB (hexadecyl
trimethyl ammonium bromide) protocol with an extra phenol/chloroform-isoamyl alcohol
(1:1) purification step (Fujimura et at, 2008). DNA extracts were cleaned using the Wizard®
PCR Preps DNA Purification System kit (Promega); cleaned extracts were resuspended in
TE buffer. PCR reactions consisted of 3 uL DNA (diluted 1:50), 10X PCR buffer, 2 mM
dNTPs, 50 uM MgCl2, 10 uM forward and reverse primers (Proligo, CO), 0.7 U Platinum
Taq DNA polymerase (Invitrogen Life Technologies), and nuclease-free water (Integrated
DNA Technologies, Inc.) to a final volume of 30 uL. The DNA Engine DYAD™
thermocycler (MJ Research, Inc., Watertown, MA) conditions were as follows: initial
denaturation for 1 min at 94°C, 35 cycles of denaturation (94°C for 45 s), annealing (52°C for
45 s) and extension (72°C for 1 min 30 s), and final extension at 72°C for 10 min. PCR
products were run on 1.2% agarose gels to confirm amplification.

PCR products (2 uL) were loaded into a CEQ™ 8000 sequencer (Beckman-Coulter Inc.)
along with CEQ 400 size standard mixture. Run conditions were 60°C separation
temperature, 4 kV voltage, and 120 min separation time. Analysis was performed using the
amplicon fragment length polymorphism (AFLP) program of the CEQ™ 8000 sequencer and
the cubic model for size standard with a bin width of 1.5 bp. Peaks less than 11% of the total
sample peak height were not included. Profiles from separate DNA extractions and PCR
reactions were compared to assess reproducibility and suitability for analysis.

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Relative abundance of genotypes was calculated by relativizing the fluorescent signal
strength of each fragment peak to the total peak area within each sample (Osborne et al,
2006). Community structure was assessed graphically with Nonmetric Multidimensional
Scaling (NMS) using PC-ORD 5.0 software (McCune and Mefford, 1999; Mills et al, 2006;
Ramette, 2007). NMS was calculated on the basis of a Sarensen distance measure with 50
runs with real and randomized data and a maximum of 500 iterations to assess stability
(instability criterion was 0.00001). A stepwise reduction in dimensionality (6D-1D) was
used to minimize stress along with a random starting configuration (user-provided seeds).
When possible (i.e. for balanced analyses), multivariate differences were tested statistically
with nested permutational multivariate ANOVA (NPMANOVA) (Anderson, 2001);
otherwise, univariate differences were tested with Multi-Response Permutation Procedures
(MRPP) (McCune and Grace, 2002) to examine effects between treatments.

Community level physiological profiles
The metabolic potential of mycorrhizosphere bacteria was assessed by community level
physiological profiles (CLPP) based on broad differences in C-substrate use patterns
(Hofman et al, 2004; Palmroth et al, 2005). Mycorrhizosphere soil solutions (100 uL) were
inoculated into Biolog EcoPlates™ (Biolog, Hayward, California) containing triplicate wells
of 31 substrates commonly found in the rhizosphere (at least nine are considered to be
constituents of plant root exudates) along with a redox dye (tetrazolium violet) that is
reduced to a purple color during substrate oxidation. The plates were incubated in the dark at
~20°C. Reaction patterns (optical density at absorbance of 590 nm) were analyzed with a

154

microplate reader (Biolog Microstation plate reader and Biolog MicroLog 3 4.01 A software)
at 24-h intervals for five days.

For each plate, optical densities (OD590) of triplicate wells were averaged and values for the
water control subtracted from each averaged well value; negative values were adjusted to
zero. Average OD590 values were calculated for substrate guilds: including amino acid (Larginine, L-asparagine, L-phenylalanine, L-serine, L-threonine, glycyl-L-glutamic acid),
amide/ amine (phenylethylamine, putrescine), carbohydrate (D-cellobiose, a-D-lactose, Pmethyl-D-glucoside, D-xylose, I-erythritol, D-mannitol, N-acetyl-D-glucosamine),
carboxylic acid (D-glucosaminic acid, D-galactonic acid, y-lactone, D-galacturonic acid, 2hydroxy benzoic acid, 4-hydroxy benzoic acid, y-hydroxybutyric acid, itaconic acid, aketobutyric acid, D-mallic acid), polymer (tween 40, tween 80, a-cyclodextrin, glycogen),
and miscellaneous (pyruvic acid methyl ester, glucose-1-phosphate, D,L-a-glycerolphosphate) C substrates (Preston-Mafham et ah, 2002). Kinetic analysis of OD590 values by
substrate guild was performed by calculating the area under each substrate use curve from 24
to 120 hours, which corresponded to the log (linear) phase of bacterial growth.

Area-under-curve values for PHC-treated and untreated communities were compared by 1 way ANOVA (a = 0.05) with either plant or organic soil treatments pooled. Principal
components analysis (PCA) was used to assess treatment and substrate effects on variation in
bacterial community profiles. All analyses were conducted using Statistica 6.1 (StatSoft Inc.,
USA).

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Results
PHC quantification from soils
Not all plant - organic soil combinations germinated in sufficient quantities to allow for a
full factorial treatment with an equal number of replicates; thus, treatments with n<3 were
not included in PHC analyses. For the single-plant systems (2005-06), triplicate samples
were obtained at the two harvest dates (t = 1, 16 weeks) for P-FH, P-CWD, P-FHoil, B-FH,
L-FH, L-CWD, L-FHoil and the unplanted controls (N-FH, N-CWD, N-FHoil), for a total of
30 pots (each with three soil horizons) per harvest. For the double-plant systems (2006-07),
triplicate samples were obtained at the two harvest times for PL-FH, PL-CWD and the
unplanted controls (N-FH, N-CWD), for a total of 12 pots per harvest.

Total PHC concentrations (sum of F2, F3a, F3b, F4 fractions) in organic layers of PHCtreated soil systems averaged over 150,000 ppm at 1 week (Figure 3.2). The high ppm was
due, in part, to the lower particle density in organic compared to mineral layers. PHCs
extracted from organic (FH, CWD and FHoil) soil layers accounted for 90% of the total
PHCs extracted from each soil system; PHC levels in Ae layers were typically greater
(~7.3%) than in Bf layers (~1%). In organic layers, PHC concentrations generally decreased
over 16 weeks, with significant losses in FH (p<0.001) and CWD (p=0.02) soils only. The
levels of PHC-like chemicals extracted from untreated (control) FH and CWD layers were
very low, indicating that the GC-FID chromatograms largely represented the PHCs added to
these systems (and not other soil constituents such as solvent-extractable soil organic matter).
In FHoil layers, up to 39% (mean of 15%) of PHCs extracted from PHC-treated soils
represented residual PHCs from the in situ soil contamination event 8 months earlier. No

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significant changes in PHC concentration were apparent in mineral soil layers after 16 weeks,
although levels in Ae layers tended to increase with time.

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H

50O0
2500

Bf

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.JHIHkzzEzzzi—

_^H^^^.

EH

CWD

EHdl

Organic Soil Layer

Figure 3.2: Concentration of total PHCs (ppm) in three organic soil layers (FH, CWD and
FHoil) for PHC-treated and untreated controls at 1 and 16 weeks (data pooled for plant
treatment). Bars represent standard errors of the means. Significant losses of PHCs within
organic soil treatment groups are indicated by *.

Analysis of PHC fractions within plant — organic soil systems
Planted and unplanted systems were included in this analysis (i.e. P-FH, P-CWD, P-FHoil,
B-FH, L-FH, L-CWD, and L-FHoil single-plant systems, PL-FH and PL-CWD double-plant
systems, and N-FH, N-CWD, and N-FHoil no-plant systems). The F2 (nC10-nC16) fraction
157

PHCs from organic soil layers averaged 35,000 ppm after 1 week and 6,000 ppm after 16
weeks. Significant decreases in F2 PHCs were found in most planted and unplanted organic
layers, including the P-CWD (p=0.002), B-FH (p=0.02), L-CWD (p=0.03), L-FHoil
(p=0.001), N-FH (p=0.02), and N-CWD (p=0.02) treatments. The F3a (nC16-nC23) and F3b
(nC23-nC34) fraction PHCs (containing straight-chain, branched and cyclic alkanes as well
as small PAHs) were generally present between 40,000 and 80,000 ppm, but values varied
greatly both within and between treatment groups (Figure 3.3). The F4 (nC34-nC50) fraction
PHCs (containing some larger PAHs) were usually present in organic layers at <20,000 ppm.
Although levels of larger PHC (>C16) chemicals also tended to decrease with time in organic
soil layers, only P-CWD and B-FH systems showed consistently significant decreases in F3a
(p=0.001 for both treatments), F3b (p-0.004 and 0.001, respectively) and F4 (p=0.002 and
0.009, respectively) fraction PHC concentrations over the study period.

In contrast to the organic layers, PHC concentrations tended to increase with time in Ae
layers (perhaps due to leaching) (Figure 3.4). PHC levels in Ae layers under the CWD layer
were extremely variable compared to levels in FH systems. F2, F3a, F3b and F4 fraction
PHC levels were usually between 1,000 and 8,000 ppm; not high enough to account for all
PHC loss from the organic layers above. Again, F4-fraction PHCs were the lowest (-1,000
ppm). For the Bf layers, PHC levels averaged about 1,000 ppm (Figure 3.5). PHC levels
tended to increase most in the lingonberry compared to other plant systems.

158

.

m

-

+ F3a
~ * T F3b
f
F4 <

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80, 000

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L

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16 wk

Figure 3.3: Concentrations of F2 (nC10-nC16), F3a (nC16-nC23), F3b (nC23-nC34), and F4
(nC34-nC50) PHCs (ppm) in organic layers of plant - soil systems at 1 and 16 weeks after
PHC treatment. Plant treatments include: pine (P), birch (B), lingonberry (L), pine and
lingonberry (P+L), and no plant (N). Bars represent standard errors of the means (n=3);
significant differences (1-way ANOVA) within treatment groups are indicated by *.
159

Ae (FH)

* i

j T F2
~ F F3a
T F3b
37. F4

1

i
i.

L

«

i

T
1

(

1

\

tt

.4 i-

i

Ae (CWD)

Ae (FHoil)

-1
L

PL

N

J
L

1 wk

PL

16 wk

Figure 3.4: Concentrations of F2 (nC10-nC16), F3a (nC16-nC23), F3b (nC23-nC34), and F4
(nC34-nC50) PHCs (ppm) in Ae layers of plant - soil systems at 1 and 16 weeks after PHC
treatment. Plant treatments include: pine (P), birch (B), lingonberry (L), pine and
lingonberry (P+L), and no plant (N). Bars represent standard errors of the means (n=3);
significant differences (1-way ANOVA) within treatment groups are indicated by *.
160

Bf(FH)

F3a
F3b
F4

+*

\i &

i

ti

Bf(CWD)

i r *

*

Bf(FHoil)

4$

L

PL

L

PL

16 wk

1 wk

Figure 3.5: Concentrations of F2 (nC10-nC16), F3a (nC16-nC23), F3b (nC23-nC34), and F4
(nC34-nC50) PHCs (ppm) in Bf layers of plant - soil systems at 1 and 16 weeks after PHC
treatment. Plant treatments include: pine (P), birch (B), lingonberry (L), pine and
lingonberry (P+L), and no plant (N). Bars represent standard errors of the means (n=3);
significant differences (1-way ANOVA) within treatment groups are indicated by *.
161

Soil properties
The C and N contents were much greater in organic (FH, CWD, or FHoil) compared to
mineral (Ae or Bf) soil layers (Table 3.2). In organic layers, C content ranged from
approximately 19-48% (mean C content of mineral soils was <2%) and was significantly
greater (p<0.001) in PHC-treated soils compared to controls at both 1 and 16 weeks. The
CWD layers contained significantly more C (p<0.001) than either of the FH or FHoil layers.
The C content tended to be greater in FHoil compared to FH layers, presumably due to C
inputs from the earlier PHC contamination event. Similar C levels were observed for all
mineral soils at 1 week, but after 16 weeks, C levels tended to be greater in mineral soils
below the PHC-treated organic layers, including the FHoil controls.

Total N ranged from approximately 0.28-0.64% in organic layers and generally did not vary
with PHC treatment or over time. In contrast, mean N content of mineral soils was usually
<0.1%. For both PHC-treated and control systems, N was significantly lower (pO.OOl) in
CWD soils compared to FH and FHoil soils at 1 and 16 weeks; mean N content of CWD was
approximately half that of FH and FHoil soils. The combination of greater C and lower N
content in CWD resulted in high C:N ratios that ranged from approximately 135 tol83 and
were significantly greater (p<0.001) than the C:N ratios in FH and FHoil soils, which were
similar (i.e. ranging from approximately 30 to 65). Overall, C:N ratios did not change much
between 1 and 16 weeks.

Comparison of planted and unplanted groups (data not shown) generally showed no
differences in N and C content in organic or mineral soil layers, although, in Ae soils, C

162

tended to increase from 1 to 16 weeks in planted compared to unplanted systems (as well as
in PHC-treated compared to untreated systems). There were no C differences in the Bf soils
for the different plant treatments.

Soil pH (water) was generally lower in the organic and Ae soils (means of 5.3 and 5.4,
respectively) than the Bf soils (mean of 5.7) at 1 week, but tended to increase after 16 weeks;
this trend was greater in the untreated controls compared to the PHC-treated soils. In FH and
CWD controls, pH significantly increased in all three soil layers (organic, Ae, and Bf), but no
significant changes occurred in the FHoil controls. In the FHoil soil systems, pH was
generally lower than in the FH and CWD systems, particularly in the Bf layer. Comparisons
of plant treatments revealed no significant differences.

