FUNGI IN LOWLAND RIVER FLOODPLAIN ECOSYSTEMS296481/JL...FUNGI IN LOWLAND RIVER FLOODPLAIN ECOSYSTEMS...

FUNGI IN LOWLAND RIVER FLOODPLAIN ECOSYSTEMS

Submitted by

Janice Laraine Williams

B. App. Sc. (Env. Anal.), B. Sc. (Hons)

A thesis submitted in total fulfilment

of the requirements for the degree of

Doctor of Philosophy

Department of Environmental Management and Ecology

School of Life Sciences

Faculty of Science, Technology and Engineering

La Trobe University

Bundoora, Victoria 3086

Australia

November 2010

ii

Frontispiece: Sexual reproductive structures of an aquatic Oomycete growing on

Eucalyptus camaldulensis leaves submerged in a laboratory mesocosm.

iii

Table of Contents

Table of Contents iii

List of Figures vi

List of Tables xviii

Abstract xix

Statement of Authorship xx

Acknowledgements xxi

Chapter 1 1

General Introduction 1

1.1 Ecological Studies of Fungi in Australia 1

1.2 Floodplain Wetland Ecosystems 1

1.3 Carbon Cycles in Wetlands 2

1.4 Aim 2

1.5 Approach 2

1.6 Organisation of the Thesis 3

Chapter 2 5

Carbon Cycling in Lowland River Floodplain Wetlands 5

2.1 Introduction 5

2.2 The Breakdown and Decomposition of Plant Detritus 7

2.3 The Influence of Environmental Conditions 10

2.4 The Wetland Biota 21

2.5 Wetland Carbon Cycles 24

2.6 Plant detritus as a carbon resource 36

2.7 Fungal Ecology 38

2.8 Summary 52

Chapter 3 54

Pilot Study: Fungal Community Structure on a variety of substrates from within a

floodplain wetland. 54

3.1 Introduction 54

3.2 Materials and Methods 56

3.3 Results 60

iv

3.4 Discussion 64

3.5 Conclusions 69

Fungal activity on Eucalyptus camaldulensis leaves under aquatic conditions is enhanced

by terrestrial aging. 70

4.1 Introduction 70

4.2 Methods 72

4.3 Results 77

4.4 Discussion 99

4.5 Conclusions 108

Fungi in Drying Wetland Sediments 110

5.1 Introduction 110


5.3 Results 119

5.4 Discussion 127

5.5 Conclusions 131

Fungal biomass accumulation on E. camaldulensis leaves differs between rivers and

wetlands 133


6.2 Methods 136

6.3 Results 147

6.4 Discussion 166

6.5 Conclusions 177

Synchrotron FTIR micro-spectroscopy reveals chemical changes accompanying aquatic

leaf decay. 179



7.3 Results 191

7.4 Discussion 225

7.5 Conclusions 235

Chapter 8 237

General Discussion 237

8.1 Carbon Cycling in Australian Floodplain Wetlands 237

v

8.2 The Importance of Fungi in Australian Floodplain Wetlands 240

8.3 Additional Observations 242

8.4 Knowledge Gaps 247

8.5 Future Directions 249

8.6 Concluding Remarks 250

Appendix 1: Optimising Noise Reduction 251

References 252

vi

List of Figures

Figure 2.1: A stylised representation of the carbon compartments and fluxes found in

most wetland systems. Based on Webster and Benfield, 1986. 26

Figure 3.1: A map of Normans Lagoon, showing the position of sample sites. The inset

shows the location of Albury-Wodonga, situated on the Murray River, which

forms the border between the states of New South Wales and Victoria. Normans

Lagoon is found on the north side of the Murray River, south-east of Albury,

NSW. 56

Figure 3.2: The western site at Normans Lagoon. The riparian vegetation at this site is

dominated by Eucalyptus camaldulensis. Outside this zone, the vegetation is

influenced by rural land use. 57

Figure 3.3: MDS plot of all T-RFLP data collected from Normans Lagoon. Sediment

samples are shown in black, floc samples are shown in grey and plant detritus

samples are shown in white. Symbols, as shown in the legend, have been used to

differentiate between substrate types. 61

Figure 3.4: MDS plot of T-RFLP data from sediment and floc samples. Samples from the

west lagoon site are shown in black, samples from the east lagoon site are shown

in grey, and samples from the river at Doctors Point are shown in white. Symbols

indicate sediment moisture class, i. e. dry (star), damp (diamond), water’s edge

(squares), inundated (hexagons), deep (circles) and reed bed (triangles). Overlaid

circles are intended to indicate site groups and have no statistical meaning. 62

Figure 3.5: MDS plot of T-RFLP data from sediment and floc samples from the Normans

Lagoon west site. Symbols indicate sediment moisture class, i. e. dry (stars), damp

(diamonds), water’s edge (squares), inundated (hexagons), deep (circles), and reed

bed (triangles). Dry samples are shown in white, damp samples in grey and wet

sediments in black. Overlaid circles illustrate samples are grouped by sediment

moisture content, but have no statistical meaning. 63

Figure 3.6: MDS plot of T-RFLP data from plant detritus samples from Normans Lagoon.

The west site is shown in black, while the east site is shown in grey. Overlaid

circles indicate plant species groupings but have no statistical meaning. 64

Figure 4.1: A schematic diagram of mesocosm design. All mesocosms were comprised of

fish tanks, with a 2cm layer of wetland sediment in the bottom. This was covered

vii

with nylon mesh, and leaf packs were placed upon the mesh. Leaf packs were

covered with weighted wire. In wet treatments, filtered wetland water was added

to a depth of 15cm. 73

Figure 4.2: Mass loss in the dry (i), moist (ii), and wet (iii) treatments over 140 days.

Fresh leaves are indicated by closed symbols while aged leaves are indicated by

open symbols. Mass loss, on the y-axis, is shown as percentage of the mass of the

leaf pack prior to incubation. Error bars indicate the standard error of the data

point. 80

Figure 4.3: Changes in organic matter content of leaf packs over 140 days are shown for

the dry (i), moist (ii), and wet (iii) treatments. Fresh leaves are shown as closed

symbols while aged leaves are shown as open symbols. Organic matter content (y-

axis) is expressed as a percentage of the leaf dry mass. Error bars indicate the

standard error of the data point. 81

Figure 4.4: Fungal biomass (mg.cm-2

) on eucalypt leaves, as calculated from ergosterol

concentration. Differences in fungal biomass between aged (open symbols) and

fresh (closed symbols) leaves are shown for the dry (i), moist (ii) and wet (iii)

treatments. Error bars indicate the standard error of the data point. 83

Figure 4.5: A three dimensional MDS plot based on a 2-stage resemblance matrix for

oxidative and hydrolytic enzyme data for all times, treatments and leaf types.

Fresh leaves are represented by solid symbols, while aged leaves are shown as

open symbols. Treatments are symbolised by stars (dry treatment), diamonds

(moist treatment) and circles (wet treatment). Arrows indicate that the direction of

change in enzyme activity with different treatments differs between fresh leaves

(solid arrow) and aged leaves (dashed arrow). Fresh and aged leaves in the wet

treatment have similar enzyme activity. 86

Figure 4.6: The activity of cellobiohydrolase on aged (open symbols) and fresh (solid

symbols) E. camaldulensis leaves in the dry (i), moist (ii) and wet (iii) treatments.

Incubation time is indicated in days on the x-axis, while enzyme activity rate

(µmol substrate converted per hour, per gram leaf dry weight) is shown on the y-

axis. Error bars indicate the standard error of the data point. 87

Figure 4.7: The activity of β-D-glucosidase on aged (open symbols) and fresh (solid



viii



Figure 4.8: The activity of α-D-glucosidase on aged (open symbols) and fresh (solid





Figure 4.9: The activity of β-D-xylosidase on aged (open symbols) and fresh (solid





Figure 4.10: The activity of phenol oxidase on aged (open symbols) and fresh (solid





Figure 4.11: The activity of peroxidase on aged (open symbols) and fresh (solid symbols)

E. camaldulensis leaves in the dry (i), moist (ii) and wet (iii) treatments.




Figure 4.12: An MDS plot of T-RFLP data from fresh (closed symbols) and aged (open

symbols) E. camaldulensis leaves prior to incubation indicates that different

fungal communities are present on the two leaf types (2D stress is 0.11). Circles

are added to clearly define different groups shown in ANOSIM analysis. 97

Figure 4.13: MDS plots of T-RFLP data showing changes in fungal community structure

in the dry (i), moist (ii), and wet (iii) treatments. Closed symbols represent fresh

leaves while open symbols represent aged leaves. Incubation time is indicated by

symbols. Pre-incubation samples are shown as circles, squares indicate 8 days

incubation, the inverted triangle indicates 16 days incubation and upright triangles

indicate 100 days incubation. 98

ix

Figure 4.14: An MDS plot of T-RFLP data from E. camaldulensis leaves incubated in

under dry, moist and wet conditions for 100 days. Clusters overlaid on the MDS

plot show that the fungal community structure differs between fresh and aged

leaves in the dry treatment, while communities on fresh and aged leaves in the

moist and wet treatments are similar to each other. 99

Figure 5.1: Quat Quatta East lagoon is located north of the Murray River, approximately

6 km west of the township of Howlong, NSW. 113

Figure 5.2: Positions were marked along a transect in the exposed sediments. Position 1

was exposed at the commencement of the 35 day period, while position 6 was

inundated. Panel (a) shows the water level after 8 days while panel (b) shows the

water levels after inflows began on Day 35. The wetland was frequently visited by

grazing cattle (c) and dead fish and algal blooms were observed as the wetland

approached dryness (d). 115

Figure 5.3: The water level in Quat Quatta East Lagoon as it approached dry conditions

(a) and after it had received inflows from the Murray River (b). In the

background, the dominant tree species, E. camaldulensis, can be observed, while

reed beds and immature E. camaldulensis are seen in the foreground. 116

Figure 5.4: The mean moisture content of wetland sediments over a period of wetland

drying and subsequent re-wetting. Position 1 (indicated by a white upright

triangle) is furthest from the centre of the water body, while position 6 (indicated

by a black inverted triangle) is closest to the centre of the water body. Positions 5

and 6 were submerged until exposure on day 22. All samples were exposed on

Day 22, while all samples were inundated on Day 35. Error bars indicate the

standard error of each data point, with maximum standard error being 2.8%. 119

Figure 5.5: The organic matter content of wetland sediments. There is a trend towards

decreasing organic matter with drying, and increasing organic matter with re-

wetting, but this pattern is not consistent across all sediment positions. Error bars

indicate standard error about the mean of each data point. 120

Figure 5.6: Hyphae observed in sediment samples. Shown are aseptate hyphae that may

be Chytridiomycetes, Zygomycetes or Oomycetes (a , b and c), and a septate

hypha with a clamp connection (CC) indicating a species belonging to the

Basidiomycetes (d). 400x magnification. Scale bar = 50µm. 121

x

Figure 5.7: Fungal propagules found in sediment samples (a, b and c) and a fungal hypha

enclosed within a diatom frustule (d). The propagule shown in panel (a) is an

encysted Oomycete propagule and a similar specimen is seen germinating in panel

(b). The organism adjacent to the propagules (arrow) appears to be an amoeba.

The presence of a prominent oil droplet (arrow) suggests the propagule in panel

(c) is a chytrid sporangium. 400x magnification. Scale bar = 50µm. 122

Figure 5.8: The trajectory of change in fungal community structure at position 1 over

time. Error bars indicate standard error about the mean of three points for each

time. Axes are relative positions in data space and have no units. 124

Figure 5.9: MDS plot of sediment fungal community structure shows that there is some

separation of groups by sediment moisture content. Symbols indicate different

moisture content classes (see legend). Overlaid circles are intended to illustrate

grouping of samples by moisture class, but have no statistical meaning. 124

Figure 5.10: MDS plot of T-RFLP samples from Quat Quatta sediments on Day 90. Dry

sediments are shown by open symbols; wet sediments are shown by solid

symbols. Sediments that had not been wet (Bank samples – see legend) separate

well from sediments in the reed bed that had recently been submerged (circles),

and those from positions 1 and 2, that had been submerged for 55 days (upright

and inverted triangles, respectively). The circles are added to illustrate grouping

only, and do not indicate statistical similarity. 125

Figure 6.1: A diagrammatic representation of the experimental design. Litter bag

positions are represented by asterisks. 137

Figure 6.2: Location of field sites within Australia (inset) and within the state of Victoria.

139

Figure 6.3: The study site on the Kiewa River at Killara showing the wetland (a) and the

river (b). River red gums (E. camaldulensis) and exotic species dominate the

riparian vegetation, and understorey vegetation is dominated by pasture grasses.

140

Figure 6.4: The study site on the Ovens River at Peechelba East showing the wetland (a)

and the river (b). River red gums (E. camaldulensis) dominate the riparian

vegetation, and understorey vegetation is dominated by native grasses. 141

Figure 6.5: The Murray study site showing Quat Quatta East lagoon (a) and the Murray

River west of Howlong (b). River red gums (E. camaldulensis) dominate the

xi

riparian vegetation along this stretch of the river and around wetlands, and

understorey vegetation is dominated by grasses. 142

Figure 6.6: Change in the mean organic matter of leaf packs over a 107 day incubation

period is shown for the Ovens (i), Murray (ii) and Kiewa (iii) sites. Samples

incubated in the river indicated by open symbols while samples from wetlands

and indicated by solid symbols. The x-axis shows time in days, while the y-axis

shows the organic matter content of leaf discs as grams per square centimetre.

Error bars indicate the standard error of the data point. 149

Figure 6.7: The nitrogen content of leaves taken from the Ovens (i), Murray (ii) and

Kiewa (iii) sites. Samples incubated in the river indicated by open symbols while

samples from wetlands and indicated by solid symbols. Ovens samples are shown

as circles, Murray samples as triangles, and Kiewa samples as diamonds. The x-

axis shows time in days, and the y-axis shows nitrogen content in mg.cm-2

. Error

bars show standard error of the mean of each data point. 150

Figure 6.8: Carbon to nitrogen ratio of leaf litter from litter bags in the river and wetland

at the Ovens (i), Murray (ii) and Kiewa (iii) sites. The x-axis shows the incubation

time in days, while the y-axis indicates the ratio of mean carbon content (mgC.cm-

2) to mean nitrogen content (mgN.cm

-2). Wetland sites are indicated by solid

symbols, river sites are indicated by open symbols. The red line indicates the

approximate value below which a resource is considered nutritious to

invertebrates. At day 30 C:N in the Ovens River and wetland were similar. 152

Figure 6.9: Fungal biomass from leaves submerged at the Ovens (i), Murray (ii) and

Kiewa (iii) sites. Open symbols indicate river samples while solid symbols

indicate wetland samples. Samples at the Ovens sites are represented by circles,

samples at the Murray site are represented by triangles and samples at the Kiewa

site are represented by diamonds. Un-dotted (white and black) symbols indicate

biomass determined by biovolume, while dotted symbols (White and grey)

indicate biomass measured by ergosterol concentration. Error bars indicate

standard error about the mean at each data point. 156

Figure 6.10: MDS ordination plot of T-RFLP results showing fungal community structure

changes over time. Hexagons as shown in the legend indicate community

structure on leaves prior to being submerged (Day 0), and after 2, 15, 30, 60 and

107 days submerged in the water. Circles indicate the grouping of samples from

xii

30 days and before on the left, and from 60 days and after on the right. X- and Y-

axes indicate relative position in multivariate space and have no units. 158

Figure 6.11: This plot indicates the mean position of MDS points from each sampling

occasion, with error bars indicating standard error about that mean. The data

indicate that fungal community structure changed significantly between Day 0 and

day 30, then changed significantly in a different manner between day 30 and Day

107. X- and Y-axes indicate relative position in multivariate space and have no

units. 158

Figure 6.12: An MDS plot of T-RFLP from the Kiewa System showing variation in

fungal community structure between river and wetland sites at days 30, 60 and

107, but not at earlier times. X- and Y-axes indicate relative position in

multivariate space and have no units. Circles indicate samples taken at the same

time and have no statistical meaning. Numbers indicate sample time for individual

data points. 159

Figure 6.13: An MDS plot of T-RFLP from the Ovens System showing only small

differences in fungal community structure between river and wetland sites within

sampling times. X- and Y-axes indicate relative position in multivariate space and

have no units. Circles indicate samples taken at the same time and have no

statistical meaning. 159

Figure 6.14: An MDS plot of T-RFLP from the Murray System showing small

differences in fungal community structure between river and wetland sites within

sample times, although the ordination will be influenced by the small number of

replicates for Murray wetland samples. X- and Y-axes indicate relative position in

multivariate space and have no units. Numbers indicate sample time (days). 160

Figure 6.15: An MDS plot of T-RFLP data from the wetland samples showing differences

in fungal community structure between wetland sites and clustering by sampling

time. Circles are intended to indicate groupings by time and do not indicate a

statistical difference. X- and Y-axes indicate relative position in multivariate

space and have no units. Numbers indicate sample time (days). 161

Figure 6.16: An MDS plot of T-RFLP from the Day 60 showing that the fungal

community structure tends to cluster by river and wetland sites, and wetland also

cluster by river system. Circles are intended to indicate groupings by time and do

not indicate a statistical difference. X- and Y-axes indicate relative position in

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multivariate space and have no units. An outlying sample from the Ovens River

may have been periodically exposed due to low flows in the river. 161

Figure 6.17: MDS ordination plot showing Oomycete community structure with time.

Hexagons as shown in the legend indicate community structure on leaves after 2,

15, 30, 60 and 107 days submerged in the water. X- and Y-axes indicate relative

position in multivariate space and have no units. 163

Figure 6.18: The mean position of all points from each time as shown in Figure 6.17. The

axes represent dimensions of ordination space and have no labels. Error bars

indicate the standard error of the data point. This graph shows a clear change in

the composition of the Oomycete community moving from left to right, driven by

increases in the number of T-RF lengths. 163

Figure 6.19: MDS ordination plot of the Oomycete community structure showing a low

degree of separation between sites, and between rivers and wetlands. River sites

are indicated by open symbols while wetland sites are indicated by solid symbols.

Symbols indicate the Kiewa (diamonds), Murray (triangles) and Ovens (circles)

sites. 164

Figure 6.20: MDS ordination plot of Oomycete T-RFLP data from 15 days submerged

leaf packs. River sites are indicated by open symbols while wetland sites are

indicated by solid symbols. Symbols indicate the Kiewa (diamonds), Murray

(triangles) and Ovens (circles) sites. 165

Figure 6.21: MDS ordination plot of Oomycete T-RFLP data from leaf packs submerged

in the Kiewa River and an associated wetland. Samples from the wetland are

shown as solid diamonds, while samples from the river are shown as open

diamonds. 166

Figure 7.1: Floodplain aged leaves were inserted into mesh litter bags (a), then submerged

in the river and floodplain wetland (b). The wetland was an ox-bow lake on the

floodplain of the Kiewa River, in north-eastern Victoria, Australia, with

vegetation dominated by Eucalyptus camaldulensis (c). 185

Figure 7.2: 8 µm cross sections of the mid-vein of Eucalyptus camaldulensis leaves that

have been stained with periodic acid/Schiff’s (PAS). The left panels show a fresh

leaf (a), and the right panels show a leaf that has been aged on the floodplain then

submerged in a wetland for 2 days (b). Images are shown at 100x (i) and 400x (ii)

magnification. While no fungal tissue can be observed in the fresh leaf, in the

xiv

partially decomposed leaf, darkly staining fungal hyphae have replaced the

phloem tissues and much of the mesophyll. Bundle sheath cells appear to have

collapsed in the partially decomposed leaf. Fungal hyphae can be seen on either

side of the fibre sheath and within the mesophyll of the partially decomposed leaf

(b(ii)). X = xylem; P = phloem; FS = fibre sheath; E = epidermis; BS = bundle

sheath; F = fungal tissue; M = mesophyll; OV = oil vesicle. 187

Figure 7.3: Anatomical Structure of the mid-vein of Eucalyptus camaldulensis. Above we

see a line drawing (a) and an image of an 8µm cross section of the midvein of an

E. camaldulensis leaf (b). The leaf has been stained with periodic acid/Schiff’s

(PAS) and is shown at 200x magnification. X = xylem; P = phloem; FS = fibre

sheath; E = epidermis and cuticle; BS = bundle sheath; PM = palisade mesophyll;

SM = spongy mesophyll; C= collenchyma. 193

Figure 7.4: Compound microscope images leaf sections. Images are shown at 200×

magnification and stained with PAS. The fresh leaf (a), the leaf that was leached

in autoclaved water (b), and the leaf that was leached in non-sterile river water (c)

show no sign of fungal colonisation. The leaf that was found on the floodplain (d)

is extensively colonised with fungal hyphae, which is observed as darkly staining

tissues surrounding the xylem and fibre sheath, and replacing the phloem and

mesophyll. Fungal hyphae (H) are networked throughout the mesophyll region of

the leaf found on the floodplain. 195

Figure 7.5: Broad scale images of the leaf mid-vein, acquired using the Focal Plane Array

FT-IR micro-spectrometer (x15 magnification). Here we show four micrometer

sections through E. camaldulensis leaves that were (a) removed from a living tree

(fresh); (b) leached in autoclaves river water; (c) leached in untreated river water;

and (d) found on the surface of the floodplain (floodplain aged). Bright field

images of the unstained leaf section are shown in the left-most column, while

images showing the spatial distribution of absorbance in the following bands are

shown in subsequent columns: (i) 1700-1637 cm-1; (ii) 1530-1486 cm-1; (iii)

1284-1185 cm-1; (iv) 1729-1713 cm-1; and (v) 1180-900 cm-1. Lightest pink

indicates highest absorbance while darkest blue indicates lowest absorbance. 196

Figure 7.6: Loss of polysaccharides on leaching. Here, the same leaf sections that were

shown in Figure 7.4 (Bright field) and Figure 7.5 are viewed with portions of the

leaf mesophyll. FPA-FTIR spectral images show absorbance in the polysaccharide

band (1800-900 cm-1) of a fresh leaf (a), a leaf that has been leached abiotically

xv

(b) and biotically (c) in the laboratory, and of a leaf found on the floodplain.

Maximum absorbance is fixed at 120 au (lightest pink) and minimum absorbance

is 0 au (darkest blue). The highly absorbing are to the left of the midvein in image

(d) is a minor vascular trace. 197

Figure 7.7: S-FTIR spectral maps of sections through the leaf mid vein. 201

Figure 7.8: Fungal features in wetland and river decomposed leaves. 202

Figure 7.9: Contour map images of absorbance in amide bands I, II and III. 204

Figure 7.10: Fungal reproductive structure with contours indicating the distribution of

absorbance from amide bands and aromatics. This feature was located in the

section of the leaf that had been submerged in the river for 15 days (240*220 µm).

Contour maps of absorbance in amide I (a), amide II (b), amide III (c) and

aromatic bands (d) are shown. 205

Figure 7.11: Compound microscope images of sections of terrestrially aged leaves that

have partially decomposed in a wetland. Images are shown at 200x magnification

and stained with PAS. After 2 days in the wetland (a) fungal hyphae (H) can be

observed around and within the vascular bundle and throughout the remaining

mesophyll (M) tissue. After 15 days (b) fungal distribution is similar to day 2, but

fissures are developing in the mid-vein. After 30 days (c), 60 days (d), and 107

days (e) leaf disintegration was continuing, and the fibre sheath had decayed.

Xylem (X) and the epidermis (E) remained intact. 210

Figure 7.12: Broad scale images of the leaf mid-vein, acquired using the Focal Plane

Array FT-IR micro-spectrometer (x15 magnification). Here are shown four

micrometer sections through floodplain aged E. camaldulensis leaves that have

been submerged in a floodplain wetland for (a) 2; (b) 15; (c) 30; (d) 60; and (e)

107 days. Visible light images of the unstained leaf section are shown in the left-

most column, while images showing the spatial distribution of absorbance in the

following bands are shown in subsequent columns: (i) 1700-1637 cm-1; (ii) 1530-

1486 cm-1; (iii) 1284-1185 cm-1; (iv) 1729-1713 cm-1; and (v) 1180-900 cm-1.

Lightest pink indicates highest absorbance while darkest blue indicates lowest

absorbance. 212

Figure 7.13: FPA-FTIR images of the leaf midvein from leaves submerged in the Kiewa

River. 213

xvi

Figure 7.14: Compound microscope images of sections of terrestrially aged leaves that

have partially decomposed in a river. Images are shown at 200x magnification and

stained with PAS. After 2 days in the river (a) fungal hyphae can be observed

around and within the vascular bundle and throughout the mesophyll, some

mesophyll tissue is still present. After 15 days (b) fungal distribution is similar to

day 2. Loss of integrity in is greater than in the wetland, possibly due to sand

blasting effects from suspended river sediments. Sand grains in the samples cause

sections to disintegrate as the microtome makes its cut. After 30 days (c), 60 days

(d), and 107 days (e) leaf disintegration is continuing, and the fibre sheath has

decayed. Xylem and the epidermis remain intact and fungal reproductive

structures become abundant. 215

Figure 7.15: Spectra extracted from a segment of the S-FTIR map of the 2 day aquatically

decomposed sample, shown here at the left, with z-values representing absorbance

in the amide I band. Each spectrum represents a pixel with an area of 5 x 5 µm,

which overlaps the adjacent pixel by 3 µm, to give a total transect length of 19

µm. Spectra are presented in a colour similar to that shown in the S-FTIR map.

Pink - orange spectra are derived from the fungal tissue, blue spectra are derived

from xylem and the green spectra are derived from the interface between these

two tissue types. Chemical gradients radiating away from fungal tissues can be

observed for polysaccharides and lignin. Protein is evident as amide peaks 1 and 2

in the first 2 spectra (pink and red). Carbonyl peaks indicating aldehydes &

ketones and lipids (and lignin) are also illustrated. 218

Figure 7.16: The second derivative of three replicate spectra from fresh leaves, and three

spectra from leaves submerged in the wetland for 2 days. Only wave numbers

corresponding to the amide I peak are shown. Peak maxima represent different

structural features of proteins such as extended chains (low components) at 1624-

1637 cm-1

, α-helices at 1654 cm-1

, unstructured components at 1645 cm-1

, and

turns and bends at 1663-1694 cm-1

. Peak broadening and shifts to higher

frequencies indicate weaker hydrogen bonding in the molecule. 219

Figure 7.17: PCR analysis of extracted spectra. Spectra extracted from regions of the S-

FTIR maps that had strong absorbances in the amide I band were normalised and

smoothed. The second derivative of the absorbance within the amide I band was

then taken, as this region is indicative of protein structure. Principle components

xvii

analysis indicated significant differences in protein structure between the two

sample groups. 220

Figure 7.18: Fine scale relationships between protein and lignin distribution 223

Figure 8.1: A schematic diagram representing the temporal patterns observed when a leaf

substrate is colonised by fungi and Oomycetes (top) and an explanation for this

temporal pattern in terms of the spatial distribution of fungal propagules within

the wetland and floodplain ecosystem (bottom). Time periods are indicated by

roman numerals, indicating the biomass accumulating on the substrate (top), and

the likely position of the substrate when these organisms colonise and grow to

maximum biomass levels (bottom). 245

Figure 8.2: This graph illustrates the number of data points remaining for each T-RFLP

sample after noise was removed. The percentage of noise reduction is shown on

the x-axis, while the number of taxonomic units represented is shown on the y-

axis. 251

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List of Tables

Table 5.1: The moisture status of sediments at 6 positions over a 35 day period. .......... 115

Table 6.1: Summary of the environmental conditions in the river at each site over the

study period. Data are based on the mean value of mean daily readings, with

maximum and minimum mean daily readings going to comprise the stated range.

Data were obtained from the Murray-Darling Basin Authority website

(http://www.mdba.gov.au/water/live-river-data). ................................................ 142

Table 7.1: Bands of interest in the FT-IR spectra obtained from decomposing leaf

material in the range 1800-900 cm-1. The column labelled “Wave Number”

indicates the frequency limits between which each band was integrated, “Band

Assignment” indicates the molecular vibration responsible for absorbance in the

stated frequency range (υ = stretch; υs = symmetric stretch; υas = asymmetric

stretch δ = deformation (bend)). The reference from which this assignment was

sourced is listed in the third column, and an indication of the probable source of

these vibrations within leaf material is indicated in the “notes” column............. 189

Table 7.2: The 2nd

derivative peaks that gave positive loadings for principal components

1, 2 and 3. The protein structure associated with these peaks, as reported by Byler

and Susi (1986), is shown in the column labelled “Protein structure”. ............... 220

xix

Abstract

Fungal decomposition processes are recognised for their importance in terrestrial

ecosystems, but their role in aquatic ecosystems is not as well established. In particular,

the function of fungi in wetland carbon cycles is poorly understood. Indeed, one of the

few studies of fungi in Australian wetlands suggested that fungi played no role in carbon

cycling, based on an absence of fungal phospholipid biomarkers in sediments. In this

thesis I examine the community dynamics of aquatic fungi and their functional role in

floodplain wetlands.

An initial survey of fungal diversity in a floodplain wetland (Chapter 3) clearly showed

that fungi were present on a range of submerged and exposed substrates. A mesocosm

experiment then examined the influence of leaf age and moisture availability on fungal

dynamics (chapter 4). A period of terrestrial aging was found to promote fungal

colonisation of Eucalyptus camaldulensis leaves during aquatic decomposition,

potentially impacting wetland food chains, water quality and carbon dynamics.

DNA from a range of fungal species was extracted from the sediments of a floodplain

wetland undergoing drawdown (Chapter 5), while fungal biomass was below detection

limits. The results suggest that fungi do not grow in wetland sediments. Rather, the

sediments act as “seed bank” that stores fungal propagules until the wetland refills.

Aged E. camaldulensis leaves undergoing aquatic decomposition in river and wetland

ecosystems were examined to compare fungal and Oomycete community dynamics

(Chapter 6) and changes in leaf chemical composition (Chapter 7). Fungal biomass was

consistently lower in wetlands, when compared to streams and Oomycete biomass

comprised a substantial proportion of total fungal biomass at various times in both

ecosystems. Initial increases in fungal and Oomycete biomass were correlated with

temporary improvement in leaf food value. Infra-red mapping of these partially

decomposed E. camaldulensis leaves (Chapter 7) showed changes in the distribution of

polysaccharides, protein and lignin over time, and changes in lignin distribution occurred

earlier in the wetland than in the stream. The activity of oxidative fungal enzymes was

indicated as an important mechanism in the transformation of refractory carbon

compounds in aquatic ecosystems.

xx

Statement of Authorship

Except where reference is made in the text of the thesis, this thesis contains no material

published elsewhere or extracted in whole or in part from a thesis submitted for the award

of any other degree or diploma.

No other person's work has been used without due acknowledgment in the main text of

the thesis.

This thesis has not been submitted for the award of any degree or diploma in any other

tertiary institution.

Research procedures reported in the thesis carried out in the Ovens River Nature

Conservation Reserve were undertaken under Department of Sustainability and

Environment Permit No: 10004708 / File No:FF383072.

Signed: Date:

xxi

Acknowledgements

Carrying out a PhD project and writing the thesis is indeed a marathon. I could not have

succeeded without the help, support, guidance and forbearance of numerous people, and I

will attempt to name them all now.

Initially, my project supervisors were Darren Baldwin and Gavin Rees of the Murray-

Darling Freshwater Research Centre (MDFRC), and Roger Croome from La Trobe

University. Roger retired in 2009 (although he has maintained an interest in my project)

and was most ably replaced by Phil Suter. To you gentlemen, my most humble and

sincere gratitude. Darren, who was also my supervisor when I worked with MDFRC,

should also be acknowledged for his role in formulating the original research question,

after a project investigating blackwater events revealed that there were no ecological

studies of fungi from Australian wetlands.

As it takes a village to raise a child, it takes a department to nurture a PhD project. This

project was wholly undertaken at the Albury-Wodonga campus of La Trobe University in

the Department of Environmental Management and Ecology and was supported

throughout by its staff. So, thanks to DEME, but in particular I would like to

acknowledge the guidance and assistance of Susan Lawler, who was the post-graduate co-

ordinator throughout most of this project and more recently became head of department.

Rachel Bahrij handled the buying and receiving, booking and managing of all manner of

logistical concerns. To Pettina Love, who shared my office for a number of years and

assisted with field work, thanks for your advice, your friendship, your patience and your

honesty. Thanks also to Peter Pridmore, Martin Fussell, Warren Paul, and Ewen Silvester

for advice, stimulating conversation and friendship, and to the Department for project and

conference funding.

Thank you to the staff and board of MDFRC, where the majority of the lab work done for

this project was carried out. In particular, I would like to thank Garth Watson and Tara

Pitman for help in the lab. Garth provided valuable guidance with PCR and DNA

extractions and Tara kept the lab organised, very graciously washed my glassware on

several occasions, and kept me on my toes. Shaun Meredith assisted me in locating

suitable field sites. This help was greatly appreciated. MDFRC also provided assistance

with conference funding on several occasions, and in kind support in terms of computers,

xxii

journals and administration, so thanks to Ben Gawne, Rosie Busuttil, Rhonda Sinclair and

Julie Newell. To the rest of the MDRFC staff, thanks for your friendship and support.

Deep and most heartfelt appreciation should go to my family, all of whom supported and

helped me in some way throughout the project. John, Giselle and Alex all helped with

field work and were patient with my moods and constant work load. Thanks also to Mum,

Dennis, Karen, Johnny, Karina, Aunty Lola, Laurie, Mavis, Paul, Mandy, Mark, Ann,

Mick, Nikki, Ant and Cathrine for your constant support and encouragement over the

years. Though you’re not here to receive it, thanks to Dad and Keith for encouraging me

to do my best, and then do a little better. Thanks also to Ali Mitchell, Nicole Costigan,

Sally Hladyz and Dale McNeil for emotional support, guidance and understanding when

it was hard find.

La Trobe University provided a scholarship for three and a half years, and this was

augmented with a top-up scholarship from the eWater CRC, who also provided research

funds, training opportunities, and the chance to share my research with professionals in

the field of aquatic ecology. The eWater CRC, and its CEO, Gary Jones, took great

interest in their post-graduate students, and provided us with the chance to gather, learn,

and share our research at an annual student meeting. Thanks to the other eWater post-

grads for the support and friendship I found during these meetings and kudos to Gary

Jones for maintaining his sense of humour when others may have lost theirs. Very special

thanks to Anne Taylor, the eWater student co-ordinator, for her role in managing

scholarships, research and conference funding and the annual student meeting. Her efforts

to help us extended well beyond her responsibilities, and the post-graduate students

developed a true affection for her.

The Australian Society for Limnology (ASL) and the British Mycological Society (BMS)

provided funding to assist me in attending conferences. ASL members have encouraged

my research with the student prize at their 2009 conference, and many helpful

conversations over the past few years.

I also wish to acknowledge the assistance of Parklands, Albury-Wodonga, who manage

the site on the Kiewa River where some of the field work was conducted. The staff there

could not have been more helpful.

xxiii

I could not have completed the analysis at the Australian Synchrotron without the

assistance of a large group of people. Firstly, I would like to thank Ewen Silvester, who

conceived the approach to the synchrotron experiment and assisted and supported

throughout its execution. Ian Boundy and Stefania Tombs at the histology lab at Monash

University, and Sally Caine, also of Monash University, assisted in the preparation of leaf

sections. Mark Tobin, Ljiljana Puskar and Danielle Martin of the Australian Synchrotron

were indispensible in their assistance with sample analysis and data analysis. Kerry

Whitworth and John Pengelly of Murray Darling Freshwater Research Centre and Nicole

Costigan, also assisted with sample analysis. Roger Croome advised me in matters of

plant anatomy and Phil Suter gave advice on the design of the field work component,

invertebrate feeding and wetland ecology.

Lastly, I would like to acknowledge the line of teachers who inspired my interest and

passion for science. Firstly, thanks to Phillip Hitchcock, my 5th

class teacher, who

encouraged me to think for myself. Thanks also to Brian Copeland, who taught me

science throughout high school and nurtured my enquiring mind (patience of a saint!).

During my undergraduate years, David Robertson, Ken Page and Robin Watts from

Charles Sturt University were inspirational. Andrew Bray and Gordon Tribbick from

Mimotopes taught me a lot about chemistry, proteins and debating and showed me that a

PhD was not beyond my grasp. Lastly, but by no means least, my Honours supervisor,

Kim James of Deakin University inspired my interest in floodplain wetlands and taught

me that I still have a lot to learn.

In the absence of any of the above people, I may not be where I am today.

To anyone who I may have overlooked, thanks to you too.

http://www2.synchrotron.org.au/content.asp?document_id=5537

1

Chapter 1

General Introduction

The role of micro-organisms in nutrient cycles has been widely recognised, and the

integrated study of micro-organisms and the chemical changes they initiate in the

environment has become known as biogeochemistry (Mitsch and Gosselink 1993a). In

wetlands, studies of biogeochemistry have been focused primarily of the activities of

anaerobic bacteria in the sediments (Boon and Sorrell 1991; Gutknecht, Goodman et al.

2006; Sorrell and Boon 1992b), and benthic and pelagic aerobic bacteria (Howitt,

Baldwin et al. 2007; Wetzel 1992). Very little research exists that examines the role of

aquatic fungi in the cycling of carbon in wetlands. Given the known importance of fungi

in terrestrial systems, we might intuitively expect fungi to have some role in carbon

cycling in wetlands, but this role had not been explored, much less quantified. As carbon

dynamics take on increasing importance in the light of the enhanced greenhouse effect

and global climate change, an examination of this role is timely.

1.1 Ecological Studies of Fungi in Australia

The most important previous study of the ecology of aquatic fungi in Australia was the

doctoral thesis by Ken Thomas (Thomas 1992), and the subsequent publications

(Thomas, Chilvers et al. 1989; Thomas, Chilvers et al. 1992). However, this study dealt

with fungal dynamics in an upland stream, an ecosystem quite different to floodplain

wetlands. Other studies have investigated fungal dynamics in the litter layer of eucalypt

forests (Cabral 1985; Eicker 1973; Macauley and Thrower 1966; Orsborne 1987), fungal

endophytes (Bertoni and Cabral 1988; Bettucci and Saravay 1993) and the effect of

pathogenic fungi on eucalypts (Abbott, Vanheurck et al. 1993). There are no published

studies of fungal dynamics from Australian floodplain wetlands, and very few from

floodplain wetlands in other countries (Baldy, Chauvet et al. 2002; Chergui and Pattee

1993; Pattee and Chergui 1995; Singer 1988).

1.2 Floodplain Wetland Ecosystems

In Australia’s Murray-Darling Basin, floodplain wetlands are important cultural sites.

Both aboriginal people and those arriving over the last two centuries have utilized them

2

for water, food gathering, forestry, flood mitigation and biodiversity conservation (Lloyd,

Atkins et al. 1992). These wetlands also have spiritual significance for many people.

Ecologically, floodplain wetlands are important for a variety of reasons. They influence

water quality (Mulholland and Kuenzler 1979), are biodiversity and productivity hot

spots (Hillman and Quinn 2002) and ground water fed wetlands may act as a drought

refuge for a range of different species (Baldwin and Humphries 2001). In many ways, the

value of floodplain wetlands in Australia lies in their heterogeneity, each of more than

7000 wetlands on the Murray River floodplain (Pressey 1990) harboring a unique

community (Lloyd, Atkins et al. 1992). Fungi are likely to be important components of

these ecosystems, even though their interactions with other organisms have not yet been

investigated.

1.3 Carbon Cycles in Wetlands

The concentration of carbon dioxide in the atmosphere has increased by one third in the

last 150 years, while methane concentrations have increased by 140% (Reddy and

DeLaune 2008). Wetlands are one of the major non-anthropogenic sources of these

emissions, with natural wetlands producing methane in the range of 100-300 x 1012

grams of carbon per year (Chynoweth 1996; Reddy and DeLaune 2008). Carbon

emissions from wetlands tend to increase with ecosystem productivity (Whiting and

Chanton 1993), and floodplain wetlands can be highly productive systems (Briggs and

Maher 1985). Wetlands are also known to store more carbon per hectare than most other

ecosystem types (Houghton 1995; Schlesinger and Melack 1981; Whittaker and Likens

1973b). It is important that we understand what contribution aquatic fungi might make to

these emissions, and how their presence might influence emission from other sources.

Such an understanding would contribute to the design of wetland management practices

that reduce carbon emissions and increase carbon storage.

1.4 Aim

The aim of this thesis was to examine the role of fungi in the decomposition of various

carbon sources in the floodplain wetlands of Australian lowland rivers.

1.5 Approach

For this thesis it was deemed appropriate to take an exploratory approach, and to

examine the fungal community of floodplain wetlands as a whole, rather than identify

3

fungal species present. So while it is acknowledged that identifying fungal taxa is

important to determine the impacts of finer scale interactions within an ecosystem, only

broad taxonomic and functional groups of fungi have been discussed in this project, and

no attempt has been made to definitively identify fungal species.

1.6 Organisation of the Thesis

The scientific literature discussing carbon cycling in wetlands and, within this context,

the activities of fungal decomposers are reviewed (Chapter 2). This review suggests that

it is reasonable to assume that, of the total fungal community within wetlands, the

saprobic fungi are most likely to have the strongest influence on wetland carbon cycles.

However, as no study has investigated fungal dynamics in lowland river floodplain

wetlands, it is important to first confirm that fungi are present, and which substrates they

are colonising.

A pilot study was devised where fungal DNA was extracted from a variety of substrates

within a floodplain wetland (Chapter 3). These substrates included wetland sediments

under various moisture conditions, sediments from the nearby river, wetland macrophyte

litter, and the abscised leaves of Eucalyptus camaldulensis (the river red gum) and Salix

alba (white willow). These data indicated differences in fungal community structure

between sediments and plant detritus. Changes were also observed with different

sediment moisture contents, between sites in the wetland, and between sediments in the

river and wetland. The fungal community structure on eucalypt leaves differed from that

on other plant detritus.

Given these preliminary findings, a mesocosm experiment was designed to investigate

fungal dynamics on E. camaldulensis leaves under different moisture regimes that may

exist within the filling and drawdown cycle of floodplain wetlands (Chapter 4). As the

seasonality of this cycle influenced the quality of the eucalypt leaf as a fungal substrate,

both fresh leaves and leaves that had aged under terrestrial conditions on the floodplain

were used. This experiment showed that fungal dynamics differed markedly between

fresh and aged eucalypt leaves, demonstrating that leaching (either by submergence or

aging on the floodplain), was an important process that removes leaf components that

inhibit fungal colonisation. Importantly, a significant increase in fungal biomass that

occurred on aged leaves immediately after being submerged was absent on fresh leaves.

4

After 60 days, the fungal community structure on fresh and aged leaves was similar, but

biomass was significantly lower on fresh leaves. For all mesocosm treatments, fungal

biomass was approximately ten times less than that reported from leaf litter in streams.

Importantly, much of the fungi observed growing on leaves in the mesocosms was not

true fungi but belonged to the Oomycetes (see Frontispiece). The biomass of these

organisms is not assessed when the ergosterol method is used to estimate fungal biomass.