163

Table 3.2: Means (± standard errors) for soil properties (C, N, pH) in control and PHCtreated plant - organic soil systems at 1 and 16 weeks. Percent C and N are reported on an
air-dry basis.
Weekl
Total C %
Org
Ae
Bf
Total N %
Org
Ae
Bf
C:N Ratio
Org
Ae
Bf
pH
Org
Ae
Bf

noPHC

FH
PHC-treated

noPHC

CWD
PHC-treated

noPHC

FHoil
PHC-treated

19.07 ±2.040
1.04 ±0.235
1.09 ±0.189

33.80 ±3.751
1.68 ±0.235
1.75 ±0.189

40.9 ±1.761
1.46 ±0.210
1.37 ±0.169

48.4 ± 2.544
1.90 ±0.210
1.34 ±0.169

23.1 ±3.397
1.07 ±0.210
2.27 ±0.169

36.92 ±1.911
1.68 ±0.210
2.18 ±0.169

0.60 ± 0.062
0.055 ± 0.003
0.062 ± 0.004

0.59 ± 0.034
0.058 ± 0.003
0.071 ±0.004

0.31 ±0.024
0.045 ± 0.003
0.067 ± 0.004

0.28 ± 0.029
0.040 ± 0.003
0.060 ± 0.004

0.60 ± 0.094
0.054 ± 0.003
0.11 ±0.004

0.57 ±0.031
0.061 ± 0.003
0.10 ±0.004

31.9± 1.142
18.7 ±3.873
17.4 ±1.755

56.9 ±5.291
28.7 ±3.873
24.6 ±1.755

134.9 ±11.44
31.7 ±3.464
20.4 ±1.569

182.0 ±26.46
46.7 ±3.464
22.8 ±1.569

40.3 ±1.768
19.6 ±3.464
20.8 ±1.569

64.4 ± 0.840
27.6 ±3.464
21.5 ±1.569

5.5 ±0.1
5.2 ±0.1
5.5 ±0.1

5.4 ±0.1
5.0 ±0.1
5.9 ±0.1

4.9 ±0.1
5.3 ±0.1
5.8 ±0.1

5.0 ±0.1
5.2 ±0.1
5.5 ±0.1

5.5 ±0.1
5.3 ±0.1
5.4 ±0.1

5.4 ±0.1
5.3 ±0.1
5.4 ±0.1

Week 16
Total C %
Org
Ae
Bf
Total N %
Org
Ae
Bf
C:N Ratio
Org
Ae
Bf
pH
Org
Ae
Bf

noPHC

FH
PHC-treated

noPHC

CWD
PHC-treated

noPHC

FHoil
PHC-treated

22.4 ±1.866
0.89 ±0.235
1.42 ±0.189

29.3 ± 5.228
2.32 ± 0.235
1.92 ±0.189

47.8 ±2.275
1.09 ±0.235
1.53 ±0.189

53.8 ±1.750
2.83 ± 0.235
2.05 ±0.219

21.7 ±2.354
1.41 ±0.235
2.11 ±0.189

37.2 ±3.596
2.080 ±0.210
2.18 ±0.169

0.644 ± 0.073
0.049 ± 0.003
0.073 ± 0.004

0.563 ± 0.063
0.051 ±0.003
0.081 ±0.004

0.29 ± 0.047
0.045 ± 0.003
0.080 ± 0.004

0.30 ±0.021
0.050 ± 0.003
0.079 ± 0.004

0.558 ±0.056
0.064 ± 0.003
0.104 ±0.005

0.574 ± 0.041
0.062 ± 0.003
0.101 ±0.004

35.3 ±1.807
18.0 ±3.873
19.3 ±1.755

51.7 ±6.547
45.7 ±3.873
23.7 ±1.755

174.1 ±28.83
24.4 ±3.873
19.2 ±1.755

183.3 ±20.33
56.4 ±3.87
25.7 ±2.03

38.9 ±1.358
22.2 ±3.873
20.3 ±1.755

64.4 ±3.320
33.1 ±3.464
21.4 ±1.569

5.6 ±0.2
5.8 ±0.1
5.9 ±0.1

5.4 ±0.1
5.3 ±0.1
6.1 ±0.1

5.4 ±0.3
5.7 ±0.1
6.0 ±0.1

5.1 ±0.2
5.2 ±0.1
6.0 ±0.1

5.3 ±0.1
5.6 ±0.1
5.5 ±0.1

5.0 ±0.1
5.6 ±0.1
5.6 ±0.1

164

Bacterial community structure
Overall, PHC treatment of the plant - soil systems had no effect on genotypic richness or
structure (composition) of bacterial communities. Pairwise comparisons (MRPP) of PHC
treated systems at 1 and 16 weeks revealed that genotype richness increased significantly
(p=0.007) over time. Community structure varied significantly (p=0.036) between PHCtreated and control groups after 16 weeks.

Analysis (NPMANOVA and MRPP) of the single-plant (i.e. pine, birch and lingonberry)
systems showed significant differences (p=0.002) in bacterial community structure in the
different organic soil layer treatments. Pairwise comparisons indicated that the differences
were between FH and FHoil groups (p<0.001), and CWD and FHoil groups (p=0.009), but
not between FH and CWD groups. There were no interaction effects between organic soil
layer and PHC groups.

The type of plant (i.e. pine, birch or lingonberry) appeared to have less of an effect on
genotype richness and community structure than the complexity (i.e. single- or double-plant)
of the systems (Figure 3.6). The single-plant systems did not significantly differ with respect
to either genotype richness or community structure. However, community structure varied
significantly (p<0.001) for both pine and lingonberry in double- compared to single-plant
systems; community structure also varied between pine and lingonberry within the doubleplant systems. These patterns were consistent for the richness data, with an additional
significant difference (p<0.001) between pine and lingonberry in the single-plant systems.
There were no interaction effects between plant and either PHC, harvest, or organic soil layer

165

groups. The overall effect of plant on bacterial community structure is shown in the NMS
ordination (Figure 3.7). The figure shows tight clustering (in the lower left quadrant) of
communities from the double-plant systems within the more dispersed distribution of singleplant system communities.

Seven DNA fragments corresponded to 16S fragment lengths for Pseudomonas (343, 344 bp)
and Sphingomonas (317, 318, 319, 320, and 335 bp) species (Mills et ah, 2003). The
frequency of these fragments (nested one-way ANOVA) varied with plant, but did not vary
over time or with organic soil layer or PHC treatment, although some genotypes increased or
decreased in frequency with different plants.

B i r c h

Pine

single" /

*• /

Pine

^ubie

single

\ * Lingonberry single
«

\

t

» Lingonberry double

Figure 3.6: Schematic diagram showing plant effects on bacterial genotype richness and
community structure. Arrows (solid, genotype richness and community structure; dashed,
richness only) between plant treatments indicate significant differences. Treatments not
connected by arrows are not significantly different.

166

•

I H Pine (single)
9

Birch (single)
Lingonberry (single)

|

| Pine (double)
Lingonberry (double)

•
• B>

•

•
•

9%

/•"•*

•

©

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II

t

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Axis 1 (r2 = 59.6%)

Figure 3.7: NMS of bacterial community structure associated with single- (pine, birch or
lingonberry) and double- (pine + lingonberry) plant systems (stress = 16.42; instability =
0.06).

167

Bacterial community level physiological

profiles

Areas under C substrate oxidation curves for bacterial communities growing on EcoPlates™
were consistently greater for amino acids, carbohydrates and carboxylic acids in all
treatments; lower values were observed for amides/ amines, polymers and the miscellaneous
substrates. No differences in substrate use were found between PHC-treated and untreated
communities or within the different organic soil groups. In the unplanted FH and CWD
systems, areas under the substrate use curves were lower in the PHC-treated compared to
untreated systems (which were similar to the planted system profiles); in the unplanted FHoil
systems, areas under the curves were virtually identical in the PHC-treated and control soils
and both treatments produced profiles similar to the planted system profiles.

Principal components analysis (PCA) of the areas under substrate use curves (24-120 h) for
the various plant - organic layer - PHC treatments generated the ordination shown in Figure
3.8; the two principal components axes explained 85.02% of the variation between samples.
The N-containing substrates (amines/ amides and amino acids) showed the greatest relative
effect on the plot, followed by carbohydrates and carboxylic acids. Of the treatment
variables, plant exhibited a greater effect than either organic soil or PHC. There was a
tendency for unplanted systems to group away from the planted systems (i.e. towards the
right side of the plot); the lingonberry treatments also grouped slightly away from pine, birch
and alder (i.e. towards the lower quadrants of the plot).

168

•

Pine

#

Birch

"•"-' Alder
Lingontaerry

•

No Plant

Carbohydrates
Amino acids

°1
r-i
Carboxylic acids

Plant
Amines/Amides
Axis 1 (72.08%)

Figure 3.8: PCA ordination of areas under C substrate curves showing the relative influence
of PHC, organic soil and plant treatment variables. Plant effects are indicated by different
symbols: pine (squares), birch (circles), alder (stars), lingonberry (triangles), and no plant
(crosses).

169

Discussion
PHC Biodegradation
The process of PHC biodegradation in reconstructed sub-boreal forest soil systems was more
closely related to qualities of the organic soil layer than to genotypic or broad metabolic
structure of the bacterial communities in the mycorrhizosphere. We found significant
reduction of total PHC levels following (1-16 weeks) addition of crude oil to pristine forest
floor (FH) and coarse woody debris (CWD) soil systems. These lignin-rich and
metabolically active soil layers form substantial surface components of northern forest
landscapes (Lundstrom et al, 2000; Prescott et al, 2000). Their tendency to retain PHCs in
the soil organic matter was evident in our study, as PHC concentrations were far greater than
those extracted from the underlying mineral soils in each system. Soil analyses showed a
significantly lower pH and higher C:N ratio in CWD compared to FH layers at both 1 and 16
weeks; however, these differences did not appear to reduce biodegradation capacity of these
soil communities. The intrinsic ability for PHC biodegradation existed within soil microbial
communities of the pristine organic layers.

Prior in situ treatment of the forest floor with PHCs did not appear to have enhanced
microbial biodegradation capacity for subsequent PHC treatment in these soils, as was
expected (Miller and Herman 1997; Braddock et al, 2003; Seghers et al, 2003; Diaz 2004).
The lack of significant PHC reduction over the study period found in the previously
contaminated (FHoil) soil layer may be partly explained by the presence of residual PHCs.
Residual PHCs likely consisted of a greater proportion of more recalcitrant compounds that
contributed to a more chemically complex and heterogeneous environment. Aging of

170

contaminants in soil generally reduces bioavailability to decomposer organisms (Alexander,
2000). This may have increased variation in biodegradation patterns within treatment groups
and masked any significant PHC loss over the 16-week duration of the experiment. The C:N
ratio of FHoil soils was not significantly greater than FH soils; neither differed substantially
from the C:N ratios reported for the forest floor measured in situ (about 50) (Arocena and
Sanborn, 1999) or from oil-contaminated soils in Alaska (52.7) (Braddock et al, 2003). The
pH of FHoil was similar to CWD; both were lower than FH, particularly after 16 weeks.
These physico-chemical soil properties were therefore not related to the differences in
biodegradation observed among the different organic soil layers. Although FH and CWD
exhibited significantly different bacterial community structure patterns compared to FHoil
layers, no differences in C use profiles were found.

Processes that reduce PHC concentrations in oiled soils over time (i.e. biodegradation,
evaporation, water washing, or photooxidation) lead to distinct chemical profiles in soil; a
decrease in n-alkane (<nC17) concentration relative to larger compounds and PAHs is typical
of biodegradation profiles (Braddock et al, 2003). In general, we found a trend of reduced
F2 (nC10-nC16) PHCs for all treatment groups by 16 weeks. Several plant - soil systems
exhibited significant decreases in F2 concentrations, including both planted (birch -FH, pine CWD, lingonberry -CWD and -FHoil) and unplanted (-FH and -CWD) systems. This finding
was not surprising as many genera of soil bacteria are known to completely mineralize
aliphatic compounds (e.g. alkanes <nC16) through central metabolic pathways such as 0oxidation and Kreb's cycle (McGill et al, 1981; Miller and Herman 1997). Our results
suggest that biodegradation capacity for F2 PHCs exists within all the soil systems tested.

171

Also, unplanted soils, which originated as sub-boreal forest rhizosphere, continued to support
biodegradative microbial communities for at least eight months after soil collection.

In contrast, significant decreases of F3a (nC16-nC23), F3b (nC23-nC34) and F4 (nC34nC50) PHCs occurred only in pine -CWD and birch -FFf systems. Enhanced PAH
degradation in rhizosphere compared to non-rhizosphere soils was also reported by Joner et
al. (2006). Microbial utilization of PHCs often creates demand for other nutrients such as N
and P (Miller and Herman, 1997). Thus, it is likely that the energy (C) and nutrient (N and
P) resources necessary to support biodegradation of these larger and more chemically
complex PHCs were limited in the unplanted soil systems that lacked inputs of C-rich
substrates through exudation from mycorrhizal roots. Strong sorption of these larger PHC
compounds to soil components may have also reduced their bioavailability to degrading
microorganisms. Alternatively, the lack of significant change in levels of larger PHCs may
have been due to slow decomposition of these compounds (i.e. perhaps 16 weeks was not
long enough to detect changes in PHC concentrations).

Mycorrhizosphere

effect

The mycorrhizosphere generally supports greater microbial biomass and activity (i.e.
mycorrhizosphere effect) than non-rhizosphere soils, which increases the biodegradative
potential of planted over unplanted systems (Linderman, 1988; Ingham and Molina, 1991;
Rygiewicz and Anderson, 1994). In PHC-contaminated soils, plant-fungal exudates are also
expected to support proliferation of functional groups of organisms with enhanced capacity
for cometabolic biodegradation of PHCs. With respect to pyrene biodegradation, Mueller

172

and Shann (2007) showed that the quality of C in root exudates was more important than C
quantity, suggesting that specific mycorrhizosphere communities may influence
biodegradation. Several studies have reported elevated densities of PHC-degrading bacteria
in contaminated soils (Lindstrom et al., 1999; Delille et al, 2004). Braddock et al. (2003)
found that culturable populations of total heterotrophic and crude oil emulsifying bacteria
were elevated in soils 25 years after an Alaskan oil spill compared to soils from an adjacent
reference plot. Our comparisons of bacterial community C use profiles distinguished
unplanted from planted soils on the PCA ordination. Palmroth et al. (2005) also showed C
utilization differences between vegetated (pine and poplar) and unvegetated soil communities
using similar methods. In our study, nested analyses within organic soil and PHC treatment
groups showed lower metabolic activity in only the untreated FH and CWD systems;
preliminary plate counts (data not shown) also indicated lower culturable bacterial density in
untreated (i.e. no PHC added) FH and CWD (104-105 CFUs ml/ 1 soil solution) compared to
either the untreated FHoil or the PHC-treated FH, CWD and FHoil (107-108 CFUs mL"1 soil
solution) systems. These findings suggest that C inputs by both plants and PHCs support
greater bacterial density; however, a plant appears to enhance biodegradation of >nC16
PHCs.

The finding that statistically significant reductions of the larger PHC (nC16+) fractions
occurred in only planted (pine and birch) systems provides indirect evidence for a
stimulatory role of ECMs in the biodegradation process. ECMs may simply provide habitat
and C substrates that enhance bacterial cometabolism, as described earlier (Sarand et al,
1998; 2000; Heinonsalo et al, 2004). In addition, ECMs may secrete oxidative enzymes that

173

open aromatic ring structures, thereby overcoming the thermodynamically limiting step in
PAH metabolism (Burke and Cairney, 2002; Sen, 2003). Several recent studies have
reported that mycorrhizas inhibited biodegradation of naphthalene, fluorene (Genney et al,
2004), anthracene, anthraquinone, chrysene, dibenz[a,h]anthracene (Joner et al, 2006), and
pyrene (Koivula et al, 2004) when individually spiked into microcosms. Genney et al.
(2004) suggested that ECMs inhibit PHC biodegradation in situations of C limitation. Thus,
in these studies, PAH treatment in the absence of cometabolic substrates may have led to
increased competition between fungi and bacteria for the same energy resources, resulting in
decreased PAH mineralization and accumulation of dead-end metabolites in soil (Gadgil and
Gadgil, 1971).

Differences in experimental design may explain our findings in context with these studies. In
our study the establishment of diverse ECM communities on the fine roots of pine and birch
was observed to be virtually ubiquitous, which likely reduced free-living soil fungi in the
rhizosphere and thus limited this group of competitors (Lindahl et al., 2007). Incorporation
of soil depth (i.e. increased three-dimensional space in miniature soil systems that contained
horizons) in our mycorrhizal systems was also expected to reduce competition between ECM
fungi with saprotrophic activities and the associated bacterial community (Cairney, 2005).
Also, addition of reduced C substrates in PHC mixtures probably fueled the large
heterotrophic biomass associated with the pine and birch mycorrhizospheres (Nannipieri et
al., 2003; Morgan et al., 2005), resulting in a generally stimulatory effect on biodegradation.
This explanation is supported by Koivula et al. (2004), who found that inhibition of PHC
biodegradation (due to C limitation) was eased in oiled soils where alternate C substrates

174

were available. However, an overall inhibitory effect on biodegradation could explain why
most mycorrhizal systems (including ERM systems) did not exhibit significant losses of
larger PHCs in the current study.

Although ERMs have been reported to exhibit oxidative enzyme activity (Burke and Cairney,
2002), the lack of significant loss of larger PHCs indicates that they did not contribute to
biodegradation in our lingonberry systems. One explanation may be that the fine root
systems were shallow and lacked extensive ERM mycelia that did not sufficiently fill the
larger pots into which the single lingonberry seedlings were planted; secreted enzymes and
exudates may have been diluted with distance from the ERM roots. Joner and Leyval (2003)
reported that the (mycor)rhizosphere effect on PAH degradation sometimes does not extend
more than 1 mm from roots colonized by arbuscular mycorrhizas. Corgie et al. (2003)
described bacterial gradients with increased densities of heterotrophs and PAH degraders
closest to the roots (0-3 mm) that corresponded to a gradient of phenanthrene biodegradation.
In a molecular study, Corgie et al. (2006) showed different bacterial communities were
selected by rhizosphere depending on distance from the roots; bacteria exhibited different
activity profiles and values for biodegradation. Alternatively, it is possible that lingonberry
roots were not mycorrhizal (i.e. cell colonization, which was confirmed by microscopy, may
not have included ERM fungi), although we assumed that ERM fungi accounted for at least
some of the endophytic community. Very little is known regarding biodegradation capacity
in ERM systems, and further investigation is warranted.

175

In the single-plant systems examined here and by Palmroth et al. (2005), C use profiles
showed little differentiation between bacterial communities associated with the
mycorrhizosphere of specific plants, although lingonberry appeared to cluster slightly away
from pine, birch and alder. Bacterial genotype richness was significantly greater in
lingonberry compared to pine and birch systems. It is unknown whether this greater richness
is somehow related to the significant biodegradation for the F2 PHC fraction observed in
lingonberry -CWD and -FHoil systems. Bacterial community structure varied significantly
for both pine and lingonberry in the double-plant systems compared to any of the single-plant
systems. Furthermore, bacterial community structure varied significantly between
lingonberry and pine when the two plants shared the same pot. The NMS analysis
graphically reaffirms that growth conditions between double- (tight clustering) and single(more dispersed distribution) plant systems are different enough to influence bacteria
community structure. The soil conditions being the same, we presume that a double-plant
system augments the level of competitive interactions (perhaps by providing more
specialized niches owing to different root systems) compared to a single-plant system, and
that is reflected in the bacterial diversity. Although significant PHC biodegradation was not
observed in either of the double-plant (pine and lingonberry -FH or -CWD) systems tested,
this may represent another example of increased environmental complexity where withingroup variation was too great to detect PHC reduction over 16 weeks.

It was unfortunate that difficulties with seed germination resulted in incomplete datasets for
the alder treatment. Associations with N-fixing Frankia were expected to enhance PHC
biodegradation via a gradual and continuous delivery of N and C to the rhizosphere

176

environment (Selmants et al, 2005; Roy et al, 2007). A hint of the effect of greater N
availability on mycorrhizosphere communities (i.e. altered amino acid requirements) may
have slightly influenced the C use distribution of alder compared to the other ECM systems.
Interactions between this additional functional group and the rest of the mycorrhizosphere
community should be investigated in future PHC biodegradation studies.

Overall, surface application of crude oil appeared to have minimal impact on the diversity
(genotype richness and relative abundance) or metabolic capacity (community physiological
profiles) of bacterial communities in the mycorrhizosphere. Huertas et al. (2000) suggested
that heterogeneity in the soil matrix may provide a protective effect against the solvent shock
associated with initial PHC contamination. Other studies have also reported the inability to
distinguish communities from PHC-contaminated and reference soils on the basis of broad
physiologies such as C mineralization; bacterial communities surviving in oiled soils have
been described as metabolic generalists (Lindstrom et al, 1999; Palmroth et al, 2005).
However, the physiological capacity for biodegradation of various organic substrates, which
is controlled by the functional redundancy of the community originally present (Setala et al,
2000; Delille et al, 2003), seems to remain intact in PHC-contaminated soils for some time
in the absence of further disturbance.

The methods used in this study did not distinguish the PHC-degrading component from the
overall composition of bacterial communities. However, common hydrocarbon degraders
such as Pseudomonas sp. grow well on Biolog EcoPlates

and thus were expected to be

represented as part of the C-utilizing community assessed. The molecular dataset was also

177

expected to include known PHC-degraders (e.g. pseudomonads and sphingomonads) whose
corresponding fragment lengths were identified (i.e. 342-344 bp and 317-320, 334-335 bp,
respectively) by Mills et al. (2003) using the same rDNA primer pair. These fragments
appeared in both PHC-treated and control mycorrhizospheres of pine, birch and lingonberry
systems, with no differences in frequency observed from 1-16 weeks.

Conclusions
We found that mycorrhizosphere communities enhanced PHC biodegradation in
reconstructed sub-boreal forest soils, particularly in ECM systems. The ECM and ERM
fungal communities associated with these systems and their roles in the biodegradation
process were investigated in Chapter 2. The level of PHC contamination used in this study
appeared to have minimal impact on soil bacterial community structure or broad metabolic
functions associated with PHC biodegradation.

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Chapter 4: Root-associated microbial communities differ with Pinus contorta var.
latifolia and Vaccinium vitis-idaea co-inhabiting sub-boreal forest soils
Abstract
Rhizosphere communities, including mycorrhizas and closely associated heterotrophic
microorganisms, represent critical functional groups for decomposition and carbon/ nutrient
cycling processes in northern forest ecosystems. The spatial heterogeneity of the soil
environment contributes to the great biodiversity of soil microorganisms. However, the
complexity of the system and the multifunctional nature of many microorganisms have made
it difficult to comprehend linkages between communities and ecosystem processes and also
to predict how ecosystems may respond to environmental disturbances such as soil
contamination. In this study, we used a bioassay approach to assess the relative contributions
of plant and soil properties to spatial distribution patterns of ecto- (ECM) and ericoid (ERM)
mycorrhizal fungi as well as root-associated bacterial communities inhabiting the shared
rhizosphere of pine (ECM host) and lingonberry (ERM host). Soil systems were either
untreated or treated with petroleum hydrocarbons (PHCs), simulating contamination events.
Surface-sterilized pine {Pinus contorta var. latifolia) seeds and lingonberry {Vaccinium vitisidaea) seedlings were planted into Conetainer

pots containing an organic layer (mor

humus [FH] or coarse woody debris [CWD]) overlying sandy mineral horizons (Ae and Bf)
of field-collected forest soils obtained from central BC, Canada. After 4 months, BC light
crude oil (219 mg cm"2) was applied to the soil surface around the seedling stem; systems
were destructively sampled at 1 and 16 weeks following treatment. Soils from each layer
were analyzed for PHC concentration (not presented here), pH and total C and N content.
The composition, relative abundance and vertical distribution (i.e. variation with soil layer)

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of eight frequently occurring ECMs on pine roots were assessed using light microscopy.
Community profiles (i.e. based on the relative abundance of all genotypes) were generated
for all root systems using length heterogeneity PCR and primers targeting the ITS (fungi) and
16S (bacteria) regions of ribosomal DNA. We found that the main components of ECM
communities were consistent with those described from field-based studies. Genotype
analysis by non-metric multidimensional scaling (NMS) and multi-response permutation
procedures (MRPP) revealed that both plant and soil properties influenced the structure of
root-associated fungal and bacterial communities; however, patterns of community structure
varied among the different functional groups. Fungal communities were distinctly different
on pine (ECM) and lingonberry (ERM) roots; only ECM fungal communities were structured
vertically in the three layers of soil, representing direct interactions between fungi and soil in
the ectomycorrhizal association. In contrast, ERM communities appeared to vary more
between soil systems (i.e. FH-Ae-Bf and CWD-Ae-Bf) than between soil layers. Bacterial
community structure varied between mycorrhizal root systems and between soil layers,
indicating that differences between the ECM and ERM root environment and soil properties
are both important with respect to bacterial niche differentiation. PHC contamination
appeared to have little effect on the composition of root-associated microbial communities.

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Introduction
Rhizosphere communities, including mycorrhizas and closely associated heterotrophic
microorganisms, represent critical functional groups for decomposition and carbon/ nutrient
cycling processes in northern forest ecosystems. The spatial heterogeneity of the soil
environment contributes to the great biodiversity of soil microorganisms and promotes
species co-existence through greater resource partitioning (Ettema and Wardle, 2002;
Ramette and Tiedje, 2007). However, the complexity of the system and the multifunctional
nature of many microorganisms have made it difficult to comprehend linkages between
communities and ecosystem processes and also to predict how ecosystems may respond to
environmental disturbances such as soil contamination (Barrios, 2007). Disturbances may
directly impact microbial functions, lead to loss of a functional group, change rates of
ecosystem processes or alter resource availability (Balser et ah, 2006). The question arises
as to how much detail is necessary to understand system behaviour and to mitigate for effects
of environmental disturbance (Surridge, 2006). To answer this question may require that we
step back from studying individual system components in favour of studying functional
groups of soil biota that have tight linkages to functions that underpin soil-based ecosystem
services (Balser et al., 2006; Smithwick, 2006; Barrios, 2007).

Northern forests are dominated by trees in the families Pinaceae, Betulaceae, Fagaceae and
Salicaceae that typically form ectomycorrhizal (ECM) symbioses with soil fungi that exhibit
variable distribution patterns across forest landscapes according to host specificity (Molina et
ah, 1992) and soil heterogeneity (Horton and Bruns, 2001). ECMs occupy the structural and
functional interface between the plant and soil environment and regulate plant nutrient uptake

186

as well as carbon release to soil (Smith and Read, 1997). These mechanisms of carbon and
nutrient exchange are understood to some extent (i.e. at genetic and biochemical levels) in
ECM systems, but not in an ecological context, which involves interactions with other
microbial communities sharing the rhizosphere. Although interactions with heterotrophic
bacterial communities associated with the ECM mantle and extraradical mycelia are known
to be important for mobilization, uptake and translocation of nutrients (Burke and Cairney,
1998), an ecological understanding is impeded by that lack of integration between ECM
fungal and bacterial studies. Additionally, roots of understory vegetation such as Ericaceae
(e.g. Vaccinium, Rhododendron, Gaultheria, species) commonly share ECM rhizosphere
space (Read and Perez-Moreno, 2003). These plants usually form ericoid mycorrhizas
(ERMs) with fungi that are often unidentified, but also appear to exhibit high diversity at
small scales (Berch et ah, 2002; Perotto et al., 2002) and are expected to contribute to
nutrient cycling and decomposition processes in forests (Cairney, 2000; Read and PerezMoreno, 2003). Interactions between different mycorrhizal communities, as well as potential
sharing of fungal symbionts, may contribute to ecological processes in important ways that
we do not yet understand.