The presence of fungi on natural substrates was then examined in a field setting. In

Chapter 5 the biomass and community structure of fungi in wetland sediments is

explored, while in Chapter 6 fungal and Oomycetes biomass and community structure

are examined on submerged floodplain aged leaves, and compared between river and

wetland environments. Sections of leaves that had been “conditioned” in the river and

wetland were then examined using Fourier transform infrared micro-spectroscopy

(Chapter 7). Here the chemical changes occurring in the leaf substrate as fungal

decomposition proceeded were examined over a time series. As these results are spatially

explicit, the sequence in which various leaf components were degraded was revealed, and

differences were observed between leaves incubated in the river and the wetland.

Finally, the experimental results are discussed in the context of other research into

aquatic fungi in freshwater ecosystems (Chapter 8). While there is a growing body of

knowledge regarding fungal dynamics on plant detritus in river ecosystems, our

understanding of aquatic fungi in wetland and floodplain ecosystems is less complete.

The areas of study important for a more complete understanding of the role of aquatic

fungi in wetland carbon cycles are discussed, and research approaches are suggested.

5

Chapter 2

Carbon Cycling in Lowland River Floodplain Wetlands

2.1 Introduction

Carbon is life. Every carbon atom on planet Earth spends eternity passing into and out of

living organisms. This is the essence of the carbon cycle, and the food chains and

decomposition pathways that drive it. The rate at which carbon moves along these

pathways has changed as a result of human activities including river regulation, flood

mitigation and wetland drainage. Wetlands are highly productive systems capable of

storing large amounts of carbon (Brinson, Lugo et al. 1981; Houghton 1995; Mitsch and

Gosselink 2000; Whittaker and Likens 1973a), so processes that accelerate the

transformation of stored wetland carbon into gaseous forms have the potential to

contribute substantially to global climate change (Chynoweth 1996; Reddy and DeLaune

2008; Whiting and Chanton 1993). For example, over 10% of global carbon dioxide

emissions each year result from the degradation and destruction of peatlands (Parish,

Sirin et al. 2008). Thus, it is important that we understand the mechanisms of carbon

transformations in wetlands and adjacent systems.

The floodplain wetlands of lowland rivers in Australia are often ephemeral, experiencing

both terrestrial and aquatic conditions over time. While they are commonly highly

productive in terms of aquatic vegetation, they also receive large amounts of plant litter

from outside of the water body (allochthonous litter). As much of this litter is not directly

grazed upon by fauna (Brinson, Lugo et al. 1981; Newman 1991), decomposition

processes will determine how much carbon is stored in a wetland, and how much is

released into the atmosphere. It is therefore essential to fully understand aquatic

decomposition processes in these systems.

Floodplain wetlands are found adjacent to streams that, at least occasionally, flood

beyond their banks and have an associated floodplain (Mitsch and Gosselink 1993b).

These systems are generally characterized by a cycle of flooding and drying that entails

alternation between terrestrial and aquatic conditions (Mitchell 1992). In Australia,

examples of floodplain wetlands include ephemeral and permanent ox-bow lakes, and

river red gum (Eucalyptus camaldulensis, Dehnh.) forest wetlands. These water bodies

6

are commonly referred to as “billabongs” and the terms floodplain wetland and billabong

are used interchangeably.

Australia’s floodplain wetlands differ from lakes in two important ways. Firstly, lakes

generally attain a depth that allows thermal and chemical stratification to occur. This

results in anoxic bottom layers and the dominance of anaerobic decomposition processes

within the sediments. The relatively shallow depth of wetlands enables oxygenation of

the water column and the surface layer of sediment, at least during winter. This means

that 70-80% of decomposition is carried out by aerobic organisms (Kjoller and Struwe

1980). Secondly, carbon inputs to lake sediments are generally dominated by algae,

which contain no lignin, whereas in wetlands carbon inputs are usually dominated by

macrophytes, which are 20-30% lignin. Lignin is degraded slowly under anoxic

conditions. Once lignified plant detritus becomes entrained within anoxic sediments it

may be stored there for extensive periods (Westermann 1993).

The distinctive features of floodplain wetlands are that they are linear in form (often

being former stream channels), both energy and matter pass through them in large

quantities, and they maintain lateral and longitudinal connectivity to other ecosystems

(Mitsch and Gosselink 1993b). Flood events are the main mechanism through which

lateral connectivity and energy flow are achieved. These events provide a pulse of

nutrients and organic matter, while also altering pH, salinity and dissolved oxygen, and

influencing primary production. The alternation of flooded and dry conditions sets up

temporal gradients in oxygen, nutrient and carbon contents of soils and sediments (Hart

and McGregor 1980) that can affect the rate of microbial decomposition processes

(Mitsch and Gosselink 1993a).

The floodplain wetlands have important ecological functions in Australian systems. They

are able to improve the quality of water that passes through them, are biodiversity and

productivity hot spots (Kingsford, Curtin et al. 1999), and act as a seasonal drought

refuge for water birds (Kingsford and Norman 2002) and fish (Arthington, Balcombe et

al. 2005; Magoulick and Kobza 2003). They are also important fish nurseries and

waterfowl habitat due to rich phytoplankton and zooplankton communities that provide a

foundation for the food chain. Extensive macrophyte growth may provide shelter and a

refuge from predators.

7

However, each wetland is unique in terms of its physical and chemical characteristics and

may support a distinctive community (Lloyd, Atkins et al. 1992). There are more than

7000 open water wetlands on the floodplain of the Murray River alone (Pressey 1986),

covering 2200 km2, an area equal to 17% of the floodplain (Margules and Partners, P and

J Smith Ecological Consultants et al. 1990). These systems could therefore be considered

important based solely on their extensive area.

2.2 The Breakdown and Decomposition of Plant Detritus

Much of the literature that describes the processing of dead organic material refers to

both breakdown and decomposition. Here these terms are defined in the same manner as

Boon (2006): breakdown is the gradual loss of mass from organic material by means of

physical fragmentation, leaching and the conversion of organic carbon to inorganic

forms, while decomposition (and mineralisation) describes the conversion of organic

substances into carbon dioxide and methane (Boon 2006; Boulton and Boon 1991). In

wetlands, decomposition is mediated by a variety of micro-organisms, while breakdown

results from physical, chemical and biological processes. This review will focus

primarily on decomposition in wetlands, and discuss the contribution of these

decomposition processes to the overall breakdown of organic matter.

Further to this, the term “oxic” will be used to describe conditions where oxygen is

available, while the term “aerobic” will be used to denote metabolic processes that

require oxygen. Similarly, “anoxic” describes the absence of oxygen, and “anaerobic”

denotes processes that do not require oxygen.

2.2.1 Aquatic plant litter decomposition

The breakdown and decomposition of organic plant material in aquatic systems is

described as a three stage process (Bunn 1988a). It consists of:

(a) Leaching of soluble substances;

(b) Colonisation by micro-organisms and the commencement of decomposition

through the activities of extra-cellular enzymes (microbial conditioning); and

(c) Physical fragmentation, mediated by mechanical forces and the activities of

macro- (>2mm) and meio-fauna (<2mm) (Pidgeon and Cairns 1981; Webster and

Benfield 1986).

8

Leaching is responsible for the largest mass loss from submerged litter in the initial

stages of decay (Day 1983). In aquatic ecosystems the rate of leaching is not limited by

precipitation, but is controlled by species specific litter quality and plant tissue type

(wood, bark, leaves or other structures)(Baldwin 1999; O'Connell, Baldwin et al. 2000).

Water soluble organic substances such as sugars, amino acids and organic acids, begin to

leach from plant detritus immediately after immersion. Up to 30% of the organic carbon

can be lost from litter after just 4 days in the water (Brinson 1977). The leaves of E.

camaldulensis become leached within 72 hours (Glazebrook 1995), while leaching of

soluble substances from woody debris can take more than a year (Thomas, Chilvers et al.

1992). The amount of DOC released from submerged twigs by leaching in the first 48

hours is proportional to the mass of wood immersed (O'Connell 1998), but for large

woody debris, decay is confined to thin surface layers (O'Connor 1992). Relatively small

amounts of organic substances are leached from particulate organic matter (POM)

(O'Connell, Baldwin et al. 2000). Carbon that is leached from submerged litter into the

water column is known as dissolved organic carbon (DOC).

Following leaching the litter is composed of refractory organic compounds. These

compounds are largely insoluble plant structural polymers including cellulose,

hemicellulose and lignin (Chamier 1985; Esau 1977). The leaves are colonised by

microorganisms and decomposed at various rates, depending on the physical and

chemical characteristics of the water body (Westermann 1993). The microbial

community composition also changes over time as the substrate composition changes,

utilising soluble carbon, cellulose and lignin sequentially (Moorhead and Sinsabaugh

2006). This process is known as “leaf conditioning”. Together, leaching and leaf

conditioning may be responsible for up to 70% of the mass loss from submerged plant

detritus (Findlay, Howe et al. 1990).

The decomposition rate is, by definition, the amount of carbon consumed from a resource

over time. Carbon consumption by micro-organisms is influenced by factors such as

moisture availability, temperature, the quality of the resource and the availability of

oxygen (Baldwin 1999; Mackensen, Bauhus et al. 2003). While temperature, moisture

and resource quality influence the metabolic rates of the microbial community, the

availability of oxygen influences decomposition rates chiefly via changes to the

microbial community composition. Under oxic conditions, submerged leaves are

9

colonised by communities dominated by aerobic fungi, while under anoxic conditions,

anaerobic bacteria are dominant (O'Connell 1998). As anaerobic decomposition tends to

proceed more slowly (Boon 2006; McLatchy and Reddy 1998; Webster and Benfield

1986), anoxia slows decomposition rates.

In aquatic systems, leaf conditioning improves the food value of the leaves for

invertebrate consumers (Bärlocher 1982; Bärlocher 1985; Bärlocher and Corkum 2003).

During conditioning, concentrations of nitrogen and phosphorus increase as structural

polysaccharides are metabolised and microbial biomass accumulates (Suberkropp 1992;

Suberkropp, Godshalk et al. 1976). In streams, fungi comprise the majority of the

microbial biomass in the period prior to the point where leaves have maximum food

quality (Findlay and Arsuffi 1989).

Fungi improve the food value of leaf detritus by adding fungal biomass (Bärlocher 1985;

Bärlocher and Kendrick 1975), and enzymatically converting plant polymers into more

digestible forms (Suberkropp 1992; Zimmer, Oliveira et al. 2005). While it has been

suggested that these processes improve palatability to aquatic invertebrates (Kaushik and

Hynes 1971), Suberkropp et al. (1983) found no relationship between fungal biomass and

palatability. However, palatability is not the same as food value. Insects require sterols

and long chain linoleic (ω3) polyunsaturated fatty acids for metamorphosis and pupation.

These compounds are not available from plants or bacteria but can be provided by fungal

tissues (Boulton and Boon 1991). Furthermore, fungi may provide nutrients to

invertebrates that are critical for growth, maturation and reproduction (Albariño,

Villanueva et al. 2008; Cargill, Cummins et al. 1985).

In aquatic systems, conditioned litter is consumed by a group of aquatic invertebrates

functionally defined as “Shredders” (Merritt and Cummins 1978). These are mainly the

larvae of invertebrates that shred and chew coarse particulate organic matter (CPOM),

such as plant detritus. They include stoneflies, caddisflies, crane flies and midges

(Cargill, Cummins et al. 1985; Reid, Quinn et al. 2008; Suberkropp, Arsuffi et al. 1983).

This group of organisms is known to affect litter breakdown and decomposition rates in

streams by digestion and respiration of the organic carbon (Golladay and Sinsabaugh

1991; Pidgeon and Cairns 1981), and mechanical fragmentation (Van Wensem, Van

Straalen et al. 1997). However, invertebrate communities can differ markedly between

10

streams and wetlands (Sheldon, Boulton et al. 2002; Suter and Hawking 2002). While

invertebrate communities in wetlands also rely on allochthonous carbon (Batzer and

Wissinger 1996), shredders that are specifically adapted to process plant litter are rare in

wetlands (Balcombe, Closs et al. 2007; Mulholland 1981). The influence of shredders on

leaf breakdown can not be assumed to be the same for rivers and wetlands.

In a forested floodplain wetland on an Australian lowland river, fragmentation dominated

the latter half of aquatic leaf break down and decomposition (Glazebrook 1995), a

process known to take about 4 months (Briggs and Maher 1983). While fragmentation

was the most important factor affecting leaf mass loss in streams (Golladay and

Sinsabaugh 1991), a study in the littoral zone of a lake has suggested that fragmentation

and microbial processes are of equal importance in non-flowing environments

(Kominkova, Kuehn et al. 2000). Similarly, Janssen and Walker (1999) concluded that

physical fragmentation and the proportion of carbohydrates were of equal importance in

the control of leaf break down rates in a floodplain wetland on an Australian lowland

river. As structural carbohydrates are the main energy source for microbes in submerged

leaves, these studies suggest that invertebrate feeding and microbial activity gain equal

importance in determining the rate of leaf breakdown in the absence of flow.

In aquatic ecosystems abiotic carbon loss, carbon mineralisation and mechanical

fragmentation are important factors contributing to the overall rate of breakdown and

decomposition of submerged leaf litter. Carbon loss is largely due to leaching, which is

rapid and immediate. The composition of the decomposer community responsible for

carbon mineralisation is heavily dependent on environmental factors such as dissolved

oxygen, and varies between water bodies. Invertebrate activity, in combination with

abiotic factors including flow, dominates litter breakdown rates, but microbial activity

may be of equal importance to breakdown rates in the absence of flow.

2.3 The Influence of Environmental Conditions

Floodplain wetlands are highly dependent upon seasonal flooding. In a survey of 58

wetlands on the Murray River floodplain, Roberts (2006) reported that 86% were

categorised as “lentic channel forms”. This wetland type includes ox-bow lakes and

meander cut-offs, and former channels of various lengths in low-lying areas of the

floodplain that are connected, or potentially connected, to the river at or above minimum

11

regulated flow (Pressey 1986). This wetland type accounted for 50% of all 6433 wetlands

identified by Pressey in 1986. Other wetlands which are in higher areas of the floodplain

receive water when river flows are above maximum regulated flow (Pressey 1986), and

may dry out intermittently. However, minimum regulated flows have changed in recent

years, and many wetlands in low-lying areas may now also dry periodically. Roberts

(2006) reported that only 81% of lentic channel forms were conneted at minimum

regulated flow.

The physical and chemical characteristics of a floodplain wetland can have a strong

influence on the rate of plant litter decomposition, particularly in terms of the biological

processes involved. In floodplain wetlands, one of the strongest influences on plant litter

decomposition is the flooding and drawdown cycle, which is accompanied by seasonal

temperature changes. These processes affect the decomposer community (invertebrates

(Balcombe, Closs et al. 2007), fungi and bacteria) by influencing the availability of

organic substrates (food), the availability of suitable habitat, and opportunities for

reproduction. The influence of the water regime and seasonal changes is reflected in the

measurable physical parameters. These include the hydrological regime, temperature,

turbidity and sedimentation, salinity, pH, oxygen availability and redox potential,

nutrients and litter quality. These factors have been shown to strongly influence

decomposition rates and pathways (McLatchy and Reddy 1998).

2.3.1 The hydrological regime

Annual flooding supplies floodplain wetlands with carbon in the form of leaf litter, large

woody debris (LWD) and leachates from the soil. Floodwaters also deliver nutrients

including nitrogen and phosphorus (Baldwin and Mitchell 2000; Robertson, Bunn et al.

1999). Flood events flush and refill floodplain wetlands, creating the potential for high

levels of primary productivity (Bayley 1991; Junk, Bayley et al. 1989). A significant

amount of dissolved organic carbon (DOC) may also be returned to the river channel

during a flood event from reservoirs on the floodplain such as wetlands (Hillman 1995;

Sedell and Dahm 1990). While billabongs and other lentic water bodies on floodplains

generally receive water rapidly during flood events, water is lost gradually by

evaporation. Thus, while water quality may be similar to that of the river when the

wetland is full, water quality parameters begin to vary from the river as soon as the

connection is lost (Hillman 1986).

12

The onset of flood in a floodplain wetland brings a number of changes to the physical

environment. These include soil/sediment anoxia, the importation and exportation of

organic matter, nutrients and sediment, and shifts in pH and oxidation states (Baldwin

and Mitchell 2000; Boon and Mitchell 1995; Mitsch and Gosselink 1993a). However,

periodic flooding alternating with dry periods can increase plant productivity, enhance

soil nutrient cycles, re-oxygenate root zones and flush toxins and wastes from the system

(Mitsch and Gosselink 1993a). Flooding may influence plant litter breakdown differently

depending on its position in the wetland. Neckles and Neill (1994) found that flooding

increased litter mass loss rates where litter was lying above the sediment, but slowed

decomposition below the sediment surface. These changes will have ramifications for

wetland decomposition.

The timing and/or season of flooding also have a significant impact on the activity of the

decomposer community in Australian wetland sediments. Flooding of wetlands in

summer, as opposed to autumn/winter, can cause rapid increases in methane emissions

from the sediments (Boon, Mitchell et al. 1997) and accelerated decomposition of

floodplain leaf litter (Glazebrook and Robertson 1999). River regulation has altered the

timing and season of flooding in many Australian river systems. Under regulated

conditions the majority of small and medium floods are captured by headwater dams in

winter/spring, and minor to moderate flooding occurs in summer due to irrigation

releases (Frazier and Page 2006). The change in the timing of these flooding events from

winter/spring to summer coincides with higher water temperatures. This affects both the

metabolic activity of micro-organisms and the ability of the water and soils to retain

oxygen. As a consequence, there are changes in the types of decomposition processes

that predominate, and in the rate at which they proceed. Changes in flood seasonality also

affect the primary production in floodplain wetlands (Robertson, Bacon et al. 2001).

Decomposition is faster in wetlands that are flooded for at least part of the year than at

sites that are never flooded (Neckles and Neill 1994). This is because flooding

accelerates leaching and the microbial conditioning phases of decay (Baldwin 1999;

Glazebrook 1995). When flooding alternates with drying, both aerobic and anaerobic

decomposer communities are able to contribute to organic matter decomposition.

However, overseas research indicates that alternating oxic and anoxic conditions slows

13

decomposition (Mitsch and Gosselink 1993a). The duration of wet and dry periods and

the depth of inundation are also likely to affect the decomposition of leaves in wetlands.

However, the impact of these factors has not been specifically addressed. Sediment

drying has also been reported to induce the formation of acidic conditions where sulfidic

sediments are present (McCarthy, Conallin et al. 2006), which is likely to impact all

wetland biota in profound ways, including the decomposer community.

2.3.2 Temperature

Temperature affects the decomposition of plant material in wetlands through its influence

on the activity of the microbial community. The metabolic rate of micro-organisms

approximately doubles with each 10°C rise in temperature, then begins to plateau before

the temperature reaches a critical value where the organisms’ proteins begin to denature .

This relationship has been described as the “Q10 factor” (Kirschbaum 1995; Kirschbaum

2006). Many authors have demonstrated faster rates of leaf breakdown under warmer

conditions (e. g. Coûteaux, Bottner et al. 1995; Glazebrook and Robertson 1999; Guo

and Sims 2001; Wood 1974) and others have found higher microbial biomass in warmer

weather (Goncalves, Graca et al. 2007; Scholz and Boon 1993a). Temperature changes

can also trigger reproductive events in aquatic micro-organisms (Thomas, Chilvers et al.

1989). High levels of DOC in combination with high temperatures are known to cause

“black water events”. A hypoxic black water event is a catastrophic decrease in dissolved

oxygen caused by a sharp increase in oxygen demand by bacteria in the water column

(Howitt, Baldwin et al. 2007). Serious ecological consequences, including fish kills, may

ensue.

Temperature was the main driver of coarse woody debris decomposition on the floor of

Australian eucalypt forests (Mackensen, Bauhus et al. 2003), explaining more than 30%

of the variability in its decomposition rate (Campbell, James et al. 1992b). This is in line

with findings that refractory forms of organic matter from soils and litter are more

sensitive to changes in temperature than labile organic matter (Conant, Drijber et al.

2008).

2.3.3 Turbidity and sedimentation

Relative to their associated stream, floodplain wetlands are generally more turbid and

more highly coloured (Hillman 1986). Turbidity may be further increased by the activity

of livestock and feral animals (Hart and McGregor 1980). In shallow lakes particles such

14

as sediments and algal cells may go through a cycle of sedimentation and re-suspension

(Scheffer 1998). This continuous deposition of sediments in floodplain wetland

environments results in the burial of organic detritus, such as leaf litter, after it settles on

the sediment surface.

The continued deposition of sediments over leaf litter can alter the quality of the leaves

as a food resource for both invertebrates and micro-organisms. Sedimentation can reduce

the availability of oxygen and nutrients, and limit opportunities for reproduction and

dispersal. Janssen and Walker (1999) found that burial of decomposing leaves affected

breakdown rates only where the leaves were considered to be fast decomposers. The

breakdown of “slow” species was not inhibited by burial.

2.3.4 Salinity and pH

Where connectivity with the parent stream is relatively frequent, floodplain wetlands

have water which is extremely fresh, varying from 50 to 2250 μS.cm-1

(Goonan, Beer et

al. 1992; Hillman 1986). Salinity begins to influence the macro-biota at levels around

1000 μS.cm-1

(Goonan, Beer et al. 1992; Hart, Bailey et al. 1991; Nielsen, Brock et al.

2003). Where connectivity is less frequent, in terminal wetlands or where there are

inflows of saline groundwater, high salinity (James, Hart et al. 2009) and hypersaline

(Reid and Brooks 2000) conditions may occur.

Billabongs exhibit a wide range of pH values, and large changes can occur on a daily

basis. This range is so large that it is certain to affect chemical equilibria within the water

column and sediments, however, more than 80% of wetlands have mean pH values close

to neutral (Hillman 1986). Rapid changes in pH have been recorded in association with

decomposition, and short-lived increases in photosynthesis (Tan and Shiel 1993).

Flooding may also lead to reduced oxidation states and shifts in pH that can reduce the

availability of some nutrients (Mitsch and Gosselink 1993a).

The presence of sulfides in the sediments of floodplain wetlands provides the potential

for the development of acid sulfate sediments. Where wetlands have been inundated for

extended periods and where source water is of poor quality, salinity is high, soil pH is

low or oxidised iron is visible on the sediment surface, oxidation of the sediments as a

result of drying can result in the development of acid (Baldwin, Hall et al. 2007). This

15

has catastrophic effects on wetland biota (McCarthy, Conallin et al. 2006). It is believed

that the ingress of saline groundwater inhibits activity by methanogenic bacteria and

favours sulfate reducing bacteria. Where sulfate reducing bacteria consume the bulk of

sediment carbon, hydrogen disulfide can build up in the sediments. Oxidation leads to the

formation of sulfuric acid (Hall, Baldwin et al. 2006).

2.3.5 Oxygen availability and redox potential

Oxygen demand in wetlands can be as high as 2.7 gO2.m-2

.day-1

(Robertson, Healey et al.

1997), leading to low dissolved oxygen concentrations in the water column and

sediments (Ford, Boon et al. 2002). This may limit aerobic decomposition in the

sediments to a surface layer of less than 1cm thickness (Cappenberg 1988), or exclude it

entirely. Iron oxide mediated photochemical degradation of organic carbon in the water

column also contributes to oxygen demand (Howitt, Baldwin et al. 2004). As discussed

earlier, high oxygen demand may accompany floods in floodplain forest wetlands, as the

inundation of floodplain litter leads to sudden increases in DOC.

It is generally accepted that decomposition in wetlands occurs more slowly under anoxic

conditions (Brinson, Lugo et al. 1981; Neckles and Neill 1994). In most cases, however,

there are many factors involved in determining decomposition rates, and the importance

of oxygen is variable. Godshalk and Wetzel (1978b) found slower breakdown of POM

under anoxic conditions but concluded that the effect of dissolved oxygen was minor in

comparison with temperature and the initial nitrogen concentration of the substrate. This

was in contrast to findings that DOC decomposition was strongly influenced by oxygen

concentrations (Godshalk and Wetzel 1978a). While anoxia slows the degradation of

plant detritus on inundated floodplains, the rate of DOC uptake from the water column is

unaffected by anoxic conditions (O'Connell 1998).

While fungi may be present in anoxic environments, there is no published work

demonstrating that they are able to participate actively in decomposition under these

conditions. Fungal growth is severely limited below 4% oxygen saturation in

waterlogged soils and 2% saturation in decaying wood (Swift, Heal et al. 1979), and

activity ceases with the onset of anoxia (Lopez-Real and Swift 1977). Research has

shown that submerged E. camaldulensis leaves are colonised by aerobic fungi under oxic

conditions, while under anoxic conditions they are colonised by anaerobic bacteria

16

(O'Connell, Baldwin et al. 2000). Therefore, the alleviation of sediment anoxia by

wetland drawdown and drying may lead to more complete decomposition of plant

material in wetlands, while continuing sediment anoxia will restrict decomposition to

anaerobic pathways.

2.3.6 Nutrient availability

Murray River billabongs are nutrient rich environments in terms of nitrogen and

phosphorus (Goonan, Beer et al. 1992; Gribben 2000), and billabongs in other areas have

also been shown to be nutrient rich (Hart and McGregor 1980; Leahy, Tibby et al. 2005).

Nutrient enrichment accelerates leaf breakdown in streams (Suberkropp 1995;

Suberkropp and Chauvet 1995; Webster and Benfield 1986), and higher rates of litter

decomposition in a Murray River floodplain wetland have recently been correlated with

higher nutrient concentrations (Watkins, Quinn et al. 2010). In streams, higher nutrient

concentrations may result in decomposing leaves becoming nutrient enriched without

observed changes in mass loss (Elwood, Newbold et al. 1981) and fungal activity,

respiration and sporulation rates may also increase (Suberkropp 1991b). Micro-

organisms are also known to use solid substrates and DOC differently in waters of

differing trophic status (Baldy, Gobert et al. 2007).

The decomposition of leaf material results in net losses of nitrogen, phosphorus, calcium,

magnesium and potassium (Day 1983). Losses of nitrogen have been observed within 3

days of immersion (Glazebrook and Robertson 1999). Inundation of wetland sediments

after a period of drying results in an initial flush of nitrogen from the sediments to the

water column (Baldwin and Mitchell 2000), and additional nitrogen leached into the

water from plant litter may lead to high concentrations of dissolved nitrogen.

The nitrogen content of the substrate can also influence decomposition rates (Gessner

and Chauvet 1994). The leaves of sclerophyllous plants such as eucalypts have C:N

ratios higher than 100:1 (Boon 2006; Glazebrook and Robertson 1999). When using

these leaves as a substrate, large amounts of external nitrogen may be needed to sustain

microbial metabolism (Boon 2006). Aquatic hyphomycete fungi are capable of utilizing

dissolved inorganic nitrate and ammonium from the water column (Thornton 1963) to

address this shortfall. Thus, submerged leaf litter colonised by aquatic fungi may become

enriched with nitrogen (Glazebrook and Robertson 1999).

17

While bioavailable forms of phosphate are leached from leaves when they are submerged

(Baldwin 1999), fungi colonising leaves are able to take up phosphate from the water

column. This can result in observed relative increases in leaf phosphorus content as

leaves undergo aquatic decomposition (Glazebrook and Robertson 1999). Leaf break

down proceeds faster where phosphate concentrations are higher (Chauvet 1987; Meyer

1980; Rosset, Bärlocher et al. 1982; Suberkropp 1991b; Suberkropp, Godshalk et al.

1976), and this is likely to be due to a stimulating effect of phosphorus on microbial

activity.

The re-wetting of wetland sediments can release a pulse of phosphorus into the water

column (Baldwin and Mitchell 2000). While the origin of this phosphorus is likely to be

plant litter (Qiu, McComb et al. 2003), increased moisture availability in dry sediments

leads to increases in microbial biomass, with a corresponding increase in phosphorus

derived from microbial sources (Qiu, McComb et al. 2003). Thus, the flux of phosphorus

from wetlands sediments on rewetting is regulated by microbial activity.

Dissolved organic carbon can undergo photodegradation reactions. Leachate from E.

camaldulensis leaves can be chemically degraded upon exposure to light, a reaction

which results in CO2 production and reduced concentrations of aromatic structures and

double bonds (Howitt, Baldwin et al. 2004). Metal ions in solution can increase the rate

at which naturally occurring DOC undergoes photolysis (Howitt, Baldwin et al. 2004;

Zepp, Callaghan et al. 1998; Zepp, Faust et al. 1992). DOC photodegradation rates

increase when iron oxide is present in the water column (Howitt, Baldwin et al. 2004).

Humic and fulvic substances in solution are able to reduce Fe(III) to Fe(II), a reaction

which also produces CO2 (Miles and Brezonik 1981; Voelker, Morel et al. 1997). As

freshly produced iron oxides are most reactive, photodegradation is especially relevant in

wetlands and shallow lakes where anoxic layers in the water column may form in warmer

weather. The mixing of oxic and anoxic waters is likely to generate fresh iron oxides,

which may further increase oxygen consumption (Howitt, Baldwin et al. 2004). Under

the anoxic conditions found in submerged sediments and reduced soils, fungi with

metabolic requirements for manganese may also be present (Aguiar, de Souza-Cruz et al.

2006; Jennings and Lysek 1996; Kapich, Steffen et al. 2005; Keishi, Yasoo et al. 2006;

Miyata, Tani et al. 2004; Thompson, Huber et al. 2005; Wainwright 1992).

18

2.3.7 Litter type / substrate quality

The quality of organic matter has a strong influence on decomposition rates, and on the

decomposer community structure (Gulis 2001). Decomposition rate varies with the

proportions of labile and refractory organic substances in plant litter. For example,

decomposition rate has been found to vary with proportions of hemicellulose, cellulose

and lignin in wetland macrophytes (Gessner and Chauvet 1994; Godshalk and Wetzel

1978b; Suberkropp, Godshalk et al. 1976; Webster and Benfield 1986) and in the leaves

of riparian trees (Janssen and Walker 1999). High lignin content also confers slower

decomposition, with twigs decomposing more slowly than leaves (Shearer and von

Bodman 1983). The seasonality of litter fall may also be a critical factor in litter

decomposition. This will affect the length of time litter is air dried on the floodplain

before inundation, the temperature at inundation and whether leaves are still green when

they are inundated (Boon 2006). Leaf age has been shown to strongly influence the

speciation and bioavailability of organic matter leached from the leaves of Eucalyptus

camaldulensis (Baldwin 1999).

Litter may be derived either from within the wetland (autochthonous carbon) or from

sources outside the wetland, i.e. the floodplain (allochthonous carbon). Autochthonous

carbon sources are largely derived from macrophytes and algae, while allochthonous

carbon consists largely of the litter of riparian vegetation. Autochthonous carbon sources

in Australian floodplain wetlands, such as Phragmites sp and Typha sp., have leaves

which senesce in situ and begin the decomposition process in what is known as the

“standing dead” position (Aseada, Nam et al. 2002; Gessner 2001; Kominkova, Kuehn et

al. 2000). This group of plants forms a large proportion of the biomass produced in

wetlands, and on the shores of lakes and rivers (Kuehn and Suberkropp 1998a;

Pieczynska 1993). Fungi play a dominant role in the decomposition of standing dead leaf

litter (Gessner 2001; Kuehn, Lemke et al. 2000; Kuehn and Suberkropp 1998a; Newell

2001; Newell, Moran et al. 1995; Van Ryckegem, Gessner et al. 2007). Rates of standing

dead litter decomposition have been reported to be in the same order of magnitude as

plant production rates, leading to minimal litter accumulation (Jackson, Foreman et al.

1995).

Algae are another significant autochthonous carbon source in wetlands. In contrast to

macrophytes, algal production is often consumed directly by zooplankton and

19

invertebrates in wetlands (Boon 2000; Bunn and Boon 1993). During algal blooms, large

amounts of algal biomass may precipitate to the sediment surface. These cells are quickly

decomposed because they lack the structural carbohydrates that are characteristic of

higher plants.

The leaves of Eucalyptus sp. comprise a large proportion of the allochthonous carbon

reaching floodplain wetlands in Australia. In the aquatic phase they are colonised by both

fungi and bacteria. The activity of fungal decomposers on the leaves of E. globulus has

been compared with that on European leaf species (Bärlocher and Graça 2002; Canhoto

and Graça 1996; Chauvet, Fabre et al. 1997). The main difference between these leaf

types is the composition of the fungal community. While European species support an

‘autumn’ community, E. globulus leaves support ‘summer’ species, and these

communities correspond with the season of leaf accession (Bärlocher and Graça 2002). A

delay in colonisation relative to European leaf species (Canhoto and Graça 1996), and the

use of the stomata as a point of access to leaf mesophyll have also been reported

(Canhoto and Graça 1999).

2.3.8 Carbon compounds inhibiting biological processes

Plant tissues can also contain substances that are capable of inhibiting growth and

activity in micro-organisms (Bärlocher and Oertli 1978). These include tannins, phenolic

compounds and essential oils (Boon and Johnstone 1997). These substances may act

synergistically to regulate microbial distribution and community structure. Antifungal

compounds provide plants with resistance to fungal infection, especially in woody tissues

(Cromack and Caldwell 1992; Mackensen, Bauhus et al. 2003). Leaching removes many

of these inhibitory substances from dead plant material, and may enhance fungal

colonisation (Bärlocher, Canhoto et al. 1995; Canhoto and Graça 1996; Dix and Webster

1995). Some fungi are able to neutralise these compounds with laccases and phenol

oxidases in order to degrade lignin (Dorado, van Beek et al. 2001; Jönsson, Palmqvist et

al. 1998; Leonowicz, Cho et al. 2001).

Tannins are a heterogeneous group of phenolic compounds defined by their ability to

precipitate proteins and react with ferric chlorides (Boulton and Boon 1991). They are

also able to complex with cellulose and hemicellulose (Zucker 1983), making them more

resistant to decomposition (Boulton and Boon 1991). The tannin content of plant detritus

20

influences the rate of leaf processing due to fragmentation by invertebrates and microbial

conditioning (Campbell and Fuchshuber 1995; Canhoto and Graça 1995; Gessner and

Chauvet 1994; Suberkropp, Godshalk et al. 1976).

Tannins may be either hydrolysable or condensed. While both types have inhibiting

effects on fungal colonisation, they differ in the mechanism of inhibition. Condensed

tannins are able to inhibit both enzyme activity and growth, while hydrolysable tannins

inhibit growth only (Nichols-Orians 1991). Their inhibitory effects can be attenuated in

the presence of Al3+

, Fe3+

or Mn2+

ions (Marschner and Kalbitz 2002). However,

Juntheikki and Julkunen-Tiitto (2000) found that while condensed tannins are able to

bind with hydrolytic enzymes and precipitate them, enzyme activity may remain

unaffected.

The role of polyphenols as defensive compounds against fungi in aquatic environments is

generally accepted (Bärlocher, Canhoto et al. 1995; Bunn 1988b; Canhoto and Graça

1999). Leaching of soluble phenols from leaves can result in faster microbial

colonisation (Bunn 1988b). The polyphenolics in eucalypt leaves have a deterrent effect

on invertebrate shredders and micro-organisms (Canhoto and Graça 1995; Canhoto and

Graça 1999; Graça, Pozo et al. 2002). This is thought to be due to their ability to bind

fungal and invertebrate digestive enzymes and block digestion (Graça, Pozo et al. 2002).

Fungal sensitivity to phenols seems to be species specific (Canhoto and Graça 1999),

partly explaining why distinct fungal communities develop on eucalyptus leaves, as

opposed to deciduous European species (Alnus glutinosa, Castanea sativa and Quercus

faginea)(Canhoto and Graça 1996).

Terrestrial, wetland and littoral plants have been found to be a major source of

polyphenols in temporary wetlands (Serrano 1992). While large influxes of polyphenols

to wetlands due to leaching from terrestrial litter have been correlated with rainfall events

(Serrano 1992), polyphenols are also a by-product of the decomposition of lignin (Kirk

and Farrell 1987) and tannins (Serrano 1992). Oxidative breakdown of lignin by fungal

enzymes produces quinones (6-membered rings with 2 carbonyl groups), which are able

to complex with proteins, polypeptides, amino acids and amino sugars via the amino

group (Suberkropp, Godshalk et al. 1976). This complexation provides resistance to

decomposition (Dix and Webster 1995).

21

Leaf essential oils are used by the plant as a defence against the activities of

microorganisms and herbivores (Stone and Bacon 1994; Terzi, Morcia et al. 2007).

Eucalypt essential oils may depress or completely inhibit the growth of selected aquatic

hyphomycetes, even more effectively than polyphenols (Canhoto and Graça 1999; Graça,

Pozo et al. 2002). This is probably due to their interference with fungal enzymes

(Canhoto, Bärlocher et al. 2002). In Australian systems, high concentrations of essential

oils have also been shown to inhibit the decomposition of leaf litter (Bailey, Watkins et

al. 2003; Boon and Johnstone 1997). The anti-fungal ingredient of eucalyptus oils is 1,8-

cineole (Terzi, Morcia et al. 2007). The leaves of Eucalyptus camaldulensis have a 1,8-

cineole content in the range of 12.7 to 71.7% of total terpenoid yield (the overall mean

terpenoid yield was 1.86% , w/w, leaf dry wt). Leaves with the higher proportion of 1,8-

cineole were found to be more resistant to attack by grazing insects (Stone and Bacon

1994).

The oils within Eucalyptus leaves can remain intact well after senescence and abscission

because they remain locked within oil vesicles. These vesicles become leaky over time,

so these oils can continue to inhibit microbes and invertebrates for some time after

immersion (Canhoto, Bärlocher et al. 2002). Chironomid larvae have been observed to

recognise and avoid oil vesicles, while Tipula larvae showed no indication of similar

abilities (Canhoto and Graça 1999).

2.4 The Wetland Biota

Biodiversity in floodplain wetlands (billabongs) may exceed that found in the adjacent

channel (Bornette, Amoros et al. 1998). High densities of pelagic and epiphytic algae are

common, and the phytoplankton can be diverse (Gell, Sluiter et al. 2002; Reid and

Ogden 2009; Shiel 1983). Similarly, these water bodies carry distinctive and diverse

zooplankton communities (Hillman 1986; Nielsen, Hillman et al. 2002; Ning, Nielsen et

al. 2008; Tan and Shiel 1993). The billabong environment favours macrophyte growth,

and the banks are usually well vegetated and lightly grazed when compared with stream

banks (Hillman 1986). Macrophyte communities exhibit considerable diversity, and vary

widely between wetlands (Brock 1994; Reid and Quinn 2004; Robertson, Bacon et al.

2001).

22

Wetland macro-invertebrate communities differ substantially from those found in

streams (Balcombe, Closs et al. 2007; Boulton and Lloyd 1991; Hillman and Quinn

2002), and often display high diversity (Goonan, Beer et al. 1992). Communities vary

between wetlands, e.g. between billabongs with different hydrological regimes (Quinn,

Hillman et al. 2000; Sheldon, Boulton et al. 2002), but are similar where wetland

characteristics are similar (Marchant 1982).

Wetland fish communities tend to be dominated by the European carp (Cyprinus carpio)

other introduced species such as Redfin (Perca fluviatilis), Tench (Tinca tinca) and

Mosquito fish (Gambusia affinis) and small native species such as Western Carp

Gudgeon (Hypseolotris klunzingeri) and Flathead Gudgeon (Philypnodon grandiceps)

(Ellis and Meredith 2004; Hillman 1986; McNeil and Closs 2007).

Billabongs and other floodplain wetlands are also home to waterfowl in large numbers

and considerable diversity (Kingsford and Norman 2002), frogs (Healey, Thompson et

al. 1997), reptiles (e.g. red bellied black snake (Pseudechis porphyriacus) (Shine 1987)

and long necked tortoise (Chelodina longicollis) (Kennett, Roe et al. 2009)), platypus

(Ornithorhynchus anatinus) and water rats (Hydromys chrysogaster) (Grant and Temple-

Smith 1998; Hillman 1986; Reid and Brooks 2000; Woollard, Vestjens et al. 1978).

2.4.1 River red gums

In temperate regions of eastern Australia, the dominant tree species on the margins of

floodplain wetlands is the river red gum (Eucalyptus camaldulensis). River red gums

generally occur as woodlands (Bacon, Stone et al. 1993), but may form dense forests in

flood prone areas adjacent to rivers. Thus, the plant litter on floodplains in these regions

is often dominated by the leaves of this species (Glazebrook and Robertson 1999;

Robertson, Bunn et al. 1999). In addition, wood production from floodplain forests can

yield up to 900 gC.m-2

.yr-1

(Briggs and Maher 1983).

E. camaldulensis litter, composed of leaves, twigs, bark and fruit, often in large

quantities (Glazebrook 1995), is one of the major sources of DOC in Australian lowland

rivers (O'Connell, Baldwin et al. 2000). This litter can account for up to 290 gC.m-2

.yr-1

(Briggs and Maher 1983; Robertson, Bunn et al. 1999). The wood of this species

decomposes slowly (O'Connor 1992), with an estimated life on the floodplain of 375

23

years (Mackensen, Bauhus et al. 2003). This creates a large pool of woody debris on the

floodplain and within aquatic environments (MacNally, Parkinson et al. 2002).

Physico-chemically, wood differs substantially from leaves. Wood has a higher carbon to

nitrogen ratio, higher density, lower concentrations of pectins, lower surface to volume

ratio and lower parenchyma to sclerenchyma ratio (Erickson, Blanchette et al. 1990c). As

a carbon resource, submerged wood is variable in size, while leaves have more or less

uniform size. The input of wood to the aquatic environment is episodic, while leaf inputs

are seasonal. Maximum litter inputs occur in summer, although leaf accession occurs

throughout the year in lower amounts (Glazebrook and Robertson 1999). Woody

substrates also have a much longer residence time in aquatic habitats (Shearer 1992).

Thus, wood and leaves are vastly different substrates for the aquatic decomposer

community.

In terrestrial and aquatic environments wood pieces decay from the outer layer inwards,

and decay rates increase exponentially towards centre layers as the wood begins to crack

and split (O'Connor 1992). However, overall decay is relatively slow, and woody debris

is consistently present on the floodplain over time. Thus, wood serves as a reservoir of

microbial propagules. For example, the period required for twig break down can exceed

390 days (Thomas 1992). Over this period, newly available resources can be quickly

colonised from pre-existing populations, and seasonal changes in resource availability

have little influence on the establishment of new populations.

Leaves represent a more transient resource than wood. E. camaldulensis leaves take 2-4

months to break down in the aquatic phase, with a half life of 80 days (Briggs and Maher

1983). The leaves decay according to an exponential decay pattern at rates similar to

other eucalypt and temperate hardwood species (Briggs and Maher 1983). Results

comparing decay rates in lentic and lotic systems have been varied. Glazebrook (1995)

observed slower break down in lentic sites of a forested wetland. Contrary to this,

Janssen and Walker (1999) found that the break down rate of river red gum litter did not

differ between stream and wetland sites. These results may be influenced by season,

which corresponds with differences in dissolved oxygen, temperature and moisture

availability. As rate of decay increases with temperature, maximum E. camaldulensis leaf

inputs generally coincide with maximum temperatures, and therefore maximum decay

24

rates. However, higher temperatures may also limit dissolved oxygen concentrations and

lead to wetland drying. Litter inputs may also increase if eucalypt trees experience

moisture stress (Stone and Bacon 1994; Stone and Bacon 1995).