Petroleum hydrocarbon (PHC) contamination is a type of environmental disturbance that can
lead to considerable changes in physical and chemical soil properties on spatial scales
ranging from square meters to hectares. Initially, spilled PHCs spread laterally within the
organic layer of forest soils; lighter fractions eventually move down the soil profile, along the
paths of roots and fissures. Fragmentary patterns of PHC constituents that persist in mobile
form, fixed in soil pores and fissures, adsorbed on the surface of organic and mineral soil

187

constituents, or forming a free-phase continuous cover on the soil surface (Trofimov and
Rozanova, 2003) increase soil heterogeneity and have unknown effects on soil microbial
communities. The major impacts appear to be related to altered soil water, nutrient and
oxygen regimes (Tarradellas and Bitton, 1997). At the same time, input of labile C substrates
(and subsequent release of soil communities from C limitation) may promote metabolic
activity on the part of all the microorganisms not directly inhibited by the PHCs and provide
the potential for them to play a major role in decomposition and nutrient mobilization (Read
and Perez-Moreno, 2003). The unevenness of PHC distribution in different soil layers may
impact different microbial communities in contrasting ways that could reveal new features of
functional diversity in forest soils.

In this study, we used a bioassay approach to assess the relative contributions of plant and
soil properties to spatial distribution patterns of ecto- (ECM) and ericoid (ERM) mycorrhizal
fungi as well as root-associated bacterial communities inhabiting the shared rhizosphere of
pine (ECM host) and lingonberry (ERM host). Soil systems were either untreated or treated
with PHCs to gain some understanding of how communities may change in response to PHC
contamination events. Using a combination of morphological and molecular methods to
describe patterns of community structure along with multivariate analysis techniques to
correlate soil properties, we aimed to contribute to the understanding of ecophysiological
functioning of this group of microorganisms that is ubiquitous on the northern forest
landscape.

188

Materials and methods
Field site
In September of 2006, forest soils were collected from the Kenneth Creek field site, located
about 100 km east of Prince George (53°34'N, 122°47'W) in the wet, cool subzone of the
sub-boreal spruce (SBSwkl) biogeoclimatic zone of central British Columbia. In 1982, the
forest was logged and burned, then subsequently planted with lodgepole pine {Pinus contorta
Dougl. Ex Loud. var. latifolia Engelm.); currently, the site is a mature, even-aged pine stand
with small hybrid white spruce {Picea glauca x engelmannii Parry ex Engelm.) and a few
western hemlock (Tsuga heterophylla (Raf.) Sarg.). Young subalpine fir {Abies lasiocarpa
(Hook.) Nutt.) and sitka alder (Alnus crispa var. sinuata (Reg.) Rydb.) are present at the edge
of the forest, along the access road; western redcedar {Thuja plicata Donn) and trembling
aspen {Populus tremuloides Michx.) are also present in an unlogged stand across the main
road. The site has a thick understory of oval-leaved blueberry {Vaccinium ovalifolium Sm.);
mosses and lichens (e.g. Peltigera) cover the forest floor, with some Lycopodium species.

The soils on this site were described by Arocena and Sanborn (1999). Soils are classified as
Eluviated Dystric Brunisols (Soil Classification Working Group, 1998), and consist of sandy
parent material with low clay content and few coarse fragments. The forest floor is mor
humus, from 2-5 cm thick with copious fungal mycelia present. The C:N ratio of the forest
floor is approximately 50 and the pH (water) is approximately 4.2. Gray Ae horizons are
generally 1-2 cm thick, with thicker pockets in some areas. Red Bf horizons extend to almost
30 cm, beneath which are Bm (27-60 cm), BC (60-100) and C (> 1 m) horizons. The C:N
ratios of the Ae and Bf horizons are about 20 and 13, respectively. The pH (water) of the Ae

189

horizon is 4.2 while the pH of the Bf horizon is about 4.8, increasing with depth to about 6.0
at the transition to the C horizon. Fine roots are found at depths greater than 1 m. Large
coarse woody debris (i.e. downed trees), the legacy of past forest management, is abundant
all over the site.

Bioassay, PHC treatment and sampling
Forest soil layers were reconstructed (i.e. Ae [~1 cm] and Bf [-15 cm] mineral soil layers
beneath organic [FH or CWD] soil layers [~2 cm]) in Cone-tainer™ pots (3.8x21 cm,
Stuewe and Sons, Corvalis, Oregon), as described previously. Lingonberry (Vaccinium vitisidaea L.) seedlings were transplanted into these soils and surface-sterilized seeds of
lodgepole pine {Pinus contorta var. latifolia), collected from the SBS and obtained from the
Ministry Tree Seed Center, Surrey, BC (seed lot DWD20050009A (location 079-B-008)),
were then planted into the organic layer of each pot. All pots were kept in the greenhouse
(22°C day temperature, 15°C night temperature, and 16 h photoperiod) and watered two or
three times per week for the duration of the experiment. The plants were fertilized once a
month (5 mL of NPK fertilizer; providing 100 ppm each of NPK) for the first four months
during seedling and mycorrhizal (ECM or ERM) establishment. At four months, 3 mL crude
oil (219 mg cm"2) was pipetted onto the organic soil surface of each pot, around, but not
touching, the seedling stem.

Plant - soil systems were destructively sampled at 1 and 16 weeks following PHC treatment.
Individual pots were emptied into trays, and the two plants (pine and lingonberry) were
gently removed from the soil with as little disturbance as possible to the three soil layers

190

(organic, Ae and Bf). Soil layers were collected from each pot, combined within treatment
groups (Table 4.1), and then air-dried for soil nutrient analysis and pH. PHC concentrations
in soil layers were also determined and are presented in Chapter 3. Root systems (with the
pine or lingonberry seedling excised) were shaken free from the soil and washed in sterile
dHaO. All root systems were examined under a dissecting microscope to ensure that they
were free of other roots or hyphae and soil particles; the distribution of ECM morphotypes
was also described for pine root systems. Root systems were then divided into three samples
(organic, Ae and Bf soil layers) and stored in 2 mL tubes at -20°C until DNA extraction.

Table 4.1: Plant, organic soil layer, and PHC treatment variables (n=3 for each combination).
Plant
Pine [P] + Lingonberry [L]

Organic Soil Layer
Forest floor [FH]
Coarse woody debris [CWD]

PHC
PHC
No PHC (control)

Soil Analysis
Organic (FH and CWD) and mineral soil (Ae and Bf) samples were analyzed for total C and
N content using <100-mesh samples (air-dried, then ground in a Model MM200 ball mill;
Retsch, Haan, Germany) by dry combustion using a Model 1500 NC Elemental Analyzer
(Fisons, Milan, Italy). The pH of organic soils was measured in a 1:2 soil to deionized H2O
suspension while 1:1 suspensions (in deionized water) were used for mineral soils (Kalra and
Maynard, 1991).

191

LH-PCR andfragment analysis
Frozen root systems were crushed in liquid nitrogen and DNA was extracted using a CTAB
(hexadecyl trimethyl ammonium bromide) protocol with an extra phenol/chloroform-isoamyl
alcohol (1:1) purification step (Fujimura et al., 2008). DNA extracts were further cleaned
using the Wizard® PCR Preps DNA Purification System kit (Promega) to remove phenolics
and other oily contaminants. These cleaned extracts were resuspended in TE buffer.

Fungal communities were characterized by amplicon length heterogeneity PCR (LH-PCR),
which provides an estimate of relative abundance of genotypes in a community (Martin and
Rygiewicz, 2005). For fungi, the ITS2 region of ribosomal DNA was amplified using the
forward primer ITS3 (5'GCATCGATGAAGAACGCAGC) (White et al, 1990) and the D3
fluorescent dye-labeled reverse primer NL4B (5'GGATTCTCACCCTCTATGAC) (Martin
and Rygiewicz, 2005). ITS3 is a universal primer that binds in a conserved domain 128 bp
from the 3' end of the 5.8 S rDNA whereas NL4B binds in the large subunit (28S) at
basidiomycete- (and ascomycete-) specific sites (Martin and Rygiewicz, 2005); PCR
products are expected to be vary from approximately 400 to >600 bp in length. PCR
reactions consisted of 10X PCR buffer, 2 inM dNTPs, 50 uM MgCl2, 10 uM forward and
reverse primers (Proligo, CO), 0.7 U Platinum Taq DNA polymerase (Invitrogen Life
Technologies), and nuclease-free water (Integrated DNA Technologies, Inc.) to a final
volume of 27 uL, to which 3 uL DNA (diluted 1:10) was added. The DNA Engine DYAD™
thermocycler (MJ Research, Inc., Watertown, MA) conditions were as follows: initial
denaturation for 4 min at 94°C, annealing for 1 min at 48°C, and extension for 2 min,
followed by 35 cycles of denaturation (94°C for 30 s), annealing (48°C for 30 s) and

192

extension (72°C for 1 min 30 s) and final extension at 72°C for 6 min 30 s. All PCR
products were run on 1.2% agarose gels to confirm amplification.

For bacteria, the D4 fluorescent dye-labeled forward primer 27F
(5'AGAGTTTGATCMTGGCTCAG) and unlabeled reverse primer 355R
(5'GTCGCCTCCCGTAGGAGT) were used to amplify 16S rDNA (Mills et al, 2003). PCR
reactions consisted of 3 uL DNA (diluted 1:50), 10X PCR buffer, 2 mM dNTPs, 50 uM
MgCl2, 10 uM forward and reverse primers (Proligo, CO), 0.7 U Platinum Taq DNA
polymerase (Invitrogen Life Technologies), and nuclease-free water (Integrated DNA
Technologies, Inc.) to a final volume of 30 uL. The DNA Engine DYAD™ thermocycler
(MJ Research, Inc., Watertown, MA) conditions were as follows: initial denaturation for 1
min at 94°C, 35 cycles of denaturation (94°C for 45 s), annealing (52°C for 45 s) and
extension (72°C for 1 min 30 s), and final extension at 72°C for 10 min. All PCR products
were run on 1.2% agarose gels to confirm amplification.

Fungal and bacterial samples were analyzed separately, but following the same procedure.
PCR products (2 uL) were loaded into a CEQ™ 8000 sequencer (Beckman-Coulter Inc.)
along with CEQ 600 bp (for fungal fragments) or CEQ 400 bp (for bacterial fragments) size
standard mixture. Run conditions were 60°C separation temperature, 4 kV voltage, and 120
min separation time. Analysis was performed using the amplicon fragment length
polymorphism (AFLP) program of the CEQ™ 8000 sequencer and the quartic (for fungi) or
cubic (for bacteria) model for size standard with the minimum relative peak height set at 1%
and a bin width of 1.5 bp.

193

The relative abundance of genotypes in each sample (i.e. fungal community) was calculated
by relativizing the fluorescent signal strength of each fragment peak to the total peak area
within each sample (Osborne et ah, 2006). Nonmetric Multidimensional Scaling (NMS) was
calculated on the basis of a S0rensen distance measure with 50 runs with real and
randomized data and a maximum of 500 iterations to assess stability (instability criterion was
0.00001) using PC-ORD 5.0 software (McCune and Mefford, 1999). A stepwise reduction in
dimensionality (6D-1D) was used to minimize stress along with a random starting
configuration (user-provided seeds). Stress (i.e. goodness of fit measure) and instability (i.e.
measure of change in stress at each iteration) were used to evaluate the structure of the
ordination results. The final solution for NMS was accepted after comparing 50 runs with
real to randomized data using Monte Carlo simulations (McCune and Grace, 2002). Pairwise
comparisons between groups were tested statistically with Multi-Response Permutation
Procedures (MRPP), a non-parametric method that provides a statistic of the magnitude of
differences between groups (i.e. effect size), given as the chance-corrected, within group
agreement (A) and a p-value (McCune and Grace, 2002).

194

Results
Soil properties
As expected, C and N contents were much greater in organic (FH, CWD) compared to
mineral (Ae, Bf) soil layers (Figure 4.1). In organic layers, mean C content in control (i.e. no
PHC treatment) FH soil layers was 12.1% compared to 50.4% in CWD layers. In PHC
treated soils, the amount of C increased to 21.6% in FH layers, but did not significantly
change in CWD soil layers, where it was 51.6%). In Ae and Bf layers, C content did not vary
in FH and CWD soil systems: levels for Ae and Bf layers were 1.6% and 1.2%, respectively,
in controls compared to 3.5% and 1.5% in PHC treated systems.

N content was 0.30% in control FH layers and 0.25%) in control CWD layers. In the PHC
treated FH layer, %>N increased to 0.45% whereas %N decreased to 0.18% in PHC treated
CWD layers. N content was 0.08% and 0.07% in Ae and Bf layers, respectively; these
values did not vary between soil systems or in PHC treated and control soils.

Soil pH (water) was generally lower in PHC treated compared to untreated soil systems, and
was not different in FH and CWD systems. In control soil systems, pH was about 5.8 in the
organic layer, 5.3 in the Ae layer, and 5.6 in the Bf layer. In PHC treated systems, pH was
5.3 in the organic layer, 5.0 in the Ae layer, and 5.6 in the Bf layer. There were no
differences in soil properties at 1 and 16 weeks.

195

ECM morphotypes
Four months after seed germination (i.e. week 0, time of the PHC treatment), virtually all
pine root tips were colonized with ECM fungi. ECM richness ranged from three to seven
(mean of five) ECM morphotypes per root system. In general, ECMs, extraradical mycelia,
and rhizomorphs were more extensively developed by the 16-week sampling time. The
density of root tips was greater in the organic layer of the soil profile, but most root tips
occurred in the B flayer (i.e. greater relative abundance). As shown in Figure 4.1, ECMs
were often associated with specific soil layers and varied in terms of relative abundance and
spatial distribution on different parts of the root systems.

Eight ECM morphotypes were frequently identified on root systems, including Cenococcum,
MRA, E-strain, Amphinema, two Russulaceae (including Lactarius), and two RhizopogonSuillus types. Rare ECM types were also observed, but were not included in the present
analysis. Morphological descriptions of pine ECMs are provided elsewhere (Appendix A).

Clusters of Cenococcum ECMs dominated the upper roots of pine in the FH layer, but were
never observed in the CWD layer of these double-plant systems. In PHC-treated systems,
Cenococcum morphotypes appeared to be less abundant after 16 weeks and root tips
appeared dry and less robust. Amphinema ECMs and external mycelia were often associated
with Cenococcum tips near the organic-Ae layer interface, but were also occasionally
observed in the Bf layer. The other ascomycetes, MRA and E-strain, were commonly found
throughout the soil profile, but in low relative abundance.

196

Two members of the Russulaceae (one Lactarius and one Russula type) dominated most root
systems (i.e. ECMs observed on 80-90% of root tips), particularly by the 16-week sampling
time; at least one ECM was identified as a species of Lactarius based on the presence of
laticifers. Root tips varied in colour from yellow to orange-brown and were generally
unbranched and smooth or appeared velvety due to the few short undamped hyphae that
resembled cystidia early in their development. Although mainly found in the Bf layer,
Russulaceae ECMs were also common on the upper lateral roots, particularly in the CWD
layer.

Two Rhizopogon-Suillus types were found associated with root tips exclusively in the Bf
layer. The first was a dark brown, coralloid morphotype. The second was white with a
slightly tuberculate form. By the 16-week sampling time, both ECMs had developed
extensive rhizomorph networks and hyphal fans that extended away from root tips into the Bf
soil.

197

Soil Properties

%C

%N

ECM Morphotypes

PH

# Fungal
Genotypes

# Bacterial
Genotypes

35

32

44

41

sfiiiiiL

organic layer
(FH or CWD)

19.1/51.0* 0.38/0.22* 5.6

2.6

0.08

5.2

33

23

45

41

1.4

0.07

5.6

34

33

42

45

Total 58

46

48

45

* FH/CWD organic soil layers
Total

Cen" - Cenococcum
Amp- - Amphinema
ftus- Russulaceae
Lac- Lactams
RhS1 - Rhizopogon-Suilius 1
RhS2 - Rhizopogon-Suilius 1
MRA
E - E-strain
' only in FH systems

Figure 4.1: Vertical distribution of soil properties (C, N, pH), ECM morphotypes (Cen,
Cenococcum; Amp, Amphinema; Lac, Lactarius; Rus, Russulaceae; Rh-Sl, RhizopogonSuilius 1; Rh-S2, Rhizopogon-Suilius 2; MRA; E-strain), and total number of ECM/ ERM
fungal and bacterial genotypes associated with pine (P) or lingonberry (L) in organic (FH or
CWD), Ae and Bf soil layers of a shared-rhizosphere system.

198

Fungal community

structure

The complete fungal dataset included 112 root samples and 75 amplicon fragments (400-690
bp in length). Multivariate analysis using nonmetric multidimensional scaling (NMS) gave a
three-dimensional solution with a final stress of 18.98 and instability of 0.08. Fungal
community structure varied between the 1-week and 16-week harvest times (p<0.001);
pairwise comparisons within PHC-treated and untreated (control) systems at 1 and 16 weeks
indicated that the changes in fungal community structure over time occurred in PHC-treated
(p<0.001) systems only. PHCs showed no other effects on fungal community structure.
Comparisons between fungal communities in different soil layers (organic, Ae, Bf) at 1 and
16 weeks showed significant differences (p=0.020) only between communities inhabiting the
organic layer.

Figure 4.2 shows the NMS ordination for fungal community structure within plant and soil
layer groups. Plant (i.e. pine or lingonberry) had a significant effect (p<0.001) on fungal
community structure. Fungal communities associated with pine roots differed (p<0.001)
from communities associated with lingonberry roots sharing the same soil system (i.e. with
overlapping rhizospheres). This distinction between pine and lingonberry fungal
communities is clearly visible along the 1st and 2 nd ordination axes (explained 39.9% of the
variation in the dataset) in Figure 4.2, where pine communities tended to group to the left,
and lingonberry to the right, of the ordination space. Comparisons (using MRPP) of pine and
lingonberry root communities by soil layer (organic, Ae, Bf) also revealed significant
differences (p<0.001) in community structure for all pairwise analyses.

199

•
#

Pine - org

H

Pine - Ae
Pine - Bf

"

Lingonberry- org
T

f

Lingonberry- Ae

m

jr

A • •
09

11

•

T

•

A

•

«

•.,

•

Lingonberry - Bf

ri

«

y
•

Al
*•

3
W
T

•
4 •

m

•

*"
*

•

A

•
• k"

T

A

Axis 1 (r2 = 21.4%)

Figure 4.2: NMS ordination of fungal community structure by plant (pine or lingonberry)
and organic (FH and CWD) and mineral (Ae, Bf) soil layers (stress = 18.98; instability =
0.08).

200

Soil properties also contributed to fungal community structure. Figure 4.3 shows the NMS
ordination for fungal community structure within the three soil layers (organic, Ae, Bf) of the
two soil (FH and CWD) groups along the 1st and 3rd ordination axes (explained 41.6% of the
variation in the dataset). The most visible distinction appears to be between the FH and
CWD systems, which tend to separate vertically. Within the layered soil systems, MRPP
analysis revealed significant differences (p=0.027) between communities in organic and Bf
layers, but not between Ae and either organic or Bf layers. Comparison of three soil layers
for FH-Ae-Bf and CWD-Ae-Bf systems showed significant differences (p<0.003) between
all combinations of the two systems, but no significant differences between layers within the
same type of system (i.e. overlap of fungal community structure in all 3 layers).

201

17

4
•4

FH-org
f

FH-Ae
FH-Bf
CWD-org
CWD-Ae

T

•

V

V

/

•

4

*

V

CWD-Bf

•jV

a

f

I

•

m

">?

-

#

.-.'.•
•
•

•

,:4

v

•
#

Axis l(i* = 21.4%)
Figure 4.3: NMS ordination of fungal community structure by soil layer in the two soil
systems (FH-Ae-Bf and CWD-Ae-Bf) (stress = 18.98; instability - 0.08).

202

Separate analyses of pine and lingonberry root systems were based on 61 root samples (58
DNA fragments ranging from 400-690 bp in length) and 51 root samples (46 DNA fragments
ranging from 400-630 bp), respectively. Within plant groups, significant differences
(p=0.002) in fungal communities were found only between the organic and Bf layers of pine
(Figure 4.4a). An even closer examination of pine-fungal communities in soil layers of FHAe-Bf and CWD-Ae-Bf systems showed that significant differences were associated with the
CWD systems and occurred between all layers: CWD-Ae (p=0.030), CWD-Bf (p=0.018),
and Ae-Bf (p=0.006) (Figure 4.4b). Comparisons of FH-Ae-Bf systems with CWD-Ae-Bf
systems showed significant differences in pine-associated fungal communities only between
the Ae layers of the two systems. For lingonberry, no differences were found between soil
layers, but significant differences in fungal community structure were found between all
layers of FH-Ae-Bf and CWD-Ae-Bf systems (Figure 4.4).

203

Pine (ECMJ

Ungonberry (ERM)
organic

^ organic

3)

•

^g

+ Ae

:

Bf

I Bf I

1 ptzQ.002

b)

CWD

FH

cwp
Ae

+4
•

•

4-

Ae I

Ae

Ae j

Bf

of

Bf "
Bf
' p<0,001

f

pmO.037
2 pwO.,030
3 pMQ.QQg

2

p<0.0O1

3 p=0.006

* pssQ.OfB

Figure 4.4: Pairwise comparisons of fungal community structure (genotypes) in pine (ECM)
and lingonberry (ERM) groups: a) within soil layers; b) within and between layers of FH and
CWD soil systems. Significant differences (MRPP) are represented by dark arrows and
corresponding p-values; light (slashed) arrows represent no differences between groups.

204

Both plant systems supported the trends associated with harvest time and PHCs that were
described for the complete fungal dataset. Differences in fungal community structure at 1
and 16 weeks post-PHC treatment were significant only in PHC-treated systems and there
were no significant differences between PHC-treated and control communities at either 1 or
16 weeks.

Bacterial community structure
The same pine and lingonberry root systems used to assess fungal community structure were
used for bacterial community structure analyses. This dataset consisted of 119 root samples
and 49 DNA fragments, ranging between 300 and 420 bp. Pine roots accounted for 61 root
samples (48 DNA fragments); lingonberry systems accounted for 58 root samples (45 DNA)
fragments. The PHC and harvest time variables had little effect on bacterial communities,
which appeared to follow the same trends in distribution as fungal communities.

NMS analysis resulted in a two-dimensional solution with a final stress of 15.96 and
instability of 0.06 (Figure 4.5). As with fungi, plant (pine and lingonberry) appeared to be
the dominant variable in structuring root-associated bacterial communities, but soil layer and
type of soil system (i.e. FH-Ae-Bf compared to CWD-Ae-Bf) were also important variables.
The figure shows clear separation of the pine from the lingonberry bacterial communities
along the two ordination axes (explained 83.7% of the dataset), and some distinction between
organic and mineral soil layers associated with each plant.

205

m

Pine-FH org

A

pjne-FH min

D

Pine-CWD,org
Pine-CWD m i n
Lingonberry- FH org
Lingonberry-FH min
Lingonberry- CWD org
Lingonberry- CWDmin

(ft

Axis 1 (r= = 51.2%)

Figure 4.5: NMS of bacterial community structure associated with plant (pine or
lingonberry) in organic (FH or CWD) and mineral soil layers (stress - 15.96; instability:
0.06).

206

Nested pairwise comparisons between bacterial communities of different plant and soil
treatment groups are summarized in Figure 4.6. In combined plant groups (P+L), MRPP
analysis revealed significant differences between organic and Bf layers (p=0.005), but not
between Ae and either organic or Bf layers (i.e. same as for fungi). In separate pine and
lingonberry analyses, Bf layers varied from both organic (p<0.001) and Ae (p=0.005) layers,
which did not differ from each other (Figure 4.6a). Bacterial community structure varied
significantly (p<0.001) between pine and lingonberry systems in each soil layer.

Overall, comparisons of three soil layers within FH-Ae-Bf and CWD-Ae-Bf systems showed
significant differences (p<0.001) in bacterial community structure between all combinations
of soil layers (Figure 4.6b). Within FH and CWD system types, significant differences were
found between organic-Bf and Ae-Bf layers of only the FH systems (p<0.001 for both).
Differences (p<0.001) in bacterial community structure in all soil layers were also observed
between FH-Ae-Bf and CWD-Ae-Bf systems. Separate analyses of pine and lingonberry
communities showed that the organic and Bf layers varied significantly in the CWD-Ae-Bf
systems of pine (p=0.030) and in the FH-Ae-Bf systems of lingonberry (p<0.001). Pine
CWD systems also differed between Ae and Bf layers (p=0.037). Differences (p<0.001) in
bacterial community structure between soil systems were significant only in lingonberry
systems.

207

a)

P+L

Pine

Lingonberry

organic

organic

*

[ Ae

T Ae

f

organic
Ae

i

Bf

Bf
' p=0.005
p<0.001

p=0.005

b)

FH

QA/D

FH

Bf
H

CWD

FH

CWD

Ae

4

'( t
Ae

Ae

Bf *""**

Bf

p<0.001

'Ae

Bf

1

Ae

V

Bf

Bf

1
p=0.037
i

Ae j

T

t

"Bf

1

p<0.001

p=0.030

Figure 4.6: Pairwise comparisons of bacterial community structure (genotypes) in all plant
systems (P+L), as well as in pine (ECM) and lingonberry (ERM) groups: a) within soil
layers; b) within and between layers of FH and CWD soil systems. Significant differences
(MRPP) are represented by dark arrows and corresponding p-values; light (slashed) arrows
represent no differences between groups.

208

Discussion
This study is the first to describe the spatial distribution patterns of microbial communities
from a shared rhizosphere perspective, encompassing fungal and bacterial communities
associated with pine (ECM host) and lingonberry (ERM host) roots interacting within the
same soils. Our results showed that plant and soil properties both contributed to community
structure of root-associated communities, but that community patterns varied among the
different guilds of microorganisms (i.e. ECM and ERM fungi and associated bacteria) at
different spatial scales within the mycorrhizosphere. Overall, PHC contamination had little
effect on the composition of root-associated microbial communities.

ECM and ERM root communities
Within our soil systems, fungal community structure differed distinctly between plants (pine
and lingonberry), reflecting the specificity of host root systems to form ECM or ERM
symbioses. It has long been presumed that ECM and ERM fungal communities were both
spatially and functionally separate (Smith and Read, 1997). We also found that bacterial
community structure varied between the different mycorrhizal root systems, indicating that
differences between the ECM and ERM root environment are also important with respect to
bacterial niche differentiation. Mycorrhizal interactions with heterotrophic bacterial
communities associated with the mantle and extraradical mycelia of ECM fungi and ERM
roots are important for mobilization, uptake and translocation of nutrients required for fungal
and, ultimately, plant growth (Burke and Cairney, 1998). As the establishment of ECM
symbiosis may alter root morphology (mantle and extraradical mycelia) and physiology (e.g.
membrane permeability, exudation patterns) in different ways than ERM symbiosis,

209

particularly with respect to quality and quantity of root exudates (Rygiewicz and Anderson,
1994), it is not surprising that mycorrhizal root-influenced differences in habitat supported
distinct bacterial communities.