2.5 Wetland Carbon Cycles

On a global scale, wetlands are important in the biogeochemical cycling of carbon

(Westermann 1993). Wetlands can be extremely productive systems but grazing of this

primary production is limited because dead plant material is comprised primarily of

complex plant polysaccharides and lignins (Benner, Maccubbin et al. 1984; Gessner,

Schieferstein et al. 1996; Wetzel 1990; Wetzel and Howe 1999) and is not easily

digested. This means that much of the carbon incorporated into plant biomass is not

consumed by higher trophic levels, but passes into the detrital food web (McLatchy and

Reddy 1998). For example, Phragmites australis (Cav.) Trin. Ex Streud. is high in ligno-

cellulose and reed stands of this species may have net above ground production in excess

of 1 kg dry mass per square metre of surface area (Gessner 2001). As a consequence

most of the biomass from these plants is shed as litter. This litter enters the detrital pool

where it is decomposed by fungi, bacteria and invertebrates (Jackson, Foreman et al.

1995; Wetzel 1990). The insoluble fraction of plant detritus that is not mineralised is

stored as sediment organic carbon.

Wetland soils contain about one third of all of the organic matter that is stored in the

world’s soils (Mitsch and Gosselink 1993a). If one accepts the Ramsar definition of

wetlands (Convention on Wetlands (Ramsar)), 40% of global terrestrial carbon may be

stored in wetlands. Peatlands and forested wetlands are particularly important as carbon

sinks. Deforestation and agricultural use of these wetlands accelerates the degradation of

the soil humus, causing significant flux of greenhouse gases to the atmosphere (Aiken,

McKnight et al. 1985).

The most common conceptualisation of the wetland food web is that of the “grazing food

chain”, where phytoplankton are the primary producers that are consumed by

zooplankton, which are then consumed by fish (Elton 1927). While this is occurring,

however, the metabolic activities of micro-organisms are utilising much of the available

carbon in the wetland. These resources are in the form of living and dead phytoplankton

and plants and non-living carbon compounds. As these resources are mineralised gaseous

25

carbon compounds are produced, and released into the atmosphere. Residual carbon, that

carbon that is not respired or incorporated into biomass, is stored in the sediments and

water column. This carbon is referred to as humic and fulvic substances. Residual

biomass in plant detritus, and the biomass of decomposer organisms, is now known to be

as important as primary production as a basal resource in aquatic food webs (Hessen

1998; Pace, Cole et al. 2004; Reid, Quinn et al. 2008).

Biomass undergoes both physical and chemical changes as it undergoes decomposition.

It is partitioned by particle size into POC and DOC. POC may be further classified as

either fine (FPOC) or coarse (CPOC) (e.g. leaf litter and woody debris). Decomposition

of DOC and FPOC in floodplain wetlands is largely mediated by bacteria (Baldwin 1999;

Boon 2006), while fungi dominate the aquatic decomposition of CPOC (Suberkropp

1991a; Suberkropp and Klug 1976) under oxic conditions (O'Connell 1998).

The fluxes and storages in a typical wetland carbon cycle are illustrated in Figure 2.1,

which is based on a diagram published by Webster and Benfield (1986). It shows how

inorganic carbon from the atmosphere is incorporated into biomass via photosynthesis.

This plant biomass also produces carbon dioxide and acts as a conduit for methane to

pass from the sediments into the atmosphere. As this plant biomass senesces and dies,

carbon is leached into the water column. Carbon is also leached from allochthonous litter

and floodplain soils. The resulting DOC is mineralised by aerobic or anaerobic bacteria

in the water column, depending on the availability of dissolved oxygen. Carbon dioxide

respired by these bacteria is either dissolved, escapes to the atmosphere, or is converted

into other inorganic carbon ions, depending on the pH of the water. Insoluble carbon

compounds from plant, algal and animal detritus are mineralised by fungi and bacteria

under submerged conditions. Residual carbon passes into the surface sediments, thence

to anoxic sediments below the surface where carbon mineralisation is dominated by

aerobic and anaerobic bacteria, respectively. These processes are discussed in greater

detail below.

26

Figure 2.1: A stylised representation of the carbon compartments and fluxes found in most wetland

systems. Based on Webster and Benfield, 1986.

2.5.1 Primary and secondary production

The major route via which carbon dioxide is transformed into organic carbon is

photosynthesis (by both plants and algae). Aquatic macrophyte production in wetlands

may be very high (Bott, Barnes et al. 1983; Briggs and Maher 1985; Roberts and Ganf

1986) and is often stimulated by wetland drying and re-wetting (Briggs and Maher 1985).

Litter production by E. camaldulensis in Australian floodplain wetlands is in the order of

120 - 700 g dry weight.m-2

.yr-1

(Briggs and Maher 1985; Campbell, James et al. 1992a).

This represents a substantial contribution of carbon to the wetland systems and floodplain

soils. The presence of aquatic macrophytes may also influence the flow of methane from

the sediments to the atmosphere (Boon 2000; Boon and Mitchell 1995; Boon and Sorrell

1991; Sorrell and Boon 1992a).

The river red gum (E. camaldulensis) is the dominant tree species around many

Australian floodplain wetlands. The total litterfall from the river red gum is similar to

other eucalypt species (~400 g.m-2

.yr-1

), except there is a lower proportion of leaves

(~25%) (Briggs and Maher 1983). Reid et al.(2008) measured direct inputs of litter into

27

streams which had riparian vegetation dominated by E. camaldulensis. They found litter

inputs were 550 – 617g AFDW.m-2

.yr-1

, with lateral litter movement accounting for

<10% of total inputs. This species sheds more litter in dry periods, and this generally

corresponds with periods when wetlands are dry. Standing stocks of river red gum litter

on a reach of the Darling River, NSW, were found to be 2.7 kg/m2 for twigs, 1.5 kg/m

2

for bark, 1.5 kg/m2 for leaves, and 0.8 kg/m

2 for gum nuts (Francis and Sheldon 2002).

The productivity of decomposer micro-organisms themselves can also be considered as

an important carbon sink (Buesing and Gessner 2006), although less persistent than plant

carbon (Findlay, Howe et al. 1990). In a freshwater marsh in Switzerland, Buesing and

Gessner (2006) found that the annual productivity of bacteria on submerged plant litter

under aerobic conditions far outweighed fungal production. Annual productivity was

18.8 and 2.4 mg C.g-1

C for bacteria and fungi respectively. Together, bacteria and fungi

accounted for ~1500 g C.m-2

.yr-1

, while plant productivity in the same wetland was ~600

g C.m-2

.yr-1

. This indicates that the microbial community are important producers in

wetlands.

Food chains represent the dominant pathway for carbon transformation in many systems.

In wetlands, trophic relationships are complex. Bunn and Boon (1993) used stable

isotopes to investigate food webs in Australian floodplain wetlands (billabongs or ox-

bow lakes). Based on the premise that consumers are slightly (~1‰) 13

C-enriched when

compared to their food source, Cherax sp. (a freshwater crayfish) was identified as a

major consumer of detrital carbon. However, they found that all the major consumer

groups were 13

C-depleted when compared to the riparian and macrophyte vegetation,

implying that an unknown 13

C-depleted carbon source was being consumed

simultaneously with vegetation. Methanotrophic bacteria and planktonic Chlorophyta

were suggested as possible sources of this carbon. While fungi tend to be 13

C-enriched

relative to their substrate, this varies with substrate (Mayor, Schuur et al. 2009), between

species and with growth stage (Henn, Gleixner et al. 2002), and between respiration and

fermentation metabolic pathways (Henn and Chapela 2000). This means that fungal

decomposers remain as possible candidates for this unknown carbon source. Moreover,

Reid et al. (2008) found that biofilms (comprised of fungi, bacteria and algae) on

submerged leaves of Eucalyptus sp. were an important carbon source in lowland streams.

Given that fungal tissues provide nutrients vital to the diets of aquatic invertebrates (see

28

section 2.2.1), aquatic saprobic fungi provide a conduit for carbon flow between

submerged litter and wetland consumers.

2.5.2 Sources of DOC

Vegetation is an important source of dissolved carbon (DOC) in aquatic systems. Of the

particulate ligno-cellulose derived from wetland plants, 10-40% is converted to DOC

prior to being consumed by micro-organisms (Benner and Opsahl 2001). Up to 25% of

the carbon in river red gum leaves is readily leachable upon inundation (Glazebrook and

Robertson 1999). Francis and Sheldon (2002) calculated that red gum litter had the

potential to release 235g/m2 of DOC, 80% of which would be derived from the leaves.

Large amounts of DOC are also leached from these leaves on the floodplain prior to

entering the aquatic environment (Baldwin 1999). Both terrestrial and aquatic leaching

result in losses of polyphenols and essential oils from leaves (Serrano and Boon 1991).

These compounds inhibit enzymatic processes in invertebrates and micro-organisms

(Bengtsson 1992; Canhoto and Graça 1995; Canhoto and Graça 1999; Serrano and Boon

1991). For this reason, floodplain aging may be an important factor influencing the rate

of microbial colonisation of leaf litter once submerged.

In the Murray-Darling Basin, biomass and concentrations of DOC in wetlands may be up

to three orders of magnitude greater than in the river channel (Boon 1991; Briggs, Maher

et al. 1993; Briggs and Maher 1985; Briggs, Maher et al. 1985; Hillman and Shiel 1991)

and DOC has been found to be negatively correlated with water level (Briggs, Maher et

al. 1993; Lloyd, Atkins et al. 1992), indicating evaporative concentration.

Laudon et al. (2004) showed that the export of DOC from boreal catchments was

positively correlated with the proportion of each catchment designated as wetlands. In

particular, the total area of wetlands had a strong effect on the seasonality of DOC

exports. Results such as these suggest that the timing and extent of DOC imports into

aquatic environments may be altered as a response to global climate change (Tranvik,

van Hees et al. 2004).

In the stationary waters of wetlands pulsed litter inputs and the associated rapid leaching

can result in profound changes in water quality. The most important of these changes is

the rapid decrease in dissolved oxygen, as aerobic organisms in the water column

29

metabolise DOC and use up oxygen in the process. Leaf aging on the floodplain prior to

flood events reduces their contribution to DOC with flood onset (Baldwin 1999).

Permanent inundation of litter accelerates decomposition relative to permanently dry

conditions (Langhans and Tockner 2006). However, cyclic flooding and drying leads to

faster decay relative to permanent flooding in warmer climates (Battle and Golladay

2001; Ryder and Horwitz 1995), while no difference was found in cooler climates

(Inkley, Wissinger et al. 2008). Accelerated decay in winter relative to summer has also

been reported (Langhans and Tockner 2006). This suggests interplay between inundation,

temperature and dissolved oxygen concentrations that ultimately dictates aquatic litter

breakdown rates.

The concentration of DOC in unpolluted river water is commonly 1–5 mgC.L-1

, but may

reach 10 mgC.L-1

. Concentrations of labile DOC in this range may be turned over twice a

day by planktonic bacteria (Findlay and Sinsabaugh 1999). Previously, about 25% of

DOC was considered biodegradable (Volk, Volk et al. 1997). However, O’Connell et al.

(2000) found that 40% of the DOC released from river red gum leaves was used rapidly

by planktonic bacteria and there is evidence that bacteria are also able to use higher

molecular weight compounds including polyphenols and ligno-cellulose (Findlay and

Sinsabaugh 1999). High extracellular enzyme activity in the water column of Australian

billabongs has been attributed to the metabolic activity of bacteria metabolising DOC

(Boon 1989; Boon 1990; Boon and Sorrell 1991; Scholz and Boon 1993a; Scholz and

Boon 1993b).

In addition, litter and woody debris from riparian eucalypt forests and woodlands, which

is slow to decompose (Hutson and Veitch 1985; Mackensen, Bauhus et al. 2003; Nagy

and MaCauley 1982; Presland 1982), can build up on the floodplain and dry wetland

sediments. These wetlands consequently hold significant reservoirs of carbon, in both

living and non-living forms. During a flood event 25-30% of the leaf mass can be

transformed into DOC (Glazebrook and Robertson 1999; O'Connell 1998).

2.5.3 Utilisation of DOC in the water column

Research conducted in marine environments has shown that DOC is utilised by bacteria.

This has become known as the microbial loop (Azam and Cho 1987; Meyer 1994;

Pomeroy and Wiebe 1988), and bacteria utilising DOC may account for up to 50% of

30

oceanic production (Azam, Fenchel et al. 1983; Cole, Findlay et al. 1988). Mann and

Wetzel (1995) found that the microbial loop was operating in a riverine wetland system,

particularly in the surface waters. Enzyme studies in Australian floodplain wetlands

(Boon 1989; Boon 1990; Boon and Sorrell 1991; Scholz and Boon 1993a; Scholz and

Boon 1993b) support this finding. Bacterial production is known to be grazed by

zooplankton in marine and freshwater systems (Calbet and Landry 1999; Sanders, Porter

et al. 1989; Sherr and Sherr 2002; Sherr and Sherr 1991; Thorp and Delong 2002), and

this links the microbial loop into the microbial food web and classical aquatic food

chains (Lenz 1992).

Substances leached from submerged plant litter and soils are retained in the aquatic

environment in the form of DOC in the water column and pore-water of the sediments.

This DOC is comprised largely of humic and fulvic substances. The bio-available

fraction, mainly low-molecular weight compounds, is rapidly metabolised by bacteria

(Baldwin 1999; Findlay and Sinsabaugh 1999; Fischer, Mille-Lindblom et al. 2006;

O'Connell 1998). Burns and Ryder (2001b) showed that microbial enzyme activity in

river and wetland sediments increased sharply in response to pulses of DOC from flood

waters. Where bacteria are most active, high molecular weight refractory DOC tends to

accumulate, while the activity of fungi leads to the accumulation of DOC of intermediate

molecular weight (Fischer, Mille-Lindblom et al. 2006). These refractory components of

the DOC undergo photodegradation (Howitt, Baldwin et al. 2004; Moran and Zepp

1997), a process that tends to produce more biologically available carbon compounds.

This may stimulate further microbial activity (Howitt 2003), but photodegradation is

limited to the photic zone (Moran and Zepp 1997).

DOC is comprised largely (75%) of humic acids, with the remainder being composed of

carbohydrates, fulvic acids, low molecular weight organic acids and amino acids (Hood,

Williams et al. 2005; Thurman 1985) and is the dominant form of organic matter in the

water column. When the water column is well oxygenated, both bacteria and fungi are

capable of exploiting DOC, with phenyl oxidase producing fungi being particularly

important in the degradation and transformation of aquatic humic substances (Claus and

Filip 1998). However, fungal biomass in the water column is very low (Libkind, Brizzio

et al. 2003; Wurzbacher, Bärlocher et al. 2010), and comprised mostly of yeasts and

parasitic forms (Lefevre, Bardot et al. 2007). Thus, on the balance of available

31

information, bacteria are more important in utilising DOC in the water column of

wetlands than fungi.

2.5.4 Transformations of inorganic carbon

Dissolved inorganic carbon is usually defined as the total concentration of carbon

dioxide, carbonate and bicarbonate. In aquatic environments, transformations between

these inorganic carbon species are highly dependent upon pH, and the chemical processes

involved have been outlined in several texts (e.g. Manahan 1994). Carbon dioxide,

bicarbonate and carbonate may also be reduced by methanogenic bacteria under anoxic

conditions.

2.5.5 Mineralisation of plant detritus

In streams, particulate organic carbon (POC) is consumed by animals such as

invertebrate shredders after first being colonised by micro-organisms including fungi and

bacteria. According to the river continuum concept (Vannote, Minshall et al. 1980)

detritus is processed sequentially by a series of invertebrate groups that utilise it in

different ways. This system relies on the sequential processing of organic matter as it

moves downstream with current. When not connected to the main channel, wetlands are

not influenced by this unidirectional flow of carbon, so POC is consumed and

decomposed in situ. Where wetlands are oligotrophic and aerobic, invertebrates may

consume POC. However many wetlands suffer from anoxia, eutrophication and chemical

extremes such as high salinity and low pH. In these cases, it is likely to be benthic micro-

organisms that will utilise POC.

The oxic zone in wetlands usually consists of the water column and the top layer of the

sediments (a few millimetres) (Alverez, Rico et al. 2001; Boon 2006; Buesing and

Gessner 2006; Mitsch and Gosselink 1993a). The sediment associated with living plant

roots will also be oxygenated. Aerobic micro-organisms require oxygen to act as an

electron receptor in the oxidation of organic compounds, and as a reactant in the

enzymatic breakdown of organic polymers (Brune, Frenzel et al. 2000; Reddy and

DeLaune 2008). Also, when oxygen takes part in a redox reaction, a relatively high

positive reduction potential is achieved.

For a single organism, aerobic decomposition of organic substances yields more energy

than anaerobic decomposition because the organic substrates can be completely oxidised

32

to carbon dioxide rather than an intermediate compound. During the aerobic

decomposition, complex carbohydrate polymers are cleaved by the extracellular enzymes

of micro-organisms into monomers such as glucose. These are then transported into the

organism’s cell where they undergo glycolysis and oxidative phosphorylation. This

process yield 38 ATP molecules per glucose molecule (Lehninger, Nelson et al. 1993)

and micro-organisms are able to capture approximately 30-50% of this energy. By way

of comparison, the anaerobic fermentation of glucose to alcohol and carbon dioxide by

yeasts yields 4 ATP molecules. The various end products of anaerobic carbohydrate

metabolism may be degraded further by a series of anaerobic organisms using a range of

metabolic pathways, ultimately yielding carbon dioxide and/or methane.

Fungi have been characterised as the most important group of micro-organisms

responsible for the aerobic decomposition of plant material in agricultural soils, forest

soils, freshwater sediments and peat lands (Collins, Sinsabaugh et al. 2008; Cornut, Elger

et al. 2010; Freeman, Ostle et al. 2004; Gutknecht, Goodman et al. 2006; Wainwright

1992).

2.5.6 Mineralisation of sediment organic matter under oxic conditions

In wetlands, anoxic and oxic conditions can occur in extremely close proximity. Reduced

oxygen diffusion into soils due to flooding, and high biological and chemical oxygen

demands result in the development of 2 distinct soil/sediment layers; the oxidised surface

layer where aerobic metabolic processes are possible and the reduced lower layer where

respiration is anaerobic (Reddy and DeLaune 2008). Anaerobic respiration is carried out

by sediment bacteria, but it is possible that fungi in the sediments, such as yeasts and

chytrids, may generate energy by fermentation under these conditions (Wurzbacher,

Bärlocher et al. 2010). The water column may also have an anoxic zone, particularly if

temperatures, salinity or biological oxygen demands are high.

In hydrologically dynamic wetlands, such as floodplain wetlands that flood and dry, the

vertical limit with an oxic zone above and an anoxic zone below varies in both space and

time (Boon 2006), moving up and down with water levels. Plant roots and animal

burrows can also create pathways through which oxygen can diffuse. Thus a mosaic of

oxic/anoxic zones can develop in the sediments, and also in the water column, and may

vary daily (Boon 2006).

33

POC on the sediment surface tends to be colonised by anaerobic decomposer organisms,

which are predominantly bacteria. This is, in part, due to the high oxygen demand of

wetland sediments (McLatchy and Reddy 1998).

In ephemeral systems, drawdown may lead to desiccation of sediment organic matter. As

well as influencing microbial activity, desiccation can promote oxidative chemical

reactions (Baldwin and Mitchell 2000). In the presence of molecular oxygen, carbon

substrates are completely degraded to carbon dioxide via the aerobic respiration of

bacteria, fungi, protozoans and wetland fauna. As oxygen is a very strong oxidant, almost

all organic compounds represent possible substrates or food sources.

In the wetland environment oxygen is consumed in preference to other electron acceptors

because aerobic metabolism releases more energy. Even organisms capable of utilising

substrates via other metabolic pathways will use aerobic pathways if possible. Facultative

anaerobes, capable of aerobic respiration and fermentation, are able to use the organic

substrate itself as an electron acceptor in the absence of oxygen. However, not only is

less energy obtained by fermentation, but it results in the production of a by-product that

can not be further metabolised.

Twice the amount of carbon dioxide is generated by decomposer organisms under oxic

conditions as under permanently anoxic conditions (Godshalk and Wetzel 1978a; Reddy

and Patrick 1975). This gas accounts for most of the organic material decomposed under

oxic conditions, with methane emissions being minor. As wetlands are inundated, there is

a flush of inorganic carbon from the sediments, mostly in the form of carbon dioxide.

With ongoing inundation, progressively more methane is evolved (Boon, Mitchell et al.

1997). This may be due to the decreasing availability of oxygen in the waterlogged

sediments and water column due to biological and chemical processes, and the relatively

poor solubility of oxygen in water, especially at elevated temperatures. Shifts from oxic

to anoxic conditions in floodplain wetlands can occur rapidly (Boon and Mitchell 1995).

2.5.7 Mineralisation of sediment organic matter under anoxic conditions

Oxygen depletion is common in wetland sediments (Mitsch and Gosselink 1993a) and

inside wood (Cooke and Whipps 1993). Where oxygen consumption exceeds oxygen

34

diffusion in the sediments, they will become anoxic. This change results in a shift in

microbial metabolism from aerobic to anaerobic pathways. Under anoxic conditions,

bacteria are considered to be responsible for the bulk of biologically mediated

mineralisation processes, and fungi and protozoa are of minor importance.

While fungi are generally regarded as strictly aerobic heterotrophs, a number of studies

have shown that they are able to survive, grow and participate in fermentation reaction

under anoxic conditions. Fungi have been isolated from micro-aerobic conditions in litter

(Reith, Drake et al. 2002), inside wood (Erickson, Blanchette et al. 1990b), and from

blackened leaves in aquatic environments (Field and Webster 1983; Pattee and Chergui

1995).

Many fungal species are facultative anaerobes (Gibb and Walsh 1980). For example, the

facultative anaerobic fungus Aqualinderella fermentans has been isolated from stagnant

ponds and marshes that have low oxygen concentrations (Emerson and Natvig 1981).

The alternating oxic/anoxic conditions common in wetlands may enable facultative

anaerobes to store nutrients required for anaerobic metabolism during their aerobic

growth phase (Cooke and Whipps 1993). Field and Webster (1983) demonstrated that

fungi are able to survive under anoxic conditions in wetlands, but they are unable to

grow.

Apart from lower energy yields, anoxic conditions cause oxidative enzymes to become

inactive. This limits the range of substrates available for metabolism. Lignin, in

particular, is mainly degraded via oxidative pathways. However, long term incubation

experiments have revealed that lignin may be decomposed very slowly by anaerobic

bacteria (Benner, Maccubbin et al. 1984). Moreover, the most reactive and bioavailable

carbon is used in oxic zones (Hulthe, Hulth et al. 1998). The more refractory carbon that

enters the anoxic zone must therefore be used at a slower rate (O'Connell, Baldwin et al.

2000).

Fermentation is carried out by either obligate or facultative anaerobes. Fermentation is

the major anaerobic pathway whereby high molecular weight compounds are broken

down into smaller compounds that can be utilised by other organisms (Mitsch and

Gosselink 1993a; Reith, Drake et al. 2002; Van Laere 1994). Fermentative organisms

35

can utilise substrates such as organic acids, sugars, alcohols and amino acids. End

products can include carbon dioxide, hydrogen, acetate, formate, butyrate, lactate,

succinate, volatile fatty acids, and alcohols. These products may be further oxidised, but

the process of oxidation will depend on the availability of electron receptors, the type of

organic substrates available, and the microbes competing for the substrates. Further

oxidation is carried out by denitrifiers, iron, sulfate and manganese reducers, and

methanogens (Reddy and DeLaune 2008). In general, the process that yields the most

energy from the oxidation of the organic substrate will dominate.

2.5.8 Gaseous carbon emissions

Wetlands may be a carbon source or a carbon sink (Mitsch and Gosselink 1993a). So the

proportion of carbon flowing through the various decomposition pathways is important

in determining the wetland’s net contribution to greenhouse gas emissions. As wetlands

are generally open systems, the rates of organic matter import, export and storage are

also significant.

Occasionally in some wetlands, a balance between methanogenesis and methanotrophy

results in negligible methane emissions (Boon, Mitchell et al. 1997). Boon (2000) has

discussed the importance of methanogenesis, methanotrophy and methane emission in

wetland carbon cycles. He reported that methane emissions from a range of different

floodplain wetlands in Australia ranged from the limits of detection to 8mmol.m-2

.hr-1

.

Emissions from permanent natural wetlands showed high variability, with winter maxima

below the limit of detection and summer maxima reaching 3 mmol.m-2

.hr-1

(Boon 2000;

Sorrell and Boon 1992a). Methane emissions were also high in areas supporting

emergent macrophytes, and daytime emissions were double nightly rates. Emissions from

ephemeral wetlands fell within the same range of values as permanent wetlands, but were

dependent on the timing and duration of flooding (Boon 2000). Occasionally in some

wetlands, a balance between methanogenesis and methanotrophy resulted in negligible

methane emissions (Boon, Mitchell et al. 1997). Boon (2000) has estimated that the bulk

of carbon emissions from wetland sediments are the result of methanogenesis, with

aerobic decay having a secondary role in gas emission.

In summer months, the water column in many shallow floodplain wetlands becomes

stratified with respect to oxygen concentration (Ford, Boon et al. 2002). As a result,

36

surface waters are oxygenated but contain virtually no methane during the day, while de-

oxygenated water overlying the anoxic sediments is high in methane. This situation

places severe restriction on the oxidation of methane by methanotrophic bacteria (Ford,

Boon et al. 2002). However, nightly overturn of the water column mixes the stratified

layers and promotes methanotrophy in the early morning. This suggests that stratification

is an important factor contributing to elevated methane emissions where shallow

wetlands hold water over the summer period.

2.6 Plant detritus as a carbon resource

The rate at which plant detritus is decomposed, and the micro-organisms that mediate

decomposition, are strongly influenced by both the chemical composition of the substrate

and the distribution of the substrate through space and time. Simple and easily

metabolised plant carbohydrates such as sugars typically occur at low concentrations in

the environment because they are rapidly turned over by common decomposer micro-

organisms (Suberkropp and Klug 1976; Tranvik, van Hees et al. 2004). More complex

compounds, such as the structural carbohydrates, build up in the environment as a result

of extensive biogeochemical transformations (Suberkropp and Klug 1976). They resist

rapid microbial decomposition and are not generally known or described as discrete

molecules (Tranvik, van Hees et al. 2004).

2.6.1 Plant structural carbohydrates

Structural carbohydrates of plant origin include cellulose, hemicellulose and lignin.

These are complex polymers composed of repeating sub-units of sugars in cellulose and

hemicellulose, and phenyls in lignins. In order to be utilised metabolically, these

polymers must be broken down enzymatically.

2.6.1.1 Cellulose

Cellulose is a linear polymer composed of around 14 000 repeating β-1,4-glucose

residues, with cellobiose (a disaccharide) being the basic repeating unit. It is the most

abundant organic resource on Earth, with approximately 4 x 109 tons being formed

annually (Aro, Pakula et al. 2005; Melillo, McGuire et al. 1993). Cellulose comprises 30-

50% of plant cell walls (Esau 1977) and 40-50% of the dry weight of temperate woods

(Erickson, Blanchette et al. 1990c; Shearer 1992). In wetlands, 10-20% of leaf litter and

40-50% of mature wood is composed of cellulose (Boulton and Boon 1991).

37

As a component of plant cell walls, cellulose is found in association with pectin and

hemicellulose, and it is additionally cross-linked to lignin in structural tissues (Evert and

Esau 2006). The proportion of cellulose in leaf material remains fairly constant

throughout decomposition, paralleling mass loss (Suberkropp and Klug 1976). This may

be due to its gradual exposure to microbial activity as lignin is slowly removed. It may be

that cellulose and hemicellulose are metabolised preferentially, and that lignin

decomposition is a secondary metabolic process triggered by nutrient limitation

employed primarily as a means of further exposing cellulose (Cooke and Whipps 1993;

Cullen and Kersten 1992; Suberkropp, Godshalk et al. 1976; Van Laere 1994).

2.6.1.2 Hemicellulose

Hemicellulose is the second most abundant polysaccharide in nature and generally takes

the form of an heterogeneous polymer composed of beta-linked sugar units. These sugars

include β-1,4-D-xylans, β-1,4-D-mannans, and β-1,4- and β-1,3-galactans. The xylans

are most common in cereals and hard woods and have a backbone composed of β-1,4-

linked D-xylose. (Galacto)glucomannans are found in soft woods and hard woods and

have a backbone of β-1,4-linked D-mannose. Arabinan and galactan are less abundant,

and consist of α-1,5-linked L-arabinose and β-1,3-linked D-galactose units respectively

(Aro, Pakula et al. 2005).

The percentage of hemicellulose in leaf material has been found to remain fairly constant

over the period of decomposition, paralleling mass loss (Suberkropp, Godshalk et al.

1976). It is degraded at a rate equal to, or faster than, cellulose decomposition in most

wetlands, and is degraded five times faster than lignin (Suberkropp, Godshalk et al.

1976). The rate of hemicellulose decomposition is lowered by anoxic conditions (Ford

1993).

2.6.1.3 Lignin

Lignins are complex aromatic polymers formed from phenyl propanoid subunits, such as

coniferyl, synapyl and ρ-coumaryl alcohols, joined by carbon-carbon and ether linkages.

In cell walls, they are also extensively cross-linked (Aro, Pakula et al. 2005). Strong and

random bonding between phenyl-propane subunits, and the diversity of chemical

structures found in lignins from different plants, make this polymer resistant to microbial

degradation (Dix and Webster 1995).

38

Lignins comprise 20-30% of woody tissue and wetland macrophytes (Ford 1993; Kirk

and Farrell 1987; Shearer 1992), and 10% of leaf litter mass (Suberkropp, Godshalk et al.

1976). They form a physical barrier that protects cellulose and hemicellulose from

enzymatic attack (Aro, Pakula et al. 2005).

The lignin content of submerged wood is a strong influence on its decay rate (Melillo,

Naiman et al. 1984). In leaf material, lignin content has been found to increase over the

period of aquatic decomposition. Moreover, the absolute magnitude of the lignin fraction

in submerged leaves also increases, remains stable for a time and then subsequently

decreases (Suberkropp, Godshalk et al. 1976). Suberkropp et al. (1976) suggested that

this increase in total lignin content was a result of the formation of complexes composed

of lignin and nitrogen compound.

2.7 Fungal Ecology

It is well established that fungi are important in the decomposition of dead plant material

under terrestrial (Swift, Heal et al. 1979) and aquatic (Gessner, Gulis et al. 2007)

conditions. In many forest ecosystems, fungi dominate the microbial biomass in both the

litter and soil layers (Cromack and Caldwell 1992; Eicker 1973; Kjoller and Struwe

1992; Wainwright 1992). In these systems they are fundamental to the transformation

and translocation of complex detrital molecules (Cromack and Caldwell 1992).

As multicellular decomposer organisms, fungi have several metabolic advantages over

bacteria. They are capable of indeterminate growth and can spread throughout large

areas; they are able to translocate nutrients and metabolites through their mycelia; and

fungi are able to tolerate stressful environmental conditions (Jennings and Lysek 1996).

Regardless of these advantages, the decomposer community is usually comprised of

fungi, bacteria, protozoa and invertebrates (Schulze and Walker 1997). It is the

interaction of these organisms that determines the rate and mechanisms of plant detritus

break down and decomposition (Hieber and Gessner 2002; Mille-Lindblom, Fischer et

al. 2006a).

Fungi are able to colonise a wide range of substrates rich in organic compounds. These

include living plants and animals, dead plants and animals, soil organic matter and

dissolved organic matter. There is also evidence that fungi are capable of fixing carbon

39

dioxide from the atmosphere (Parkinson, Wainwright et al. 1989). Fungi have been

found to degrade plant detritus in aquatic environments including salt marshes (Buchan,

Newell et al. 2003; Weinstein, Kreeger et al. 2002), freshwater marshes (Gessner and

Van Ryckegem 2003; Kuehn and Suberkropp 1998b), peat bogs (Thormann, Currah et

al. 2003; Trinder, Johnson et al. 2008) and streams (Cornut, Elger et al. 2010; Gulis,

Kuehn et al. 2009; Hieber and Gessner 2002; Pascoal, Cássio et al. 2005). The aquatic

hyphomycetes in particular, are of high importance in the breakdown of leaf litter in

streams (Bärlocher 1992; Bärlocher, Canhoto et al. 1995; Bärlocher and Graça 2002;

Graça, Pozo et al. 2002; Sridhar, Duarte et al. 2009; Suberkropp 2001). The importance

of fungi in floodplain wetlands has not yet been established.

There is a limited amount of research into the ecology of fungi in Australian freshwater

wetlands (Scholz and Boon 1993b) and rivers (Suter 2009; Swart 1986; Thomas 1992;

Thomas, Chilvers et al. 1989; Thomas, Chilvers et al. 1992). However, over the last

decade, advances have been made in techniques for detecting and characterising fungal

populations (e.g. Andjic, Hardy et al. 2007; Borneman and Hartin 2000; Boulton and

Boon 1991; Collado, Platas et al. 2007; Fackler, Schwanninger et al. 2007; Glen,

Tommerup et al. 2001; Guglielmo, Bergemann et al. 2007; Kumar and Shukla 2005;

Lawley, Ripley et al. 2004; Martin and Rygiewicz 2005; Ritz 2007; Rodriguez, Cullen et

al. 2004; Singh, Nazaries et al. 2006). These new techniques now provide the means to

further investigate the contribution of the fungi to floodplain carbon cycling.

2.7.1 Fungi as decomposers of plant material

Saprotrophic fungi (saprobes) are fungi that obtain their nutrition from dead organic

matter. The organic matter is known as their “substrate”, and serves as both habitat and

food source. Fungi invade the plant tissue and obtain nutrition by secreting extracellular

enzymes. These enzymes break down plant polysaccharides into monosaccharides that

can be readily absorbed across the fungal cell wall. Not all plant cell components can be

broken down and absorbed by all fungal species, although it is a competitive advantage

to be able to use a range of carbon sources (Wainwright 1992). For this reason most

species of fungi elaborate a range of different extracellular enzymes, but the suite of

enzymes produced will differ between species. The enzymes that a species produces will

dictate which substrates it can successfully colonise.

40

2.7.2 Aquatic fungi

The fungi known to inhabit stream, lakes and wetlands include representatives of the

Eumycota (true fungi), Stramenopiles (fungus-like protists) and Labyrinthulomycetes

(slime moulds). Within the Eumycota, all major phyla are represented (Ascomycota,

Chytridiomycota, Zygomycota and Basidiomycota) (Nikolcheva and Bärlocher 2004;

Shearer, Descals et al. 2007). Saprotrophic fungi found colonising submerged wood and

leaves include the aquatic hyphomycetes and aero-aquatic hyphomycetes. These are

functionally defined groups. The aquatic hyphomycetes reproduce via sigmoid or tetra-

radiate conidia (Ingold 1976), while aero-aquatic hyphomycetes reproduce via buoyant

conidia (Premdas and Kendrick 1991). Stramenopiles are represented by the Oomycetes,

which reproduce asexually via flagellated zoospores (Dick 1969). Aquatic orders include

the Saprolegniales, Leptomitales, Lagenidales and Peronosporales (Dick 1976b).

Labyrinthulomycetes are not generally considered as saprotrophic fungi, and they are not

addressed further in this thesis.

Gulis et al. (2009) recently discussed the role of fungi in fresh water environments, while

Jobard et al. (2010) reviewed the importance of fungal organisms in pelagic food webs

and Wurzbacher et al. (2010) reviewed the role of fungi in lake ecosystems.

2.7.2.1 Aquatic hyphomycetes

The aquatic hyphomycetes are a sub-group within the Deuteromycotina, a group defined

by the absence of a sexual life stage (Hawksworth and Rossman 1997). The name

“hyphomycetes” is derived from the fact that they produce conidia directly from the

mycelium, not from a fruiting body. Genetically, the majority of aquatic hyphomycetes

are classified with the Ascomycota, but some are members of the Basidiomycota.

Aquatic hyphomycetes are the dominant fungal group on tree leaves in fast flowing

streams (Bärlocher 1992; Ingold 1942) and their spores are adapted to rapidly disperse

longitudinally in the stream (Iqbal and Webster 1973). These fungi are absent in silted or

muddy areas where leaves turn black due to lack of oxygen. However, they may be found

in terrestrial habitats (Webster 1977), or colonising living plants in streams (Sati and

Belwal 2005; Sridhar and Bärlocher 1992).

2.7.2.2 Aero-aquatic hyphomycetes

41

Aero-aquatic fungi are a functionally defined group that are most abundant in shallow,

stagnant waters (Field and Webster 1983; Fisher and Webster 1979) where they produce

vegetative growth on submerged plant litter. They are also known to occur in streams

(Bärlocher 1992; Goos 1987; Willoughby and Archer 1973). Sporulation only occurs

when the substrate becomes exposed to air (Pattee and Chergui 1995; Shearer, Descals et

al. 2007) and the resultant propagules are buoyant and float on the water’s surface.

Taxonomically, most aero-aquatic fungi are classified as mitosporic Ascomycetes

(Ascomycetes lacking a sexual phase in development). Four aero-aquatic species have

been classified as Basidiomycetes, and one as Oomycete (Shearer, Descals et al. 2007).

2.7.2.3 Oomycetes

The Oomycetes (Peronosporomycetes), also known as water moulds, are aquatic

organisms common in waterlogged soils (Dick 1962; Dick 1963) and stagnant waters

such as ditches, ponds (Park, Sorenson et al. 1978), the littoral zones of lakes (Dick

1976b; Hallett and Dick 1981; Nechwatal, Wielgoss et al. 2008), wetlands (Czeczuga,

Muszyńska et al. 2007), and streams (Shearer and von Bodman 1983). Common genera

include Achlya, Saprolegnia, Pythium, Phytophthora, Aphanomyces, Dictyuchus and

Leptolegnia (Hallett and Dick 1981; Nechwatal, Wielgoss et al. 2008; Park, Sorenson et

al. 1978). The Achlya and Saprolegnia are abundant in near-shore areas, while Pythium

and Aphanomyces are found in oxic pond sediments (Park, Sorenson et al. 1978).

However, the abundance of Oomycetes and their propagules is usually much lower in

sediments than in surface waters (Hallett and Dick 1981).

The Oomycetes are taxonomically distinct from the Eumycota (true fungi), being more

closely related to brown algae and diatoms and possibly evolved from a photosynthetic

ancestor (Lamour, Win et al. 2007). Morphologically, however, they are similar in form

and habitat to the Chytridiomycetes (see below). When colonising submerged plant litter,

Oomycetes are observed early, then rapidly decline (Bärlocher 1990; Bärlocher and

Kendrick 1974). Peaks in the abundance of Oomycete propagules have been observed to

coincide with the seasonal influx of fresh substrate in spring, summer and autumn

(Hallett and Dick 1981), further suggesting a preference for newly submerged litter.

These organisms prefer neutral to acid pH (Dick 1963), and may be limited by calcium

concentration (Czeczuga, Muszyńska et al. 2007), which is a cofactor for the enzyme

pectin lyase (Suberkropp 2001).

42

Many species of Oomycetes are pathogenic to plants and animals. This includes Pythium

phragmitis, which is known to infect the common wetland reed, Phragmites australis

(Nechwatal, Wielgoss et al. 2008). Most of the species belonging to Achlya, Dictyuchus

and Saprolegnia are opportunistic pathogens to fish (Leclerc, Guillot et al. 2000), taking

advantage of minor injuries to infect surface tissues (Singhal, Jeet et al. 1987). They may

also infect crustaceans, mosquito larvae and the eggs of fish and invertebrates (Hulvey,

Padgett et al. 2007).

2.7.2.4 Chytrids

Chytrids (Chytridiomycota) are an important component of the fungal decomposer

community in fresh water environments. They are also parasites of both plants and

animals, able to digest proteins, chitin, and cellulose with high efficiency. Chytrids

reproduce asexually via the production of zoospores. These zoospores may be an

important food source for zooplankton. The ecology of Chytrids in aquatic ecosystems is

reviewed by Gleason et al. (2008).

2.7.3 Fungal communities

While more than 600 species of fungi are known to colonise the leaves of Phragmites

australis (Gessner and Van Ryckegem 2003), less than twenty species generally coexist

on the substrate at any one time. Three processes interact to determine which species will

be successful. Firstly, their ability to use the available resources (i.e. the enzymes they

can secrete) (Chamier and Dixon 1982b). Secondly, environmental factors such as

temperature, water flow and oxygen availability (Findlay and Arsuffi 1989; Pattee and

Chergui 1995) select for a pool of species that are tolerant of local conditions and can

potentially colonise a new substrate. Thirdly, competition for resources on a discrete

substrate such as a submerged leaf initiates a succession of species over time (Fryar,

Booth et al. 2005; Mille-Lindblom, Fischer et al. 2006a).

The fungal communities on submerged litter substrates in streams are usually composed

of four to ten abundant or common species of aquatic hyphomycetes (Chamier, Dixon et

al. 1984; Thomas, Chilvers et al. 1992). These communities vary seasonally (Nikolcheva

and Bärlocher 2005; Thomas, Chilvers et al. 1989). In Australia, distinct communities

colonise the submerged leaves of Eucalyptus viminalis in summer /autumn and

spring/summer, although some species show no seasonal variability (Thomas, Chilvers et

43

al. 1992). While seasonal changes in fungal community composition may be correlated

with seasonal litter inputs (Thomas, Chilvers et al. 1989), Suberkropp (1984) found that

many species were found predominantly in summer or winter regardless of the time litter

was introduced to the stream.

The communities of aquatic fungi found in flowing and standing water habitats are

distinctive (Wong, Goh et al. 1998). Broadly speaking, flowing waters are dominated by

aquatic hyphomycetes, while standing waters are dominated by aero-aquatic

hyphomycetes (Pattee and Chergui 1995). Many species of aero-aquatic fungi are also

able to withstand prolonged periods of drying (Field and Webster 1983), which is

advantageous in ephemeral habitats such as floodplain wetlands. Standing water has

lower oxygen concentration, higher temperatures, and higher concentrations of nutrients

and DOC relative to flowing water and wetland fungi tend to be adapted to these

conditions.

Reduced oxygen availability will select species able to metabolise carbon anaerobically

(fermentation), disadvantaging species that oxidise lignin. Fungi are able to ferment

glucose (Cochrane 1956), starch (Oda, Saito et al. 2002) and gelatine (Roberts 1899),

and grow anaerobically (Davies, Theodorou et al. 1993; Theodorou, Lowe et al. 1992;

Trinci, Davies et al. 1994), but anaerobic fungi comprise a small proportion of anaerobic

saprobes in terrestrial litter (Reith, Drake et al. 2002). The metabolic pathways used by

fungi under aquatic anoxic conditions require investigation. However, many aero-aquatic

hyphomycetes are able to survive anoxic conditions for up to 12 months as vegetative

hyphae (Field and Webster 1983), although they may not grow under these conditions

(Fisher and Webster 1979). Aquatic hyphomycetes can survive for limited periods under

low oxygen conditions (Field and Webster 1983) but they experience reduced levels of

function in terms of leaf decomposition, diversity, biomass and reproduction (Medeiros,

Pascoal et al. 2009b). Pattee and Chergui (1995) found a gradient from the well-

oxygenated waters of the Rhone River where aquatic hyphomycetes were dominant, to

the anoxic conditions of an ox-bow lake on the Rhone floodplain where aero-aquatic

hyphomycetes dominated.