High genotype richness (i.e. 58 and 46 DNA fragments, respectively for ECM and ERM
fungi, and 49 DNA fragments for root-associated bacteria) was observed for all three guilds
of microorganisms. Although high richness has been related to ecological resiliency, there is
no current sense of this threshold, particularly with respect to molecular assessment methods.
For ECMs, genotype richness was much greater than morphotype richness, which consisted
of eight common (and several rare) morphotypes. Discrepancies between morphological and
molecular analyses of ECMs have been reported in many studies and reflect the dynamic
character of mycorrhizal systems (Rosling et al, 2003). High genotype richness could be
due to the presence of inconspicuous or rare ECM fungi or other root-associated fungi such
as ascomycetous double colonizers (Rosling et al, 2003), saprotrophs (Lindahl et al, 2007)
or dark septate endophytes (Mandyam and Jumpponen, 2005) that may coexist with ECM
fungi on the roots. Their DNA was likely amplified along with the ECM fungal DNA
extracted from the washed root systems and included in the total genotype count following
LH-PCR. Secondary colonization (i.e. succession) of root tips already colonized by ECM
fungi also may have occurred; we noted a trend of increasing richness of ECM morphotypes
and genotypes over the 16 weeks of the study that was generally taken as a reflection of
community development with time on the young root systems. Succession in ECM root
colonization was also reported by Massicotte et al. (1999). In addition, an enhancement of
genotype richness could result from intraspecific variation within fungal taxa, which could

210

increase the number of DNA fragments based on variables characters of ITS rDNA that are
unrelated to functional diversity (Horton, 2002). The same factors that contributed to
genotype richness in ECM systems also likely contributed to enhanced richness in ERM
systems.

We also found substantial overlap in fragment lengths (genotypes) occurring in pine (ECM)
and lingonberry (ERM) communities, both for fungi and bacteria. For fungi, this could be
due to similarities in communities of non-mycorrhizal fungi (i.e. saprotrophs, dark septates,
etc.) associated with pine and lingonberry roots in the shared rhizosphere. These nonmycorrhizal fungi may not express host specificity to the same extent as ECM and ERM
fungi; communities may be structured more by the quantity rather than quality of available
substrates (i.e. exudates in the rhizosphere). Alternatively, the presence of shared fungal
genotypes in the two plant systems could be simply coincidental (i.e. as fragment lengths do
not reflect taxa in LH-PCR), or may indicate sharing of mycorrhizal symbionts between root
systems. In resynthesis experiments, Vralstad et al. (2002) showed that several strains of
ERM fungi comprising the Rhizoscyphus ericae aggregate formed true ECMs with conifer
(spruce and pine) and angiosperm (birch) species, although no isolates formed both ECMs
and ERMs. Villarreal-Ruiz et al. (2004) reported the ability of a fungus from the R. ericae
aggregate to form simultaneously both ECMs and ERMs in culture with Pinus sylvestris and
Vaccinium myrtillus seedlings; however, any sharing of mycorrhizal symbionts has yet to be
demonstrated in soil systems. Although we observed close interactions between pine and
lingonberry roots in the soils in this study, the resolution of the community fingerprinting

211

method (i.e. LH-PCR) was not sufficient to demonstrate sharing of mycorrhizal fungal
symbionts (i.e. only sharing of amplicon fragment lengths).

In studies relating microbial communities to ecological processes at larger spatial scales (e.g.
rhizosphere, stand, etc.), an understanding of spatial patterns of functional guilds of
microorganisms may be more important than the taxonomical details. Community
fingerprinting methods such as LH-PCR and terminal restriction fragment length
polymorphism (TRFLP) provide little taxonomic resolution of microbial communities
compared to methods such as sequencing, but are expected to provide sufficient resolution to
separate communities based on broad variables (Kuyper and Landeweert, 2002). Mills et al.
(2003) found no difference in resolution between LH-PCR and TRFLP in assessments of
PHC-degrading bacterial communities. The coarse filter approach used here to describe
patterns of root-associated communities in the shared rhizosphere potentially offers
ecologically relevant information at the expense of some fine-scale resolution.

Soil properties
Soil properties of the reconstructed systems used in this study generally resembled soil
properties at the field site. Exceptions included a slightly lower C:N ratio (about 40) in the
FH layer of the bioassay and less acidity in all three layers compared to the forest site. PHC
contamination altered soil properties by increasing C content, particularly in the organic (FH)
soil layers, and generally lowering pH in organic and Ae layers. Lower pH could be partly
due to production of metabolic acids and may reflect enhanced microbial activity in response
to addition of readily metabolizable C substrates present in PHCs. Lindahl et al. (2007)

212

found increased saprotroph activity in the vicinity of mycorrhizal root tips with increased C
availability. Nitrogen levels increased in FH layers, decreased in CWD layers, but remained
the same (i.e. much lower than organic soil) in both mineral soil layers. Due to the high
porosity in organic soil layers, PHCs, particularly more hydrophobic compounds, tend to be
retained in these layers as compared to mineral soils. Hydrophobic compounds also have a
greater tendency to adsorb to organic soil components than to mineral ones (Xing et al.,
1994). Thus, microbial communities in organic layers must inhabit PHC-contaminated
habitat to a greater extent than communities inhabiting the mineral layers below, which
consisted mainly of sand particles with few binding sites for the PHC chemicals that
eventually made their way down the profile along the roots.

However, changes to soil properties related to PHC contamination appeared to have little
effect on community composition of root-associated microorganisms (fungi and bacteria), at
least within the 16-week duration of the experiment. Setala et al. (2000) found that soil
microbial communities appeared to exhibit high resilience to environmental disturbances
when soil organic layers (e.g. humus, woody debris, etc.) were not severely disrupted. The
mycorrhizosphere may offer some physical protection from potential toxic effects (e.g.
solvent shock) of PHC treatment, surfaces for biofilm formation (themselves, protective
structures), as well as a steady supply of C substrates supporting ongoing microbial
metabolism. The extent to which these protective factors extend into the rhizosphere remains
unknown and was not investigated in this study.

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In the shared rhizosphere systems studied here, soil properties exerted two kinds of effects on
microbial community structure: contrasting properties of the FH-Ae-Bf and CWD-Ae-Bf soil
systems, which are potentially important during (mycor)rhizosphere development, and
vertical segregation in the soil profile (reflection of broad changes in physical soil
properties).

FH and CWD systems
The planting of pine seeds into FH and CWD organic layers provided opportunities for
potentially different indigenous microorganisms to colonize the newly germinating roots in
these two systems. The greater C:N ratio of the CWD layer could potentially enhance
different communities of ECM fungi as well as saprotrophic communities. However,
differences between root-associated communities (ECM fungi and bacteria) inhabiting the
different soil layers were generally not observed in pine systems, except in Ae layer
comparisons. This difference may be due to percolation of different metabolic products from
CWD decomposition (i.e. compared to FH) downward in the soil profile, which could
influence microbial communities in the Ae layer below.

Lingonberry communities appeared to vary more between soil systems (i.e. FH-Ae-Bf and
CWD-Ae-Bf) than between soil layers; the same trend was observed for both fungal and
bacterial communities associated with lingonberry roots. It is possible that the soil habitat in
and below CWD was different in ways (that we did not measure) from FH systems that
supported different groups of microorganisms associated with lingonberry roots. Unlike
pine, lingonberry cuttings (rooted in reconstructed forest soils) were transplanted into the soil

214

systems, which may have influenced fungal and bacterial colonization in unknown ways (e.g.
introduced some competition). Further interpretation of this finding is difficult in light of the
current limited understanding of ERM fungi.

Vertical segregation
ECM fungal communities exhibited vertical community structure, reflecting the direct
interactions between fungi and soil in the ectomycorrhizal association. In general, ECM
fungal communities in organic soil layers were distinctly different from Bf-layer, but not Aelayer, communities. In our soil systems, the boundaries above and below the Ae layer were
not well defined and ECM fungal communities often overlapped in these zones. Rosling et
al. (2003) found that ECM community composition varied between organic, eluvial and
deeper mineral horizons of a Swedish podzol, and that most taxa described from the organic
layer were also found in the eluvial horizon below. Genney et al. (2006) found that the
layered nature of the soil substrate was more likely to encourage horizontal rather than
vertical extension of mycelia and that ECMs and extraradical mycelia often occupied
adjacent layers of organic and upper mineral soils. A comparison of FH and CWD soil
systems revealed that differences between soil layers that influenced community structure
were mainly attributable to CWD systems. The overlap in fungal communities between
layers again reflects the continuity and dynamics of ECM systems.

In contrast, ERM fungal communities of lingonberry did not vary in the different soil layers.
Due to their endomycorrhizal nature, ERM fungal communities may be less influenced by
differences in soil properties compared to ECM fungi that are in direct contact with soil on

215

the outer root surfaces and extraradical mycelia. In addition, compared to ECM fungi, there
are relatively few ERM fungi, which may occupy fewer niches. As epidermal root cells
appear to be composite structures that are colonized by a variety of fungal symbionts, each
cell potentially functions as a separate unit (Berch et al, 2002; Perotto et al., 2002). Thus,
variation in ERM community structure may occur at the level of individual hair roots rather
than the level of the whole root system (i.e. rhizosphere), as for ECM communities.

Bacterial communities that inhabit the outer surfaces of the ECM mantle and mycelia, as well
as the ERM roots, also directly interact with the soil environment, and were found to exhibit
complex patterns of vertical segregation similar to ECM fungal community patterns.
Depending on the resolution of analysis, bacterial community structure was found to vary
between Bf and organic and Bf and Ae layers. In pine systems, differences were associated
with CWD systems, whereas in lingonberry systems, differences were associated with FH
systems. Changes in soil chemical, mineralogical, and structural properties with depth create
complex and variable sets of microbial habitats over very small distances, to which freeliving bacterial cells may be particularly sensitive (Genney et ah, 2006). Due to their small
size and structural vulnerability to environmental stresses, bacterial communities may be
expected to vary along much smaller spatial scales compared to the fungal communities with
which they are associated.

The vertical distribution patterns of ECM morphotypes described on the root systems of pine
seedlings in this study were consistent with those described from field-based studies in
European forests (Rosling et al, 2003; Tedersoo et al., 2003; Baier et al, 2006; Genney et

216

ah, 2006; Lindahl et al., 2007). In our soil systems, the density of ECM root tips was greater
in organic compared to mineral soil layers, but this was not related to either morphotype or
genotype richness (genotype richness was also not different between soil layers for ERM
fungi and bacteria). Most of the root system (i.e. much higher abundance of mycorrhizal
roots) occurred in mineral (i.e. Bf layer) soil layers. Some ECMs {Cenococcum, Amphinema,
Russulaceae and Rhizopogon-Suillus types) dominated certain parts of the soil profile, while
other ECMs (MRA and E-strain) occurred throughout the profile, usually at low relative
abundance. These community patterns are not always revealed from studies using soil cores
where a fairly arbitrary and shallow sample of the belowground habitat is taken. Our
rhizosphere approach had the advantage of sampling entire root systems (albeit small and
young), which represent ecological units on the landscape. For seedlings, at least, this
approach provided a more precise picture of the abundance component of diversity, which
may be important in terms of ecological functions.

Similar ECM partitioning patterns in temperate forests (USA) were attributed to ECM
specificity (or generality) for resource use in the different soil layers (Dickie et ah, 2002).
For example, soil organic matter (SOM) is an important source of substrates for
decomposition and nutrient cycling in forest ecosystems (Read and Perez-Moreno, 2003).
The prevalence of ECM fungi in well-degraded litter and humus layers of organic soils
supports the hypothesis that they play a significant role in mobilizing N from the more
recalcitrant organic matter in boreal forest soils (Lindahl et al., 2007). Linkages between
ECM community structure and function have been inferred using ECM exploration types (i.e.
groups of ECM morphotypes defined by the amount, organization and extent of the

217

extraradical mycelia) as the units for analysis (Agerer, 2001). Patterns are thought to indicate
differential resource utilization, which suggests that ECMs have distinct foraging strategies
and different capacities for resource acquisition (Agerer, 2001; Baier et al., 2006). The
relative abundance of hyphae of various ECMs may strongly influence host and ecosystem
function (Dickie et ah, 2002).

In our study, forest floor (FH) layers were dominated by Cenococcum, which has often been
associated with high humus content and a large C:N ratio (Rosling et al, 2003; Baier et al.,
2006). Cenococcum is a short distance exploration type (Agerer, 2001) characterized by a
dense cover of emanating hyphae, suitable for making multiple contacts with the loose
organic cover of the forest floor (Baier et al., 2006). Although Cenococcum tends to be
tolerant of changing environmental conditions (e.g. temperature, moisture) typical of organic
soil layers in forests, some morphotypes in the current study appeared dry, less robust, and
less abundant after 16 weeks in the PHC treated systems compared to untreated controls
(Chapter 2). Cenococcum is often associated with Amphinema, which tends to inhabit the
more environmentally stable interface between the organic and Ae layers (Rosling et ah,
2003; Baier et al, 2006). Amphinema (as well as Piloderma, which did not colonize root tips
in this study, but whose distinctive yellow hyphae were frequently observed in organic soil
layers) has been described as medium-distance (fringe) exploration type characterized by
fans of emanating hyphae with extended contact to the soil, as well as rhizomorphs that
ramify and interconnect repeatedly (Agerer, 2001).

218

Whereas Cenococcum dominated the FH layer, it was consistently absent from the CWD
layer, which was characterized by a greater C:N ratio due to the prevalence of lignified
substrates as well as other physical differences. Instead, the two common Russulaceae
morphotypes tended to dominate root systems in CWD (dead wood <2.5 cm diameter).
Lactarius (and other Russulaceae) ECMs are typically associated with acidic, low-N soils,
and may also contribute directly to decomposition via oxidative enzyme activity (Lilleskov et
ah, 2002). CWD is also an important habitat for resupinate thelephoroid and athelioid fungi,
as well as Sebacinaceae (Tedersoo et ah, 2003). We found that thelephoroid ECMs
represented a rare component of the ECM fungi described in this study; these fungi tend to be
inconspicuous (and also may not establish with young seedlings), but they likely contributed
to genotype richness from roots in CWD.

Pine roots in mineral soils were frequently dominated by the Russulaceae ECMs, which often
accounted for more than three quarters of the total number of roots tips per seedling. These
contact exploration types had smooth mantles with few emanating hyphae and were well
equipped to explore dense soil horizons with narrow pores (Agerer, 2001; Baier et ah, 2006).
They also tended to have preference for higher bulk density of mineral compared to organic
soil layers (Genney et ah, 2006).

The two Rhizopogon-Suillus ECMs described in this study were exclusively found in the Bf
layer. Though not usually dominant on individual root systems, extensive extraradical
mycelia and rhizomorphs of these ECMs likely accounted for a high proportion of biomass.
Suillus and Paxillus morphotypes have been studied extensively in the lab (including with

219

respect to PHC biodegradation), but they have not been frequently described from field
studies (Genney et al., 2006). The lack of field observations may be an artifact of the
tendency to focus studies on upper soil layers only, missing yet another functional
component of the system; Rosling et al. (2003) found Suillus luteus in mineral layers in 2 (of
3) 52-cm columns of soil. Both Suillus and Rhizopogon have long-distance exploration
strategies and produce highly visible thick rhizomorphs from tuberculate ECM tips (Agerer,
2001). Proliferation of hyphal fans towards suitable metabolic substrates in microcosm soils
has been reported by Bending and Read (1997); in this study, we frequently observed hyphal
fans extending into both PHC treated and control mineral soils. Fungal species present in
small numbers as mycorrhizas but that form extensive extraradical mycelia may be
functionally very important for C and N cycling (Genney et al, 2006).

Conclusions
Mycorrhizas, along with the heterotrophic microorganisms tightly associated with the
mycorrhizosphere, occupy the structural and functional interface for carbon and nutrient
cycling between the aboveground and belowground food webs in northern forests.
Interruptions in community functions through environmental disturbance such as PHC
contamination could potentially disrupt essential ecosystem processes at a landscape scale.
For the guilds of microorganisms at this trophic level (i.e. ECM fungi, ERM fungi and
bacteria), PHCs did not appear to exert many detrimental impacts. However, it is unknown
whether ongoing community development will occur along the same trajectory over the
longer term in PHC contaminated systems compared to uncontaminated systems or whether
this is functionally meaningful in terms of ecosystem processes. Furthermore, detrimental

220

impacts to microbial groups that are spatially abundant or possess more specialized functions
could lead to community changes over time, so longer term studies are warranted. Analysis
of variation in community structure at the rhizosphere scale gives a broad sense of
community function by relating changes to habitat properties. Our finding that community
patterns varied with both plant and soil characteristics and at different scales for the three
groups assessed contributes to the greater understanding of forest soil ecology and
sustainability.

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Chapter 5: Conclusions and future considerations for microbial ecology and
sustainability management in PHC-contaminated northern forest ecosystems
This research was based on the premise that oil spills occur in boreal forest ecosystems with
unknown scope and frequency. In addition, the fate and ecological impacts of petroleum
hydrocarbon (PHC) contaminants in forest soils are not well understood from either ecotoxicological or microbial ecological perspectives. Mycorrhizal systems (i.e. plant-fungal
symbioses and associated heterotrophic bacterial communities) represent the dominant
microbial biomass in northern forest soils and may be most vulnerable to the environmental
impacts of direct PHC exposure. Conservation of the integrity of mycorrhizal systems in
contaminated forests may be key for management for both ecological resilience and
remediation in a sustainability context.

The purpose of this research was to examine interactions between PHCs and mycorrhizal
communities in the rhizosphere, where different guilds of microorganisms interact within the
larger trophic group tightly linked to decomposition, carbon and nutrient cycling, and
primary production. This work was both experimental and exploratory in nature, as
hypotheses were both tested and generated. Several methodological approaches were
simultaneously used to gain a better understanding of the physical, chemical and biological
changes that occurred in the rhizosphere of sub-boreal forest soils following contamination
with ecologically relevant (i.e. equivalent to several tonnes per hectare) levels of oil. Insights
along a number of themes are presented in the following paragraphs.

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Rhizosphere model for mycorrhizal systems
Our model system was comprised of the root system of a single plant (pine, birch or
lingonberry), germinated (or transplanted, in the case of lingonberry) into verticallystructured, field-collected, sub-boreal forest soils. These plant-soil systems attempted to
incorporate some of the complexities of soil and microbial community structure in the
rhizosphere, but within the confines of the bioassay (i.e. limited dispersal, etc.). These
systems captured a sub-set of the soil microbial community diversity expected in situ, which
included ecto- (ECM) and ericoid (ERM) mycorrhizal fungal propagules capable of forming
symbioses with different root systems in different soil types, under greenhouse conditions.
Incorporation of soil vertical structure and depth expanded microbial habitat and niche, and
revealed features of ectomycorrhizal (ECM) fungal diversity (e.g. extensive extraradical
mycelia of the relatively abundant Rhizopogon-Suillus 1 morphotype) that have generally
been overlooked in the deeper mineral soils.

The rhizosphere represents a discrete functional unit that repeats over northern forest
landscapes. A single rhizosphere obviously does not represent the full complexity of the
forest soil environment, where dynamic interactions between many different types of
rhizospheres exhibit emergent properties (i.e. the community possesses capabilities that its
individuals lack) (Allen et ah, 2003). However, concurrent assessment of plant, soil and
microbial community properties at the rhizosphere level had the advantage of allowing some
extrapolation to broad spatial scales over landscapes exhibiting high levels of environmental
heterogeneity, a necessity for application to sustainable land management. At the same time,
details of microbial communities at greater levels of taxonomic, genetic, or physiological

226

resolution could be assessed at the level of the individual root tip. Thus, the rhizosphere
scale is intermediate to species-based approaches for understanding functional diversity and
ecophysiology on smaller scales, and approaches for assessing spatial distribution and
ecological functions on larger scales. In this baseline study, we focused on structure,
distribution, and broad ecological functions of microbial communities at the rhizosphere
scale.

A characteristic of more ecologically inclusive studies is greater variation within treatment
groups, which can impair the ability to detect significant differences between them. Here, we
used several methodological approaches and multivariate statistical analyses to assess
changes in different system components and to look for consistent trends potentially pointing
to biological significance. The remaining unexplained variation is likely the result of noise
from unmeasured environmental and spatial variability (Ramette and Tiedje, 2007).

Ecological integrity and resilience
Environmental disturbance provides a means for understanding forest stability/ resilience
arising from complex interactions within ecosystems (Amaranthus, 1998). We assessed
environmental changes associated with soil fertility and physical structure that were expected
to impact community structure and function of microorganisms and higher trophic-level
organisms, hi our systems, surface contamination with crude oil led to increased
concentrations of potentially toxic PHC chemicals ranging from <nC10 to >nC50 in size
(GC-FID analysis). Soil C analysis confirmed that C content increased in PHC treated
compared to untreated (control) soils, whereas N content remained unchanged. Thus, PHCs

227

may potentially inhibit some microorganisms (due to toxicity) and stimulate others (due to C
availability). The more acidic pH that was generally associated with PHC treated soils
supports the hypothesis that microbial activity was enhanced, although this was not explicitly
tested. Although we found no differences in community structure (based on relative
abundance of microbial components, not just richness) between PHC treated and control
systems, this may be a reflection of the relatively low-resolution method used (i.e. LH-PCR),
relative simplicity of the plant-soil (rhizosphere) systems, and relatively short period of study
(16 weeks). Clearly there is a need for future field-based studies incorporating older and
deeper root systems, different types of plant hosts and soil types, and on greater scales of
space and time to more fully understand these interactions.

Mycorrhizal communities were generally resilient to ecologically relevant levels of PHC
contamination (i.e. equivalent to -7-22 tonnes per hectare, which is within the potential range
of contamination levels expected from a burst pipeline, etc.) when the integrity of the system
was conserved (i.e. no further disturbance to mycorrhizal roots or soil structure). In intact
systems, most PHCs were retained in the organic soil layers (GC-FID analysis), which may
have limited PHC exposure of mycorrhizas and associated heterotrophic communities at
greater depths in the soil profile. Although microbial communities inhabiting the organic
layers were exposed to far greater concentrations of PHCs than deeper communities,
morphological examinations of individual ECMs and community fingerprinting (LH-PCR)
studies revealed few changes in community structure (based on relative abundance of the
fungal and bacterial components of each rhizosphere community) attributable to PHC
contamination. We hypothesize that the mycorrhizosphere may also have served a protective

228

function for both mycorrhizal fungi and associated heterotrophic communities, possibly
through exclusion of potentially toxic PHCs (i.e. due to properties of fungal cell walls,
biofilm formation, and solvent flow along the mycelial surface). However, it was difficult to
separate the effects of chemical toxicity from habitat changes (e.g. altered carbon/ nutrient
supply, pH, water holding capacity, etc.) in PHC-contaminated soil systems.

Morphological and DNA-based analyses do not directly address questions concerning PHC
impacts on ecological functions because they reveal little of what the organisms are doing.
Using biochemical (enzyme-based) methods to assess C metabolism, we found that PHC
treatment had no effect on ECM laccase activity (for catalyzing the opening of aromatic ring
structures) or on bacterial community C use profiles for substrates commonly found in the
rhizosphere. More detailed functional analyses are needed to better understand relationships
between PHC contamination and microbial functions, but these coarse-filter findings also
point to general system resilience.

PHC biodegradation and functional redundancy
PHC concentrations (GC-FID analysis) generally decreased in all plant-soil systems over 16
weeks, indicating an inherent capacity for PHC biodegradation. Biodegradation was
particularly evident in pine and birch (ECM) systems that were grown in soils with FH and
CWD organic layers providing the initial microbial inoculum. Both the forest floor (FH) and
coarse woody debris (CWD) organic layers were expected to contain microorganisms well
adapted for decomposition/ biodegradation of C substrates typically found in these soils. The
structure of ECM fungal and bacterial communities varied between these plant-soil systems,

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indicating functional redundancy for PHC biodegradation. Standing et al. (2007) suggested
that functional redundancy among widespread soil processes such as decomposition/
biodegradation may be so large that relationships between microbial diversity and ecosystem
function may lose relevance at larger spatial scales. Our results indicated redundancy for C
substrate metabolism for individual ECMs (laccase assay) as well as for bacterial
communities (CLPP analysis). Insights into functional heterogeneity in biodegradation
processes (i.e. specific metabolic reactions along catabolic pathways) within these
communities require higher levels of resolution than offered in this study and should be the
focus of future research.

ECMs appear to enhance biodegradation via at least two mechanisms. Our results showed
that greater densities of microorganisms capable of PHC biodegradation were supported in
the mycorrhizosphere compared to non-rhizosphere soil (i.e. mycorrhizosphere effect). This
large heterotrophic biomass is likely fueled through addition of reduced C substrates in PHC
mixtures, resulting in a generally stimulatory effect on biodegradation. In unplanted soils,
we found greater bacterial densities in PHC-treated compared to untreated (control) soils,
indicating that availability of readily metabolizable C substrates is at least one factor
influencing the density of microbial communities in the mycorrhizosphere. In addition, we
observed that the ECM extraradical mycelia greatly expanded the surface area for potential
bacterial colonization and biofilm formation, which could also enhance PHC biodegradation
through expansion of habitat and niche of biodegrading communities.

230

In the absence of mycorrhizosphere (i.e. in unplanted systems), levels of the smaller (<nC16)
PHC chemicals were significantly reduced by indigenous bacterial communities, but levels of
the larger (nC16+) PHCs did not change substantially. This provides indirect evidence for a
stimulatory role of ECMs in the biodegradation process. More direct evidence was inferred
from our finding that many ECMs exhibited the potential to open aromatic rings via laccase
secretion, thus contributing to the metabolic synergy for PHC biodegradation that appears to
exist within the mycorrhizosphere community. Laccase secretion by ECMs is one of many
potential functions that may impact PHC biodegradation in forest soils for which we have no
clear understanding with respect to ecological significance. Many of the dominant ECMs
(e.g. Rhizopogon-Suillus 1, Russulaceae 1, and Lactarius) described in our study exhibited
high laccase activity, both at the root tip and along the extraradical mycelia/ rhizomorphs.
These morphotypes tended to inhabit the deeper, mineral soil layers and have not been well
studied. The dominant ECMs may represent the most functionally relevant community
component (i.e. greater biomass and greater contribution to nutrient uptake and C demand),
with the numerous minor (less frequently occurring) ECMs representing functional
equivalents of the dominant types, but with different environmental requirements and
tolerances (Allen et ah, 2003). Thus, the minor types may contribute to ecosystem resilience
in changing environmental conditions (i.e. disturbance) that are capable of replacing the
dominant ECMs should they decline (i.e. insurance effect). This hypothesis remains to be
specifically tested for mycorrhizal fungal communities, particularly with respect to
contaminated soil management.

231

We did not find significant PHC biodegradation in the systems where additional complexity
was incorporated (i.e. double-plant systems with ECM and ERM hosts or FHoil soil systems
that contained residual PHCs from the previous contamination event) into the experimental
design. The additional complexity may have overwhelmed the spatial and temporal
limitations of the bioassay, resulting in greater within-group variation. In the double-plant
systems, we found greater diversity (richness and relative abundance) of fungal and bacterial
communities than in single-plant systems, and each root system supported both shared and
distinct components of microbial communities. We hypothesized that double-plant systems
augmented the level of competitive interactions (perhaps by providing more specialized
niches owing to properties of the different root systems) compared to single-plant systems,
which may have inhibited the overall biodegradation process at key steps along the
biodegradation pathway. In the FHoil soils, residual PHCs probably consisted of a greater
proportion of larger (>C16) PHCs that may be degraded more slowly and be less available to
microbial metabolic systems. In terms of biodegradation potential based on community
profiles, FHoil soil system fungal and bacterial communities varied no more from FH and
CWD systems than these latter systems varied from one another.