Leaf species is often the most important controlling factor in the composition of fungal

decomposer communities (Thomas 1992; Thomas, Chilvers et al. 1992). Gulis (2001)

44

found that at least some species show preference for specific substrates, with

communities on wood and grass being distinct from those found on tree leaves. The

chemical composition of the substrate differs between species, and will also change as

decay progresses. The proportion of cellulose, hemicellulose and plant defensive

compounds decreases with decay, while the proportion of lignin increases (Golladay and

Sinsabaugh 1991; Rosset, Bärlocher et al. 1982; Suberkropp, Godshalk et al. 1976). This

confers an advantage on species able to degrade lignin, and encourages a change in

community composition. As the substrate becomes more fragmented the decomposer

community becomes increasingly dominated by bacteria (Baldy, Gessner et al. 1995;

Golladay and Sinsabaugh 1991; Suberkropp and Klug 1976).

Mille-Lindblom et al. (2006b) found that substrate quality was more important than

water chemistry in regulating the composition of the microbial community on submerged

litter in lakes, while Fischer et al. (2009) found water chemistry had the stronger

influence on stream communities. Schoenlein-Crusius et al. (1999) found that the

occurrence of aquatic hyphomycetes, and to a lesser extent zoosporic fungi, was

correlated with high concentrations of calcium in the substrate.

Competition is an important structuring agent for all ecological communities, including

decomposer communities (Fryar, Booth et al. 2005; Mille-Lindblom, Fischer et al.

2006a; Mille-Lindblom and Tranvik 2003; Wicklow 1981). For example, under aquatic

conditions, terrestrial fungi do not generally compete successfully with aquatic

hyphomycetes (Bärlocher and Kendrick 1974; Suberkropp and Klug 1976). This is likely

to be because they are less efficient at acquiring and metabolising resources under these

conditions. However, some species of aquatic hyphomycetes have been shown to

produce antibiotic substances that are able to inhibit the growth of bacteria and other

fungi (Gulis and Stephanovich 1999). Conversely, bacteria (Fischer, Mille-Lindblom et

al. 2006; Moller, Miller et al. 1999; Romani, Fischer et al. 2006), invertebrates

(Bärlocher and Kendrick 1975) and viruses (Jackson and Jackson 2008) may inhibit

fungal growth, and bacteria may benefit from the presence of fungi (Romani, Fischer et

al. 2006).

Colonisation of aquatic detritus is random and patchy, and the composition of the fungal

community on submerged litter may be difficult to predict (Fischer, Bergfur et al. 2009).

45

On substrata introduced into streams, species richness is initially low, rises to a peak

(within weeks or days) then remains stable or declines (Nikolcheva, Cockshutt et al.

2003). Those fungal species that are first to colonise a substrate have a strong influence

over the composition of the fungal community that subsequently develops (Bärlocher

1992; Sanders and Anderson 1979). This effect remains in place even if environmental

conditions change markedly (Sridhar, Duarte et al. 2009). However a change from aerial

to aquatic conditions leads to a gradual change in community composition (Van

Ryckegem, Gessner et al. 2007).

2.7.4 Biomass

The abundance of fungal organisms on submerged plant substrates is generally measured

in terms of biomass. This is commonly estimated by determining the content of the

fungal biomarker ergosterol in the leaf matrix. Biomass is influenced more strongly by

substrate type than is community structure (Nikolcheva and Bärlocher 2005), but it is

also influenced by water quality parameters (Sridhar and Bärlocher 2000; Suberkropp

and Chauvet 1995).

When new substrates enter the aquatic environment, fungal biomass tends to steadily

increase to a maximum and then decline (Das, Royer et al. 2007). The time taken to

reach maximum biomass may vary with leaf species and environmental conditions such

as nitrate concentrations (Das, Royer et al. 2007). During the period leading up to “full

conditioning”, fungal biomass generally exceeds bacterial biomass (Findlay and Arsuffi

1989; Kaushik and Hynes 1971) and sporulation and mycelial production continue even

as fungal biomass declines (Baldy, Gessner et al. 1995).

Where a leaf substrate has been colonised by fungi in a terrestrial or aerial environment,

fungal biomass may decrease markedly when the substrate is submerged. Fungal biomass

on decaying standing leaf blades of Phragmites australis reached 548 μg ergosterol /g

AFDM, but decreased to about one third of that value (191 μg ergosterol/g AFDM) when

the leaf blade collapsed into brackish marsh water (Van Ryckegem, Gessner et al. 2007).

2.7.5 Enzyme activity

The synergistic activity of microbial extracellular enzymes is the main driver of plant

litter decomposition. While no single enzyme can be used to predict litter mass loss,

multiple linear regression of enzyme activity shows that mass loss is strongly related to

46

the combined activity of a range of ligno-cellulose degrading enzymes (Jackson,

Foreman et al. 1995; Sinsabaugh, Moorhead et al. 1994). This implies that the rate of

litter decomposition is controlled by the same factors that influence enzyme activity.

These factors include the chemical composition of the substrate, and environmental

conditions such as leaching, temperature and pH (Chróst 1990).

Soluble organic matter is leached from plant litter within days of immersion (See 2.5.3).

Thus, only the more refractory components of the middle lamella and primary and

secondary cell walls (Chamier 1985) are available to fungi after leaching. The middle

lamella is rich in pectic polysaccharides, while the primary cell wall is composed of

cellulose, hemicellulose, pectic polysaccharides and glycoprotein. The secondary cell

wall is mainly composed of cellulose but does not contain pectic compounds. If

lignification occurs, it begins in the middle lamella and proceeds through the primary and

secondary cell walls (Evert and Esau 2006). Thus, enzyme activity on submerged litter is

primarily towards cellulose and hemicellulose, as these compounds are most abundant.

Activity towards lignin, pectin and minor plant components will also be present. While

the rate of fungal mineralisation of organic substrates is governed by the production of

enzymes, enzyme production is controlled by the availability of nitrogen and phosphorus.

Where nutrients become limiting, fungi are required to expend energy to use enzymes to

sequester more nutrients from their substrate. Thus, enzyme expression is largely

controlled by the substrate quality (Sinsabaugh, Gallo et al. 2005).

Enzyme activity may be influenced by temperature, pH and salinity (Chróst 1990). It is

well established that increases in temperature stimulate enzyme activity (Sinsabaugh,

Moorhead et al. 1994). Thus, optimum temperatures for enzyme activity are usually well

above those occurring in natural aquatic environments, so many microbes show no

adaptation to temperature (Chróst 1990). However, pH varies widely between aquatic

ecosystems, and microbes show preference for acid or alkaline pH ranges similar to those

occurring in natural systems, 4.0-5.5 and 7.5-8.5 respectively (Chróst 1990). Enzymatic

litter decomposition is also known to be inhibited by salinity. In marine environments,

both low and high salinities are inhibitory (King 1986).

Aquatic hyphomycetes are able to secrete enzymes that are active towards all the major

leaf structural polysaccharides (Abdullah and Taj-Aldeen 1989; Suberkropp, Arsuffi et

47

al. 1983), although the level of activity varies with species (Chamier 1985). Two-thirds

of aquatic hyphomycetes and aero-aquatic hyphomycetes examined by Abdullah and Taj-

Aldeen (1989) were able to hydrolyse cellulose. Aquatic hyphomycetes are able to

hydrolyse hemicellulose (Chamier 1985; Thornton 1963) and oxidise lignin (Abdel-

Raheem and Shearer 2002; Abdel-Raheem and Ali 2004). The Basidiomycetes are well

recognised for their importance in lignin degradation in terrestrial plant litter (Cromack

and Caldwell 1992; Mackensen, Bauhus et al. 2003) but they have not been proven to

actively decompose litter under aquatic conditions.

2.7.6 Enzymatic breakdown of plant polymers by fungi

Virtually all fungal species are able to readily absorb naturally occurring low molecular

weight carbohydrate monomers. Most fungi are able to enzymatically degrade organic

polymers, such as protein, pectin, cellulose and starch, to monomers and thence to carbon

dioxide and methane. Fewer species are also able to degrade lignin (Dix and Webster

1995). The following is a brief summary of the mechanisms involved in the enzymatic

breakdown of the major plant cell wall polymers. The decomposition of plant litter in

aquatic environments by fungi has been thoroughly reviewed by Gessner et al. (2007),

and Gamauf et al. (2007) discuss the fungal degradation of plant cell wall polymers.

2.7.6.1 Cellulose

A minimum of three enzymes are required to degrade cellulose completely. These

enzymes act synergistically to cleave glycosidic bonds to produce smaller chains,

cellobiose dimers and finally glucose (Ander 1994; Erickson, Blanchette et al. 1990a).

Endoglucanases (endocellulases) are usually secreted first. These are enzymes that cleave

internal β-1,4-linkages, to produce oligosaccharides. Cellobiohydrolases (exocellulases)

simultaneously cleave terminal β-1,4-linkages from either the reducing or non-reducing

end of the oligosaccharide, depending on the nature of the enzyme. This liberates

cellobiose. Finally, β-glucosidases attack oligosaccharides and cellobiose to remove D-

glucose (Aro, Pakula et al. 2005; Gamauf, Metz et al. 2007). Most saprotrophic fungi

will produce a range of cellulose degrading enzymes. For example, the soft rot fungus

Trichoderma reesei produces glycoside hydrolases belonging to seven enzyme families

(Warren 1996).

Cellulose degrading enzymes may also be oxidative (Ander 1994; Ayers, Ayers et al.

1978). For example, cellobiose oxidase uses molecular oxygen to oxidise cellobiose and

48

oligosaccharides to give gluconic and cellobionic acids (Ljungdahl and Eriksson 1985).

An alternative pathway for cellulose oxidation has been observed in Sporotrichum

pulverulentum. Cellobiose:quinone oxidoreductase (cellobiose dehydrogenase) reduces

quinones and phenoxy radicals (from lignin degradation) in the presence of cellobiose

(Ayers, Ayers et al. 1978). These systems are common in white rot fungi. Brown rot

fungi may attack cellulose via a hydrogen peroxide:ferric iron system (Shimada,

Akamtsu et al. 1997) that cleaves off reducing sugars in the presence of oxalic acid.

The synthesis of cellulases is generally repressed when glucose or other readily

metabolised sugars are present. Lignin-related phenols may also repress cellulase

production. In this case, phenol oxidases (ligninases) may polymerise these phenols to

reduce the inhibitory effect (Ljungdahl and Eriksson 1985).

2.7.6.2 Hemicellulose

As hemicellulose is a heterogeneous polymer a range of enzymes is required to hydrolyse

it completely. A suite of enzymes generally works in concert, as follows. Firstly,

exoglycosidase removes side chains, and then endohemicellulase attacks the main glycan

chain at unbranched points. This reaction produces oligosaccharides. Exoglycosidase and

hemicellulase then cleave the oligosaccharides into monosaccharides, which are rapidly

taken up through the fungal cell wall. For example, the degradation of xylan involves

endo-1,4-β-D-xylanases and β-xylosidases, which attack the sugar chain forming the

back bone of the polymer. Side chain cleaving enzymes may include α-glucuronidase and

acetyl xylan esterase (Aro, Pakula et al. 2005; de Vries and Visser 2001). Alternative

enzyme systems, which act via similar mechanisms, are required to degrade xyloglucan

and galacto-glucomannan (Gamauf, Metz et al. 2007). The presence of short-chain oligo-

polymers of xylose will induce the secretion of these enzymes (Cooke and Whipps

1993).

2.7.6.3 Pectin

Pectin is comprised of polymerised α-1,4-linked galactouropyranose, which may have

methylated carboxyl groups. It may also contain α-1,2-linked rhamnose, α-1,3-linked

arabinofuranose (araban), α-1,5-linked arabinofuranose and β-1,4-linked galactopyranose

(galactan) (Cooke and Whipps 1993). Pectin degrading enzymes are hydrolytic and

include polygalacturonases and pectin lyases. Pectate lyases, which act on non-esterified

pectin, are characteristic of bacteria. Aquatic hyphomycetes exhibit polygalacturonase

49

activity at pH values above 5 (Chamier and Dixon 1982a; Chamier and Dixon 1982b).

The uptake of the released galacturonic acids stimulates fungi to release further enzymes

including hemicellulases and cellulases (Cooke and Whipps 1993). Conversely, high

concentrations of sugars can repress the production of these pectinases (Cooke and

Whipps 1993). Pectin degrading enzymes are the first to be induced on a plant based

substrate. Pectinase activity upon the middle lamella can separate cells and expose cell

walls to further degradation.

2.7.6.4 Lignin

Lignin is highly resistant to hydrolytic degradation, so lignin degrading enzymes are

oxidative, rather than hydrolytic. White rot Basidiomycetes are the only microbial group

able to mineralise lignin to carbon dioxide and water. Brown rot fungi such as the

xylariaceous Ascomycotina modify lignin by demethylation but cannot degrade it

completely. Soft rot fungi attack lignin under water logged or dry conditions (Gamauf,

Metz et al. 2007).

In lignified plant tissues, lignin must be degraded before energy can be obtained from

polysaccharides. Lignin degradation is thought to be a secondary metabolic function for

most fungal species, serving to expose cellulose (Cooke and Whipps 1993). Other

research suggests that lignin degrading enzymes are induced by conditions of nitrogen

starvation. Either way, lignin breakdown does not produce enough energy to be self-

sustaining (Jeffries, Choi et al. 1981), so expenditure of cellular energy to degrade it is

likely to be a means for obtaining additional energy or nutrients.

As lignin is a complex and heterogeneous aromatic polymer, numerous chemical

reactions would be required to complete its breakdown. A range of breakdown products

may also be degraded (Ljungdahl and Eriksson 1985). The main fungal enzymes

involved in lignin degradation are laccases (phenol oxidases), manganese peroxidases

and lignin peroxidases. Non-proteinaceous compounds, low molecular weight free

radicals, are released into the substrate to initiate decay in many cases. Additional

enzymes may be secreted to generate these molecules (Aro, Pakula et al. 2005).

Phenol oxidases oxidise benzenediols to semi-quinones with molecular oxygen. Laccases

(benzenediol: oxygen oxidoreductase) are enzymes belonging to this group, that contain

50

copper at their catalytic centre and are active against phenols and similar molecules.

They oxidise polyphenols, methoxy-substituted phenols and aromatic di-amines by

reducing molecular oxygen to water (Gamauf, Metz et al. 2007). Laccase, in the presence

of suitable substrates may also oxidatively cleave carbon-carbon linkages in lignin and

oxidise veratryl alcohol to veratraldehyde (Bourbonnais and Paice 1990).

Laccases are commonly produced by Basidiomycetes, and may be produced by

Ascomycetes. There are no reports indicating that Zygomycetes or Chytridiomycetes

produce laccases (Baldrian 2006). In water-saturated environments, laccase activity is

driven by the availability of oxygen. For example, laccase activity is generally low in

peatlands (Pind, Freeman et al. 1994; Williams, Shingara et al. 2000), but when oxygen

becomes available as a result of drawdown, laccase activity increases sharply. Increased

laccase activity can lead to the depletion of compounds that inhibit the activity of other

oxidative and hydrolytic enzymes (Freeman, Ostle et al. 2004). Under water, laccase may

be involved in the degradation of wood and humic substances (Claus and Filip 1998;

Hendel and Marxsen 2000).

Peroxidases are a group of enzymes which catalyse the oxidation of substrate by a

peroxide to give phenols. Lignin peroxidases resemble other peroxidases in that they

contain a heme group and operate via a catalytic cycle. That is, the enzyme is oxidised by

hydrogen peroxide and two electrons are removed. The enzyme returns to its resting state

by oxidising donor substrate, removing one electron from each of two molecules. These

substrates usually include phenol, anilines, aromatic ethers and non-phenolic lignin

structures (Hammel 1997).

Manganese peroxidases are similar to other peroxidases, except that Mn(II) is the

electron donor (Mn(III) is generated). The reaction requires the presence of organic acid

chelators such as glycolate or oxalate, which stabilise Mn(III) and allow it to be released

from the enzyme. These Mn(III) chelates oxidise the more reactive structures that make

up about 10% of the lignin polymer. The activity of this enzyme on lignin may facilitate

subsequent attack by lignin peroxidase (Hammel 1997).

Fungi may also secrete additional compounds such as hydrogen peroxide and

veratraldehyde, which diffuse into the substrate away from the hyphae and penetrate into

51

the ligno-cellulose. These substances may initiate degradation. Brown rot fungi initiate

lignin degradation by producing low molecular weight diffusible compounds that are

able to act upon the wood at a distance from the hyphae. These compounds might include

hydroxyl, peroxyl and hydro-peroxyl radicals, which are generated via the Fenton

reaction (Gamauf, Metz et al. 2007). The presence of molecular oxygen and iron are

required for this reaction to proceed. Many fungi also secrete oxalic acid, which chelates

iron, but also reduces the pH, increasing enzyme activity (Gamauf, Metz et al. 2007).

Where peroxidases are active, a source of hydrogen peroxide is required to initially

oxidise the enzyme. This is achieved by extracellular oxidases which reduce molecular

oxygen to hydrogen peroxide while oxidising low molecular weight organic compounds.

Similar compounds may act as intermediaries in substrate oxidation. For example,

veratryl alcohol can be oxidised to a radical cation which is not rapidly degraded. It can

thus act as a one-electron oxidant when it encounters a second oxidisable substrate, and

in the process the veratryl alcohol molecule is regenerated (Harvey, Schoemaker et al.

1986).

Given that lignin degradation is an oxidative process, lignin has previously been

considered to be inert under anoxic conditions (Evans and Fuchs 1988; Kirk and Farrell

1987). However, long term incubation of wood blocks in anaerobic mud revealed that

they were slowly degraded by rod-shaped bacteria (Benner, Maccubbin et al. 1984; Holt

and Jones 1983). It has also been believed that aquatic hyphomycetes were not able to

degrade lignin (Boulton and Boon 1991). However, both aquatic and aero-aquatic

hyphomycetes are capable of partial lignin break down (Fisher and Webster 1979). Thus

aquatic lignin degradation by fungi is plausible, even under micro-aerobic conditions.

2.7.6.5 Other polymers

Proteins, chitin, cutin, waxes and suberin are persistent polymers that are available to a

broad spectrum of fungi for an extended period (Cooke and Whipps 1993). These

polymers tend to be minor components of plant litter in aquatic environments, so

research into aquatic decomposition has tended not to focus on them. Storage polymers

such as starch, inulin, glycogen and lipids are theoretically available to a wide range of

fungi. In practice, they are only utilised by fungal parasites and the primary colonisers of

dead material (Cooke and Whipps 1993).

52

2.8 Summary

While the body of knowledge concerning fungal ecology has established a firm

understanding of fungal metabolism and taxonomy, little work has been carried out in

aquatic environments within Australia, and while fungal specimens have been identified

from standing waters in Australia, no ecological studies have taken place in floodplain

wetlands. It is therefore necessary to explore fungal ecology in Australian floodplain

wetlands from the most basic of starting points, and ask the question, “What role do

fungi play in the ecology of ephemeral floodplain wetlands?” Fundamental aspects of

this question are examined in Chapter 3, where the fungal community structure is

compared between sediment and plant detritus substrates in a typical Australian

floodplain wetland.

54

Chapter 3

Pilot Study: Fungal Community Structure on a variety of substrates

from within a floodplain wetland.

3.1 Introduction

Research conducted on floodplains of the Rhône (Pattee and Chergui 1995) and Garonne

Rivers (Baldy, Chauvet et al. 2002) has shown that aquatic fungal communities differ

between rivers and lentic habitats on their floodplains. This has been attributed to

differences in temperature, aeration (Dix and Webster 1995), dissolved organic carbon

concentrations and flow (Pattee and Chergui 1995). However, large differences in

community structure have also been observed between submerged wood and submerged

leaves (Gulis 2001) and between different leaf species (Thomas, Chilvers et al. 1992).

Differences have also been noted between similar substrates in different locations

(Suberkropp and Chauvet 1995) and these differences have been attributed to differences

in riparian vegetation and abiotic variables.

To date, there are no published studies that have investigated fungal community structure

in Australian floodplain wetlands, although there have been numerous studies that have

investigated the importance of bacteria in these systems (Boon 1990; Boon and Sorrell

1991; Boon, Virtue et al. 1996; Bunn and Boon 1993; Burns and Ryder 2001b; Ford,

Boon et al. 2002). A number of taxonomic studies have recorded the occurrence of

aquatic fungal species in rivers (Cowling and Waid 1963; Price and Talbot 1966;

Thomas 1992; Thomas, Chilvers et al. 1989; Thomas, Chilvers et al. 1992) and non-

flowing habitats (Hyde and Goh 1998), and fungal biomarkers have been isolated from

biofilm growing on submerged wood (Scholz and Boon 1993a; Scholz and Boon 1993b).

Boon and his co-authors (1996) used phospho-lipid fatty acid (PLFA) profiles to

investigate microbial diversity in floodplain wetland sediments. They were unable to

detect fungal biomarkers and concluded that fungi were not important in wetland

sediment carbon cycling. While many aquatic fungi species are tolerant of low oxygen

environments (Field and Webster 1983; Medeiros, Pascoal et al. 2009b), and some are

facultative anaerobes capable of fermentation (Reith, Drake et al. 2002), there is no

evidence that fungi are able to dominate decomposer communities under anoxic

55

conditions. As the sediments of floodplain wetlands are known to become anoxic soon

after inundation (Baldwin and Mitchell 2000; Mitsch and Gosselink 1993b), the findings

of Boon et al. (1996) are plausible.

Plant detritus serves as a substrate for aquatic fungi in wetlands. Most of the

allochthonous plant production in wetlands is not consumed by grazers (Brinson, Lugo et

al. 1981), but is decomposed in the standing dead position or after it collapses to the

sediment surface (Buesing and Gessner 2006). Allochthonous litter, derived from

riparian vegetation, provides another possible substrate for fungal communities. In the

aerial position, in the water column and on the surface of the sediments, conditions exist

in which fungal communities might develop, although bacteria are known to dominate

submerged detritus under anoxic conditions (O'Connell, Baldwin et al. 2000). Fungi

dominate substrates such as these in rivers, particularly in the early stages of

decomposition when particle size is large (Suberkropp 1991a).

While the activities of the decomposer community in the sediments of floodplain

wetlands and fungal dynamics within wood biofilm have been investigated, there have

been no reports of investigations into fungal dynamics on plant detritus in these systems.

Indeed, the failure to detect fungal biomarkers has lead to the suggestion that fungi are

not “a significant component of the microbial community” in floodplain wetlands (Boon,

Virtue et al. 1996). Therefore it is first necessary to confirm the presence of fungi in

Australian floodplain wetlands. Following this, the micro-habitats which fungi colonise

should be determined. These microhabitats might be influenced by substrate type,

location and water depth.

In the following experiment, T-RFLP analysis was used to firstly detect the presence of

fungal DNA, then to compare fungal community structure profiles between wetland

sediments, and allochthonous and autochthonous plant substrates, in a floodplain

wetland. These profiles were then compared between sites within the wetland, and with

sediment from the associated river. The community structure in sediments from a variety

of water depths and submerged and standing dead litter were also compared. The results

of this investigation were intended to generate hypotheses for further experiments.

56

3.2 Materials and Methods

3.2.1 Study site

Normans Lagoon is a billabong adjacent to the Murray River at Albury, NSW, Australia

(146°56’06”E, 36°05’42”S; Figure 3.1). The billabong is 675m long and 40m wide, and

2-6 m deep along the central axis. Open water is interspersed with reed beds, and

populations of the floating fern, Azolla filiculoides (Lam.) are often present. On the

north-east side it is bounded by a road reserve, where the dominant riparian vegetation is

Eucalyptus camaldulensis (Dehnh.) (Figure 3.2). Exotic tree species belonging to the

genus Salix are also present. Stands of emergent macrophytes consisting of Eleocharis

sphacelata, Cyperus eragrostis and Juncus ingens are present along the shore line.

Brasenia shreberi is the dominant floating attached macrophyte with Azolla filiculoides

also floating on the water surface. On the south-western side the lagoon is bordered by

pasture. Water enters the lagoon from the Murray River at the downstream (west) end

during minor flood events (18 000 ML/day), but fills from the upstream end during major

floods (Gribben, Rees et al. 2003).

Figure 3.1: A map of Normans Lagoon, showing the position of sample sites. The inset shows the

location of Albury-Wodonga, situated on the Murray River, which forms the border between the

states of New South Wales and Victoria. Normans Lagoon is found on the north side of the Murray

River, south-east of Albury, NSW.

57

Figure 3.2: The western site at Normans Lagoon. The riparian vegetation at this site is dominated by

Eucalyptus camaldulensis. Outside this zone, the vegetation is influenced by rural land use.

3.2.2 Sample collection

Samples of sediment and plant material were collected from Normans Lagoon on

October 20th

, 2006. Triplicate sediment samples were taken from the Murray River at

Doctors Point, at the western end of the lagoon and from a site on the eastern side of the

lagoon. Vegetation was sampled only from the two lagoon sites. Samples were collected

in sterile specimen jars and frozen at -20°C on return to the laboratory, less than 1 hour

later.

Sediment samples consisted of (a) dry soil sampled at a distance of 5 m from the water’s

edge, (b) moist soils, sampled <1 m from the water’s edge, (c) recently submerged soil,

sampled ~30cm into the lagoon from the water’s edge, (d) submerged flocculent material

(floc) (from ~20cm water depth), possibly comprised of Azolla filiculoides detritus, (e)

submerged sediment (from ~50cm water depth), and (f) submerged sediment from

Cyperus sp. (east site) and Juncus sp. (west site) reed beds (20-50cm water depth).

Vegetation samples consisted of (a) Eucalyptus camaldulensis leaves from the sediment

surface, (b) Salix alba leaves, yellowed and floating on the surface, (c) Standing dead and

submerged Juncus ingens leaf blade, from the west site, (d) Standing dead and

58

submerged Cyperus eragrostis leaf blades, from the east site, and (e) submerged bark

(species unknown).

3.2.3 Terminal restriction-fragment length polymorphism (T-RFLP)

DNA was extracted from sediment samples, using an UltraCleanTM

Soil DNA Isolation

Kit (MoBio Laboratories, Inc, Solana Beach, CA, USA.) following the alternative

protocol recommended for maximising yields. Optimisation of this method for use with

both sediment and plant samples led to the following modifications: (a) The samples

were centrifuged and the supernatant was removed, then weighed into bead tubes; (b)

cells were lysed using a bead beater for 10 seconds on high, followed by 2 minutes on

ice, and these steps were repeated three times, and (c) after adding solution S2, a protein

precipitating reagent, tubes were incubated at 4°C for 1 hour rather than 5 minutes. The

DNA extracted from these samples is hereafter referred to as the “genomic” DNA.

PCR was used to amplify the internal transcribed spacer region II (ITS2) of fungal

ribosomal DNA (rDNA), found between 5.8S-coding sequence rDNA and Large sub-unit

coding sequence (LSU) of the ribosomal operon (Martin and Rygiewicz 2005). The ITS

regions (I and II) are more variable than the other subunits on the gene, and there are

sequence variations among species (Kumar and Shukla 2005). Thus, this region of the

fungal DNA is useful for developing community profiles.

The PCR reaction was carried out using Qiagen HotStarTaq® DNA polymerase

(Quiagen, Hilden, Germany) and the accompanying reaction buffer. A 50 µL reaction

volume was used, comprised of 0.5 µL Taq, 5 µL of 10x reaction buffer, 1 µL dNTP

(dATP, dGTP, dTTP and dCTP, 10 mM of each; Genesearch, Arundel, Qld), 2 µL

forward primer (0.4 µM per µL), 2 µL reverse primer (0.4 µM per µL), 1 µL genomic

DNA, and 38.5 µL sterile milliQ water. Genomic DNA was used either undiluted or

diluted to 10% in sterile milliQ water. The forward primer used was ITS3 (5'- TCG ATG

AAG AAC GCA GC – 3') and the reverse primer was ITS4 (5'-TCC TCC GCT TAT

TGA TAT GC – 3') (White, Bruns et al. 1990). Primers were synthesised by Gene Works

(Hindmarsh, S.A.). The forward primer carried a fluorescent label (5, 6 -

Carboxyfluorecein) at the 5' end. These primers produce a DNA fragment approximately

350 bp in length (length may vary between species) and are specific to fungi (Kumar and

Shukla 2005).

59

Amplification was carried out in an iCycler (Bio-Rad, Hercules, CA , USA) thermocycler

using a Touchdown protocol (Don, Cox et al. 1991). This approach has been used to

amplify fungal DNA (Green, Freeman et al. 2004; Kasuga, Taylor et al. 1999; Meyer,

Carta et al. 2005), and in conjunction with T-RFLP (Arteau, Labrie et al. 2010)

previously. The PCR protocol consisted of initial denaturation at 95°C for 15 minutes (1

cycle), then 15 cycles of denaturation at 94°C for 15 seconds, annealing for 15 seconds,

and extension at 72°C for 60 seconds. Annealing temperature was reduced by 1°C after

each cycle, commencing at 60°C and ending at 45°C. A further 20 cycles then followed

with denaturation at 94°C for 15 seconds, annealing at 48°C for 15 seconds, and

extension at 72°C for 60 seconds. A final cycle was included with denaturation at 94°C

for 15 seconds, annealing at 48°C for 60 seconds, and extension at 72°C for 6 minutes.

The PCR product was then held at 4°C until removed from the thermocycler and stored

at -20°C. PCR product was visualised under UV light on a 1% agarose gel incorporating

ethidium bromide (0.4 µg.ml-1

). Retention of the amplicons was compared with a lambda

digest size standard to determine their approximate length.

The PCR product contains primers, enzymes, salts and other substances that may become

problematic during the restriction digest or subsequent electrophoresis. These

contaminants were removed using the UltraClean PCR Clean-up Kit (MoBio

Laboratories, Inc, Solana Beach, CA, USA), following the manufacturer’s instructions.

However, when samples were combined with the Spinbind buffer the pH increased to

above 8.5, so this was adjusted to less than 7.5 by adding 10 μL of 3M Sodium Acetate

(molecular biology grade, Gene Works, Hindmarsh, S.A.).

Approximately 80-100 ng of fungal DNA was digested with 3 μL of Hin6I restriction

enzyme (1 unit / μL) at 36 – 37°C for 3 hours, using a reaction volume of 25 μL. After

this time, 75 μL of ice cold ethanol was added to each tube to halt the reaction. Then 2.5

μL of 5 M NaCl was added to each tube with gentle mixing and incubation overnight at

-20°C to precipitate the DNA. The tubes were then centrifuged at 4°C for 30 minutes

before the supernatant was removed. The DNA was dried before being dispatched for

fragment analysis. Fragment analysis was performed at the Australian Genome Research

Facility Ltd (AGRF) at The Walter and Eliza Hall Institute in Melbourne, Australia. The

60

3730 platform (Applied Biosystems, Carlsbad, CA, USA) capillary electrophoresis

system was used.

Fragment size and peak area data for each sample were tabulated, and fragment sizes

were rounded to the nearest whole number. Peaks less than 30 base pairs in length were

disregarded, to exclude possible contributions from primers. The data for each sample

were normalized, by converting each peak area to a percentage of the total peak area for

that sample. The peak rejection threshold was determined by comparing the loss in data

peaks when all data comprising less than 0.01, 0.1, 0.25, 0.5, 1 and 3% of the total

sample data were removed. The threshold was set at the level where the loss of T-RFs

per profile was minimal and sufficient T-RFs remained to analyse the community profile

(Sait et al, 2003) (See Appendix 1). Data were normalised and a minimum threshold of

0.25% of total peak area was applied. This serves to remove bias stemming from

differing amounts of PCR product being subjected to gel electrophoresis, while having a

minor effect on the overall area of the remaining T-RF peaks (Rees, Baldwin et al. 2004;

Sait, Galic et al. 2003).

Data were analysed with multi-dimensional scaling (MDS) using Primer 6 for Windows,

version 6.1.6 (Primer-E Ltd, Plymouth, UK). The data were presence/absence

transformed, and a Bray-Curtis resemblance matrix was then generated analysing

between samples. This resemblance matrix was analysed using MDS with 10 restarts.

Hierarchical Cluster analysis, in group average cluster mode, was used to generate

clusters that were then overlaid on MDS plots. Analysis of Similarity (ANOSIM) (Clarke

and Warwick 2001), was performed, and the global R statistic was calculated to

determine the significance of observed separation between groups.

3.3 Results

The method of DNA extraction from wetland sediments and plant material proved

successful in most instances. The use of the “touchdown” thermocycler program in PCR

proved suitable for amplifying fungal DNA, and the majority of the product obtained was

shown to be less than 500bp in length. This confirmed that the ITS2 section of the

ribosomal DNA was amplified. Terminal restriction fragments of 97 different lengths

were isolated. Fragments ranged in length from 35 to 510 bp.

61

3.3.1 Sediments and vegetation

The most important factor in structuring fungal communities in Normans Lagoon was

substrate (Figure 3.3). At the 25% similarity level sediment samples and flocculent

material from the sediment surface clustered into a group, while plant detritus formed a

separate group. Sediment and plant substrates were dissimilar (ANOSIM: R=0.763;

P<0.005), while floc and sediments were not distinguishable (pair-wise ANOSIM: R= -

0.067; P=0.563). The presence of outlying samples indicated high variability in the

structure of fungal communities. Fungal communities present on Eucalyptus sp. leaves

were dissimilar to those on sediments (pair-wise ANOSIM: R=0.716; P<0.005) and floc

(pair-wise ANOSIM: R=0.630; P= 0.100).

Figure 3.3: MDS plot of all T-RFLP data collected from Normans Lagoon. Sediment samples are

shown in black, floc samples are shown in grey and plant detritus samples are shown in white.

Symbols, as shown in the legend, have been used to differentiate between substrate types.

3.3.2 Site effects

Another factor influencing the structure of fungal communities was location, although

the data indicated that location was a less important factor than substrate. Sediments from

the Murray River at Doctors Point, and the east and west sites at Normans Lagoon

formed clusters at approximately 30% significance level (Figure 3.4). There was a low

level of separation between the east, west and Doctors Point sites (ANOSIM: R=0.101;

P<0.05), indicating that variability between sites was only slightly larger than variability

within sites. Differences between the east and west sites, the east site and Doctors Point,

and the west site and Doctors Point (Pair-wise ANOSIM: R=0.233, P<0.05; R=0.514,

62

P<0.05; R=0.346, P=0.063, respectively) were minor, but showed a trend towards

increasing difference with increasing distance between sites.

When sediment and floc samples were considered separately to plant substrates there was

some low level separation between groups (ANOSIM: R= 0.302; P<0.05). Sediment

samples taken from the Murray River at Doctors Point differed from samples in the east

and west (Pair-wise ANOSIM: R=0.514, P<0.05; R=0.346; P=0.063, respectively) of the

lagoon (circles overlaid on Figure 3.4). This indicated a possible difference between

fungal community structure in river and wetland sediments.

Figure 3.4: MDS plot of T-RFLP data from sediment and floc samples. Samples from the west

lagoon site are shown in black, samples from the east lagoon site are shown in grey, and samples

from the river at Doctors Point are shown in white. Symbols indicate sediment moisture class, i. e.

dry (star), damp (diamond), water’s edge (squares), inundated (hexagons), deep (circles) and reed

bed (triangles). Overlaid circles are intended to indicate site groups and have no statistical meaning.

3.3.3 Moisture effects

Samples were classified according to a relative scale as either “dry”, “damp” or “wet”.

Moisture content was not measured. All inundated samples were considered to be “wet”.

“Dry” sediments were not inundated and had been exposed for some time. “Damp”

63

sediments had been inundated recently but were exposed to the air at the time of

sampling. Some separation of sediment and floc samples according to moisture class was

evident (ANOSIM: R=0.361; P<0.05), although differences between dry and damp, dry

and wet, and damp and wet classes (Pair-wise ANOSIM: R=0.407, P=0.100; R=0.392,

P<0.05; R=0.331; P<0.05, respectively) were small. Where plant and sediments from the

west site were considered together, the level of separation remained low (ANOSIM: R=

0.382, P<0.05). However, when sediments (only) from the west site were considered

separately, moisture classes were found to have dissimilar fungal communities

(ANOSIM: R=0.773; P P<0.005) (Figure 3.5). Dry sediments differed from damp and

wet sediments (Pair-wise ANOSIM: R=0.500, P=0.333; R=0.410, P=0.091, respectively)

but the level of significance was low due to a small number of replicates for damp and

dry sediments. This suggested that variation due to substrate and site were more

important in determining the structure of fungal communities in Normans Lagoon than

moisture class, but that moisture effects may have been important within sites and

substrate types.

Figure 3.5: MDS plot of T-RFLP data from sediment and floc samples from the Normans Lagoon

west site. Symbols indicate sediment moisture class, i. e. dry (stars), damp (diamonds), water’s edge

(squares), inundated (hexagons), deep (circles), and reed bed (triangles). Dry samples are shown in

white, damp samples in grey and wet sediments in black. Overlaid circles illustrate samples are

grouped by sediment moisture content, but have no statistical meaning.

64

3.3.4 Response of fungal community to substrate species differences

Plant detritus samples showed little separation by site (ANOSIM: R=0.253, P=0.079),

with the species of the plant substrate (excluding bark and willow due to a low number of

replicates) appearing to influence fungal community structure more strongly (Figure 3.6).

The fungal community structure differed between Eucalyptus sp., Juncus sp. and

Cyperus sp. leaves as they decomposed in the wetland (ANOSIM: R = 0.573; P<0.005),

but while fungal communities on eucalypt leaf samples were variable, they differed from

Cyperus sp. (Pair-wise ANOSIM: R=0.741; P<0.05) but not from Juncus sp. (Pair-wise

ANOSIM: R=0.250; P=0.200), while Cyperus sp. and Juncus sp. were dissimilar

(R=0.661; P=0.067). Separation between detritus samples based on moisture status was

not significant (R = -0.042, P = 0.509).

Figure 3.6: MDS plot of T-RFLP data from plant detritus samples from Normans Lagoon. The west

site is shown in black, while the east site is shown in grey. Overlaid circles indicate plant species

groupings but have no statistical meaning.

3.4 Discussion

It was possible to amplify the ITS2 region of the rDNA specific to fungal communities

from sediments and plant detritus using PCR. This indicates that fungi are present in

Normans Lagoon and are thus likely to be present in similar floodplain wetlands in

65

Australia. Analysis of terminal restriction fragment length from a range of substrates

sampled from throughout the wetland and the adjacent river shows that the structure of

these fungal communities is influenced by substrate composition, site and moisture class.

3.4.1 Changes in fungal community structure with time

The fungal community composition of submerged leaves is known to change over time

(Canhoto and Graça 1996; Pattee and Chergui 1995; Suberkropp and Klug 1976). As all

fungal substrates were collected from the wetland without knowledge of their total

exposure time, exposure time is a possible source of variation in fungal community

structure on plant detritus. Specifically, this effect may partly explain the high variability

observed in the fungal community structure on Eucalyptus sp. leaves, which may float on

the water’s surface for a period, before becoming submerged and possibly buried in

sediments. Each of these changes is likely to influence the composition of the fungal

community, and succession due to the changing composition of the substrate induces

further changes in fungal community structure.

The composition of fungal communities on submerged plant litter may be strongly

influenced by randomly arriving “founder” species that tend to maintain a presence in the

community over time (Fischer, Bergfur et al. 2009; Pattee and Chergui 1995), even

though their abundance may change. The presence/absence transform used gives the

same level of importance to all species presence, regardless of abundance. These founder

species may increase variability in fungal community structure between samples. The

importance of these temporal factors can not be inferred using the current data set.

3.4.2 Fungal community structure differences in Normans Lagoon

Fungal communities found in surface flocculent material (floc), possibly associated with

detritus from Azolla filiculoides, were not significantly different from sediment samples.

This was not unexpected as floc samples may have contained some sediment. However,

fungal community structure on plant detritus differed significantly from those on

sediment and floc. These differences between fungal communities are likely to be

associated with differing requirements for nutrition, different tolerances to sediment

anoxia, and the ability to compete with other micro-organisms for resources.

Where plant detritus is decomposed in rivers, fungi dominate the decomposer community

when particle size is large. When particle size is small, bacteria are the dominant

66

decomposers (Suberkropp and Klug 1976). Fungi have been shown to selectively

colonise resources depending on their quantity, quality and distribution, and in a manner

that minimises energy expenditure (Boddy 1993; Bolton and Boddy 1993). Plant detritus

is a large, higher quality (more nutrients, less recalcitrant compounds), concentrated

resource that the fungal mycelium can consume as it extends, while organic resources in

the sediments are smaller, more dispersed, and lower quality. This suggests that aquatic

fungi would be likely to colonise plant detritus in preference to sediments. As bacteria

are not constrained within a multicellular form, they may be transported passively by

flow to positions close to the resource and obtain energy from within the sediments with

greater efficiency. Previous research investigating microbial dynamics in wetland

sediments (Boon, Virtue et al. 1996) indicates that bacteria dominate decomposition.

Inundated wetland sediments are usually anoxic (Boon and Mitchell 1995), and while the

absence of oxygen does not inhibit the metabolic activities of a range of anaerobic

sediment bacteria, few aquatic fungi are able to survive these conditions in active life

stages (Cooke and Whipps 1993). For these reasons we can expect the fungal community

characteristic of sediments and floc in Normans Lagoon to be composed of species or life

stages tolerant of low oxygen conditions and facultative anaerobes such as yeasts,

chytrids and Oomycetes. DNA derived from dormant propagules of fungal forms

inhabiting the water column or floodplain soils may also be represented by terminal

restriction fragments (Lee and Taylor 1990). Indeed, it is possible that propagules

comprise a large proportion of the sediment fungal community. If the active life stages of

species producing these propagules are associated with Azolla filiculoides communities,

this would explain similarities in fungal community structure between floc and

sediments. Alternatively, the fungal community on floc samples may have been

associated with sediment particles that comprised a proportion of the floc.

Fungal community structure on decaying leaves differs with leaf species (Canhoto and

Graça 1996; Thomas 1992). In particular, the leaves of Eucalyptus species support

different fungal assemblages to both deciduous European species (Canhoto and Graça

1996) and other Australian indigenous trees such as Acacia sp. (Thomas 1992; Thomas,

Chilvers et al. 1992). The leaves of Eucalyptus species contain high levels of tannins and

essential oils that have been shown to inhibit growth in aquatic hyphomycetes (Canhoto

and Graça 1999) and cause a lag in fungal colonisation with respect to other tree species

67

(Graça, Pozo et al. 2002). For these reasons it is likely that Eucalyptus leaves support

specialised fungal taxa (Thomas, Chilvers et al. 1992). However, all of the previous

studies have examined fungal colonisation of Eucalypt sp. leaves in rivers, and it is

possible that fungal colonisation patterns are different in wetlands. The limited results for

these leaves from Normans lagoon show high variability between replicates, but indicate

some similarity with other submerged plant detritus.

Sediment sample outliers consisted of dry and damp exposed or recently inundated

sediments that may have contained plant roots or other plant material. These samples

clustered with submerged Eucalyptus sp. leaves and bark, which are likely to be high in

lignin (Cork and Pahl 1996). It is likely that these substrates supported a distinctive

fungal community characterised by specialist fungal species capable of degrading

refractory organic material such as lignin.

The lignin content of submerged decaying leaves is also known to influence the structure

of the fungal community (Gulis 2001). Wood and bark are rich in lignin, while

Eucalyptus sp. leaves have a relatively high lignin content when compared with the

leaves of Salix sp., for example (Gessner and Chauvet 1994; Janssen and Walker 1999).

The degradation of lignified plant tissues requires specialised enzymes (Deobald and

Crawford 1992), so fungal species able to secrete these enzymes will have an advantage

on lignin-rich plant detritus, and this advantage will influence community structure.

There is a weak trend shown for fungal community structure in sediments and floc to

cluster according to location within the wetland, and in the river. This indicates some

spatial heterogeneity within the wetland. Heterogeneity may be driven, in part, by

differences in sediment texture. Visual examination suggested that sediments in the

Murray River were sandy, while those at the western side of the lagoon were silty, and

those in the eastern site were higher in clay. Sediment texture will influence gas diffusion

and will thus influence dissolved oxygen concentrations. Sedimentation rate will affect

the duration of substrate availability to aerobic decomposers.

While location influences fungal community structure it is not a simple variable, but is

composed of spatial differences in a range of environmental variables. These variables

might include vegetation, incident light, land use and prevailing winds, and their

68

interaction. Spatial differences in riparian and aquatic vegetation influence fungal

community structure in terms of the availability of substrates, but as it is a persistent

feature, it also influences the pool of available fungal propagules (Suberkropp 2001).

Shading may influence the growth of algae, which would in turn influence the structure

of fungal communities (Albariño, Villanueva et al. 2008). Land use that allows cattle to

access the wetland may result in loss of vegetation, erosion and nutrient enrichment

(Robertson, Klomp et al. 1997), which would in turn influence fungal community

structure. Prevailing winds are of significance due to the abundance of the floating fern,

Azolla filiculoides. This fern will tend to move with the wind and accumulate at the end

of the wetland, providing an abundant, nitrogen rich substrate for fungi at the down wind

end of the wetland. Thus, it is difficult to determine which aspects of a location are

influencing fungal community structure.

The effects of moisture status are difficult to discern, as substrate and site effects tend to

mask them. However, when these factors are controlled, some impacts of moisture status

are discernable. Firstly, dry and damp soil fungal community structures differ from

community structure in inundated sediments, although the depth of overlying water does

not appear to be important (although samples were collected from water depths of less

than 0.5m). This may indicate that the fungal community differs between moisture

limited terrestrial conditions and aquatic conditions, which are likely to be limited by

oxygen availability (Gribben, Rees et al. 2003). Damp soils, and sediments associated

with macrophyte stands, represent positions on a gradient between these two community

types. An oxygen availability gradient is further supported by small differences in fungal

community structure between standing dead and inundated reed (Cyperus and Juncus sp.)

detritus. The willow leaf (Salix sp.) was found floating on the surface of the wetland, and

may support a fungal community adapted to aerial conditions, hence its affinity with

standing dead reeds.

3.4.3 The ecological significance of these findings

The data presented here suggest that fungal community structure is influenced by

substrate, site factors and moisture status. This implies that fungal community diversity is

strongly dependent upon the diversity of the plant detritus available, and is vulnerable to

the environmental forces that threaten plant diversity. There are indications that elements

of the sediment fungal community are independent of those found on plant detritus, and

69

these elements may include propagules and vegetative forms of species that inhabit the

water column, parasitising algae and other organisms, or decomposing dead biota from

the water column as it encounters the sediment surface. Thus, the sediments may contain

a “seed bed” of fungal propagules that remain dormant during unfavourable conditions,

such as anoxia or desiccation, and are able to colonise suitable substrates when

conditions are more favourable. While this is a positive attribute in terms of leaf

conditioning and decomposition, it may have negative aspects associated with the

prevalence of parasitic and pathogenic fungal species.

3.5 Conclusions

It is clear from the data presented in this chapter that fungi were present in all of the

substrates examined from Normans Lagoon, and strong differences were seen between

the fungal community structures in the sediments and on plant detritus. While lack of

replication in this pilot study gives me reason to defer drawing further conclusions, the

data suggest that factors such as the species of plant material, site related variables and

the moisture status of sediments may be important structuring factors for fungal

communities in floodplain wetlands.

We have seen that the effects of substrate, moisture treatment and site interact to produce

variation in fungal community structure. It is also likely that community structure

changes with time. In chapters to follow, I will explore these effects further. In Chapter

4, I present a laboratory experiment in which I compare fungal community structure on

two leaf types between terrestrial and aquatic conditions, and follow these changes over

time. In addition, the abundance and activity of these fungi are measured. In Chapter 5, I

examine the relationship between sediment moisture content and fungal community

structure in the sediments of a floodplain wetland, and measure the abundance of these

fungi. In Chapter 6, fungal community structure and other fungal dynamics on

submerged E. camaldulensis leaves are compared between rivers and wetlands at three

locations.

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Fungal activity on Eucalyptus camaldulensis leaves under aquatic

conditions is enhanced by terrestrial aging.

4.1 Introduction

In Chapter 3 I showed that fungi present in Australian floodplain wetlands are colonising

a variety of plant substrates as well as sediments. In this chapter I further explore the

activities of these fungi by examining how the biomass and activity of the fungal

community changes over time, and how this affects the decomposition of the substrate. A

laboratory experiment was used to examine fungal dynamics on fresh and terrestrially

aged leaves of the river red gum (Eucalyptus camaldulensis, Dehnh. Myrtaceae) under

moisture conditions designed to simulate those on the floodplain and in a floodplain

wetland.

E. camaldulensis is the dominant tree species in floodplain ecosystems throughout the

Murray-Darling Basin, and is known to be important in aquatic food webs (Reid, Quinn

et al. 2008) and carbon cycles (Baldwin 1999; Francis and Sheldon 2002; O'Connell,

Baldwin et al. 2000; Robertson, Bunn et al. 1999). Under natural flow regimes, these

wetlands flood predominantly in winter / spring, and summer / autumn flooding was rare

prior to river regulation (Bren 1988). As E. camaldulensis leaves fall predominantly in

summer (Glazebrook and Robertson 1999), abscised leaves lie on the dry floodplain for

some months prior to inundation, undergoing terrestrial aging. River regulation has lead

to more frequent flooding of wetlands in summer (Bren 1988; Maheshwari, Walker et al.

1995), resulting in more frequent inundation of freshly abscised leaves.

Fungal dynamics on leaves are strongly influenced by the chemical composition of the

substrate (Canhoto and Graça 1996). Fresh eucalypt leaves have been considered to be

poor fungal substrates due to high contents of oils, tannins and polyphenols (Canhoto and

Graça 1999; Graça, Pozo et al. 2002). However, when eucalypt leaves spend time on the

floodplain as part of the litter layer, they are exposed to leaching and ultra violet

radiation, and may be colonised by microbes or grazed upon by fauna. This alters their

chemical composition (Baldwin 1999), and is likely to alter their quality as a fungal

substrate.

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Fungal dynamics have never been examined in Australian floodplain wetlands. A few

studies have examined the aquatic decomposition of plant litter under lentic conditions,

without specifically examining the activity of fungi. These studies suggest more rapid

decomposition of leaves under submerged conditions (Glazebrook and Robertson 1999)

and invertebrate activity (Briggs and Maher 1983). Differences in decomposition rates in

wetlands were attributed to leaf composition (Janssen and Walker 1999), the presence of

plant-derived oils (Bailey, Watkins et al. 2003), season of inundation (Watkins, Quinn et

al. 2010) and nutrient dynamics (Shilla, Aseada et al. 2006). Studies examining the

fungal contribution to leaf decomposition rates in wetlands in other countries have shown

that rates are not influenced by differing flooding and drying frequencies (Inkley,

Wissinger et al. 2008), and are similar in the main channel and associated floodplain

wetlands (Baldy, Chauvet et al. 2002). Anoxic conditions in standing waters and

sediments are also known to slow or halt fungal activity (Medeiros, Pascoal et al. 2009a;

O'Connell, Baldwin et al. 2000; Pattee and Chergui 1995).

Fungal community structure on allochthonous litter can be expected to differ between the

terrestrial conditions on the floodplain and aquatic conditions within the wetlands (Pattee

and Chergui 1995). While terrestrial Eucalyptus sp. leaf litter is colonised by fungi such

as Mucor sp., Rhizopus sp., Aspergillus sp., Penicillium sp., Trichoderma sp., Alternaria

sp., Cladosporium sp. and Fusarium sp. (Eicker 1973), leaves submerged in still water

are more likely to be colonised by aquatic hyphomycetes e.g. Alatospora sp.,

Anguillospora sp., Clavariopsis sp., Flagellospora sp., Lunulospora sp., Tetrachaetum

sp., Tricladium sp. (Thomas, Chilvers et al. 1989; Thomas, Chilvers et al. 1992), aero-

aquatic hyphomycetes and stramenopiles (Pattee and Chergui 1995; Voglmayr 1997).

The transition from a terrestrial assemblage to an aquatic assemblage has not been

investigated on the leaves of Eucalyptus species.

Differences in species composition on leaves in terrestrial and aquatic environments are

likely to result in differences in total fungal biomass and losses of mass and carbon from

decomposing leaves, as a result of competitive interactions (Mille-Lindblom, Fischer et

al. 2006a) and differing enzymatic capabilities (Abdel-Raheem and Shearer 2002;

Abdullah and Taj-Aldeen 1989; Valikhanov 1988). As fungal colonisation occurs in two

phases (Verma, Robarts et al. 2003), we can expect fungal dynamics to reflect

exploitation of starches and pectins by early colonisers, and the use of cellulose and

72

lignin one to two months later by secondary colonisers (Dix and Webster 1995; Pattee

and Chergui 1995; Sinsabaugh, Moorhead et al. 1994). However, this pattern of activity

will be influenced by the availability of moisture and oxygen, and by the chemical

properties of the fungal substrate.

Here, I test the hypothesis that fungal community structure, biomass accumulation and

extra-cellular enzyme activity differ between fresh and terrestrially aged E.

camaldulensis leaves, and that these differences influence mass and organic matter loss

from the leaves. I further hypothesise that fungal community dynamics differ between

terrestrial and aquatic conditions and with incubation time. Freshly abscised and

terrestrially aged E. camaldulensis leaves were placed in mesocosms simulating dry and

moist terrestrial conditions and aquatic conditions, and fungal dynamics were compared

between moisture treatments, substrate and over time.

4.2 Methods

4.2.1 Mesocosms

Fresh and aged leaves were obtained in May 2007 from the floodplain at Normans

Lagoon, Albury, N.S.W (146°56’06”E, 36°05’42”S). This site is described in Chapter 3.

Whole leaves were selected, and those damaged due to insect activity were not excluded.

Leaves removed from the litter layer surface were considered to be aged if they no longer

had any green colouration. Fresh leaves were removed from branches that had blown to

the ground in a storm 3 days earlier (sensu Baldwin et al. 1999). In the laboratory, both

fresh and aged leaves were stapled into packs consisting of three leaves, labelled with a

square of water-proof paper, weighed (DM0) and dried in a dehydration oven at 50°C for

2 weeks.

Wetland sediment and wetland water were also obtained from Normans Lagoon. Living

vegetation was removed from sediments, which were then mixed until homogenous, and

then oven dried at 30°C for 7 days. Water was filtered through muslin cloth to remove

large particles and invertebrates.

Six mesocosms were assembled that consisted of 60 x 30 x 30 cm fish tanks containing

4L of homogenised wetland sediment spread evenly over the base to a depth of 2 cm

(Figure 4.1). Sediments were covered in nylon mesh to prevent leaf burial. Leaf packs of

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fresh and aged leaves were distributed randomly between moisture treatments. Weighted

wire mesh was placed over the leaves in all treatments. This was intended to ensure

leaves in the wet treatments were submerged. Mesocosms were kept at 20°C, under

predominantly dark conditions. The tanks were loosely covered with aluminium foil to

help to maintain moisture status without impeding air flow.

Treatments are described as dry, moist and wet. In the dry treatment, no water was

added to these tanks over 140 days of the experiment. In the moist treatment, enough

wetland water was added to the sediments so that they were saturated, but no water

pooled at the surface. In the wet treatment sediments were saturated, and a water depth of

approximately 15 cm was maintained by occasional addition of filtered wetland water.

Wet tanks were aerated with a bubbler to prevent the onset of anoxia. Leaf packs were

removed from the treatment tanks after 1, 2, 4, 8, 16, 32, 64, 100 and 140 days.

Preservation methods were specific to the analytical method.

Figure 4.1: A schematic diagram of mesocosm design. All mesocosms were comprised of fish tanks,

with a 2cm layer of wetland sediment in the bottom. This was covered with nylon mesh, and leaf

packs were placed upon the mesh. Leaf packs were covered with weighted wire. In wet treatments,

filtered wetland water was added to a depth of 15cm.

Fish Tank

Water

Weight

Leaf Pack

Sediment Nylon Mesh

74

4.2.2 Leaf mass loss and loss on ignition

Dried leaf packs were weighed prior to placement in the tanks (DM1 for Day 0 leaves).

On each sampling occasion, 3 leaf packs for each treatment were transferred into pre-

fired and pre-weighed crucibles. Wet leaves were gently rinsed under running tap water

to remove sediment from the leaf surface. The wet weight (WM) was recorded prior to

drying at 105°C for 48 hours. Crucibles were brought to room temperature in a desiccator

then re-weighed to determine the dry mass (DM1) of the leaves. Following this, leaves

were ashed at 550°C for 1 hour to determine the loss of mass on ignition (AFDM). Mass

loss as a percentage of the original mass was calculated as (((DM0 – DM1) / DM0) x 100).

Loss on ignition (%), an estimate of organic matter content, was calculated as (((DM1 –

AFDM) / DM1) x 100).

Data for mass loss and change in organic matter content were analysed using two-way

analysis of variance (ANOVA) with replication (3) for each sampling time. Leaf type

(fresh or aged) and treatment (dry, moist or wet) were compared with mass loss / organic

matter as the dependent variable and time was as a covariant, pair-wise for treatments

(Bärlocher 2005). Two-tailed t-tests (α=0.05) were used to test differences between

treatments and leaf type. Data analyses were performed using SPSS Statistics 17.0 (IBM,

Chicago, USA). The slope of the mass loss curve (m) was also determined as a means of

comparing mass loss rate between leaf types.

4.2.3 Fungal biomass

Ergosterol content of leaves was determined using the method published by Gessner

(2005). Triplicate samples from all times were analysed for fungal biomass. Briefly, leaf

samples were stored in HPLC grade methanol (Merck, Darmstadt, Germany), in the dark

and at room temperature prior to analysis. A 12mm diameter disc was cut from each of

three leaves, weighed and pooled for analysis. An additional set of three leaf discs was

cut from the same leaves to determine the ratio of wet mass to dry mass and AFDM. The

lipid fraction was extracted by refluxing in alkaline methanol for 30 min at 80°C.

Extracts were filtered through 25mm diameter GF/C filters before purification with SEP-

Pak solid phase extraction cartridges (Vac RC, tC18, 500mg of sorbent, Waters, Milford,

MA, USA).

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Ergosterol was quantified using a Waters HPLC system equipped with in-line degasser

(AF), 717 plus auto-sampler, 600 pump controller and 2996 Photo diode Array detector

(Waters, Milford, MA, USA). A 250mm x 4.6mm LiChrospher 100 RP-18 5u column

(Grace Davison Discovery Sciences, Deerfield, IL, USA) was maintained at 33°C. The

mobile phase was 100% HPLC grade methanol, on an isocratic gradient with a flow rate

of 1.4 ml/min. Samples were dissolved in 100% HPLC grade propan-2-ol and 10µL was

injected onto the column, with a run time of 20 minutes. The ergosterol peak was

detected at 282nm after 12.060 minutes with a retention time window of 0.605 minutes.

Data was processed using EmpowerTM

software (Waters, Milford, MA, USA). Peak area

was integrated using a traditional algorithm, with a minimum peak height of 420 au,

minimum peak area of 4930 and a rejection threshold of 50.

Peak area was converted to ergosterol concentration based on comparison with a standard

curve. Ergosterol content was then converted to fungal biomass using an estimated

conversion factor of 5.5mg ergosterol per gram fungal biomass (Gessner and Chauvet

1993) and expressed as mg of fungal biomass per cm2 leaf area. Fungal biomass data

were analysed using 3-way ANOVA on the SPSS software package (IGM, Chicago,

USA), analysing between leaf type, treatments and times (α=0.05). Paired sample t-tests

were also carried out in SPSS.

4.2.4 Extracellular enzyme activity

4.2.4.1 Hydrolytic enzyme activity

Leaf samples were preserved by freezing at -20°C. Three leaves per sample were

homogenised in 50ml of 50mM Na2HCO3 (pH 8.0), diluted in 1:1 with 50mM Na2HCO3,

and then incubated on a shaker table for 30 minutes at 20 °C. Three replicate 150 µl

aliquots of samples and a boiled control were combined 1:1 with substrate in a 96 well

microtitre plate. Four different 4-methylumbelliferone (MUB) substrates (Sigma, St

Louis, MO, USA) were used at a final concentration of 500 µM (Hendel and Marxen, in

Graça et al., 2005). These were MUB-β-D-glucoside, MUB-α-D-glucoside, MUB-β-D-

xyloside and MUB-β-D-cellobioside.

Fluorescence (Ex:Em = 355nm:460nm), was measured at 20°C using a Fluoroskan II

plate reading fluorimeter (Labsystems, Finland), and recorded every 10 minutes at 0.5

sec/well for a 3 hour incubation period. Fluorescence was converted to nmol MUB using

76

a standard and standards + quench curve. The rate of increase in MUB concentration for

each well was determined using linear regression. The rate of change of the boiled

control was then subtracted from the mean activity rate to give the enzyme activity.

4.2.4.2 Oxidative enzyme activity

Phenol oxidase and peroxidase activity was measured using L-3,4-

dihydroxyphenylalanine (L-DOPA) (≥ 99%, Fluka Cat no. 37830, Sigma-Aldrich,

Steinheim, Germany) as the electron donating substrate (Hendel, Sinsabaugh et al. 2005).

Leaf samples were preserved by freezing at -20°C. Three thawed leaves per sample were

homogenised in 50 ml of MilliQ Water, diluted 1:10 in 50mM NaCH3COO (pH 5.0)

(acetate buffer), and then incubated on a shaker table for 30 minutes at 20°C. Samples

were filtered through a GF/C filter prior to measuring absorbance. For phenoloxidase

activity, 2 ml of filtered homogenate was combined with 2ml of 5mM L-DOPA. Four

analytical replicates and 2 blanks were used. Blanks consisted of 2ml of filtered sample

with 2 ml of acetate buffer. The assay for peroxidase was similar to the phenoloxidase,

except that 0.2ml of 0.3% hydrogen peroxide was also added. Peroxidase controls

contained 2 ml of filtered sample, 2 ml of acetate buffer and 0.2 ml of 0.3% hydrogen

peroxide. The samples were incubated in the dark for 120 minutes on a shaker table at

20°C. Absorbance of samples was read at 460nm on a Cary spectrophotometer (Varian,

Palo Alto, CA, USA). Activity was calculated according to Hendel et al., (2005).

Data for the activity of hydrolytic and oxidative enzymes were compared between

treatments and substrate types, and over time using the t-Test for paired samples for

means. Pearson’s R was used to compare fresh and aged leaves in each treatment, and for

each enzyme, over days 0-16 and days 64-140. The activity of all enzymes, in all

treatments at all times was analysed using multivariate 2-way crossed ANOSIM analysis

on a 2-stage correlation matrix (Clarke, Somerfield et al. 2006) to determine whether

consistent patterns were present in overall enzyme activity (Primer 6 for Windows,

version 6.1.6, Primer-E Ltd, Plymouth, UK). The activity of individual enzymes was then

compared between treatment, leaf type and times (α=0.05) using the SPSS software

package (IGM, Chicago, USA) to perform univariate 3-way ANOVA on normalised

data.

4.2.4.3 Fungal community structure

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Fungal community structure was assessed for samples from 0, 8 and 100 days incubation,

using T-RFLP (see Chapter 3.2.3). After removal from the mesocosms, leaf packs were

stored at -20°C in 70 ml sterile jars. Three 5mm diameter leaf discs were cut from each

of three leaves in the leaf pack and frozen at -80°C overnight. Leaf discs were ground in

a 2ml safe-lock tubes (Eppendorf AG, Hamburg, Germany) using a sterile mini-pestle,

centrifuged for 1 minute at 10 x g and any supernatant removed. DNA was then extracted

from the leaf material using the “UltraClean Soil DNA Isolation Kit” (MoBio

Laboratories, Inc. California, USA) and the “Alternative Protocol (For maximum

yields)”(Mo Bio Laboratories 2003). Extracted DNA was stored at -20°C prior to

amplification. Some DNA extracts retained colour derived from the leaf substrate, which

would interfere with amplification. These samples were further purified using the

potassium acetate purification method (McVeigh, Munro et al. 1996). Polymerase Chain

Reaction (PCR) was used to amplify the internal transcribed spacer region II (ITS2) of

fungal ribosomal DNA (rDNA), found between 5.8S-coding sequence rDNA and Large

sub-unit coding sequence (LSU) of the ribosomal operon (Martin and Rygiewicz 2005).

The PCR reaction conditions, PCR product clean-up, restriction digest, fragment analysis

and data analyses were as described in Chapter 3.2.3.

4.3 Results

4.3.1 Leaf mass loss and loss on ignition

4.3.1.1 Mass loss from leaves

Mass loss from fresh and aged leaves in the dry, moist and wet treatments over 140 days

is shown inFigure 4.2. Moist aged leaves at day 140 exhibited a gain in mass (not

shown), with a corresponding large decrease in percentage organic matter. This is

consistent with contamination of these samples with sediment, and therefore these data

points were excluded from calculations. Moist fresh leaves showed a small mass loss on

the same day, but as a decrease in the proportion of organic matter in the leaf was not

observed these data points were not excluded.

Mass loss differed significantly between fresh and aged leaves, and between treatments,

and a significant interaction between leaf type and treatment was detected (ANOVA:

P<0.005). No significant mass loss was observed in the dry treatment (Figure 4.2, (i)). In

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the moist treatment (Figure 4.2, (ii)) mass loss differed between fresh and aged leaves.

The maximum mass loss was seen in the wet treatment (Figure 4.2, (iii)).

Aged leaves showed no significant mass loss in the moist treatment (Figure 4.2, (ii),

while fresh leaves lost 23% of their initial dry mass after 100 days (t-test: P<0.05). In the

wet treatment (Figure 4.2, (iii)) the most significant differences between leaf types

occurred during the first week of incubation. Over this time, fresh and aged leaves lost

25% and 10% of their initial dry mass, respectively (t-test: P<0.005). Over the remaining

decomposition time the rate of mass loss was higher for aged leaves (m= 0.09 for fresh

leaves; m=0.18 for aged leaves).

4.3.1.2 Mass loss on ignition

When considering all sample data together, the percentage of organic matter (Figure 4.3)

differed significantly between treatments (ANOVA: P<0.005). When comparing

between treatments, changes in organic matter content over time were not significantly

different between dry and moist, but wet was significantly different to the other

treatments (t-test: P<0.05), due to larger total losses of organic matter.

When considering all sample data together differences were observed between fresh and

aged leaves, but these were not significant (ANOVA: P=0.051). When data for each

treatment was analysed individually, a significant difference between leaf types was seen

in dry and wet treatments (t-test: P<0.05).


The concentration of ergosterol in the leaves indicated that fungal biomass was higher on

aged leaves than on fresh leaves (3-way ANOVA: P<0.05), and differences between

treatments were significant (3-way ANOVA: P<0.005). While in the dry treatment

biomass on fresh and aged leaves was always significantly different (t-test: P<0.005), in

the other treatments differences decreased over time (Figure 4.4). Wet fresh leaves

differed from wet aged leaves prior to 64 days incubation (t-test: P<0.005) as fungal

biomass was very low on fresh leaves. After 64 days, this difference was not observed. In

the moist treatment biomass differed prior to 16 days (t-test: P<0.005), but no difference

was observed after that time.

79

Fresh leaves carried no fungal biomass on day 0, and very little subsequently

accumulated on fresh leaves in the dry treatment. In the moist treatment, fungal biomass

on fresh leaves increased until maxima were observed between 32 and 100 days, and

then declined. Wet fresh leaves had low fungal biomass prior to day 64. Maximum

fungal biomass was observed at day 64, and biomass decreased thereafter.

Prior to incubation, aged leaves supported approximately 3.5 mg.cm-2

of fungal biomass.

The temporal pattern of changes in fungal biomass on these leaves was similar across

treatments. Biomass accumulated in two phases, with maxima at 2-4 and 32-64 days and

minima at 16 and 140 days, but the amount of biomass at these points varied between

treatments. Overall maximum fungal biomass occurred after 60 days in the moist

treatment for both fresh and aged leaves.

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Figure 4.2: Mass loss in the dry (i), moist (ii), and wet (iii) treatments over 140 days. Fresh leaves are

indicated by closed symbols while aged leaves are indicated by open symbols. Mass loss, on the y-

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axis, is shown as percentage of the mass of the leaf pack prior to incubation. Error bars indicate the

standard error of the data point.

Figure 4.3: Changes in organic matter content of leaf packs over 140 days are shown for the dry (i),

moist (ii), and wet (iii) treatments. Fresh leaves are shown as closed symbols while aged leaves are

82

shown as open symbols. Organic matter content (y-axis) is expressed as a percentage of the leaf dry

mass. Error bars indicate the standard error of the data point.

83

Figure 4.4: Fungal biomass (mg.cm-2

) on eucalypt leaves, as calculated from ergosterol

concentration. Differences in fungal biomass between aged (open symbols) and fresh (closed

84

symbols) leaves are shown for the dry (i), moist (ii) and wet (iii) treatments. Error bars indicate the

standard error of the data point.

4.3.3 Enzyme activity

4.3.3.1 Combined enzyme activity

Multivariate analysis of combined hydrolytic and oxidative enzyme activity data

indicated differences between fresh and aged leaves (ANOSIM: R=0.346; P<0.05; see

Figure 4.5) and between treatments (ANOSIM: R=0.337; P<0.05). For individual

enzymes, significant differences were found between treatments, leaf type, with time, and

for all interaction affects (ANOVA: P<0.05).

4.3.3.2 Hydrolytic enzyme activity

Hydrolytic enzyme activity differed between fresh and aged leaves in the dry treatment

(ANOSIM: R=0.442; P<0.05), but not in the moist (ANOSIM: R=0.188; P<0.05) or wet

(ANOSIM: R=0.162; P<0.05) treatments. While activity on dry fresh and moist fresh

leaves was generally close to zero, activity on dry aged and moist aged leaves was

usually higher but decreased with time. Significant differences in activity over time on

fresh (R=0.4; P<0.05) and aged leaves (R=0.332; P<0.05) were only observed in the wet

treatment, where overall maximum activity was seen after 60 days.

The activity of cellobiohydrolase (Figure 4.6) and β-1,4-glycosidase (Figure 4.7) are

indicative of the breakdown of cellulose. Cellobiohydrolase attacks the ends of cellulose

chains removing cellobiose (a glucose dimer), while β-1,4-glycosidase attacks cellobiose

to produce glucose monomers. Significant differences were observed between treatment,

leaf type and time (ANOVA: P<0.05). The data show that significant activity towards

cellulose was confined to the wet treatment (Figure 4.6 (iii)), although low activity was

observed on aged leaves in the dry and moist treatments at earlier sampling times (Figure

4.6 (i) and (ii)). Activity towards cellobiose occurred on aged leaves in the dry and

moist treatments (Figure 4.7 (i) and (ii)), with some activity occurring in fresh leaves

after 100 days in the moist treatment. Strong activity towards cellobiose occurred in the

wet treatment for both fresh and aged leaves (Figure 4.7 (iii)).

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The activity of starch degrading enzymes is indicated by α-D-glucosidase (Figure 4.8),

while activity of β-D-xylosidase indicates activity towards hemicelluloses (specifically

xylans) (Figure 4.9). Leaf type, and leaf type x treatment significantly affected the

activity of both of these enzymes (ANOVA P<0.05), while treatment and time x leaf type

also significantly affected β-D-xylosidase activity. The strongest differences in the

activity of these enzymes within a treatment occurred between moist fresh and moist

aged leaves (ANOSIM: R=0.396; P<0.05). These data suggest that starch and

hemicellulose breakdown was occurring on aged leaves in all treatments, with maximum

degradation rates occurring in the dry treatment (Figure 4.8 and Figure 4.9, (i)). On fresh

leaves, significant activity towards starch and hemicelluloses was only seen in the wet

treatment (Figure 4.8 and Figure 4.9, (iii)), although some activity was recorded on fresh

leaves in the moist treatment after 100 days (Figure 4.8 (ii) and Figure 4.9 (ii)).

4.3.3.3 Oxidative enzyme activity

The activity of phenol oxidase and peroxidase suggested that lignin breakdown was

occurring. Oxidative enzyme activity peaked early in the incubation for all treatments

and decreased with time (Figure 4.10 and 11). In the dry and moist treatments, higher

activity was seen on aged leaves during this time. Phenol oxidase activity from day 60

onwards was low in the dry treatment, moderate in the moist treatment and higher in the

wet treatment (Figure 4.10 (i), (ii) and (iii) respectively) and similar between leaf types.

However, phenol oxidase activity on aged leaves after 100 days was lower than on fresh

leaves for all moisture treatments. Similar levels of peroxidase activity were seen in the

dry and moist treatments (Figure 4.11 (i) and (ii)), with activity being close to zero from

day 60 onwards. Activity in the wet treatment (Figure 4.11, (iii)) continued at moderate

levels from day 60 onwards, with lower activity observed for aged leaves after 100 days,

than for fresh leaves. Maximum activity towards lignin was observed in the dry

treatment at day 4 (Figure 4.10 and 11, (i)).

The activity of phenol oxidase and peroxidase (Figure 4.10 and 11) differed between leaf

type and over time, with significant interaction effects (3-way ANOVA: P<0.005).

There were also significant treatment effects for phenol oxidase activity (ANOVA:

P<0.005).

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The pattern of activity of oxidative enzymes in the wet treatment differed from the

activity of hydrolytic enzymes. Where decreasing activity after 60 days was observed for

all hydrolytic enzymes for both leaf types in this treatment, oxidative enzyme activity

maintained moderate levels of activity, although activity on aged leaves was variable.

Fresh Leaves Dry Treatment

Aged Leaves Dry Treatment

Fresh Leaves Moist Treatment

Aged Leaves Moist Treatment

Fresh Leaves Wet Treatment

Aged Leaves Wet Treatment

3D Stress: 0.12

Figure 4.5: A three dimensional MDS plot based on a 2-stage resemblance matrix for oxidative and

hydrolytic enzyme data for all times, treatments and leaf types. Fresh leaves are represented by solid

symbols, while aged leaves are shown as open symbols. Treatments are symbolised by stars (dry

treatment), diamonds (moist treatment) and circles (wet treatment). Arrows indicate that the

direction of change in enzyme activity with different treatments differs between fresh leaves (solid

arrow) and aged leaves (dashed arrow). Fresh and aged leaves in the wet treatment have similar

enzyme activity.

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Figure 4.6: The activity of cellobiohydrolase on aged (open symbols) and fresh (solid symbols) E.

camaldulensis leaves in the dry (i), moist (ii) and wet (iii) treatments. Incubation time is indicated in

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days on the x-axis, while enzyme activity rate (µmol substrate converted per hour, per gram leaf dry

weight) is shown on the y-axis. Error bars indicate the standard error of the data point.

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Figure 4.7: The activity of β-D-glucosidase on aged (open symbols) and fresh (solid symbols) E.




90

Figure 4.8: The activity of α-D-glucosidase on aged (open symbols) and fresh (solid symbols) E.


91



92

Figure 4.9: The activity of β-D-xylosidase on aged (open symbols) and fresh (solid symbols) E.




94

Figure 4.10: The activity of phenol oxidase on aged (open symbols) and fresh (solid symbols) E.




95

Figure 4.11: The activity of peroxidase on aged (open symbols) and fresh (solid symbols) E.


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4.3.4 Community structure

Multivariate analysis of T-RFLP results (Figure 4.12) indicated that fresh and aged

leaves carried different fungal communities prior to incubation (ANOSIM: R =0.937;

P<0.005), but changes in community structure over time varied with treatment. In the

dry treatment, no change was observed in either the fresh or aged community over 100

days (R = 1; P<0.05) (Figure 4.13

Figure 4.13, (i)). In the moist treatment, fungal communities on fresh and aged leaves

became more similar over time, such that over 100 days global R was reduced from

0.937 to 0.37 (P=0.20) (Figure 4.13, (ii)). The community present after 100 days was

dissimilar to that found on both fresh and aged leaves prior to incubation (R = 0.512;

P<0.005). Fungal community structure also changed over time in the wet treatment

(Figure 4.13Figure 4.13, (iii)), however, in this case we observe the development of a

new community that did not differ between fresh and aged leaves (R = 0.259; P=0.30),

but was dissimilar to both communities present before incubation began (R = 0.993;

P<0.005).

4.3.4.1 Community responses to moisture treatments

The pattern of change in fungal community structure after 100 days varied between

treatments (ANOSIM: R=0.779; P<0.005). Fresh and aged samples at day 100 differed

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in the dry treatment (Pairwise ANOSIM: R=1; P=0.1), but not in the wet (Pairwise

ANOSIM: R=0.259) or moist (Pairwise ANOSIM: R=0.370) treatments (Figure 4.14).

Fresh Leaves

Aged Leaves

Figure 4.12: An MDS plot of T-RFLP data from fresh (closed symbols) and aged (open symbols) E.

camaldulensis leaves prior to incubation indicates that different fungal communities are present on

the two leaf types (2D stress is 0.11). Circles are added to clearly define different groups shown in

ANOSIM analysis.

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Figure 4.13: MDS plots of T-RFLP data showing changes in fungal community structure in the dry

(i), moist (ii), and wet (iii) treatments. Closed symbols represent fresh leaves while open symbols

represent aged leaves. Incubation time is indicated by symbols. Pre-incubation samples are shown as

circles, squares indicate 8 days incubation, the inverted triangle indicates 16 days incubation and

upright triangles indicate 100 days incubation.

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Figure 4.14: An MDS plot of T-RFLP data from E. camaldulensis leaves incubated in under dry,

moist and wet conditions for 100 days. Clusters overlaid on the MDS plot show that the fungal

community structure differs between fresh and aged leaves in the dry treatment, while communities

on fresh and aged leaves in the moist and wet treatments are similar to each other.

4.4 Discussion

This experiment has shown that fungal dynamics differ between fresh and aged E.

camaldulensis leaves. While losses of mass and organic matter generally differed little

between fresh and aged leaves, submerged fresh leaves lost more mass possibly as a

result of leaching. Fungal biomass accumulation was generally delayed on fresh leaves,

and total biomass was lower than that on aged leaves. Hydrolytic enzyme activity on

fresh leaves was lower than that on aged leaves under terrestrial conditions, but not under

aquatic conditions. Prior to incubation, fungal community structure differed substantially

between fresh and aged leaves, and subsequent changes in community composition

differed between treatments.

Fungal dynamics exhibited several important differences in response to moisture

treatments. In the dry treatment, fungal community structure did not change, leaves lost

little mass or organic matter, and fungal biomass and hydrolytic enzyme activity on fresh

leaves were close to zero. On aged leaves fungal biomass and enzyme activity were

higher, but did not generally change over time. Where the sediments were moist, some

mass loss was observed in fresh leaves, fungal biomass was able to accumulate on both

leaf types, hydrolytic enzyme activity towards plant structural carbohydrates was higher

in aged leaves and the fungal community structure became more similar between leaf

Fresh Leaves Dry Treatment

Aged Leaves Dry Treatment

Fresh Leaves Moist Treatment

Aged Leaves Moist Treatment

Fresh Leaves Wet Treatment

Aged Leaves Wet Treatment

2D Stress: 0.14

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types over time. When leaves were inundated, losses of mass and organic matter were

significantly larger than in other treatments, fungal biomass accumulation was retarded

on fresh leaves with respect to aged leaves, and enzyme activity was substantial and

similar on both leaf types. Changes in fungal community structure in this treatment

suggested that a transition from terrestrial to aquatic species had occurred.

Fungal colonisation of leaves is spatially variable, as fungi tend to extend in a radial

pattern from a random point of contact (Chamier, Dixon et al. 1984), leading to high

spatial variability in fungal biomass both within a single leaf and between leaves. This

phenomenon has resulted in high standard deviations in metrics of fungal mass and

activity.

4.4.1 Delays in fungal colonisation of “fresh” leaves

Previous studies of fungal species richness have shown that E. globulus leaves

submerged in a stream are colonised by fungi 2 weeks after chestnut and alder leaves

(Canhoto and Graça 1996; Chauvet, Fabre et al. 1997), and only 2 fungal species were

present prior to 42 days in data presented by Canhoto and Graça (Canhoto and Graça

1996). The results presented here show that fungal colonisation of fresh E. camaldulensis

leaves in standing water was delayed by at least 32 days after immersion. This lag time in

fungal biomass accumulation on fresh leaves may be due to the presence of chemical

substances that deter fungi. Oils, waxes and polyphenols such as those found in

Eucalyptus sp. leaves protect plants against fungal colonisation (Boon and Johnstone

1997; Campbell and Fuchshuber 1995; Graça, Pozo et al. 2002) and grazing insects

(Edwards 1982; Stone and Bacon 1994). Terrestrial micro-organisms and insects have

adapted to overcome these defences (Ohmart and Edwards 1991), and in many cases

insect grazing causes leaf wounds that facilitate fungal colonisation. Results from the

moist treatment, which simulates the terrestrial floodplain environment, showed that

fungal colonisers are able to overcome the effect of inhibitory substances in fresh leaves

and accumulate biomass. However, data from the wet treatment suggest that aquatic

organisms have not developed these tolerances.

The bulk of soluble polyphenols can be removed from submerged Eucalyptus leaves

within 4 days of inundation (Baldwin 1999; Love 2005). If these polyphenols were the

main cause of low fungal activity, the colonisation of fresh leaves by fungi could

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theoretically be quicker under aquatic conditions than under terrestrial conditions.

However, a lag in biomass accumulation suggests that many of the inhibitory substances

in the leaves are poorly soluble in water (e.g. waxes and oils). This is consistent with the

findings of Canhoto and Graça (1999), who reported that essential oils derived from

Eucalyptus species were more inhibitory to aquatic fungi than tannins.

This study shows that while fungal biomass accumulation is inhibited in the weeks

following the inundation of fresh leaves, hydrolytic enzyme activity occurs at similar

levels in both fresh and aged leaves, throughout most of the incubation period. However

at day 16, hydrolytic enzyme activity was significantly lower on fresh leaves, suggesting

that enzymatic break down of starch, cellulose and hemicellulose might also be inhibited

soon after inundation (analysis of the changes occurring in substrate composition would

be required to confirm this). As enzyme activity may be repressed in the presence of

reaction end-products, enzymatic cleavage of structural polysaccharides may be inhibited

by the presence of free sugars (Baldrian and Valaskova 2008; Chróst 1990). However,

this is not likely to be the cause of inhibition in this instance, as free sugars would be

utilised quickly by bacteria (Kaplan and Newbold 2003). In fact, breakdown is likely to

be accelerated by the presence of bacteria that “mop-up” free sugars. External enzyme

inhibitors such as eucalypt oils and tannins are a more likely cause of low activity. After

64 days incubation, the fungal community on fresh leaves was able to achieve the same

level of biomass and activity as that seen on aged leaves. At this time the community

composition was similar on the two leaf types. This suggests that substances inhibiting

colonisation by aquatic fungi are removed over this period or their efficacy decreases

with time.

The static conditions and evaporative concentration in wetlands may extend the period in

which fungal growth and enzyme activity are inhibited by eucalypt leaf leachates, as

compared to flowing systems. However, studies investigating the effects of Melaleuca

alternifolia oil on wetland bacteria have found these effects to be small (Bailey, Watkins

et al. 2003; Boon and Johnstone 1997). Increases in dissolved organic carbon due to the

presence of eucalypt leachates may stimulate the activity of bacteria in the water column,

causing decreases in the dissolved oxygen concentration (Howitt, Baldwin et al. 2007).

Fungi are generally more sensitive to low oxygen concentrations than bacteria, and have

a demonstrated susceptibility to eucalypt oils (Canhoto, Bärlocher et al. 2002; Terzi,

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Morcia et al. 2007), so it is likely that these leachates would impact more profoundly

upon fungal decomposers than bacteria.

4.4.2 Fungal activity under dry conditions

Birk (1979) found that mass loss from eucalypt leaves in the litter layer was largely due

to mechanical fragmentation and insect consumption, with leaching and microbial

activity being relatively insignificant. Conditions in the litter layer are generally moisture

limited (O'Connell and Grove 1987), and results from the dry mesocosm are in support of

these findings, showing that fungal communities on dry E. camaldulensis leaves did not

grow or break down the leaf substrate over a 140 day period. It is possible that fungal

decomposers are not important in the breakdown of terrestrial litter until it has been

digested and excreted by insects (Birk 1979; Choudhury 1988), and high oil and

polyphenol content of fresh leaves may deter insect feeding (Stone and Bacon 1994).

Together these factors result in a resource of low nutritional value that breaks down

slowly, and tends to build up on the floodplain when conditions are dry.

Fungal dynamics measured in the dry treatment were consistent with the current

understanding of litter decomposition in arid environments. Where moisture is absent

from the surface, litter decomposition proceeds mainly as a result of photo-degradation

(Smith, Gao et al. 2009), and biological processes occur in pulses as a response to

rainfall (Collins, Sinsabaugh et al. 2008). Under this paradigm, biological decomposition

of surface litter ceases during the dry phase and fungal activity takes place beneath the

soil surface. No loss of mass or organic matter was observed from E. camaldulensis

under dry conditions. On the floodplain, such conditions would lead to litter

accumulation (Glazebrook and Robertson 1999; Graça, Pozo et al. 2002). This

accumulation of litter represents a reserve of carbon that can be utilised by fungi and

other organisms with the next pulse of water (rain event or flood), in line with the pulse-

reserve model of arid ecosystems (Noy-Meir 1973; Ogle and Reynolds 2004; Reynolds,

Kemp et al. 2004).

Fungal activity at rates too slow to be measured is consistent with a fungal community

that is dormant, and fungal DNA amplified by PCR may be derived either from live

mycelium or viable propagules. This suggests the fungal communities colonising fresh

and aged E. camaldulensis leaves are dormant under dry conditions, or growing at an

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extremely slow rate. Such conditions occur seasonally in the floodplain wetlands of the

Murray-Darling Basin under a non-regulated water regime, although they may be

temporarily alleviated by rain events. The terrestrial and aquatic fungal communities may

use dormancy as a mechanism for surviving these predictable periods of unfavourable

conditions (Thomas 1996), inundation and drying, respectively. As wetland drying

alleviates sediment anoxia, it is likely to stimulate the germination of terrestrial

decomposer organisms within the sediments (Neckles and Neill 1994; Williams 2006).

For aquatic species, drying may trigger reproduction, resulting in the production of

dessication-resistant spores and conidia (Jennings and Lysek 1996). High temperatures

during the dry period may also “activate” these dormant propagules, priming them for

germination when conditions are suitable for growth (i.e. inundation) (Cooke and

Whipps 1993). Sharp increases in fungal biomass on aged leaves early in the incubation

period, followed by sharp declines are consistent with growth and sporulation, and may

indicate a response to leaf drying.