Plants and soils
Soil microbial communities exhibit clear spatial (and temporal) patterns across landscapes
based on environmental heterogeneity (niche) and the legacy of past disturbance events
(Ramette and Tiedje, 2007). In this study, we found that the plant was the primary
determinant of mycorrhizal fungal and associated bacterial community structure. Both
morphological examinations of individual roots and community fingerprinting (LH-PCR)

232

studies revealed distinct fungal communities for pine, birch and lingonberry systems. This
reflects the specificity of host root systems to form ECM or ERM symbioses, which were
expected to be both spatially and functionally separate. This distinction between ECM and
ERM fungal communities was maintained in the double-plant systems where the pine and
lingonberry root systems were in close proximity. In addition, ECM fungal communities
exhibited specificities for either the coniferous (pine) or deciduous (birch) host, which may
have been due to differences in root structure or exudation patterns. Surprisingly, bacterial
community structure also varied between the different mycorrhizal root systems, indicating
that differences between ECM and ERM root environments are also important with respect to
bacterial niche differentiation. Differences in microbial community structure between plant
systems reflect the many potential plant-fungal combinations that occur in repeatable units
across landscapes (Allen et ah, 2003). How the diversity of the plant community impacts
microbial community structure, ecosystem resilience and biodegradation capacity following
PHC contamination is an important consideration in both ecological and management
contexts.

Differing properties of the soil environment exerted two kinds of effects on microbial
community structure. First, communities (morphology and LH-PCR) varied between the
three soil systems tested, which were characterized by different organic layers (FH, CWD,
and FHoil) providing the initial microbial inoculum during early plant growth and
(mycor)rhizosphere development. Second, communities exhibited vertical segregation in the
soil profile (organic, Ae, and Bf layers), which is a reflection of broad changes in physical
and chemical soil properties. Results from the shared rhizosphere study (i.e. double-plant

233

systems with both pine and lingonberry) showed that community patterns varied among the
different guilds of microorganisms (i.e. ECM and ERM fungi and associated bacteria) at
different spatial scales within the mycorrhizosphere and were related to certain
environmental variables. ECM fungal communities, which are in direct contact with soil on
the outer root surfaces and extraradical mycelia, were found to vary between different soil
layers within systems, as well as between the three organic soil layers of different systems.
In contrast, ERM communities appeared to be less influenced by vertical differences in soil
properties, but to depend more on properties of the different organic soil systems, which may
influence the initial microbial inoculum composition. Variation in ERM community
structure may occur at the level of individual hair roots consisting of epidermal cells
potentially functioning as separate units rather than the level of the whole root system (i.e.
rhizosphere), as for ECM communities. Bacterial communities varied both within and
between soil systems. Due to their small size and structural vulnerability to environmental
stresses, bacterial communities may be expected to vary along much smaller spatial scales
compared to the fungal communities with which they are associated. These findings help
form the base for a better understanding of connections, linkages, and scale-related relevance
of mycorrhizosphere/ ecosystem components that contribute to ecological processes at
landscape scales and to predict how ecosystems may respond to environmental disturbances
such as soil contamination. However, the identification of common patterns of biotic and
environmental variation remains a central issue in microbial ecology.

234

Management in PHC contaminated forests
Currently, there is little ecological foundation for remediation strategies in PHCcontaminated forest soils. Rather, remediation depends on contaminant (e.g. PAH) levels
and perceived toxicological threats to soil quality, forest productivity, and wildlife,
intertwined with the economic incentive of recovering spilled oil. There is no evidence that a
single spill event in a forest poses significant ecological threats; studies suggest that over
time scales of several years, many PHCs are biodegraded via central metabolic pathways of
the indigenous soil microbiota or are transformed via specific enzymatic pathways to
products that are incorporated into the soil organic matter. Our results suggest that surface
application of relatively high rates of crude oil to plant-soil systems had little negative impact
on mycorrhizal fungal or bacterial communities in the rhizosphere, and also showed that
significant PHC biodegradation occurred within 16 weeks of treatment.

Sustainable management strategies for northern forests need to preserve diverse
environmental, social, and economic values for large areas over long periods of time.
Mycorrhizal systems consist of functional groups of soil biota that underpin ecosystem
services such as decomposition, biogeochemical cycling, primary production, soil stability,
and C sequestration, to name a few (Barrios, 2007). Thus, maintenance of mycorrhizal
integrity, which includes both the plant community and soil environment, should be an
objective of remediation strategies for PHC contaminated soils. Ideally, remediation should
utilize knowledge of the microbial communities present in contaminated soil environments,
their metabolic abilities, and their likely response to changes in environmental conditions.

235

In situ bioremediation is considered a low-disturbance and cost-effective strategy for
managing contaminated forest soils on large scales. Our results support the hypothesis that
the intrinsic capacity for biodegradation is maintained in PHC contaminated soils in the
absence of further environmental disturbance. This intrinsic biodegradation could be
enhanced through various augmentations to the process. For example, addition of organic or
woody material to the forest floor may extend the habitat of mycorrhizal systems (i.e.
increase mycorrhizosphere volume) and provide energy-rich substrates for co-metabolism of
the more recalcitrant PHC chemicals. Addition of N fertilizer has been used to relieve soils
of the N deficiency that is often associated with PHC contamination (not found in our study);
however, fertilization may also lead to conditions where mycorrhizal symbioses become too
costly for the host plant to maintain (i.e. plants no longer require N from the mycobiont),
which could lead to loss of vital functional components of mycorrhizal communities. In
addition, the planting of mycorrhizal seedlings establishes mycelial systems in the
contaminated soils and allows gradual decontamination over time. Our results suggest that
the greatest success may come from using indigenous species of host plants (grown in site
soils) for initially establishing adapted mycorrhizal communities rather than inoculating
plants grown in nursery soil mixtures with non-site-specific fungi.

In conclusion, this systems approach addressed fundamental questions in mycorrhizal
ecology by considering PHC pollution as a form of environmental disturbance. Further
research is needed for a more detailed understanding of interactions between PHCs and
mycorrhizal systems in northern forest soils.

236

References
Allen, M.F., Swenson, W., Querejeta, J.I., Egerton-Warburton, L.M., and Treseder, K.K.
(2003) Ecology of mycorrhizae: a conceptual framework for complex interactions among
plants and fungi. Annual Review of Phytopathology 41: 271-303.
Amaranthus, M.P. (1998) The importance and conservation of ectomycorrhizal fungal
diversity in forest ecosystems: lessons from Europe and the Pacific Northwest. Portland,
OR, U.S. Department of Agriculture, Forest Service, Pacific Northwest Research Station,
Gen. Tech. Rep. PNW-GTR-431. 15 pp.
Barrios, E. (2007) Soil biota, ecosystem services and land productivity. Ecological
Economics 64: 269-285.
Ramette, A. and Tiedje, J.M. (2007) Multiscale responses of microbial life to spatial distance
and environmental heterogeneity in a patchy ecosystem. Proceedings of the National
Academy of Science 104: 2761-2766.
Standing, D., Baggs, E.M., Wattenbach, M., Smith, P., and Killham, K. (2007) Meeting the
challenge of scaling up processes in the plant - soil - microbe system. Biology and
Fertility of Soils 44: 245-257.

237

OM/IM net to
interlocking irregular
synenchyma, small cell
width (1-3 um), Hartig
net present
OM non-interlocking
irregular syenchyma, IM
net synenchyma, variable
cell width (4-8 um),
laticifers, Hartig net
present

(not observed)

frosty orange-brown,
smooth, grainy or velvety,
matte, unbranched or
dichotomous, straight or
bent (1-3 mm)
yellow-orange, smooth to
velvety, matte, unbranched
or dichotomous, straight to
bent (1-3 mm)

Piloderma

Russulaceae 1

Russulaceae 2
(Lactarius)

238

(not observed)

pale yellow, cottony, matte,
dichotomous to irregular
branching

Amphinema

very few short hyaline to
yellow cystidia (2-3 um),
slightly tapered out, no basal
clamp, 100 um long, septate, no
clamps, verrucose or no
ornamentation

hyaline hyphae (2-3 urn),
septate, clamps, verrucose
ornamentation with needle-like
crystals
very few short hyaline hyphae
(2-3 um), septate, no
(sometimes?) clamps, no
ornamentation

hyaline hyphae (2-3 um),,
septate, clamps, verrucose
ornamentation

few hyaline hyphae (5-8 um),
septate, no clamps, verucose
ornamentation

OM/IM net to noninterlocking irregular
synenchyma, very thin,
large cells (5-8 urn),
Hartig net present
OM net synenchyma, cell
width 3-4 um, Hartig net
present

E-strain

MRA

OM regular synenchyma brown-black hyphae (4-5 um),
sometimes copious, septate, no
(stellate pattern), Hartig
clamps, no ornamentation
net present
black or hyaline hyphae (2-3
OM/IM felt
prosenchyma, thin, Hartig um), septate, no clamps,
verrucose ornamentation
net present

black, wooly to grainy,
matte to shiny, unbranched,
straight (1-2 mm)
black (hyaline at root tip),
felty to wooly, matte,
unbranched, straight (1-1.5
mm)
orange-brown to dark
brown (hyaline tip), grainy,
matte, unbranched, straight
(1-3 mm)

Cenococcum

Emanating Hyphae

Mantle & Hartig Net

Gross Morphology

ECM Morphotype

Appendix A: ECM morphotype descriptions

none

none

pale yellow, loose,
undifferentiated

none

none

none

none

Rhizomorphs

dark brown or hyaline hyphae
(~2-3 um), septate, clamps, no
ornamentation

OM non-interlocking
irregular to regular
synenchyma, Hartig net
present
OM net synenchyma

white 2

white 1

black

Thelephoraceae 2

gold-brown to orangebrown, smooth, matte,
monopodial pinnate,
straight (1-3 mm)
black (white at root tip),
matte, dichotomous
branching to coralloid (.5-2
mm)
white, velvety, matte (1
mm)
patchy white, grainy to
wooly, reflective (~1 mm)

239

OM net synenchyma

OM net to noninterlocking irregular
synenchyma, cell width
~4 um, Hartig net present
OM net synenchyma

hyaline hyphae (2-5 um),
septate, no (sometimes?)
clamps, verrucose or no
ornamentation

OM prosenchyma to net
synenchyma, cell width
3-4 um

hyaline hyphae (1 um), septate,
no clamps
hyaline hyphae (2-4 um),
septate, clamps

black or hyaline hyphae (~3
urn), septate, no clamps

long and copious hyaline
hyphae (2-3 um), septate,
clamps

brown or hyaline hyphae (2-4
um), septate, no clamps,
verrucose or no ornamentation

Rhizopogon-Suillus 2 white, wooly, reflective
(shiny), unbranched or
dichotomous, bent to
straight (2 mm), thick ropelike rhizomorphs
black to dark brown,
Thelephoraceae 1
smooth, matte, unbranched,
straight (1-3 mm)

Rhizopogon-Suillus 1 rusty-brown, grainy to
velvety, matte, coralloid
(<1 mm), thick ropelike
rhizomorphs and hyphal
fans

Emanating Hyphae
hyaline cystidia (2-3 um),
septate, no clamps, verrucose or
no ornamentation

Mantle
OM net to noninterlocking irregular
synenchyma, cell width
1-2 um, Hartignet
present
OM/IM net synenchyma,
Hartig net present,
amorphous deposits on
mantle surface

Gross Morphology
creamy-yellow, smooth to
velvety, matte, unbranched
or monopodial pinnate,
straight (1-4 mm)

ECM Morphotype
Russulaceae 3

none

none

none

none

none

brown-purple pigment
(purple with KOH),
restricted point attachment,
slightly differentiated,
septate, no clamps,
amorphous globules on
surface
yellow-orange pigment
(orange with KOH),
slightly differentiated,
septate, no clamps,

none

Rhizomorphs

Appendix B: Photographs of field site, soils, bioassay, and ECM morphotypes
Plate 1: Soil Collection
1) Soil pit at the Kenneth Creek field site revealing organic, Ae, Bf, Bfi, Bm, BC, and C
horizons of a Dystric Brunisol (Arocena and Sanborn, 1999); 2) Collecting the Bf layer; 3)
Collecting the FH layer (background); lxl m2 PHC-treated plot with the FH and Ae layers
removed (foreground); 4) FHoil layer in situ 4 months after PHC treatment showing
extensive yellow mycelia (a), small conifers (b), and herbaceous plants (c) growing within
the plot; 5) Close-up of Piloderma mycelia from FHoil layer (a) and of a cluster of ECM root
tips from the FH layer (b).
Plate 2: Plant-soil systems and ECMs
1) Bioassay: pine and birch systems following PHC treatment (a); lingonberry (b) and pine
(c) single-plant systems in reconstructed soil layers; cleaned root systems of pine and birch at
the 1-week sampling time (d); pine and lingonberry seedlings growing in CWD organic layer
(e); ECM roots growing into the CWD substrate (f); 2) Cenococcum ECMs dominated the
FH and FHoil layers in pine (a) and birch (b), PHC-treated (a) and untreated (b) systems;
Cenococcum ECMs with profuse emanating hyphae (a,b) Cenococcum with very few hyphae
(c); the typical stellate pattern of the outer mantle confirmed the identity of these ECMs (d);
3) E-strain ECM on pine; 4) Profuse emanating hyphae of Amphinema ECM on pine viewed
using dissecting (a) and compound (b) microscopes; 5) Characteristic needle-like crystals
ornamenting Piloderma hyphae were often found associated with pine root tips, although
Piloderma ECMs were not observed in this study.
Plate 3: ECMs (cont)
1) Russulaceae 1 ECM on pine; 2) Russulaceae 3 ECM (center) and MRA (lower left) on
birch roots; 3) Russulaceae 2 (Lactarius) ECM on pine (a); cluster of Russulaceae 2 tips
growing into CWD (b); Russulaceae 2 ECMs often dominated the root systems of pine (c); 4)
Development of Rhizopogon-Suillus 1 ECMs on pine roots from a small, whitish cluster (a),
to a coralloid cluster with brown pigmentation and ropelike rhizomorphs (b), to a series of
clusters with networks of rhizomorphs extending into the Bf soil matrix (c); 5) Cluster of
Rhizopogon-Suillus 2 on pine roots showing extensive rhizomorph development; 6)
Thelephoraceae 1 root tips (a), irregular to regular synenchyma pattern of outer mantle (b),
and clamped hyphae (c); Thelephoraceae 2 ECM on pine.

240

241

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243