Primary metabolism involves the acquisition of nutrients from the environment to use as

energy sources or in order to produce substances such as proteins, lipids and nucleic

acids (Isaac 1997). Fungal secondary metabolism is the production of new substances

from these cellular compounds that are intended for use in interaction with the

environment (Thrane, Anderson et al. 2007). Fungi may use secondary pathways, which

may include producing agents of lignin degradation, in response to desiccation (Jennings

and Lysek 1996). Thus, peaks in oxidative enzyme activity early in the incubation period

may be a response to leaf drying, and not to the treatment conditions. As no significant

mass loss was observed under dry conditions, it is likely that lignin and cellulose were

not being mineralised to any great extent under this treatment. The observed activity

reflects a response to moisture availability, as the enzyme assay measures processes in

solution. As such, these data represent potential enzyme activity under moist conditions,

and suggest that the fungal community on aged leaves is capable of responding quickly

to moisture availability.

The drying of leaves has received critical attention (Boon 2006; Boulton and Boon 1991)

as it is known to alter the chemical (Gessner and Schwoerbel 1989; Serrano 1992) and

biological (Serrano and Boon 1991) properties of leaves. In this experiment, drying

leaves at 50°C may have been sufficient to activate dormant fungal propagules which

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later germinated during incubation. However, drying is essential to standardise leaf

moisture content for gravimetric analyses. Drying at more moderate temperatures would

therefore be the preferable approach where it is desirable to minimise impacts upon

fungal community structure and biomass.

4.4.3 Fungal activity under moist conditions - the terrestrial aging process

Under terrestrial conditions with sufficient moisture, E. camaldulensis leaves on the

floodplain are colonised by terrestrial fungi. The high levels of fungal biomass

accumulated on both fresh and aged leaves in less than 30 days in the moist treatment

suggests that the fungal communities on these leaves were undeterred by leaf defensive

compounds. However, on fresh leaves the activity of enzymes that cleave sugars from

cellulose, starch and hemicelluloses was close to zero, indicating inhibition of fungal

activity towards these plant polymers. Inhibition is likely to be due to the presence of

polyphenols, which would be rapidly leached from fresh leaves on the floodplain with

rainfall (Serrano 1992). As rainfall was absent from the artificial environment of the

mesocosm, the level of enzyme activity observed on fresh leaves may have remained low

for a longer period than might be expected on the floodplain. While it would appear that

aged leaves are a better fungal substrate than fresh leaves under moist conditions, these

two leaf types represent two different stages in the terrestrial aging process that are likely

to co-exist on the floodplain, with no adverse affects on the indigenous biota.

The process of “terrestrial aging” is known to produce chemical and physical changes in

the decomposing leaf (Anesio, Tranvik et al. 1999; Baldwin 1999; Smith, Gao et al.

2009), such as a reduction in bio-available organic molecules and the loss of inhibitory

substances (Canhoto and Graça 1996; Love 2005). Differences in fungal colonisation and

activity on fresh and aged leaves indicate that the terrestrial aging process facilitates

immediate colonisation of E. camaldulensis leaves by fungi when they enter the aquatic

phase. Terrestrial aging leads to changes in fungal community structure, higher total

fungal biomass, and enhanced enzyme activity towards cellulose in aquatic ecosystems,

but losses of mass and organic matter are not significantly affected.

Under moist terrestrial conditions leaf mass loss is driven by leaching and biological

processes (Coûteaux, Bottner et al. 1995). My results have shown that fresh leaves have

low fungal biomass in the early stages of terrestrial decomposition, low rates of enzyme

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activity towards cellulose, starch and hemicelluloses and negligible losses of mass and

organic matter. This suggests that the fungal community present on fresh leaves, while

tolerant of the leaf’s chemical defences, are not efficient in the breakdown of leaf

structural carbohydrates and may be consuming other resources such as pectin, protein

and free sugars. A change in community structure after 64-100 days brings with it

increased activity towards cellulose, starch and hemicelluloses, and increased mass loss.

Under laboratory conditions, aging of E. camaldulensis leaves to the point where fungi

are not inhibited took 30 days. Under field conditions, where sunlight, leaching and litter

organisms are more active, it is likely that this period would be shorter.

My results showed that oxidative enzyme activity towards lignin reached a peak during

the first week of incubation for all treatments and for both leaf types. Lignin is less bio-

available than other leaf components, and it is commonly accepted that litter

decomposing micro-organisms utilise lignin after more bio-available leaf components are

exhausted (Hammel, 1997). While lignin degradation is likely to be most prevalent in the

soil, or at the base of the litter layer (Melillo, Aber et al. 1989), it is shown here to occur

concurrently with hydrolytic activity towards cellulose, starch and hemicellulose.

However, it is important to note that the magnitude of peaks in activity of lignin

degrading enzymes observed early in this experiment may be partly due to responses of

the fungal community to drying at high temperatures.

4.4.4 Aquatic decomposition processes

The levels of fungal biomass accumulating on submerged fresh and aged E.

camaldulensis leaves were 3.7 and 6.2 mg.g-1

litter dry mass, respectively, after 64 days.

Pozo et al. (1998) reported that E. globulus leaves accumulated 65 mg fungal biomass.g-1

litter dry mass in a Spanish stream, and similar values on the same leaves were reported

from a stream in Portugal (Sampaio, Cortes et al. 2004). The biomass accumulating in

the mesocosms was also low when compared to values reported from the leaves of other

tree species submerged in rivers in the northern hemisphere (Gessner and Chauvet 1994;

Gessner, Robinson et al. 1998; Hieber and Gessner 2002; Suberkropp 2001; Van

Ryckegem, Gessner et al. 2007) and macrophytes in freshwater wetlands (Kuehn, Lemke

et al. 2000; Kuehn and Suberkropp 1998a). Similarly low levels of biomass (5.8 mgC,g-1

AFDM) have been recorded from macrophyte litter in a prairie wetland (USA) (Verma

et al, 2003), but fungal biomass on the leaves of Acer rubrum submerged in a woodland

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wetland was ~5 times higher (28 mg.g-1

litter dry mass) (Inkley, Wissinger et al. 2008). It

is possible that the artificial environment of the mesocosm has impeded the development

of fungal biomass.

When fresh leaves were submerged, fungal biomass was initially low and remained low

for one to two months, while aged leaves began to accumulate fungal biomass

immediately, and moderate levels of biomass were maintained for a number of months.

Lower fungal biomass implies lower food value (Canhoto and Graça 1995; Suberkropp,

Arsuffi et al. 1983) and reduced palatability to invertebrates (Hladyz, Gessner et al.

2009). Thus a lag time in the colonisation of fresh leaves may have implications for

wetland food chains, as the inundation of litter and sediments in floodplain wetlands is

also likely to trigger emergence of aquatic invertebrates (Hillman and Quinn 2002;

Jenkins and Boulton 2007). If floodplain aged eucalypt leaves are inundated when the

wetland fills, insect larvae that feed on conditioned leaves will encounter abundant food

resources. The inundation of fresh leaves will not provide a nutritious food resource for

some months. Thus, if wetlands are inundated in summer when eucalypt leaves are

freshly fallen, the conditioned leaves are likely to have lower fungal biomass, lower

nitrogen content (Reid, Quinn et al. 2008), and be less nutritious for invertebrates. This

will inhibit survival, growth and reproduction in emerging invertebrates (Albariño,

Villanueva et al. 2008; Bärlocher 1985), influencing recruitment rates and reducing food

resources for higher trophic levels. This phenomenon may play a role in changes in

wetland invertebrate assemblages previously associated with river regulation (Hillman

and Quinn 2002; Quinn, Hillman et al. 2000).

Inundation poses several challenges to fungal communities present on leaves. Oxygen

availability is reduced, nutrients are removed from the substrate and competitive

interactions between fungal species are altered. These challenges usually result in the

decline of the dominant terrestrial species soon after inundation, and their replacement by

species adapted to aquatic conditions (Boon 2006; Brinson, Lugo et al. 1981; Kuehn,

Lemke et al. 2000). In terrestrial systems Eucalyptus leaves are typically colonised by

fungal taxa such as Capnodiales and Pleosporales (Dothideomycetes), Eurotiales

(Eurotiomycetes), Hypocreales and Chaetosphaeriales (Sordariomycetes ), Mucorales

and Mortierellales (Glomeromycetes ) (Dix and Webster 1995; Eicker 1973; Macauley

and Thrower 1966), while leaves in stagnant waters such as wetlands are commonly

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colonised by Chytridiales (Chytridiomycetes) (Lefevre, Bardot et al. 2007),

Leptomitales, Saprolegniales and Peronosporales (Oomycetes) (Czeczuga, Mazalska et

al. 2005; Czeczuga, Muszyńska et al. 2007), aquatic hyphomycetes (Dick 1976a; Pattee

and Chergui 1995) and aero-aquatic hyphomycetes (mainly Ascomycetes) (Kjoller and

Struwe 1980). Leptomitales, Saprolegniales and Peronosporales are Stramenopiles which

colonise plant material soon after inundation, then decline in abundance as

decomposition proceeds (Bärlocher 1992).

Amplification of fungal DNA by PCR indicates that fungal DNA is present but the

method used is not a reliable measure of abundance, and does not distinguish between

active mycelia and viable propagules. Species that colonise leafy substrates in terrestrial

environments may die with inundation, or may remain alive but become inactive.

Alternatively they may continue to decompose the leaf resource but discontinue their life

cycle, or continue their life cycle but produce propagules that are not viable in the aquatic

environment (Thomas 1996). All of these groups, along with fungi able to germinate,

grow and reproduce in the aquatic environment, may have contributed to the total fungal

DNA extracted from the decomposing leaf material. Thus, the finding that fungal

communities present before inundation and after 100 days in the water are significantly

dissimilar strongly suggests replacement of terrestrial species with aquatic species.

Identification of the fungal species involved would be required to confirm such a

transition. Alternatively, reverse transcription PCR would amplify only the DNA from

active organisms.

Fungal community structure data, when considered with fungal biomass and enzyme

activity data, suggest a 2-stage succession of fungal species on submerged leaves, similar

to that reported by Verma et al. (2003). A terrestrial community is present prior to 16

days incubation, and is then replaced by an aquatic community. The biomass and enzyme

activity of the terrestrial community (<16 days incubation) differed between fresh and

aged leaves under aquatic conditions, but after 16-30 days the biomass and activity of the

aquatic community and substrate organic matter losses were similar on both leaf types.

Enzyme activity towards lignin was highest when the terrestrial community was

dominant, then continued at moderate levels while the aquatic community was present.

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The terrestrial fungal community exhibited higher enzyme activity towards cellobiose,

starch and hemicellulose and lower activity towards cellulose under terrestrial conditions,

as compared with aquatic conditions. However under aquatic conditions, it was the

aquatic community that exhibited the highest activity rates for all of these polymers. This

suggests that the aquatic fungal community is able to efficiently break down cellulose,

starch, hemicellulose and lignin, perhaps more effectively than the terrestrial community

on the same substrates.

The water regime has a strong impact upon litter decomposition and carbon dynamics in

wetlands (Battle and Golladay 2001; Inkley, Wissinger et al. 2008; Neckles and Neill

1994; Ryder and Horwitz 1995). The results presented here show that fungal dynamics

and mass loss due to leaching differ substantially when fresh and aged E. camaldulensis

leaves are inundated, and these differences will influence water quality. Significant loss

of mass and organic matter from fresh leaves during the first week in the aquatic phase is

consistent with previous studies showing that much of the mass loss from inundated E.

camaldulensis leaves is due to leaching (Briggs and Maher 1983; Day 1983; Glazebrook

and Robertson 1999). Carbon leached from these leaves contributes to the level of DOC

in the water column. Summer inundation of floodplain wetlands can produce DOC

concentrations of around 15ppm, which is the twice that produced by spring inundation.

These rapid increases in DOC have the potential to impact upon water quality in terms of

colour, microbial activity, dissolved oxygen (Campbell and Fuchshuber, 1995) and

nutrient concentrations (Baldwin 1999; O'Connell, Baldwin et al. 2000; Robertson, Bunn

et al. 1999; Watkins, Quinn et al. 2010) and have other adverse impacts on wetland biota

(Gehrke, Revell et al. 1993; Graça, Pozo et al. 2002; Watkins, Quinn et al. 2010).

Anoxia may further inhibit fungal activity. These impacts could be expected to be

smaller when aged leaves are inundated due to lower carbon losses via leaching, and

lower bio-availability of the leachate (Baldwin 1999).

4.5 Conclusions

This experiment has shown that fungal community structure differs between fresh and

terrestrially aged E. camaldulensis leaves, and that temporal changes in fungal biomass

and enzyme activity differ in dry and moist terrestrial conditions, and aquatic conditions.

Leaf mass loss differs between leaf types when moisture is not limiting, but the

proportion of organic matter lost does not differ between fresh and aged leaves. These

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results strongly suggest that a period of terrestrial decomposition is important for E.

camaldulensis leaves as it promotes fungal colonisation, which is known to improve the

food value of submerged leaves. Summer inundation of fresh leaves in floodplain

wetlands will lead to reduced levels of fungal colonisation and leaf conditioning,

potentially impacting upon food chains, water quality and carbon dynamics.

While the current experiment provides a model for the decomposition of eucalypt leaves

in floodplain wetlands, the conditions in these habitats are more variable than those

operating in a laboratory, with complex biological interactions. The artificial conditions

in the mesocosms may also have had some influence on the amount of fungal biomass

able to accumulate on the leaves. Thus, having demonstrated a strong response of the

fungal community to moisture availability in leaf litter, in Chapter 5 I examine the

response of fungal community structure and biomass to changing moisture availability in

the sediments of a floodplain wetland as it undergoes seasonal drying.

Later chapters expand upon our understanding of fungal dynamics and leaf break down

under field conditions. The accumulation of fungal biomass on floodplain aged E.

camaldulensis leaves is compared between standing and flowing waters of lowland

streams in Chapter 6. Chapter 6 also quantifies the contribution of stramenopilous

organisms to the biomass accumulating on submerged leaves. Furthermore, as the

enzyme activity data presented here have shown that lignin and polysaccharides are

being degraded in submerged E. camaldulensis leaves, the chemical changes taking place

within the submerged leaves as a result of this activity are examined in Chapter 7.

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Fungi in Drying Wetland Sediments

5.1 Introduction

The results from the pilot study in Chapter 3 indicated that fungal DNA was present in

the sediments of Normans Lagoon, and that the fungal community structure may change

with sediment moisture content. Previously, analysis of microbial phospholipid fatty

acids (PLFA) had indicated that fungal biomarkers were absent from the sediments of

Ryans Lagoon, a similar wetland also on the floodplain of the Murray River (Boon,

Virtue et al. 1996). In this chapter I explore fungal dynamics in the sediments of Quat

Quatta East Lagoon, again on the Murray River floodplain.

The sediments of floodplain wetlands in the Murray-Darling basin are generally anoxic

when the wetlands are holding water, and organic matter decomposition in the sediments

is dominated by anaerobic bacteria (Boon 2000). In summer, the water column may

become thermally stratified each day (Ford, Boon et al. 2002), leading to anoxia in the

bottom layers, although the water column is generally mixed overnight. These anoxic

conditions, combined with the knowledge that most filamentous fungi are aerobic and

ergosterol was absent from the sediments of a typical wetland, led Boon et al. (1996) to

conclude that fungi were not important in organic matter processing in these systems.

However, we have seen in the previous chapter that fungi are able to colonise submerged

plant detritus in standing water, and fungi are also known as parasites and endophytes of

algae and wetland macrophytes (Czeczuga, Mazalska et al. 2005; Kagami, de Bruin et al.

2007; Khan 1993; Lefevre, Bardot et al. 2007), which are abundant in these systems

(Robertson, Bunn et al. 1999). As this plant and algal detritus often becomes buried in

the sediments, it is logical to conclude that fungi might also be present in wetland

sediments.

While wetland sediments are usually anoxic, the narrow sediment water interface zone

may be oxic, and oxic zones may be created by plant roots or burrowing animals. For

example, Qiu et al. (2005) found that while the proportion of respiration in wetland

sediments due to fungi was significantly lower than in leaf litter, fungal activity was

present and was higher than that found in upland soils. However, this aerobic activity is

likely to be confined to the sediment surface, where the greatest proportion of aerobic

organic matter decomposition occurs (Adams and van Eck 1988).

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Fungal DNA has also been isolated from anoxic sediments in sulfide-rich springs and a

freshwater pond (Luo, Krumholz et al. 2005). These fungi were mainly Basidiomycete

and Ascomycete yeasts, but chytrids were also present. Luo et al. (2005) also detected

ergosterol and fungal PLFAs in these sediments. Seo and DeLaune (2010) recorded

fungal respiration under highly reducing conditions in sediments from a forested wetland.

Hallett and Dick (1981) found that the diversity of Oomycete propagules in the littoral

mud of a small freshwater ornamental lake in the United Kingdom was the same as that

in the water column, although propagule abundance was lower. Anoxia is also known to

inhibit the lignin degrading enzymes of fungi (Freeman, Ostle et al. 2001) but lignin

degradation has been shown to occur in the anaerobic sediments of a freshwater marsh

(Okefenokee Swamp, southern Georgia, USA) (Newell, Moran et al. 1995).

As sediments dry out the anoxia is relieved, water is less available and the chemical

environment of the sediments changes (Baldwin and Mitchell 2000). Such extreme

changes are likely to alter the decomposer community. Changes in fungal community

structure (Toberman, Freeman et al. 2008), increased biomass (Jaatinen, Fritze et al.

2007) and increased respiration (Qiu, McComb et al. 2005) have been reported. Rinklebe

and Langer (2006) found that fungal PLFA were absent from floodplain soils that had

been inundated for long periods, but not from soils from higher positions on the

floodplain. However, exposed sediments have different properties to soils, and maximum

microbial biomass may be concentrated in the layer 10-30cm beneath the surface, instead

of in the surface layers as is the case for soils (Qiu, McComb et al. 2003).

There are no published reports of a concurrent investigation of fungal community

structure and fungal biomass in an Australian floodplain wetland, although biomass has

been inferred from sediment induced respiration measured in a coastal freshwater

wetland under conditions that inhibit prokaryotes (Qiu, McComb et al. 2005). While the

pilot study detailed here (Chapter 3) has shown that fungal DNA is present in wetland

sediments, the taxa comprising the fungal community are unknown. It is also unknown

whether the fungal DNA is derived from active mycelium or dormant propagules.

As many of the floodplain wetlands in the Murray-Darling Basin are ephemeral, the

response of the sediment fungal community to drying and re-wetting will impact upon

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the rate of organic matter respiration and food web interactions. We have seen

differences in the fungal community structure between wet and dry sediments (Chapter

3), but the sediment moisture content required to produce a response from the fungal

community, and the time taken to respond to these changes are unknown.

In this chapter I report on a field experiment where fungal community structure and

biomass were measured in a floodplain wetland as it underwent summer drawdown. The

sediments were sampled over 35 days during drawdown, and further samples were taken

after 2 months of inundation (90 days from day 1). The moisture content, organic matter

content and fungal biomass was determined for samples taken over a 2.5m wetness

gradient in the exposed wetland sediments. Spatial and temporal changes in fungal

community structure within the transect were assessed using T-RFLP, and these changes

were related to the moisture and carbon content of the sediments. I hypothesised that a

change in the fungal community composition consistent with a change from an aquatic

community to a terrestrial community would be observed between wet and dry points on

the gradient, and at the same sediment position over time. Such a change would be

accompanied by increased fungal biomass.


5.2.1 Site description

Quat Quatta East Lagoon connects with the north bank of the Murray River

approximately 5km west of Howlong, NSW (35° 58′ 19.82″ S, 146° 34′ 44.67″ E,

Elevation 150m elevation) (Figure 5.1). The Murray River is highly regulated

(Maheshwari, Walker et al. 1995), supplying water for irrigation, stock usage and

domestic supply over the warmer months, while flows in cooler months are retained in

storages. The water level in Quat Quatta East lagoon decreased between 5 December

2008 and 26 December 2008 until the wetland had contracted to three small pools

(Figure 5.3, (a)). The drying event led to the death of a large amount of fish, and a blue-

green algal bloom (Figure 5.2, (d)). Inflows commenced in late December 2008, and all

bare sediments were inundated by March 2009 (Figure 5.3, (b)). The riparian vegetation

at Quat Quatta East Lagoon is dominated by E. camaldulensis open forest with a grassy

understory, with reeds, sedges and rushes along the margins of the lagoon. The area is

grazed by cattle (Figure 5.2, (c)) and used for camping and recreational fishing.

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Figure 5.1: Quat Quatta East lagoon is located north of the Murray River, approximately 6 km west

of the township of Howlong, NSW.

5.2.2 Sample collection

One 2.5m transect was established in the sediments of Quat Quatta East lagoon as it

underwent drying between December 2008 and February 2009. The transect was oriented

perpendicular to the water line, and positions 50cm apart were marked out using wooden

pegs. The position furthest from the centre of the water body was denoted as position 1,

and positions were numbered consecutively up to position 6 (as pictured in Figure 5.2,

(a)). The sediments were sampled on five occasions. The moisture status of the

sediments, i.e. whether they were exposed, submerged or transitional (edge), at the 5

sampling dates is shown in Table 5.1. On 12 December 2008 exposed sediments were

damp on the surface due to light rain. After 35 days increased flows in the Murray River

for the purposes of downstream irrigation led to inflows to the wetland, and all 6

positions were submerged (Figure 5.2, (b)).

On each sampling occasion, the upper 5cm of sediment at each position was collected

into a sterile 70 ml specimen jar. On 7 March 2009, additional reference samples were

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taken from the dry and vegetated soil above the high water line of the wetland (bank), the

reed bed at the margin of the wetland, and positions 1 and 2. On this occasion, all of the

original six positions were submerged beneath more than 1m of water.

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Table 5.1: The moisture status of sediments at 6 positions over a 35 day period.

Position 5/12/2008 12/12/2008 19/12/2008 26/12/2008 8/1/2008

1 Exposed Exposed Exposed Exposed Submerged

2 Edge Exposed Exposed Exposed Submerged

3 Submerged Exposed Exposed Exposed Submerged

4 Submerged Exposed Exposed Exposed Submerged

5 Submerged Submerged Edge Exposed Submerged

6 Submerged Submerged Submerged Exposed Submerged

Figure 5.2: Positions were marked along a transect in the exposed sediments. Position 1 was exposed

at the commencement of the 35 day period, while position 6 was inundated. Panel (a) shows the

water level after 8 days while panel (b) shows the water levels after inflows began on Day 35. The

wetland was frequently visited by grazing cattle (c) and dead fish and algal blooms were observed as

the wetland approached dryness (d).

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Figure 5.3: The water level in Quat Quatta East Lagoon as it approached dry conditions (a) and

after it had received inflows from the Murray River (b). In the background, the dominant tree

species, E. camaldulensis, can be observed, while reed beds and immature E. camaldulensis are seen

in the foreground.

5.2.3 Sediment moisture and organic matter content

Three analytical replicates of sediment samples were homogenised by mixing, then 10-15

ml of sediment was weighed into pre-fired porcelain crucibles (wet mass - WM) before

being dried at 105°C for 72 hours. Crucibles were cooled in a desiccator, before being re-

weighed (dry mass - DM). Following this, sediments were fired in a muffle furnace for 1

hour at 550°C, cooled, and re-weighed (ash-free dry mass - AFDM). Moisture content

was calculated as the difference between WM and DM and expressed as a percentage of

WM. Organic matter content was calculated as the difference between DM and AFDM

and expressed as a percentage of DM. Moisture content and organic matter content data

was analysed using 2-way ANOVA (without replication) on the mean values for each

position in each time. Correlation between these variables was assessed using multiple

linear regressions. Data analyses were carried out using Microsoft Office Excel (2003,

SP3, Microsoft Corporation, Redmond, Washington, USA.).

5.2.4 Microscopy

Wetland sediment from each sample position was smeared onto a glass microscope slide

and stained with lacto-phenol cotton blue for 5 minutes prior to examination. These

slides were examined at 40×, 100× and 400× magnification using an Olympus BX 51

compound microscope (Olympus Australia, Pty. Ltd., Oakleigh, Victoria, Australia)

equipped with an Olympus DP12 digital camera (Olympus Australia, Pty. Ltd., Oakleigh,

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Victoria, Australia). Images were processed using Olysia software (Soft Imaging System,

Münster, Germany).

In the absence of cultured specimens, reproductive features, and knowledge of the cell

wall composition or microstructure it is not possible to make definitive classifications of

the fungal tissues observed. However, it is possible to make a preliminary identification

based on the features of higher level taxa. Higher level identification of fungal hyphae

was made according to the key published in Fungi of Australia (Walker 1996), and with

reference to Thomas (1996). The more recent key published by Adl et al. (2005) places

the fungi within the Super-group Opisthokonta, and subdivides fungi into Ascomycetes,

Basidiomycetes, Chytridiomycetes, Glomeromycetes, Microsporidia, Urediniomycetes,

Ustilaginomycetes and Zygomycetes. The Stramenopiles are placed within the Super-

group Chromalveolata, and the group formerly referred to as Oomycetes are newly

classified as Peronosporomycetes. However, this key relies on the identification of cell

organelles, the metabolic pathways used by the organisms and the composition of the cell

walls. Therefore, the earlier classification system has been used in these preliminary

descriptions of the organisms observed.

Hyphae have been identified as having (a) regular septa with adjacent clamp connection

(Basidiomycetes, Urediniomycetes, Ustilaginomycetes), (b) septa but no clamp

connections (Ascomycetes, and some Chytridiomycetes), or (c) no septa (coenocytic)

(Zygomycetes, Chytridiomycetes, Oomycetes). Microsporidia and Glomeromycetes are

obligate internal parasites and are not expected to be found in wetland sediments.

Additional references were used for preliminary identification of zoospores and encysted

zoospores (Dick 1969; Dick 1972; Hallett and Dick 1986; Pickering, Willoughby et al.

1979).

5.2.5 T-RFLP

The procedure used to extract DNA from wetland sediments, amplify the DNA using

PCR, digest to create terminal restriction fragments, and analyse fragment length data has

been detailed in section 3.2.3.

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5.2.6 Fungal biomass - ergosterol content

Ergosterol content in wetland sediments was assessed using saponification followed by

solid phase extraction and the reversed-phase liquid chromatography isocratic method

with UV detection, similar to that described in Chapter 4 (Gessner 2005). However,

minor modifications were required to accommodate the change in substrate from leaf

discs to sediment.

Briefly, ~1g of fresh sediment was preserved in 8 ml of methanolic potassium hydroxide

(KOH/MeOH, 8g/L) and stored at 4°C in the dark. On the day of extraction, sediment

was re-suspended in the storage solution and decanted into 50ml Pyrex tubes. The

storage tube was rinsed with a further 2 x 1ml of KOH/MeOH, which was added to the

50 ml tube. A control was comprised of 1g of sediment that had been fired in the muffle

furnace at 550°C for 1 hour, and then allowed to cool. To this, 250 µL of ergosterol

standard in KOH/MeOH was added. The sediments were saponified under reflux for 30

min at 80°C, then allowed to cool for a further 20 minutes.

Sediments were then resuspended before being transferred into a disposable 10ml

polypropylene centrifuge tube and centrifuged at 3000 rpm for 5 minutes in an Allegra

X-15R centrifuge (Beckman-Coulter, Fullerton, California, USA). The supernatant was

then transferred into the Sep-Pac cartridge using a pipette. A further 1 ml of methanol

was added to the centrifuge tube, the sediment was resuspended by vortexing, and then

centrifuged at 1000 rpm for 30 seconds. The supernatant was transferred to the Sep-Pac

cartridge, and then the washing step was repeated. A total of 12ml of solution was

transferred.

Additional analysis steps were the same as for leaf disc ergosterol as detailed in Section

4.3.2.

Ergosterol detection limits are a function of the minimum peak area of 4930 used in the

integration method. This corresponds to 0.2 µg ergosterol and approximately 0.85 µg

ergosterol .g-1

sediment DM. This equates to approximately 5mg fungal biomass.g-1

sediment DM.

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5.3 Results

5.3.1 Sediment moisture content

The moisture content of wetland sediments sampled over 35 days of drawdown differed

significantly between positions and times (ANOVA: P<0.001 for both variables) (Figure

5.4). Saturated wetland sediments contained 50-60% moisture, and this figure fell to 15-

45% moisture with exposure and subsequent drying. None of the sediments could be

said to be desiccated prior to inundation after 35 days. For samples from day 90, the

mean moisture content was 1.11% for the sediment (soil) sampled from the bank, 32.25%

from the reed bed, 30.63% from position 1, and 28.03% from position 2.

Figure 5.4: The mean moisture content of wetland sediments over a period of wetland drying and

subsequent re-wetting. Position 1 (indicated by a white upright triangle) is furthest from the centre

of the water body, while position 6 (indicated by a black inverted triangle) is closest to the centre of

the water body. Positions 5 and 6 were submerged until exposure on day 22. All samples were

exposed on Day 22, while all samples were inundated on Day 35. Error bars indicate the standard

error of each data point, with maximum standard error being 2.8%.

5.3.2 Sediment organic matter content

The organic matter content of wetland sediments over the drawdown period is shown in

Figure 5.5. Organic matter content differed significantly between positions (ANOVA:

P=0.01) but not between times (ANOVA: P=0.40). While there was a trend for deeper

sediments to have higher organic matter contents, this was not consistently the case.

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Similarly, there was a trend towards decreasing organic matter upon sediment exposure,

and increases with inundation, but this was not seen consistently. Increases in the

sediment moisture content were correlated with increased sediment organic matter but

the relationship was not strong (Multiple Linear Regression: R2

= 0.47; P<0.001).

Figure 5.5: The organic matter content of wetland sediments. There is a trend towards decreasing

organic matter with drying, and increasing organic matter with re-wetting, but this pattern is not

consistent across all sediment positions. Error bars indicate standard error about the mean of each

data point.

5.3.3 Fungal biomass – ergosterol content

None of the sediment samples examined contained detectable quantities of ergosterol.

5.3.4 Microscopy

As no ergosterol was extracted from the wetland sediments, the sediment samples were

examined under the microscope for evidence of fungal organisms. Evidence of fungal

hyphae (Figure 5.6) and propagules (Figure 5.7) was found but their distribution

throughout the sediments was sparse. Closer to the centre of the wetland, the hyphae

observed were mainly aseptate, and fungal hyphae were often observed within diatoms

(Figure 5.7, (d)). While aseptate hyphae may belong to the Zygomycetes,

Chytridiomycetes or Oomycetes, the septate hypha with a clamp connection (Figure 5.6

(d) and Figure 5.7 (d)) indicates a Basidiomycete.

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Encysted propagules were similar to those in the Oomycete families Saprolegniaceae and

Pythiceae, although magnification and image resolution was not sufficient to discern the

presence or absence of hairs or spikes in order to classify these propagules to genus or

species.

Figure 5.6: Hyphae observed in sediment samples. Shown are aseptate hyphae that may be

Chytridiomycetes, Zygomycetes or Oomycetes (a , b and c), and a septate hypha with a clamp

connection (CC) indicating a species belonging to the Basidiomycetes (d). 400x magnification. Scale

bar = 50µm.

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Figure 5.7: Fungal propagules found in sediment samples (a, b and c) and a fungal hypha enclosed

within a diatom frustule (d). The propagule shown in panel (a) is an encysted Oomycete propagule

and a similar specimen is seen germinating in panel (b). The organism adjacent to the propagules

(arrow) appears to be an amoeba. The presence of a prominent oil droplet (arrow) suggests the

propagule in panel (c) is a chytrid sporangium. 400x magnification. Scale bar = 50µm.

5.3.5 T-RFLP

Changes in the fungal community structure over time were not significant between

positions (Two-way ANOSIM R=0.208; P=0.001) or times (Two-way ANOSIM

R=0.347; P=0.001). Figure 5.8 shows that the fungal community structure changed

randomly between days 1 and 35 at position 1. This area remained dry until it was

inundated at day 35. After 35 days of inundation there was a change in community

structure indicated by movement to the right of the ordination. This change corresponded

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to a decrease in the mean number of terminal restriction fragments (T-RFs) obtained,

from 24.3 at day 1 to 10.7 at day 90.

The fungal community structure changed with the moisture content of the sediments.

Figure 5.9 shows that dry samples (0-10%) were dissimilar to saturated (50-60%)

(ANOSIM Pairwise test: R=0.803; P=0.001), and very wet (40-50%) samples (One-way

ANOSIM Pairwise test: R=0.806; P=0.002) with no significant difference observed

between the intermediate moisture categories (One-way ANOSIM Pairwise test:

R<0.584). The fungal community structure did not differ significantly between inundated

and exposed sediments (One-way ANOSIM: R=0.101; P=0.019). Higher mean numbers

of T-RFs were seen in samples with 15-45% moisture content (Dry samples have 8.83 T-

RF, saturated samples had 14.45 T-RF and sediments with 15-45% moisture had 17.39

T-RF). No relationship was found between fungal community structure and sediment

organic matter content (One-way ANOSIM: R=0.184; P=0.001).

Samples taken at day 90 (Figure 5.10) show that the fungal community structure differed

between the bank (never inundated), the reed bed (recently inundated), and the sediments

at positions 1 and 2 (inundated for 55 days). The largest differences were found between

position 2 and the reed bed (One-way ANOSIM: R=0.692; P=0.018) and between

position 1 and the bank (One-way ANOSIM: R=0.611; P=0.012). These differences were

associated with moisture content and the number of fungal amplicons. The reed bed

samples yielded a larger number of T-RFs (Mean 14, max 19), compared to the deeper

sediments (Mean 9.2, max 15) and the bank (mean 8.8, max. 13).

When the T-RFLP data from positions 1, 3, 6 and the dry bank were analysed using

group average cluster analysis the exposed and inundated samples were dissimilar

(Figure 5.11). Position 1 was exposed on all days except day 35. These samples were

generally clustered together, with the exception of a day 22 replicate (when these

sediments were driest) that clustered with bank sediments, a day 35 replicate (re-wet) that

clustered with position 3 sediments and a day 22 replicate that clustered with position 6

sediments. Position 6 was wet on all days except day 22. Only one day 22 replicate

separates from other position 6 samples. Inundated samples from position 3 (Day 1 and

8) clusters with position 6, while later samples (drying, dry and re-wet) cluster with

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position 1. These data suggest that changes in fungal community structure may lag

behind sediment inundation or exposure by more than 1 week.

Figure 5.8: The trajectory of change in fungal community structure at position 1 over time. Error

bars indicate standard error about the mean of three points for each time. Axes are relative positions

in data space and have no units.

Figure 5.9: MDS plot of sediment fungal community structure shows that there is some separation of

groups by sediment moisture content. Symbols indicate different moisture content classes (see

legend). Overlaid circles are intended to illustrate grouping of samples by moisture class, but have no

statistical meaning.

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Figure 5.10: MDS plot of T-RFLP samples from Quat Quatta sediments on Day 90. Dry sediments

are shown by open symbols; wet sediments are shown by solid symbols. Sediments that had not been

wet (Bank samples – see legend) separate well from sediments in the reed bed that had recently been

submerged (circles), and those from positions 1 and 2, that had been submerged for 55 days (upright

and inverted triangles, respectively). The circles are added to illustrate grouping only, and do not

indicate statistical similarity.

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5.4 Discussion

5.4.1 Fungal organisms are present in wetland sediments

While fungal biomass in the sediments of Quat Quatta East Lagoon was below detection

limits (5mg biomass per gram sediment dry mass), fungal DNA could be amplified and

microscopic examination of the sediments showed that both hyphae and propagules were

present, but in low abundance. The organisms present consisted mainly of coenocytic

hyphae, encysted propagules, and fungal tissues encased within diatom frustules.

Together, coenocytic hyphae and encysted propagules suggest that Oomycetes are

present in the wetland sediments. These organisms and their propagules are able to

survive hypoxic conditions in the surface sediments (Field and Webster 1983; Johnson,

Seymour et al. 2002), and are able to utilise a range of substrates ranging from living and

dead fish to plant detritus (Al-Rekabi, Al-Jubori et al. 2005; Czeczuga, Kozlowska et al.

2002; Czeczuga, Muszyńska et al. 2007; Pickering, Willoughby et al. 1979). Oomycetes

do not have ergosterol in their cell walls, and thus would not be detected using the

ergosterol method to measure fungal biomass. Nechwatal et al. (2008) were able to

identify 18 Oomycete species, and found that permanently flooded sediments had a more

diverse Oomycete community than drier sediments and that only 4 species were found in

both flooded and dry sediments.

The hypha observed within the diatom frustule was septate with a clamp connection,

indicating that it belonged to the Basidiomycetes. It is not clear whether this organism

was an algal parasite or a decomposer of dead algal cells. Reports of fungi parasitising

diatoms in freshwater systems commonly refer to Chytridiomycetes species (Bruning

1991; Kagami, de Bruin et al. 2007). While there are no reports of Basidiomycetes

parasitising algae in freshwater systems, Basidiomycetes have been found to parasitise

red algae in marine systems (Porter and Farnham 1986). It is therefore likely that the

organism observed is saprobic, as Basidiomycete saprobes are known among the aquatic

hyphomycetes (Ingold 1961; Nawawi 1985), in soils (Gams 2007) and marine sediments

(Burgaud, Arzur et al. 2010).

Chytrid parasites were not observed on diatoms in the sediments. These fungi are

common in lentic systems (Czeczuga, Kozlowska et al. 2002; Ellen Van Donk 1983;

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Lefevre, Bardot et al. 2007) and are also important in the decomposition of organic

matter, and conversion of inorganic carbon into organic carbon (Gleason, Kagami et al.

2008). However, their distribution is limited to fresh water with NaCl concentrations of

<2%, and their growth is known to be inhibited by low dissolved oxygen concentration

and temperatures above 30°C (Gleason, Kagami et al. 2008). These factors indicate that

Chytrids are unlikely to be active parasites of diatoms in drying floodplain wetlands over

the summer. Even when salinity is very low, temperatures regularly exceed 30°C and

hypoxic conditions may occur (<2mg O2/L) (Smith 2010). Thus this ecological niche is

unoccupied, and the biomass of dead diatoms is available to other fungi, such as soil

basidiomycetes.

If Ascomycete or Basidiomycete yeasts were present in the sediments in any significant

way, then their biomass would have registered as ergosterol, as these organisms tend to

have higher concentrations of ergosterol in their cell walls (Pasanen, Yli-Pietila et al.

1999). Hyphae indicative of the Ascomycetes were also absent from the wetland

sediments. The aquatic and aero-aquatic hyphomycetes are comprised mainly of species

in this division, and their absence suggests that sediments are not a suitable substrate for

these groups.

5.4.2 Changes with sediment exposure and inundation

The structure of this fungal community was seen to change with the moisture content of

the sediments. While sediments remained either exposed or inundated, no change was

observed in fungal community structure over space or time. However, the diversity of

fungal amplicons decreased with both drying and saturation. Maximum amplicons

diversity at 15-45% moisture content suggests that the fungal community requires a

balance between moisture and oxygen availability. At any specified location within the

wetland, such conditions will be limited to brief periods during drying and re-wetting of

the sediments, and the response of the fungal community to sediment exposure may be

delayed by more than a week.

While fungal community structure in soils is driven mainly by seasonal factors

(Toberman, Freeman et al. 2008), community structure in wetland sediments appears to

change with inundation and drying (Rinklebe and Langer 2006). Differences in fungal

community structure between inundated wetland sediments and floodplain soil from the

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bank suggests that an aquatic fungal community exists, and that it differs substantially

from the terrestrial fungal community, with maximum amplicon diversity occurring

during the transition between these two community structures.

While changes were observed in fungal community structure, no increases in fungal

biomass were found. There are several possible reasons why this might be the case.

Firstly, wetland sediments represent a harsh environment for fungi where oxygen may be

unavailable, inhibiting their ability to degrade carbon sources with oxidative enzymes

(Jennings and Lysek 1996; Wetzel 1992). Under these conditions fungi may not compete

successfully with bacteria and fail to achieve growth (Menge and Sutherland 1987).

Secondly, it may take longer than 35 days for the sediment fungal community to

germinate and grow. Regular non-catastrophic disturbances such as drying events are

known to stimulate greater diversity (Connell 1978; Grime 1973; Lake 2000). However,

it may take some time for the biota to recover from such a disturbance (Boulton and Lake

1992). Thirdly, the propagules in the sediments may require a stimulus to germinate that

they did not receive, such as a nutrient or hormone. Lastly, the fungal organisms present

in the sediments may not contain ergosterol in their cell walls (e.g. Oomycetes and

Chytrids), and therefore their biomass would go undetected.

5.4.3 Substrates for sediment fungi

The organic matter content of the sediments of Quat Quatta East Lagoon was 2-7%,

which is low when compared with 10-26% reported in a similar wetland (Boon, Virtue et

al. 1996), and only a proportion of this will be biologically labile (Baldwin 1999).

However, in a drying wetland, there are a range of possible substrates that may

accumulate on the sediment surface. These will include phytoplankton and animal

biomass. When warm weather leads to evaporative concentration of nutrients and

wetland drying both of these carbon sources will be deposited on the sediment surface.

Dying fish can contribute a significant amount of carbon towards the productivity of

waterholes in arid environments (Burford, Cook et al. 2008). These resources will be

colonised by fungi if there is sufficient oxygen and moisture to sustain their growth.

5.4.4 Method considerations

While ergosterol concentrations in the sediments of Quat Quatta East lagoon were below

0.2 µg ergosterol.g-1

, it has been extracted from aquatic sediments in other studies. For

example, ergosterol concentration was measured at 60-70 µg.g-1

and 240 µg.g-1

in

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sulfide–rich spring and freshwater pond sediments, respectively (Luo, Krumholz et al.

2005), and 39 µg ergosterol .g

-1 AFDM was found in the sediments of Canadian prairie

wetlands (Headley, Peru et al. 2002). These results do not represent high levels of fungal

biomass, but it may be that the fungal organisms in the sediments of many wetlands are

not detectable using this method.

Oomycetes and Chytridiomycetes do not contain ergosterol, and are therefore their

biomass cannot be quantified using this method (Gessner, Bauchrowitz et al. 1991;

Newell 1992). If Oomycetes, Chytrids and their propagules comprise a significant

component of the sediment fungal community, then ergosterol analysis will

underestimate the fungal biomass present. However, these organisms will contribute

DNA to fungal community structure analysis. Similarly, silicate diatom frustules may

prevent ergosterol extraction from hyphae within them, while the procedure used to

extract DNA is likely to rupture these structures. Ergosterol concentrations can also be

expected to be lower at reduced oxygen tensions as the synthesis of ergosterol requires

molecular oxygen (Safe 1973).

Ravelet et al., (2001) demonstrated that exposure of sediments to UV light, such as

occurs with wetland drying, results in the photodegradation of ergosterol to

ergocalciferol (vitamin D2). This is also true for ergosterol contained in dead fungal

tissues (Mille-Lindblom, Wachenfeldt et al. 2004), but melanins in the fungal cell wall

protect ergosterol in living fungi from UV light (Butler and Day 1998). Therefore, the

ergosterol content of exposed sediments is a valid representation of living fungal

biomass, and if samples are stored correctly (Newell 1995) exposure to UV light is not

likely to influence sediment fungal biomass assessments.

The moisture content and organic matter content of the sediments are correlated to some

extent because saturation reduces oxygen availability, and organic matter is decomposed

at different rates in oxic and anoxic sediments. If the depth of the oxic sediment layer

varied spatially, for example with differences in sediment texture, mixing the sediments

would cause variation in the relative percentages of moisture and organic matter. In

future work, these variables may be measured more accurately by taking a sediment core

and assessing moisture and organic matter content, as well as fungal biomass and

community structure, separately for the oxic and anoxic layers.

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5.4.5 Ecological implications

As the fungal community in the wetland sediment was not found to change over 35 days

while inundated, or alternatively while exposed, but changed with distance from the

shore, then it is likely that the community was dormant (or growing very slowly) and

composed mainly of propagules. Given low ergosterol results, it is likely that the

propagules were derived from Oomycetes and Chytridiomycetes. Changes in community

composition after 90 days are likely to be due to the deposition of fungal organisms and

propagules from the inflowing water.

Many species belonging to the Chytridiomycetes are parasitic on phytoplankton, and

these infections may play a pivotal role in the collapse of algal blooms (Ellen Van Donk

1983). Fungal infection of diatom hosts is favoured under conditions of high light

intensity, high temperatures, and high host density (Bertrand, Coute et al. 2004). Some

diatom species survive dry periods and periods of low abundance by forming “seed

banks” in the sediments. However, infected algal cells have been found in the sediments

when there is no sign of infected cells in the water column (Kagami, de Bruin et al.

2007), so diatoms in the sediments may act as a seed bank for fungal organisms as well.

Fungal parasites of algal often feed upon algae that are too large for zooplankton to

consume (Kagami, de Bruin et al. 2007), but when their zoospores are released into the

water column they provide a food source for Daphnia (Kagami, Van Donk et al. 2004).

This link in the trophic relationships between wetland organisms helps to ensure that

primary production is not lost when the larger algal cells fall to the sediment surface on

senescence (Kiørboe 1993). This suggests that parasitic fungi may have an important

function in pelagic food webs in wetlands (Lefevre, Bardot et al. 2007) and has been

described as the “Mycoloop” by Kagami et al.(2007).

5.5 Conclusions

The fungal community in wetland sediments may have greatly reduced activity when (i)

sediments have been inundated for an extended period, due to low oxygen availability, or

(ii) when sediments are exposed and dry, due to low moisture availability. Oxygen and

moisture availability may be more suitable for fungal organisms during transitional

periods such as flooding and drying, in shallow water or where vegetation is established.

However, the harsh chemical conditions in a drying wetland and the short duration of

132

these conditions at any point on the sediment surface severely limit opportunities for

fungal growth. It is not yet clear what proportion of the changes observed in community

structure are a response to higher temperatures, increased concentrations of dissolved

nutrients and other substances, oxygen concentration or moisture availability. In

floodplain wetlands, the fungal community in the sediments is likely to comprise a “seed

bank” of propagules that are available to repopulate the water column after drying and re-

filling when suitable substrates, such as phytoplankton and fish, are available and

environmental conditions are optimum. In future work, the importance of fungal

parasites and saprobes in controlling algal blooms and utilising algal biomass should be

considered in Australian lentic systems.

It would be beneficial to confirm the identity of the fungal taxa present in the sediments

of floodplain wetlands, so as to better understand their ecological functions. Such

investigations might involve growing cultures of the species present and sequencing their

DNA. However, the low fungal biomass in the wetland sediments indicates that the

impact of sediment fungi upon wetland carbon cycles is likely to be minimal, even

though they may be important in sustaining pelagic food webs.

133

Fungal biomass accumulation on E. camaldulensis leaves differs

between rivers and wetlands

6.1 Introduction

In the mesocosm experiment (Chapter 4) the amount of fungal biomass accumulating on

submerged Eucalyptus camaldulensis leaves was 10-fold less than that reported on E.

globulus leaves in Spanish and Portuguese stream ecosystems. Such a large difference in

fungal biomass is likely to have significant impacts on aquatic food chains and carbon

cycles. In this chapter, the accumulation of fungal biomass on E. camaldulensis leaves is

compared between river and wetland sites, and possible relationships between fungal

biomass, fungal diversity and the food value of leaf detritus are explored.

Fungal biomass can act as an important food source for aquatic invertebrates such as

snails (McMahon, Hunter et al. 1974; Newell and Bärlocher 1993) and insect larvae

(Bärlocher 1981; Bärlocher 1982; Bärlocher 1985), and form the base of aquatic food

chains (Reid, Quinn et al. 2008). Fungal biomass is either removed from the leaf surface,

or the leaf itself is consumed. Ecological interactions such as this may limit opportunities

for fungal growth and reproduction, and influence standing biomass on submerged

leaves. Chung and Suberkropp (2009) demonstrated that caddisfly larvae were only able

to grow on leaves that were colonised by the aquatic hyphomycete Anguillospora

filiformis, and early instar caddisfly larvae relied almost entirely on consuming fungal

biomass to fuel daily growth.

The variables most likely to influence the magnitude of fungal biomass accumulation in

aquatic systems include the quality of the litter substrate, the structure of the fungal

community, environmental conditions and interactions with other organisms. The

interaction of these variables over space and through time will result in variation in the

pattern of biomass accumulation between locations, seasons, and with changes in flow

and water quality.

The influence of substrate quality on fungal biomass accumulation was discussed in

Chapter 4 and reviewed by Gulis (2001). Substrate quality for submerged plant detritus

has been defined as the chemical composition of the decomposing material (Melillo,

Naiman et al. 1984) and is often described in terms of nutrient concentrations (carbon,

134

nitrogen and phosphorus). Alternatively, the percentage composition of the major classes

of plant bio-polymers (i.e. proteins, lipids, polysaccharides and lignin) may also infer

substrate quality (Chamier 1985). These factors are also known to have a strong

influence on the rate at which a substrate is decomposed (Melillo, Aber et al. 1982;

Taylor, Parkinson et al. 1989). Fungal colonisation is known to enrich leaves with

protein and nitrogen (Bärlocher 1985), and low ratios of carbon to nitrogen are correlated

with nutritious food sources (Boyd and Goodyear 1971; Hladyz, Gessner et al. 2009;

Taylor, Parkinson et al. 1989). Eucalyptus leaves colonised by fungi have been shown to

be a potential source of nutrition for consumers (Canhoto and Graça 1996; Reid, Quinn

et al. 2008; Schulze and Walker 1997), therefore changes in fungal biomass on

submerged leaves is likely to impact upon higher trophic levels in aquatic environments.

Patterns of biomass accumulation observed in Chapter 4 suggest that biomass fluctuates

with succession between early and late colonisers of the submerged leaf. This pattern

consists of rapid growth following inundation, a reduction in biomass that may be related

to sporulation events, and a later phase of growth and biomass loss. These two growth

phases correlate temporally with early and late colonisation by different fungal

communities, which may both be present on the substrate during a period of transition.

Other studies have shown changes in biomass with change in community structure

(Medeiros, Pascoal et al. 2009b; Van Ryckegem, Gessner et al. 2007).

Fungal biomass and community composition on submerged detritus can change with

location due to changes in the riparian vegetation (Bärlocher and Graça 2002; Findlay,

Howe et al. 1990; Graça, Pozo et al. 2002; Shearer and Webster 1985) and may differ

between flowing and non-flowing aquatic environments (Baldy, Chauvet et al. 2002;

Pattee and Chergui 1995). Environmental conditions such as temperature (Bärlocher,

Seena et al. 2008), salinity (Hyde and Lee 1995; Roache, Bailey et al. 2006) and pH

(Bärlocher and Rosset 1981; Iqbal 1976) are known to influence fungal community

structure (Fischer, Bergfur et al. 2009; Nikolcheva and Bärlocher 2005) while dissolved

nutrients and oxygen influence both community structure (Field and Webster 1983;

Medeiros, Pascoal et al. 2009b) and biomass production (Gulis, Ferreira et al. 2006;

Gulis and Suberkropp 2004; Sridhar and Bärlocher 2000; Suberkropp 1998). High

concentrations of ambient nitrogen may provide an opportunity for fungi colonising

135

substrates low in nitrogen to accumulate biomass and thereby enrich the substrate with

nitrogen (Stelzer, Heffernan et al. 2003).

The fungal community within decomposing leaf material in the aquatic environment co-

exists with other organisms, including bacteria and Oomycetes. Fungi may compete with

these organisms to obtain nutrients from the leaf biomass. Therefore, the abundance of

such organisms might influence the ability of aquatic fungi to grow and accumulate

biomass (Gulis and Suberkropp 2003; Mille-Lindblom, Fischer et al. 2006a). Whilst

being biochemically distinct from fungi, Oomycetes have similar morphology, size and

habitat usage (Money 1998). In addition, they tend to colonise submerged plant litter in

standing water bodies very early in the aquatic decomposition process (Czeczuga,

Mazalska et al. 2005; Nechwatal, Wielgoss et al. 2008). It has been suggested that early

colonisation may be related to this organism’s inability to hydrolyse cellulose as true

fungi can (Dix and Webster 1995). However, research suggests that many species of the

Oomycetes possess enzymes that would enable them to degrade cell wall polymers,

including cellulose (Latijnhouwers, de Wit et al. 2003; Valikhanov 1988) as well as

proteins and lipids (Al-Rekabi, Al-Jubori et al. 2005; Mozaffar and Weete 1993). Fungi

are known to compete with bacteria (Mille-Lindblom, Fischer et al. 2006a) and other

fungal species (Fryar, Booth et al. 2005) and it is reasonable to assume that Oomycetes

are also competing with fungi for resources. These competitive interactions may be

particularly important in the relatively harsh conditions that prevail over the summer

months (Menge and Sutherland 1987; Nikolcheva and Bärlocher 2004) when stream flow

and dissolved oxygen concentrations are reduced.

Fungal biomass accumulates on leaves submerged in rivers (Sridhar and Bärlocher 2000)

and standing waters (Mille-Lindblom, Fischer et al. 2006b). Pozo et al. (1998) showed

that the leaves of Eucalyptus globulus support a similar level of fungal biomass to those

of Alnus glutinosa in Spanish streams, although on E. globulus fungal diversity was

lower and colonisation was delayed (Bärlocher and Graça 2002; Graça, Pozo et al. 2002).

In Chapter 4 it was found that E. camaldulensis leaves in standing water accumulate

fungal biomass of similar magnitude to that reported from submerged macrophytes in

standing water by Mille-Lindblom et al. (2006b), but this was approximately one tenth of

that reported by Pozo et al. (1998). The variation in these results necessitates

discrimination between the effects of the leaf substrate and those of streams and standing

136

waters. Moreover, neither fungal nor Oomycete biomass accumulation, or the influence

of this biomass on substrate food quality, has been studied in Australian rivers and

floodplain wetlands, and these environments may present unique ecological challenges

for aquatic fungi.

In this chapter, I have quantified the fungal and Oomycete biomass that accumulated on

E. camaldulensis leaves submerged in lowland rivers and floodplain wetlands at three

sites over time. I have also examined changes in the structure of the fungal and

Oomycete communities and related changes in fungal biomass to changes in community

composition, and variations in the nutritional value of the leaf detritus. To achieve this, I

devised a field study, where litter bags containing E. camaldulensis leaves were

submerged in three paired wetland and river sites. The fungal biomass as ergosterol

concentration and estimation of hyphal length were compared between the two habitats

over 107 days. Fungal and Oomycete community structure were compared over time

using Terminal Restriction Fragment Length Polymorphism, and the carbon to nitrogen

ratios of partially decomposed leaves were used to estimate their nutritional value in

aquatic food webs.

6.2 Methods

6.2.1 Experimental design

Given the differences in substrate quality between fresh and floodplain aged E. camaldulensis leaves

that were discussed in Chapter 4, only aged leaves were used as substrates in this experiment, in an

attempt to reduce variation due to substrate differences. E. camaldulensis leaves were collected from

the dry floodplain at the Ovens River site and stapled into packs consisting of three leaves then dried

at 25°C for a period of 2 weeks in a Contherm Series Five dehydration oven (Contherm Scientific

Ltd, Korokoro, New Zealand). Leaf packs were sewn into envelopes made from 500 μm heavy duty

nylon mesh with polyester thread. A river rock was included to ensure the leaf packs rested on the

surface of the wetland sediment. Sufficient litter bags for all sample times were secured together with

nylon fishing line, anchored to a star picket and submerged in 20 – 30cm of water at three positions

in each river and wetland site in November 2008. Positions were chosen to correlate with the

presence of large river red gums, so that it could be assumed that propagules capable of

decomposing these leaves would be present in the environment. A diagrammatic representation of

this design is shown in

Figure 6.1.

137

Figure 6.1: A diagrammatic representation of the experimental design. Litter bag positions are

represented by asterisks.

Suitable sites were selected on 3 rivers in north-eastern Victoria. Wetlands were selected

because they (a) held water during the experimental period; (b) had close proximity to

the stream; and (c) had riparian vegetation dominated by E. camaldulensis. Close

proximity of the wetland to the stream was intended to equalise factors such as

temperature and precipitation, and the influence of soils and vegetation.

Leaf packs were retrieved from river and wetland sites after 2, 15, 30, 60, and 107 days.

All samples were removed by 10 March 2009 (Day 107). This sampling pattern was

designed to capture temporal changes in the fungal community dynamics and substrate

quality, based on results from the mesocosm experiment (Chapter 4). Dry leaves that had

not been submerged were denoted as time zero samples. Accumulated sediment was

rinsed off litter bags in the water body in which they had been incubated, and then litter

bags were stored on ice (~4°C) and transported to the lab where leaves and leaf discs

were preserved for further analysis.

6.2.2 Field sites

All field sites were situated in north-east Victoria, Australia (Figure 6.2), within the

catchment of the Murray River. The catchments of the Ovens and Kiewa Rivers

contribute to the larger Murray River catchment with the Kiewa River joining the Murray

River upstream of the Ovens confluence. Most rivers in this region are regulated, the

Murray River itself having several major impoundments, including the Hume weir at

Albury-Wodonga. The exception is the Ovens River, one of the few remaining

unregulated rivers in Australia. All sites were used by waterfowl and accessible to cattle.

Wetland

Wetland

*

3

3

3

3

3

3

3

3

3

3

*

3

3

3

3

3

3

3

3

3

3

*

3

3

3

3

3

3

3

3

3

3

*

3

3

3

3

3

3

3

3

3

3

*

3

3

3

3

3

3

3

3

3

3

*

* = litter bag

Stream

139

Sites chosen were:

1. Kiewa River (west bank) at Killara (Vic), and an adjacent wetland (Figure 6.2);

2. Ovens River (east bank), north of Peechelba Rd, Peechelba East (Vic) and an

unnamed wetland on its eastern side; and

3. Murray River (north bank), west of Howlong (NSW), and Quat Quatta East

Lagoon (see also Chapter 5).

Figure 6.2: Location of field sites within Australia (inset) and within the state of Victoria.

6.2.2.1 Kiewa River, Killara, Victoria

The Kiewa River site and the adjacent wetland were located south of the Murray Valley

Highway at Killara, north-east Victoria, Australia (36° 08′ 24.33″S, 146°57′18.37″ E,

152 m elevation), within the catchment of the Murray River. The Kiewa River rises in

the Victorian Alps and joins the Murray River below the Hume Dam. While it receives

hydro-electric power generating discharges from Rocky Valley and Pretty Valley Dams,

the flow regime of the Kiewa River is flashy, with low flows in summer, and spates

associated with winter/spring rains. Overbank flows associated with winter and spring

rainfalls fill the floodplain wetlands. Stream discharge, water level, electrical

conductivity and temperature are summarised in Table 6.1. The wetland was full at the

140

commencement of this study and did not receive any overbank flows throughout the

study period, so it was largely dry by the end of the study period. The woodland

vegetation was dominated by the river red gum (E. camaldulensis) but included some

exotic tree species. The surrounding land use was largely pastoral.

(a) (b)

Figure 6.3: The study site on the Kiewa River at Killara showing the wetland (a) and the river (b).

River red gums (E. camaldulensis) and exotic species dominate the riparian vegetation, and

understorey vegetation is dominated by pasture grasses.

6.2.2.2 Ovens River, Peechelba East, Victoria

The Ovens River site and the associated wetland are located north of Peechelba Road at

Peechelba East, Australia (36°9′ 44.72″S, 146°14′ 16.49″E, 137m), upstream of the

Peechelba Flora reserve. The Ovens River rises in the Alps of north-eastern Victoria and

has a confluence with the Murray River at Peechelba. It floods seasonally in winter and

spring of most years (Quinn, Hillman et al. 2000). Stream discharge, water level,

electrical conductivity and temperature are summarised in Table 6.1. The wetland, which

is filled with flooding flows from a minor creek, was full at the commencement of the

study but did not receive any inflows during the study period and was dry by January 19,

2009.

This site was located within the Killawarra State Forest. The vegetation of this area is

dominated by E. camaldulensis open forest with an understorey consisting of silver

wattle (Acacia dealbata), river bottlebrush (Callistemon sieberi), Chinese scrub

(Cassinia arcuata), grey parrot-pea (Dillwynia cinerascens) and native grasses (Parks

141

Victoria Website - http://users.mcmedia.com.au/~stocky/warby.html#killawarra). It is

used for camping and recreational fishing.

(a) (b)

Figure 6.4: The study site on the Ovens River at Peechelba East showing the wetland (a) and the

river (b). River red gums (E. camaldulensis) dominate the riparian vegetation, and understorey

vegetation is dominated by native grasses.

6.2.2.3 Murray River, Howlong, New South Wales

The Murray River site was located west of Howlong, New South Wales, at the Point

where Quat Quatta East Lagoon connects with the river (35° 58′ 19.82″ S, 146° 34′

44.67″ E, Elevation 150m elevation). The Murray River is one of Australia’s longest

rivers, having its source in the Snowy Mountains of New South Wales and draining west

into Lake Alexandrina in South Australia. It is highly regulated (Maheshwari, Walker et

al. 1995), supplying water for irrigation, stock usage and domestic supply over the

warmer months, while flows in cooler months are retained in storages. The water level in

Quat Quatta East lagoon decreased until the wetland was nearly dry during the study

period, but it received inflows via an inlet from the Murray River in late December 2008,

and began to refill. Stream discharge, water level, electrical conductivity and temperature

for the Murray River downstream of the study site at Corowa (36° 0′ 25.92″ S, 146° 23′

43.08″ E) are summarised in Table 6.1. The vegetation around the Murray River and

Quat Quatta East Lagoon is dominated by E. camaldulensis open forest with a grassy

understory (Margules and Partners Pty Ltd, P and J Smith Ecological Consultants et al.

1990; Pressey 1986). The area is grazed by cattle and used for camping and recreational

fishing.

142

(a) (b)

Figure 6.5: The Murray study site showing Quat Quatta East lagoon (a) and the Murray River west

of Howlong (b). River red gums (E. camaldulensis) dominate the riparian vegetation along this

stretch of the river and around wetlands, and understorey vegetation is dominated by grasses.

Table 6.1: Summary of the environmental conditions in the river at each site over the study period.

Data are based on the mean value of mean daily readings, with maximum and minimum mean daily

readings going to comprise the stated range. Data were obtained from the Murray-Darling Basin

Authority website (http://www.mdba.gov.au/water/live-river-data).

Location Discharge

(ML/day)

Mean

(range)

Electrical Conductivity

(µS.cm-1

)

Mean

(range)

Water Level

(m)

Mean

(range)

Temperature

(°C)

Mean

(range)

Ovens River at

Peechelba

507

(106-1975)

55.83

(21.07-82.11)

1.17

(0.54-3.04)

23.8

(18.2-30.4)

Murray River at

Corowa

8879

(5076-11910)

51.98

(38.67-64.59)

2.12

(1.49-2.58)

23.3

(17.7-28.3)

Kiewa River at

Killara

535

(109-2421)

33.54

(16.98-58.93)

0.94

(0.58-2.05)

22.1

(4.1-29.6)

6.2.3 Organic matter content

The leaf mass loss on ignition was determined in the same manner as described earlier in

4.2.2.

Data for mass loss and change in organic matter content were analysed using two-way

analysis of variance (ANOVA) with replication (3) for each sampling time. Time was

used as the covariant, pair-wise for treatments (Bärlocher 2005). Data analyses were

performed using SPSS Statistics 17.0 (IBM, Chicago, USA).

144

6.2.4 Carbon to nitrogen ratio

6.2.4.1 Carbon content

The carbon content of leaves was estimated as 50% of the ash-free dry mass (Lindroth,

Kopper et al. 2001), and expressed as mgC.cm-2

leaf area.

6.2.4.2 Total nitrogen content

Leaf samples taken from litter bags were stored frozen and then dried at 80°C in a

dehydration oven for 4 days. Leaves were then ground to a coarse powder in a coffee

grinder and 5 - 50mg of leaf material was then weighed into a 30ml vial. Samples were

then analysed for Total Nitrogen Content (TPN) at the analytical chemistry lab of the

Murray-Darling Freshwater Research Centre, Wodonga, following sodium hydroxide /

potassium persulfate digestion and conversion to the diazonium salt (American Public

Health Association, American Water Works Association et al. 1989).

Leaf mass was converted to leaf area using a conversion factor derived from the Dry

Mass determination of leaf discs from each site and at each time. Nitrogen content was

then expressed as mgN.cm-2

leaf area.

6.2.4.3 Mean carbon to nitrogen ratio

The carbon to nitrogen ratio was calculated by dividing mean leaf carbon content by

mean leaf nitrogen content, where both parameters were expressed in mg.cm-2

.


6.2.5.1 Ergosterol content

Fungal biomass was calculated from the ergosterol content of partially decomposed

leaves. The method used to quantify ergosterol concentration and calculate fungal

biomass has been detailed in Section 4.2.3. However, in this instance, leaf discs were

9mm in diameter, and not 12mm, and HPLC run time was reduced to 14 minutes, from

20 minutes.

6.2.5.2 Hyphal bio-volume

The biomass of filamentous fungi and stramenopiles on and within leaf discs were

determined using the “Membrane filtration method” (Hanssen, Thingstad et al. 1974;

Robinson, Borisova et al. 1996), which uses fluorescent dyes to simplify counting.

145

Briefly, three leaf discs (9mm in diameter) from each site replicate at each time were

preserved in 10ml of 1M potassium hydroxide (KOH) and stored at 4°C. Replicate leaf

discs were weighed, and then dried at 105°C for 1 week before being re-weighed to

determine the fresh: dry mass ratio for the leaf material.

Prior to analysis, the discs in 1M KOH were heated to 100°C in a heating block for 15

minutes. Then the KOH was drained off and the discs were washed three times in 10ml

of distilled water for 15 minutes. This process clears the leaf material and improves

contrast between fungal material and the leaf matrix (Díez-Navajas, Greif et al. 2007).

The three leaf discs in each sample were then blotted dry and weighed, with the mass

recorded to 4 decimal places. These discs were then homogenised in a tissue grinder and

suspended in 50 ml of 67mM K2HPO4 (phosphate buffer, pH 9.0). This suspension was

then further diluted (1:5) with phosphate buffer. Five ml of this diluted sample, mixed

with 2 drops of 0.1% phenolic aniline blue, was then filtered through a black

polycarbonate, 25mm membrane filter (0.8µm pore size) (GE Osmonics, Minnetonka,

MN, USA). Three replicate filters were prepared for each suspension, and these were

blotted dry and mounted on a standard glass slide in “Cargille” (non-drying, low UV

fluorescence) immersion oil Type B (ProSciTech, Thuringowa, Qld). Slides were

examined under a Zeiss Axioskop2 epifluorescence microscope (Ziess, Oberkochen,

West Germany) at 200 x magnification using a Nikon UV-1A filter cube.

Hyphal length estimates were made with an eyepiece grid at five randomly selected

positions on the filter using the grid-intersection method (method 5) as reported by Olson

(1950). The diameters of all filaments in one grid of each filter were measured, and mean

hyphal diameter was then calculated. Mycelial biomass was calculated from hyphal

length following the method proposed by Frankland et al. (1978). These authors used the

following equation: B = πr2 x L x D x M (where B is the mass of oven dried mycelium in

oven dried litter (g.g-1

); r is hyphal radius in mm; L is hyphal length in meters per mg of

oven dried litter; D is the relative density of the hyphae (g.cm-3

); and M is the percentage

moisture content of fresh hyphae divided by 100). Literature values for hyphal density

range from 1.1-1.5 g.cm-3

(Van Veen and Paul 1979) and moisture content generally

ranges from 80-90% (Hawker, 1950). I used an estimated density of 1.2 g.cm-3

and an

estimated moisture content of 85%.

146

Fungal biovolume data were analysed using 3-way ANOVA on the SPSS software

package (IGM, Chicago, USA), analysing between leaf type, treatments and times

(α=0.05). The fungal biomass results from ergosterol and hyphal biovolume analyses

were standardised by dividing each value by the total of all values obtained for each

method and compared using simple linear regression (SPSS, IGM, Chicago, USA).

6.2.6 Fungal community structure

6.2.6.1 Total fungal community

Fungal community structure was assessed using terminal restriction fragment length

polymorphism (T-RFLP). This method is described in Section 4.2.5.

6.2.6.2 Oomycetes community structure

Amplification of the ITS region of the Oomycetes DNA was carried using the same

genomic DNA template as the total fungal community. Amplification of Oomycete DNA

followed the procedure reported by Nikolcheva and Bärlocher, (2004), and differed from

the method described in Section 4.2.5 in terms of the length of DNA amplified, the

primers used, and the thermocycler protocol. Briefly, primers used amplified the entire

ITS region, extending from 18S rDNA to 28S rDNA. The forward primer was ITS5 (5'-

GGAAGTAAAAGTCGTAACAAGG– 3') (White, Bruns et al. 1990), which carried a

fluorescent label (5,6 Carboxyfluorecein) at the 5' end, and the reverse primer was ITSOo

(5'-ATAGACTACAATTCGCC – 3') (Nikolcheva and Bärlocher 2004). Primers were

synthesised by Gene Works (Hindmarsh, South Australia). These primers produce DNA

fragments that were approximately 1.25 kb in length and were specific to Oomycetes.

Amplification was carried out in an “iCycler” (Bio-Rad, Hercules, CA , USA)

thermocycler. The protocol consisted of initial denaturation at 95°C for 15 minutes (1

cycle), then 35 cycles of denaturation at 94°C for 15 seconds, annealing at 49°C for 15

seconds, and extension at 72°C for 60 seconds. A final cycle was included with

denaturation at 94°C for 15 seconds, annealing at 49°C for 90 seconds, and extension at

72°C for 6 minutes. The PCR product was then held at 4°C until being removed from the

thermocycler and stored at -20°C.

147

6.3 Results

6.3.1 Mass loss and organic matter content

6.3.1.1 Organic matter content

3-way ANOVA on leaf organic matter content data showed that organic matter content

differed significantly between rivers and wetlands (P<0.005) and over time (P<0.05).

These results are shown in Figure 6.6. No significant difference was found between the

Ovens, Murray and Kiewa river systems (P=0.649) or with interaction effects. The

general pattern for all sites was for a rapid decrease in organic matter content over the

first 15-30 days. Following this, organic matter content did not change and was

consistently lower in leaves from rivers than in those from wetlands.

6.3.2 Nitrogen content

3-way ANOVA showed that nitrogen content of submerged leaves differed significantly

between sample times (P<0.005), but not between sites or between rivers and wetlands

(P= 0.397 and P=0.258, respectively). However, there was an interaction effect between

river systems and time (P= 0.054). While increases in nitrogen content were seen on

some days in some sites, the nitrogen content generally decreased over time.

The nitrogen content of submerged leaves changed over time, and these results are

illustrated in Figure 6.7. In the Ovens system mean increases of 49 and 47% nitrogen

content were seen after 2 days in the river and the wetland, respectively, although there

was large variation between replicates. By day 15, nitrogen content was 40% lower than

the initial value in the wetland, but remained similar to initial levels in the river. The

nitrogen content of leaves submerged in the Ovens system was approximately half of the

initial value for the remainder of the incubation period, mean values increased with time

but variation between replicates was high. In the Murray system nitrogen content

decreased by approximately 20-25% between the initial value and the value after 2 days

submerged. Following this, small decreases were seen over time, with little difference

between the river and wetland samples, except at 30 days submerged. At this time, leaves

in the river had mean nitrogen contents 75% higher than those in the wetland. In the

Kiewa system mean nitrogen content decreased by 30-50% between day 0 and day 15

148

and then remained constant at approximately 50% of the initial value, while mean

nitrogen content decreased slightly at day 107.

149

Figure 6.6: Change in the mean organic matter of leaf packs over a 107 day incubation period is

shown for the Ovens (i), Murray (ii) and Kiewa (iii) sites. Samples incubated in the river indicated by

open symbols while samples from wetlands and indicated by solid symbols. The x-axis shows time in

days, while the y-axis shows the organic matter content of leaf discs as grams per square centimetre.

Error bars indicate the standard error of the data point.

150

Figure 6.7: The nitrogen content of leaves taken from the Ovens (i), Murray (ii) and Kiewa (iii) sites.

Samples incubated in the river indicated by open symbols while samples from wetlands and

151

indicated by solid symbols. Ovens samples are shown as circles, Murray samples as triangles, and

Kiewa samples as diamonds. The x-axis shows time in days, and the y-axis shows nitrogen content in

mg.cm-2

. Error bars show standard error of the mean of each data point.

6.3.3 Mean carbon to nitrogen ratio

The data obtained from leaves in litter bags (Figure 6.8) indicate that C:N tends to

decrease in leaves in rivers over the first 15-30 days of aquatic incubation. After this

time, leaf packs had C:N values close to that of floodplain aged (Day 0) leaves. Leaves

incubated in the wetlands showed decreases in C:N after 2 days. Over days 15-30, leaves

taken from the Ovens and Kiewa wetlands showed C:N above that of floodplain aged

leaves, but by 60 days C:N was again similar to floodplain aged leaves. Leaves incubated

in the Murray wetland showed similar trajectory of change in C:N to river samples from

the same sample time, although C:N was generally lower in the river samples.

3-way ANOVA of C:N indicated differences between sites (P=0.067) and over time

(P=0.055), but these differences were not significant at the 95% confidence level. A

significant interaction effect was detected between site and time (P<0.005).

The carbon: nitrogen ratio (C:N) results are shown in Figure 6.8 compared with the C:N

of 29:1. C:N values below this figure have been reported to represent a nutritious food

source for invertebrates (Esteves and Barbieri 1983). Submerged E. camaldulensis leaves

exceeded a C:N of 29:1 in the first 1-2 months after inundation. Sample times when C:N

fell below 29:1 varied between sites, and between rivers and wetlands. In the Ovens

system submerged E. camaldulensis leaves showed C:N below 29:1 after 2 days, but by

15 days leaves submerged in the wetland had C:N above this value and this persisted for

the remainder of the incubation period. After 15 days submerged leaves from the Ovens

River had C:N below the 29:1 but by 30 days C:N was similar to leaves from the

wetland. In the Murray system the C:N value was below 29:1 at 15 and 30 days aquatic

incubation, but rose above this value after 60 day. The C:N was always above 29:1 in the

Kiewa wetland, but in the Kiewa River C:N was less than 29:1 at sample day 2, and

approximately equal to 29:1 at day 107.

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Figure 6.8: Carbon to nitrogen ratio of leaf litter from litter bags in the river and wetland at the

Ovens (i), Murray (ii) and Kiewa (iii) sites. The x-axis shows the incubation time in days, while the y-

axis indicates the ratio of mean carbon content (mgC.cm-2

) to mean nitrogen content (mgN.cm-2

).

Wetland sites are indicated by solid symbols, river sites are indicated by open symbols. The red line

153

indicates the approximate value below which a resource is considered nutritious to invertebrates. At

day 30 C:N in the Ovens River and wetland were similar.

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6.3.4.1 Ergosterol content

Analysis of ergosterol content data indicated that fungal biomass differed significantly

over time (3-way ANOVA: P<0.005), and differed between rivers and wetlands (3-way

ANOVA; P=0.051), but no significant difference was found between river systems, or

with interaction effects. No significant differences were found using pair-wise t-tests to

compare biomass in river and wetlands and across sites.

When considered on an areal basis, biomass tended to increase over 15-30 days then

decrease. The time when maximum mean fungal biomass was observed differed between

sites. Maximum biomass occurred after 30 days in the Kiewa River, 15 days in the Ovens

River and 2 days in the Murray River (Figure 6.9). In the wetlands maxima occurred

after 2 (Kiewa), and 15 (Ovens and Murray) days. Fungal biomass decreased thereafter,

although rates of decrease differed. While fungal biomass tended to be higher in the river

than in the wetland at the Murray and Kiewa sites, at the Ovens site biomass was higher

in the wetland.

Linear regression indicated no direct relationship between leaf nitrogen content and

fungal biomass as estimated by ergosterol content (r2=0.010; P<0.05) or hyphal length

(r2=0.052; P<0.05).

6.3.4.2 Hyphal biovolume

Fungal biomass as determined by hyphal biovolume estimation differed from estimates

by ergosterol concentration. Maximum mean fungal biomass was seen in pre-inundation

leaves at all sites, decreasing with time (Figure 6.9) and differing significantly between

times (3-way ANOVA: P<0.005). Contrary to results from the analysis of ergosterol

concentration data, no difference was found between sites or between rivers and wetlands

(3-way ANOVA: P>0.05). Some correlation was found between fungal biomass as

measured by ergosterol and hyphal biovolume (Regression: r2 = 0.57; P<0.005), but a

large amount of variation in hyphal biovolume data was not explained by ergosterol

content data.

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In general, rivers and wetlands at each site showed similar patterns of temporal change,

with lower mean biomass in wetlands than in rivers, although the difference was not

significant. This pattern is seen most clearly at the Ovens and Murray sites. At the Kiewa

site an increase in fungal biomass was seen in the wetland after 60 days, and at this time

fungal biomass in the wetland exceeded that in the river. However, there was no

significant difference in biomass (by biovolume) between sites for this time (3-way

ANOVA: P=0.302).

6.3.4.3 Biomass estimates by ergosterol content compared with hyphal biovolume

Fungal biomass values measured as ergosterol content generally decreased over time,

while the combined biomass of fungi and Oomycetes, as measured by hyphal biovolume,

showed initial decreases then remained stable. The difference between fungal biomass

estimated by ergosterol and hyphal length suggests that the biomass of the Oomycetes

was more substantial during the first few days of aquatic decomposition, and after 60

days.

The differences in the two fungal biomass data sets (ergosterol and hyphal biovolume)

resulted from differences in magnitude and in the direction of change over time. Firstly,

estimates of biomass by hyphal biovolume were generally higher than those estimated by

ergosterol content, when data were expressed per area of leaf. Secondly, while ergosterol

analysis indicated decreasing fungal biomass after a maximum at 2-15 days (30 days in

the Kiewa River), hyphal biovolume data showed that biomass decreased for 15-30 days

after inundation, then remained stable or increased.

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Figure 6.9: Fungal biomass from leaves submerged at the Ovens (i), Murray (ii) and Kiewa (iii) sites.

Open symbols indicate river samples while solid symbols indicate wetland samples. Samples at the

Ovens sites are represented by circles, samples at the Murray site are represented by triangles and

samples at the Kiewa site are represented by diamonds. Un-dotted (white and black) symbols

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indicate biomass determined by biovolume, while dotted symbols (White and grey) indicate biomass

measured by ergosterol concentration. Error bars indicate standard error about the mean at each

data point.

6.3.5 Community structure

6.3.5.1 Total fungal community

Analysis of all T-RFLP data indicates that the fungal community differed with time

(ANOSIM: R=0.550; P<0.005; see also Figure 6.10), but not between sites, or between

rivers and wetlands. Pair-wise ANOSIM showed that community structure was similar

between subsequent sampling days (R<0.48; P<0.419). However, fungal community

structure at day 30 was significantly different to the community structure prior of

immersion in the water (R=0.809; P<0.05), and after 107 days in the water (R=0.683;

P<0.005). Fungal community structure was also significantly different between Day 2

and Day 107 (R=0.754; P<0.005). The trajectory of change with time (Figure 6.11) has 2

directions. For river sites, diversity increases between days 2 and 30 then decreases. In

wetlands, changes in the number of amplicons have no apparent pattern.

When T-RFLP data from the three sites were considered independently, the river and

wetland differed at the Kiewa site (R=0.542; P<0.005) (Figure 6.12), but were similar at

the Ovens (R=-0.075; P=0.668) (Figure 6.13) and Murray (R=0.036; P=0.469)(Figure

6.14) sites. The fungal community structure was similar between sites when rivers

(R=0.091; P=0.197) and wetlands (R=0.353; P<0.005) were considered independently

and pair-wise tests showed no significant differences between river sites (R<0.145;

P>0.135). However, the Kiewa wetland differed somewhat from the Ovens (R=0.364;

P<0.05) and Murray (R=0.532; P<0.05) wetlands, while the Ovens and Murray wetlands

were similar (R=0.139; P=0.267) (Figure 6.15). When data were analysed separately for

each sampling occasion, wetland sites and river sites (R=0.596; P<0.05) and river

systems (R=0.600; P<0.005) can be distinguished on Day 60 (Figure 6.16). A sample

from the Ovens River was not clustered with other river samples. These leaves may have

been periodically exposed due to low flows in the river.

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Figure 6.10: MDS ordination plot of T-RFLP results showing fungal community structure changes

over time. Hexagons as shown in the legend indicate community structure on leaves prior to being

submerged (Day 0), and after 2, 15, 30, 60 and 107 days submerged in the water. Circles indicate the

grouping of samples from 30 days and before on the left, and from 60 days and after on the right. X-

and Y-axes indicate relative position in multivariate space and have no units.

Figure 6.11: This plot indicates the mean position of MDS points from each sampling occasion, with

error bars indicating standard error about that mean. The data indicate that fungal community

structure changed significantly between Day 0 and day 30, then changed significantly in a different

manner between day 30 and Day 107. X- and Y-axes indicate relative position in multivariate space

and have no units.

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Figure 6.12: An MDS plot of T-RFLP from the Kiewa System showing variation in fungal

community structure between river and wetland sites at days 30, 60 and 107, but not at earlier times.

X- and Y-axes indicate relative position in multivariate space and have no units. Circles indicate

samples taken at the same time and have no statistical meaning. Numbers indicate sample time for

individual data points.

Figure 6.13: An MDS plot of T-RFLP from the Ovens System showing only small differences in

fungal community structure between river and wetland sites within sampling times. X- and Y-axes

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indicate relative position in multivariate space and have no units. Circles indicate samples taken at

the same time and have no statistical meaning.

Figure 6.14: An MDS plot of T-RFLP from the Murray System showing small differences in fungal

community structure between river and wetland sites within sample times, although the ordination

will be influenced by the small number of replicates for Murray wetland samples. X- and Y-axes

indicate relative position in multivariate space and have no units. Numbers indicate sample time

(days).

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Figure 6.15: An MDS plot of T-RFLP data from the wetland samples showing differences in fungal

community structure between wetland sites and clustering by sampling time. Circles are intended to

indicate groupings by time and do not indicate a statistical difference. X- and Y-axes indicate relative

position in multivariate space and have no units. Numbers indicate sample time (days).

Figure 6.16: An MDS plot of T-RFLP from the Day 60 showing that the fungal community structure

tends to cluster by river and wetland sites, and wetland also cluster by river system. Circles are

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intended to indicate groupings by time and do not indicate a statistical difference. X- and Y-axes

indicate relative position in multivariate space and have no units. An outlying sample from the

Ovens River may have been periodically exposed due to low flows in the river.

6.3.5.2 Oomycete community structure

PCR of genomic DNA from time zero (floodplain aged) samples did not produce any

product DNA. This may indicate that Oomycetes were absent from these samples, or that

inhibiting substances in these samples interfered with the amplification reaction. Thus,

time zero samples were excluded from data analysis rather than being given a zero value.

Ordination of T-RFLP results showed that the Oomycetes community changed over time

(Figure 6.17). This was a gradual transition, with substantial overlap between data from

different times. Thus, separation was not significant when all data were considered

(R=0.344; P<0.005). However, pair-wise tests showed that dissimilarity between day 2

and subsequent days increased from day 15 (R=0.235; P<0.005), through to day 30

(R=0.436; P<0.005), day 60 (R=0.569; P<0.005) and day 107 (R=0.844; P<0.005). There

is a clear trajectory for this change (Figure 6.18) which is driven by increasing numbers

of taxonomic units, although the pattern of this change differs between sites. At the

Ovens site the number of T-RF lengths increased until day 15 in the wetland, and day 30

in the river, and then declined. At the Murray site, the number of different T-RF lengths

increased steadily over time, and at the Kiewa sites this number fell until day 30, and

then rose again.

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Figure 6.17: MDS ordination plot showing Oomycete community structure with time. Hexagons as

shown in the legend indicate community structure on leaves after 2, 15, 30, 60 and 107 days

submerged in the water. X- and Y-axes indicate relative position in multivariate space and have no

units.

Figure 6.18: The mean position of all points from each time as shown in Figure 6.17. The axes

represent dimensions of ordination space and have no labels. Error bars indicate the standard error

of the data point. This graph shows a clear change in the composition of the Oomycete community

moving from left to right, driven by increases in the number of T-RF lengths.

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Figure 6.19: MDS ordination plot of the Oomycete community structure showing a low degree of

separation between sites, and between rivers and wetlands. River sites are indicated by open symbols

while wetland sites are indicated by solid symbols. Symbols indicate the Kiewa (diamonds), Murray

(triangles) and Ovens (circles) sites.

Ordination of Oomycete T-RFLP data showed that the degree of separation between river

systems was low (Figure 6.19) (ANOSIM: R=0.25; P<0.005), and pair-wise tests showed

no significant separation between individual sites. When data were analysed separately

for wetlands, the separation between sites improved (R=0.451; P<0.005). River sites

were more dissimilar (R=0.603; P<0.005), with pair-wise tests showing that this result

was largely driven by differences between the Murray and Kiewa Rivers (R=0.674;

P<0.005).

The degree of dissimilarity among sites changed with time. Separation between sites was

lowest at day 2 (R=0.289; P=0.052), reached a maximum at day 15 (R=0.738; P<0.05),

decreased again at day 30 (R=0.401; P<0.05), then increased at day 60 (R=0.59; P<0.05).

At day 15, wetland samples were separated by site (Figure 6.20), but river samples

clustered into a single group. Pair-wise analysis showed that all sites were dissimilar

(Kiewa/Ovens R=0.701; Kiewa/Murray R=0.822; Ovens/Murray R=0.688; P<0.05).

While some separation between river and wetland samples was discernable, ANOSIM

(R=0.28; P<0.05) showed that separation was low for rivers and wetlands when all sites

and times were considered. When a single site was considered, ANOSIM showed greater

dissimilarity between rivers and wetlands in the Murray (R=0.573; P<0.005) and Kiewa

systems (R=0.62; P<0.005) (Figure 6.21).

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Figure 6.20: MDS ordination plot of Oomycete T-RFLP data from 15 days submerged leaf packs.

River sites are indicated by open symbols while wetland sites are indicated by solid symbols. Symbols

indicate the Kiewa (diamonds), Murray (triangles) and Ovens (circles) sites.

Circles are drawn to illustrate groupings and do not imply statistical similarity.

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Figure 6.21: MDS ordination plot of Oomycete T-RFLP data from leaf packs submerged in the

Kiewa River and an associated wetland. Samples from the wetland are shown as solid diamonds,

while samples from the river are shown as open diamonds.

6.4 Discussion

This study has shown that the food quality of submerged leaves and the biomass and

community structure of fungi and Oomycetes changes over time in the aquatic

environment. Food quality indicators and fungal biomass (ergosterol) generally

decreased over time, while the combined biomass of fungi and Oomycetes (hyphal

biovolume estimates) showed initial decreases then remained stable. The diversity of

Oomycetes also increased over 107 days in the aquatic environment. Changes observed

in the structure of the fungal and Oomycete communities, biomass and leaf resource

quality are consistent with succession in the decomposer community as resources within

the leaf are consumed, altering its chemical composition.

6.4.1 The nutritional value of submerged E. camaldulensis leaves

Fresh E. camaldulensis leaves have been reported to have a high nitrogen content of

1.29% dry weight (Cork and Pahl 1996; Taylor, Parkinson et al. 1989), and this work

shows that floodplain aged leaves have a similar nitrogen content (1.28 + 0.60% dry

weight). This suggests that while nitrogen is undoubtedly being used by terrestrial micro-

organisms, most is being retained within the leaf. This nitrogen is likely to be either in

the form of microbial biomass or complexed with lignin. These results do not imply that

there has been no loss of nitrogen, but that relatively more carbon has been lost through

processes such as respiration. The relative loss of carbon is illustrated by changes in C:N

with litter decomposition. While surface litter has a C:N of around 100:1, the ratio falls

to 30-15:1 in partially decomposed litter and 10:1 in humus (Griffin 1972). This study

shows that when floodplain aged E. camaldulensis leaves were inundated the C:N

decreased, and the change in C:N was driven by decreases in organic matter content and

short-term increases nitrogen content. However, lower C:N values were limited to the

first 30 days after inundation, and the pattern of change differed between sites.

The C:N of a food resource is an indication of its food value to consumer organisms

(Boyd and Goodyear 1971; Cummins and Klug 1979; Hladyz, Gessner et al. 2009), and

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C:N of 29:1 has been nominated as the nutrient balance suitable for invertebrate

consumers (Esteves and Barbieri 1983). A previous study (Glazebrook and Robertson

1999) indicated that floodplain aged E. camaldulensis leaves had a C:N of 95:1, and

when these leaves were inundated on the floodplain C:N fell to 45:1 after 49 days;

subsequently C:N increased to 132:1 after 112 days. The current study has further shown

that this increase in food value is temporary and site specific, with C:N being lower than

29:1 for 2 days in the Kiewa River and Ovens wetland, 15 days in the Ovens River and

30 days in the Murray River and the associated wetland. C:N on leaves in the Kiewa

wetland was never less than 29:1. As the substrate was the same in all cases, these

differences in C:N are likely to be driven by environmental factors such as nutrient

concentrations. Change in C:N as leaves are inundated could also be expected to differ

between tree species (Kominoski, Hoellein et al. 2009). After 60 days of inundation, C:N

values were similar to those found by Glazebrook and Robertson (1999) after 49 days.

Inundated E. camaldulensis leaves break down quickly relative to the leaves of other

eucalypt species (Schulze and Walker 1997), and leaching that occurs during the first 30

days in the water has a strong influence on breakdown rates (Janssen and Walker 1999;

Liakos 1986; O'Connell, Baldwin et al. 2000). Fast decomposition rates have previously

been associated with low C:N values (Enriquez, Duarte et al. 1993), but this study has

shown low C:N vales are found in E. camaldulensis leaves for a limited period. This

period corresponds with maximum organic matter losses due to leaching, and differs

between rivers and wetlands.

Periods when loss of organic matter from submerged leaves was most rapid also

corresponded with periods of higher fungal biomass. Negligible changes in organic

matter content after 30 days of inundation similarly corresponded to lower fungal

biomass. This pattern was consistent with carbon consumption and respiration by the

decomposer community, but the change in biomass may indicate a response by fungal

decomposers to the decreased availability of specific, readily metabolised leaf

components, rather than organic matter generally. Furthermore, microbial activity and

leaching interact during aquatic litter decomposition (Bunn, De Deckker et al. 1986;

Campbell, James et al. 1992b), so the contribution of leaching, respiration and fungal

reproduction to losses of organic matter from E. camaldulensis leaves should be explored

further (cf. Pascoal and Cassio 2004).

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The nitrogen content of floodplain aged E. camaldulensis leaves was 0.045 mg.cm-2

and

reached a maximum value of 0.067 mg.cm-2

after 2 days submerged in the Ovens River.

This suggests low levels of nitrogen incorporation into E. camaldulensis leaves. This

period of nitrogen enrichment during the early phase of aquatic decomposition

corresponded with periods of high estimated Oomycetes biomass (see section 6.4.2). As

these organisms are known to be able to enrich their substrates with nitrogen

(Willoughby and Redhead 1973), they may be important in improving the nutritional

value of submerged eucalypt leaves. It should be possible to test this hypothesis using

nitrogen isotopes to trace nitrogen from the water column through to its incorporation

into proteins in leaves colonised by Oomycetes and aquatic hyphomycetes.

Carbon and nitrogen in fungal biomass are generally found in the ratio of 12-8:1, and the

optimum C:N for an artificial fungal substrate has been reported to be around 15:1

(Griffin 1972). A substrate C:N above 40:1 can inhibit the production of conidia (Jackson

and Bothast 1990), and a substrate with C:N >30:1 would be considered to be nitrogen

limited (Melillo, Aber et al. 1982). As the floodplain aged leaves of E. camaldulensis

have a C:N of around 40:1, they could be considered to be a nitrogen limited substrate.

On such substrates, biomass accumulation may be limited (Stelzer, Heffernan et al.

2003) and fungi may recycle nitrogen internally in order to maintain growth by extending

hyphal tips (Boswell, Jacobs et al. 2002; Cowling and Merrill 1966; Paustian and

Schnurer 1987). Under these circumstances fungi will allocate most of their nitrogen to

cell wall synthesis (Paustian and Schnurer 1987). This would result in increased fungal

biomass measurements, whether estimated by ergosterol content or hyphal length, with

no corresponding net increase in the nitrogen content of the substrate / mycelial complex.

Thus, in the absence of a bioavailable external nitrogen source fungal colonisation of

submerged E. camaldulensis leaves may not result in reduced C:N or increased

nutritional value. Nitrogen limitation may be significant in the role of fungi in aquatic

carbon cycles as low nitrogen availability is known to stimulate lignin degradation by

fungi (Kirk and Farrell 1987).

Over the experimental period, total nitrogen in the water column was highest in the

Murray River and lowest in the Kiewa River (0.6mg/L and 0.2mg/L, respectively, in

March) (Victorian State Government, EPA Victoria et al. 2010). This represents

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moderate to high levels of dissolved nitrogen. While aquatic fungi are capable of utilising

inorganic nitrogen from the water column (Suberkropp and Chauvet 1995), nitrogen

enrichment may stimulate production by benthic and pelagic algae, and fungi may then

utilise algal proteins as a nitrogen source (Artigas Alejo 2008). Either way, higher

ambient nitrogen in the Murray system is correlated with lower C:N, and conversely

lower ambient nitrogen in the Kiewa system is correlated with higher C:N ratios. In

support of this notion, the C:N of aged E. delagatensis submerged in a nitrogen limited

upland stream did not record values lower than 29:1 over a 64 day incubation period

(Love 2005).

6.4.2 Fungal and Oomycete biomass

The differences in the timing of fungal biomass maxima between sites and in the

trajectory of change in biomass in the two fungal biomass data sets (ergosterol and

hyphal biovolume) may be related to the flow regime. At day 60 conditions were hot, the

Ovens and Kiewa wetlands were drying and flows in these rivers were low. Low flows

may encourage increased Oomycete biomass and discourage increases in eumycota

biomass. At the same time the Murray River had high flow rates (due to irrigation flows)

and the wetland was receiving inflows. At this site, there was no suggestion of a

significant increase in Oomycete biomass.

Ergosterol concentration reflects the biomass of filamentous fungi and yeasts (Eumycota)

(Gessner and Chauvet 1993), including spores, conidia and vegetative cells in addition to

the growing mycelium (Newell 1992). Biomass estimated by biovolume (hyphal length)

reflects the abundance of Oomycetes (Stramenopila) as well as filamentous fungi, but

excludes yeasts, spores, conidia and non-filamentous fungi. Therefore, significant

differences in biomass between rivers and wetlands by ergosterol concentration, but not

by hyphal length determination are not inconsistent. Together these data suggest that

differences between rivers and wetlands were driven by the abundance of true fungi

(Eumycota), and that the combined biomass of Eumycota and Oomycota did not vary

spatially.

Aquatic hyphomycetes invest a significant proportion of their biomass into the

production of conidia which are subsequently released into the water and lost from the

decaying leaves (Suberkropp 1991b). The highest rates of sporulation occur at the end of

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the growth phase, and this has been shown to occur after approximately 18 days when

leaves are submerged in a stream (Suberkropp, Gessner et al. 1993). Fungal biomass on

submerged leaves increased for 15 days in the Ovens River and wetland, Murray wetland

and Kiewa River using the ergosterol method. These increases were not reflected in the

biovolume data. This difference probably represents the production of conidia because

these propagules contain ergosterol but are not included in biovolume estimates.

Subsequent declines in biomass were consistent with the loss of fungal biomass due to

sporulation. Gradual decreases in biomass in the Kiewa wetland and Murray River are

consistent with the gradual loss of mycelial tissue, and may indicate the absence of a

reproductive event.

Biomass estimated by biovolume indicated an increase in mycelial mass after 60 days in

the Ovens River and wetland, and after 30-60 days in the Kiewa River and wetland.

These increases were not reflected in ergosterol data. Thus, this increase probably

indicates increasing Oomycete biomass as flows in the rivers decreased and water levels

in the wetlands declined. This vegetative growth event was absent from the Murray site,

where flows were increasing in the river and the wetland was receiving inflows. Biomass

estimated from biovolume was also relatively high on Day 0 and Day 2. This would also

indicate a significant proportion of the biomass was composed of Oomycetes, and is

consistent with literature suggesting they are common during the early stages of aquatic

decomposition (Bärlocher 1990; Bärlocher and Kendrick 1974), and in terrestrial plants

and litter (Neher 1999). Biovolume biomass is consistently higher than ergosterol

biomass during the later phase of aquatic decomposition. This may be due to the

presence of non-living hyphae which would lose their ergosterol content soon after death

but remain visible by epifluorescence microscopy.

Pozo et al. (1998) found that the leaves of Eucalyptus globulus (blue gum) support

similar amounts of fungal biomass to Alnus sp. (alder). The maximum fungal biomass

found on E. camaldulensis leaves in this study was 3.69% of AFDM, while maximum

biomass reported for Alnus sp. ranged from 5.8-9.8% AFDM (Ferreira, Gulis et al. 2006;

Gessner and Chauvet 1994; Gessner, Robinson et al. 1998; Hieber and Gessner 2002)

and for Salix sp. ~8% AFDM (Baldy, Gessner et al. 1995; Hieber and Gessner 2002).

This indicates that the fungal biomass found on E. camaldulensis leaves in Australian

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streams and wetlands in this study was approximately half of that seen on deciduous tree

leaves in streams and wetlands of temperate zones of the northern hemisphere.

Possible causes of lower fungal biomass on eucalypt leaves in Australian systems include

substrate quality, and seasonal and site differences. The chemical composition of the

freshly abscised E. globulus leaves used by Graça et al. (2002) will have been different

to the floodplain aged E. camaldulensis leaves used in this study because Eucalyptus

species differ in respect to concentrations of nutrients (Cork and Pahl 1996), essential

oils (Gilles, Zhao et al. 2010), and anatomical features (Pereira and Kozlowski 1976),

while floodplain aging is also known to alter the chemical character of eucalypt leaves

(Baldwin 1999). Season is known to influence fungal biomass, with higher biomass

reported in winter (8.8% detrital mass) than in summer (4.4% detrital mass) (Suberkropp

1997) in a stream at a similar latitude to those examined in this study. However, in higher

latitudes, peak fungal biomass on submerged litter occurs in both spring and autumn

(Verma, Robarts et al. 2003). The values reported from this study in the Murray River

system over the summer months fall within the range of summer biomass results reported

by Suberkropp (1997). Site differences such as flow rate, the concentration of dissolved

oxygen (Charcosset and Chauvet 2001), the probability of burial due to sedimentation

(Cornut, Elger et al. 2010) and the concentration of nutrients and inhibiting substances in

the water column (Stelzer, Heffernan et al. 2003; Suberkropp and Chauvet 1995) will

also affect fungal biomass accumulation. The abundance of Oomycetes has also been

reported to vary seasonally (Hallett and Dick 1981), with maxima in early spring, early

summer and autumn. The peak abundance of the conidia of aquatic hyphomycetes in an

Australian upland stream occurred in late summer (Thomas, Chilvers et al. 1989).

While there are no fungal biomass results published for Australian lowland rivers in

winter, Suter (2009) found maximum spring/summer fungal biomass on E. pauciflora

and E. delagetensis leaves to be 38.6 mg/g and 104 mg/g dry mass, respectively, after 13

weeks in an Australian alpine stream. As maximum biomass on E. camaldulensis leaves

in summer was 45-50 mg/g dry mass, Suter’s (2009) data suggest that fungal biomass on

eucalypt leaves is higher under cooler conditions, but winter biomass in lowland rivers

and wetlands will need to be measured directly to determine how fungal biomass varies

seasonally.

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6.4.3 Fungal and Oomycete community structure

Several authors have found that the fungal community that colonises submerged eucalypt

litter is distinct from the community colonising other litter types (Bärlocher, Canhoto et

al. 1995; Bärlocher and Graça 2002; Chauvet, Fabre et al. 1997; Thomas, Chilvers et al.

1992). My study has found that the fungal community on submerged E. camaldulensis

leaves does not differ between rivers and wetlands at sites on three rivers. This suggests

that the specialised fungal community colonising submerged eucalypt leaves is

ubiquitous in regions where these species are indigenous (Thomas, Chilvers et al. 1992).

There have been no previous studies investigating the diversity of Oomycetes colonising

submerged eucalypt litter. My study has shown that the Oomycete community shows

greater variability than the broader fungal community, differing between rivers and

wetlands, and between sites. As these organisms not only degrade submerged litter but

also have the potential to cause disease in fish, crayfish and amphibians (Kiesecker,

Blaustein et al. 2001; Singhal, Jeet et al. 1987; Söderhäll, Dick et al. 1991; van West

2006), these differences in community composition are potentially of ecological

significance. Oomycetes are also potentially strong degraders of lipids (Beakes and Ford

1983; Mozaffar and Weete 1993), suggesting a role in attacking the oils and waxes found

in eucalypt leaves.

While fungal community structure data presented here indicate the presence of fungi

belonging to the Ascomycetes, Basidiomycetes, Chytridiomycetes, Zygomycetes and

Oomycetes, the Oomycete community structure data represent a subset of this

community (Nikolcheva and Bärlocher 2004). It was hypothesised that Oomycetes would

have a stronger influence on the composition of the fungal community in standing waters

than in flowing environments, and it was indeed possible to discriminate between rivers

and wetlands using Oomycete community structure data, but only at the Kiewa and

Murray sites. This suggests that Oomycete community structure potentially differs

between rivers and wetlands but that it is influenced by site specific factors.

The structure of Oomycete communities is known to be influenced by factors such as

oxygen concentration (Dix and Webster 1995), pH (Lund 1934), season (Khallil, El-

Hissy et al. 1995; Nikolcheva and Bärlocher 2004; Roberts 1963), salinity (Padgett,

Kendrick et al. 1988) and the composition of the substrate (Bärlocher 1992). The

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requirement for oxygen and the ability to utilise specific plant components varies among

taxa, with some species (e.g. Leptomitus lacteus) being unable to utilise sugars, instead

consuming organic acids, fatty acids and amino acids. In general, Oomycetes show a

preference for substrates that are rich in chitin and cellulose. These include dead insects,

hemp seeds and snake skin (Park, Sorenson et al. 1978). This diversity and the ability of

many species to tolerate harsh environmental conditions helps to explain their ability to

utilise submerged plant detritus as a substrate in both early and late phases of

decomposition, as substrate composition and environmental conditions change.

Changes in fungal community structure observed on submerged E. camaldulensis leaves

are consistent with succession as the leaf substrate is consumed and its composition

changes (Frankland 1982). For example, fungi adapted to consume sugars may be

replaced by those able to utilise cellulose, and these fungi are then replaced by those able

to utilise lignin (Kjoller and Struwe 1992), although some species of fungi may be able to

utilise all of these substrates and maintain a presence throughout leaf decomposition.

Moreover, fungi and stramenopiles have been found to coexist on submerged plant litter

in standing water (Czeczuga, Mazalska et al. 2005; Czeczuga, Muszyńska et al. 2007;

Nikolcheva and Bärlocher 2004) and maximum species diversity is seen 3-6 weeks after

the leaves are immersed in a water body (Fischer, Bergfur et al. 2009).

6.4.4 Ecological implications

Results from this experiment indicate that aged E. camaldulensis leaves submerged in

lowland rivers and floodplain wetlands during summer only represent a nutritious food

source for invertebrates for about 1 month after inundation. The temporal limitations on

the food value of this detritus exist because the floodplain aged leaves are low in

nitrogen, and low substrate nitrogen combined with low concentrations of nitrogen in the

water column compel fungal colonisers to use nitrogen conservatively, but nitrogen will

be lost from the leaf when fungi reproduce and release propagules. As E. camaldulensis

loses most of its leaves in summer (Glazebrook and Robertson 1999), the continual

supply of leaves would ensure that nitrogen enriched leaves do not become scarce, but

better food quality where ambient nitrogen concentrations are higher will profoundly

influence the growth and reproduction of shredder communities (Gulis, Ferreira et al.

2006). Stable isotope studies of food webs in wetland systems (cf Bunn and Boon 1993;

Hladyz, Gessner et al. 2009; Reid, Quinn et al. 2008) would help to resolve these issues.

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The amount of fungal biomass accumulating on submerged E. camaldulensis leaves in

summer is consistent with previous studies (Suberkropp 1997; Suter 2009). It can

therefore be assumed that similar levels of fungal biomass are a normal component of

food webs in these systems and the composition and abundance of other aquatic taxa

have adapted under these conditions. However, it is likely that in situ fungal biomass

values will be lower on leaves in rivers and wetlands as it will be exposed to

consumption by invertebrates such as snails (Newell and Bärlocher 1993) and caddis fly

larvae (Bärlocher and Kendrick 1975; Chung and Suberkropp 2009). Grazing by

invertebrates is also likely to alter the composition of the fungal community (Sabetta,

Costantini et al. 2000), although, there are no published studies showing that

invertebrates are actually consuming conditioned eucalypt leaves in Australian floodplain

wetlands. Given the C:N of these leaves, wetland invertebrates may consume other types

of detritus preferentially.

While similarities can be expected between floodplain wetland environments by virtue of

the absence of flow, and similar riparian vegetation, floodplain wetlands can vary widely

in terms of their water quality (Goonan, Beer et al. 1992; Hall, Baldwin et al. 2006).

Summer draw down may exaggerate these differences by concentrating dissolved

substances in the water column, and this will strongly influence the composition of

aquatic fungal communities. Water quality issues currently facing Australian floodplain

wetland ecosystems include anoxic “black water” events (Howitt, Baldwin et al. 2007),

increased salinity (Nielsen, Brock et al. 2003) and decreasing pH (Baldwin, Hall et al.

2007). While salinity, for example, has been found to reduce fungal biomass in standing

water (Czeczuga, Muszyńska et al. 2007), it is important to determine how fungal

biomass and community composition in Australian systems will be impacted by these

disturbances and how this might affect the decomposition of plant detritus and detritivore

food chains.

Oomycetes are a little studied group of aquatic organisms (Newell 1992), that are

important elements of the decomposer community in lowland rivers and floodplain

wetlands. Their presence equalises total mycelial biomass between rivers and wetlands

and they may be instrumental in enriching leaf detritus with nitrogen. They are also

candidates for the 13

C-depleted primary producer that Bunn and Boon (1993) postulated

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formed the basis of floodplain wetland food chains. However, many Oomycete species

are inhibited by saline conditions (Lord and Roberts 1985; Padgett, Kendrick et al.

1988). It would therefore be of interest to investigate the decomposer community on

submerged detritus in saline wetlands and determine whether this detritus becomes

nitrogen enriched.

6.4.5 Method considerations

In this study, mesh bags were used to exclude invertebrate shredders from the leaf packs,

so that fungal accumulation in the absence of grazing could be assessed. It should be

noted that exclusion of invertebrates results in slower breakdown rates, and litter bags

may also alter the micro-environment of the leaf detritus. In particular, fungal community

structure will be influenced if propagules are prevented from landing on the leaf surface.

Here, it is possible that accumulation of biofilm on litter bags may even have prevented

new species from colonising the submerged leaves in the later phases of decomposition.

This will have influenced both biomass and community structure results. Future work

should compare fungal dynamics between leaf packs fixed together with buttoneers

(Boulton and Boon 1991) with results from large and small mesh litter bags to determine

if these effects are significant.

Loss-on-ignition has been used frequently to estimate the carbon content of soils (Hutson

1985; Wang and Bettany 1995) with the use of conversion factors specific to soil type

(Bent Tolstrup and Per Åkesson 1982; Konen, Jacobs et al. 2002). This method has been

used less commonly for determining leaf carbon content as more accurate methods are

available, but loss-on-ignition is a cheap and rapid method to estimate carbon content

given suitable conversion factors. A 50% conversion factor has been used here and

elsewhere (Lindroth, Kopper et al. 2001; Robertson, Bunn et al. 1999), and is in line with

Wood’s (1974) finding that the carbon content of decomposing E. delegatensis leaves

was 49.9% prior to incubation, and remained stable at 47.4-52.1% after 12 months of

incubation in a forest litter layer. For the purposes of this study, this method provides a

valid estimate of carbon content.

The biovolume method used to determine the mycelial biomass of fungi and Oomycetes

is time consuming, and may underestimate fungal biomass due to difficulties in

separating it from the substrate (Frankland, Lindley et al. 1978; Newell 1992). In

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addition, the procedure used to clear leaf tissue may have removed hyphae attached to the

leaf surface. However, this method has been used to estimate fungal biomass by several

authors (Findlay, Howe et al. 1990; Findlay and Arsuffi 1989; Newell and Hicks 1982;

Newell and Statzell-Tallman 1982) and provides an opportunity to simultaneously

identify the fungi present. While it has shortcomings, a more satisfactory method for

determining the biomass of Oomycetes is not currently available.

Aquatic Oomycetes were traditionally sampled from aquatic environments using baiting

methods (Nechwatal, Wielgoss et al. 2008; Park, Sorenson et al. 1978; Pettitt, Wakeham

et al. 2002), but these methods measure the number and species of propagules, not

mycelial biomass. More recently, quantitative PCR has been used to determine the

abundance of Phytophthora capsici infecting plant material (Silvar, Diaz et al. 2005 ) as

well as species belonging to Eumycota (Landeweert, Veenman et al. 2003), but in their

current form these techniques are species specific. It may be possible to develop these

techniques by using less specific primer pairs to quantify the biomass of the Oomycete

community in environmental samples.

Both methods used to measure fungal biomass depend on standard conversion factors,

and while it is valid to compare values within the data sets, caution should be used when

comparing biomass between data sets that have used different methods. For example, a

high abundance of yeasts in a fungal community can lead to an overestimation of fungal

biomass by ergosterol concentration because yeasts have an ergosterol content (37-42

µg.mg-1

(Pasanen, Yli-Pietila et al. 1999)) significantly higher than that of aquatic

hyphomycetes (2.3-11.5 µg.mg-1

), on which the ergosterol to fungal biomass conversion

factor used is based (Gessner and Chauvet 1993). As yeasts are commonly found

associated with plant detritus in lakes (Lefevre, Bardot et al. 2007), it is likely that the

ergosterol method overestimates fungal biomass in lentic systems. However, the

ergosterol content of mycelial tissue can be substantially lower under reduced oxygen

conditions (reduced from 85% to 37% of total sterols)(Nout, Bonants-Van Laarhoven et

al. 1987; Safe 1973), as the biosynthesis of ergosterol requires molecular oxygen, thus

the tendency toward lower oxygen concentration in standing water would place

downward pressure on ergosterol concentrations. Notwithstanding error due to

conversion factors, fungal colonisation of leaves is patchy, and biomass estimates can be

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highly variable within a single leaf and between leaves (Chamier, Dixon et al. 1984).

This spatial variability may exceed errors due to inaccurate conversion factors.

6.5 Conclusions

In this chapter I have shown that fungal and Oomycete colonisation of submerged E.

camaldulensis leaves results in a temporary improvement in food value. This

improvement coincides with periods of high Oomycete biomass in the Ovens and Kiewa

systems, and higher biomass of the Eumycota in the Murray system. These spatial

differences correlate with low and high flow conditions and ambient nitrogen

concentrations, respectively, while temporal differences correlate with changes in the

fungal community structure. Fungal biomass differed between rivers and wetlands, and

results suggested that Oomycete biomass was significant during the early phase of

aquatic decomposition, and after 2 months in drying wetlands and during low-flow

conditions in a stream.

Exploration of aquatic fungal ecology in Australian systems is a recent endeavour and

there are many questions still to be answered. For example, the relative contribution of

leaching and fungal production to carbon loss from eucalypt leaves is unknown, as are

the organisms mediating nitrogen enrichment in submerged leaves. While influence of

ecophysiological parameters such as season and water quality is known for European and

North American systems, their effects have not been measured in Australian systems. It

will be particularly important to determine how resilient fungal communities are to

disturbances such as salinity, black water and acidification. In wetlands, it is important to

determine whether detritivores actually consume conditioned E. camaldulensis leaves,

given the rarity of shredders in floodplain wetlands and the low nutritional value of these

leaves. Leaves may simply become entrained within wetland sediments (Janssen and

Walker 1999) where they are degraded by anaerobic bacteria (Boon 2006). Further

refinement of methods is also needed, particularly with respect to the use of litter bags

and the development of a molecular technique to quantify Oomycete biomass in

submerged plant detritus.

Relative increases in the nitrogen content of decomposing leaves have been assumed to

be due to increasing protein as fungal biomass accumulates. However, it is possible that

leaf nitrogen is being transformed into non-bioavailable refractory organic compounds as

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a result of enzymatic reactions while bioavailable carbon is lost. This would also result in

reducing C:N but would reduce the food value of the leaf resource in practical terms. If

this is the case, there are potential implications for detritivore food chains and carbon

dynamics in lowland rivers and floodplain wetlands. In the following chapter, changes in

the chemical composition of submerged E. camaldulensis leaves are tracked through the

aquatic decomposition process in a lowland river and an associated wetland using

spatially explicit analyses. Such spatially precise data allow chemical changes to be

correlated with zones of enzyme activity around fungal tissues and fungal succession on

submerged leaves. In particular, the distribution of organic nitrogen within decomposing

leaves is mapped over time.

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Synchrotron FTIR micro-spectroscopy reveals chemical changes

accompanying aquatic leaf decay.

7.1 Introduction

In chapter 4, fungal dynamics on submerged Eucalyptus camaldulensis leaves were

explored using enzyme activity to indicate the breakdown of leaf components. These data

suggested that the rate of breakdown of different structural carbohydrates in these leaves

differs between aquatic and terrestrial conditions and with fungal community structure.

The fungal community present on these leaves changes over time and differs between

rivers and wetlands (Chapter 6). As these differences will influence rates of carbon

mineralisation and storage within a wetland ecosystem, it is important to determine how

the chemical composition of E. camaldulensis leaves changes after colonisation by

aquatic fungi. By knowing how fungi alter their substrate we can infer the functional

significance of saprotrophic fungi colonising submerged plant material in rivers and

wetlands.

In Australia’s Murray-Darling Basin, Eucalyptus camaldulensis is the most common tree

species in riparian zones and on floodplains, and its leaf litter is an important and

abundant carbon resource for aquatic ecosystems. The leaves are exposed to terrestrial

decomposition processes on the floodplain before they are inundated or washed into

aquatic systems. During this terrestrial phase, water soluble leaf components, including

soluble polyphenols (Serrano 1992), are lost through leaching and refractory leaf

components may be altered by microbial enzymes, UV light and mechanical processes.

At inundation, 70-80% of the remaining soluble organic substances, including free sugars

and starches, are lost within a few days (Baldwin 1999; Campbell and Fuchshuber 1995;

Glazebrook and Robertson 1999; Kaushik and Hynes 1971; Suberkropp, Godshalk et al.

1976), so it is the more refractory structural carbohydrates derived from plant cell walls

that provide energy to aquatic fungi (Chamier 1985). Plant cell walls are comprised

mainly of cellulose, hemicellulose, pectin and lignin, and these polymers are extensively

cross-linked.

Aquatic fungi, such as aquatic and aero-aquatic hyphomycetes, are known to produce

enzymes which breakdown cellulose (Abdullah and Taj-Aldeen 1989), hemicellulose

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(Chamier, Dixon et al. 1984; Suberkropp and Klug 1980), starch (Abdullah and Taj-

Aldeen 1989), pectin (Chamier and Dixon 1982a; Chamier and Dixon 1982b), lipids

(Abdullah and Taj-Aldeen 1989) and protein (Abdullah and Taj-Aldeen 1989). Lignin,

however, is attacked by oxidative enzymes, and it has been suggested that fungi in low

oxygen aquatic environments such as wetlands are not capable of degrading it, and that

lignin degradation is primarily a terrestrial process (Cooke and Whipps 1993). However,

fungi from both of these groups, when grown on lignin agar, have demonstrated some

lignin degradation capabilities (Abdullah and Taj-Aldeen 1989; Bergbauer, Moran et al.

1992; Fisher, Davey et al. 1983). Oxidative degradation of aquatic humic substances by

fungal enzymes has also been demonstrated (Claus and Filip 1998; Filip and Claus

2002).

Increases in both relative and absolute lignin content with decomposition have been

noted in studies of terrestrial (Gallardo and Merino 1992) and aquatic (Suberkropp,

Godshalk et al. 1976) leaf decay. It is not possible for new lignin to be made in dead

leaves, but compounds which behave chemically similar to lignin may be formed. Lignin

break down products that react spontaneously with each other and with carbohydrates in

their immediate environment are the most likely source of this “new” lignin (Melillo,

Naiman et al. 1984).

As submerged leaves are decomposed, there is often an increase in their absolute

nitrogen content (Kaushik and Hynes 1971; Pidgeon and Cairns 1981; Triska and Sedell

1976), consequent reduction in the C:N ratio (Moran, Benner et al. 1989) and increase in

energy content (Liakos 1986). It has been proposed that this increase in nitrogen is due to

the accumulation of microbial protein (Bärlocher, Mackay et al. 1978; de la Cruz and

Gabriel 1974). This was supported by the finding that fungi are able to absorb nitrogen

from the water column and assimilate it into their biomass (Suberkropp and Chauvet

1995), and increases in ambient nitrate concentrations lead to accelerated leaf break

down (Liakos 1986). However, studies have shown that microbial protein comprises less

than 30% of the total leaf nitrogen (Odum, Kirk et al. 1979), and much of the remaining

nitrogen content is not in the form of protein (Rice 1982).

Several mechanisms have been proposed to explain the absolute increases in nitrogen and

lignin content observed in decaying leaves. Suberkropp et al. (1976) suggested that leaf

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polyphenols complex with protein during the early stages of aquatic decay, and that these

complexes are isolated in the lignin fraction. This was later supported by findings that

increases in the lignin content were correlated with losses of tannins (Gallardo and

Merino 1992). Odum et al. (1979) demonstrated that nitrogen enrichment is only in part

due to the presence of microbial protein. For example, nitrogen contained in fungal

biomass comprised only 16% of total immobilised N on submerged wood (Melillo,

Naiman et al. 1984). The non-proteinaceous nitrogen fraction (~70%) is likely to be

comprised of amino sugars (such as chitin and glucosamine), nitrogen containing humic

acids and nitrogen rich complexes of phenol-protein, lignin-protein, chitin-protein and

amino-clay minerals (Melillo, Naiman et al. 1984). Up to 50% of non-proteinaceous

nitrogen in leaf detritus may be derived from chitin (Odum, Kirk et al. 1979).

Rice (1982) explained increases in nitrogen and lignin in terms of the humification of

organic matter. He proposed that, following leaching, leaf material is colonised by micro-

organisms, and the activity of their enzymes produces reactive carbohydrates and

phenols. These molecules condense with peptides and amino acids to produce

nitrogenous geo-polymers. Microbes assimilate nitrogen from the water column into their

biomass and exudates, and these compounds later decompose and condense with reactive

carbohydrates and phenols. As humification proceeds, nitrogen is gradually transformed

from relatively labile amino-sugar, amino-phenol and polypeptide forms into chemically

recalcitrant heterocyclic, aromatic forms typical of humic material. Melillo et al. (1984)

showed that nitrogen accumulation is proportional to the concentration of water soluble

polyphenols, and that nitrogen immobilisation often corresponds to increases in absolute

lignin content. They proposed a cycle of enzymatic de-polymerisation and

recondensation that continues until de-polymerisation becomes inefficient in terms of

energy yields (Melillo, Naiman et al. 1984).

Bärlocher et al. (1989) combined leaf leachate with fungal exo-enzymes, and

demonstrated that insoluble polyphenol/protein complexes are formed. They further

showed that invertebrates having alkaline gut pH or surfactants in the gut fluids were

able to extract amino acids from these complexes. Gallardo and Merino (1992) suggested

that nitrogen from microbial biomass could be precipitated by leaf tannins in the forest

litter layer, and that this process represented the first step in the formation of humic

substances.

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The changes in leaf components found in previous research have been observed at the

“whole leaf” scale. That is, the leaf is ground and homogenised and the chemical

constituents of the homogenate are determined. However, fungal activities take place at a

microscopic scale, and different components of the leaf’s anatomy may be used

preferentially by decomposer organisms. So to understand the chemical changes that take

places as a result of fungal activity, it is necessary to examine the chemical changes at a

microscopic scale, and in a spatially explicit context.

Fourier transform infra-red spectroscopy allows us to infer the chemical composition of

an organic sample by measuring the infra-red absorbance resulting from the vibration of

chemical bonds. With the addition of a microscope, very small amounts of organic

material can be characterised. When infra-red detectors are arranged in an array (FPA–

FTIR), or where the detector scans sequential positions across the surface of the sample

(FTIR mapping), a spatially explicit image of the chemical components of the sample can

be compiled. The use of a synchrotron light source (S-FTIR) improves spatial and

spectral resolution so that we can determine chemical changes at the micron scale, a scale

similar to that of fungal tissues. While this technique has been used to chemically

characterise Eucalyptus leaves (Heraud, Caine et al. 2007), and fungal tissues (Jilkine,

Gough et al. 2008; Kaminskyj, Jilkine et al. 2008; Szeghalmi, Kaminskyj et al. 2007) it

has never been used to investigate decomposition processes.

In this chapter, I investigate the use of infra-red spectro-microscopic imaging as a means

of examining the distribution of proteins, polysaccharides, lipids and lignin in the

decomposing leaves of Eucalyptus camaldulensis. Changes in distribution are tracked

from a fresh leaf, to one that has been decomposing on the floodplain, then through

several stages of aquatic decomposition. I was also able to show differences in the

protein structures present in fresh and decomposing leaves. Given that fungal community

structure and biomass are likely to differ between wetland and stream environments (see

chapter 6), I compared chemical changes in leaves decomposing in these contrasting

environments. The higher spatial resolution of the FTIR spectro-microscopy using a

Synchrotron light source was then used to examine the distribution of these compounds

within the immediate vicinity of fungal hyphae.

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7.2.1 Plant material

Fresh leaves were collected from a mature E. camaldulensis tree on the bank of the

Kiewa River at Killara, Victoria, Australia (-36.140093S, 146.955185E, 152 m

elevation). Leaf discs of 9mm diameter were cut from the leaves and immediately

preserved in 4% formaldehyde.

For the abiotic leaching treatment, several fresh leaves were placed a sealed 500ml

polycarbonate jar with autoclaved water taken from the Kiewa River. In the biotic

leaching treatment, leaves were placed in a jar with untreated water from the river. These

samples were stored in the laboratory at room temperature (approximately 20°C) for 12

weeks. Leaf sections were then stored in 4% formaldehyde.

Aged leaves were collected from the leaf litter on the Ovens River floodplain

(-36.162422S, 146.237914E, 137m elevation), and were dried at 25°C for a period of 4

weeks in a dehydration oven. A selection of floodplain aged leaves were then stored in

formaldehyde to await histological processing, while the remaining aged leaves were

transferred into litter bags made from 500 μm heavy duty nylon mesh (to limit attack by

aquatic fauna) (Figure 7.1,a). Leaves were submerged at a depth of 20-30 cm at three

replicate positions in the Kiewa River and in an adjacent wetland at Killara (Figure 7.1,

b), and then retrieved at time intervals of 2, 15, 30, 60 and 107 days. Litter bags were

stored on ice after sampling, and transported to the lab where 9 mm leaf discs were cut

from partially decomposed leaves and stored in 4% formaldehyde at 4°C while awaiting

histological processing.

7.2.2 Histological method

Leaf discs were dehydrated in a series of ethanol baths (70%, 90% and three lots of

100%) for 12 hours in each bath. This was followed by clearing in vegetable oil xylene

for 12 hours, which was repeated three times. After this, leaf sections were soaked in

paraffin wax at 58°C for 12 hours, embedded in a paraffin blocks and set prior to

sectioning on a rotary microtome (Boundy, undated; Heraud, Caine et al. 2007).

Transverse sections of leaf specimens, 4μm thick were mounted onto tin-oxide coated,

glass slides with silver layers to provide reflectance (MirrIR low e slides: Kevley

Technologies, Chesterland, OH, USA). Transverse sections of 8μm thickness were

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mounted onto calcium fluoride windows (Crystran Ltd., Poole, UK) that had been coated

with poly-D/L-lysine. Paraffin was removed from the sections by immersion in three

xylene baths for 2 minutes each, immediately prior to infrared analysis.

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Figure 7.1: Floodplain aged leaves were inserted into mesh litter bags (a), then submerged in the river and floodplain

wetland (b). The wetland was an ox-bow lake on the floodplain of the Kiewa River, in north-eastern Victoria,

Australia, with vegetation dominated by Eucalyptus camaldulensis (c).

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Additional adjacent sections were prepared for examination under a compound

microscope. These sections were stained with periodic acid / Schiff reagent (PAS), which

binds to glycogen and carbohydrates, and crystal violet, which penetrates cell membranes

and stains DNA and cytoplasm purple. Images of these sections indicating the

organisation of leaf tissues and the absence of fungal hyphae in a fresh leaf, and the

presence of fungal hyphae in a partially decomposed leaf are shown in Figure 7.2.

7.2.3 Infrared spectroscopy

Broad scale infrared images of leaf midvein and adjacent mesophyll (FPA-FTIR) were

captured in both transflectance and transmittance modes using the 15× objective of an

Hyperion 3000 infrared microscope (Bruker Optik GmbH, Ettlingen, Germany), with a

total path length of 8 μm for both methods. The infrared microscope was equipped with a

64 × 64 element focal plane array (FPA) detector coupled to a Vertex 70 FT-IR

spectrometer (Bruker Optik GmbH, Ettlingen, Germany) operating in rapid scan mode,

and located at the Australian Synchrotron (Clayton, Victoria, Australia). Opus 6.5

software (Bruker Optik GmbH, Ettlingen, Germany) was used both to control the

instrument and to process the spectra. Most spectra were acquired in the “transflectance”

mode whereby the IR beam passes through the sample and reflects off the supporting

slide and trough the sample again to the detector. The remaining spectra were acquired in

“transmittance” mode, whereby the IR beam passes through the sample and the

supporting calcium fluoride window to the detector above. Total path length in both

modes was 8 μm. 2×2 pixel binning was used for image frame collection, and

apodization was performed using Blackman-Harris 3-term function. Data were obtained

with 64 co-added scans at 4 cm-1

spectral resolution. Each data set consisted of 16384

data points, collected over 16 blocks in 4×4 and 8×2 array comprising ~0.46 mm2 of leaf

area. For images including the leaf mesophyll, two 8×2 images were collected and are

presented with the common edges abutting. Reference (background) spectra with 64 co-

added scans were measured before each sample image frame.

Higher spatial resolution FT-IR images of leaf sections were acquired using an infrared

microscope with a synchrotron light source (S-FTIR). Absorption spectra maps were

aquired in transflectance mode using the 36× objective of an Hyperion 2000 infrared

microscope (Bruker Optics GmbH, Ettlingen, Germany) with a jacketed stage purged

with dry air to maintain an environment of stable humidity. A motorised stage enabled

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point by point raster scanning of the sample. The microscope was coupled to a Vertex

80V infra-red spectrometer and was equipped with a narrow-band liquid nitrogen cooled

MCT detector (Bruker Optik GmbH , Ettlingen, Germany) and connected to the infrared

beam line 2BM1 at the Australian Synchrotron (Clayton, Victoria, Australia). This

system was also controlled using Bruker’s Opus 6.5 software. The microscope has a

glass knife-edge aperture which was used to define a sample area of 5×5 μm. Images

used in this report had an area of less than 300×80μm (See figure captions for exact

dimensions).

Figure 7.2: 8 µm cross sections of the mid-vein of Eucalyptus camaldulensis leaves that have been stained with

periodic acid/Schiff’s (PAS). The left panels show a fresh leaf (a), and the right panels show a leaf that has been aged

on the floodplain then submerged in a wetland for 2 days (b). Images are shown at 100x (i) and 400x (ii) magnification.

While no fungal tissue can be observed in the fresh leaf, in the partially decomposed leaf, darkly staining fungal

hyphae have replaced the phloem tissues and much of the mesophyll. Bundle sheath cells appear to have collapsed in

the partially decomposed leaf. Fungal hyphae can be seen on either side of the fibre sheath and within the mesophyll of

the partially decomposed leaf (b(ii)). X = xylem; P = phloem; FS = fibre sheath; E = epidermis; BS = bundle sheath; F

= fungal tissue; M = mesophyll; OV = oil vesicle.

In Figure 7.7, (a) and (b), the specimen was mapped in 5 μm steps, whereas 2 μm steps

were used in Figure 7.7, (c). Data were collected with 64 co-added scans, spectral

188

resolution of 4 cm-1

, and with Happ-Genzel apodization function. Reference spectra were

collected after each 8 sample data points with 128 co-added scans.

For both FPA-FTIR and S-FTIR data were collected in the spectral region of ~3800 to

750 cm-1

.

7.2.4 Band assignments

The amide bands (amide I (1700-1637 cm-1

), amide II (1543-1527 cm-1

) and Amide III (

1330-1290 cm-1

)) are generally considered to be indicative of the presence of proteins.

Absorbance in the amide I band indicates vibration of carbonyl (C=O) bonds from

peptide bonds in proteins, but may also include absorbance due to carbonyl groups in

esters and aromatic compounds such as quinones.

Cellulose and hemicellulose consists of pyranose monomers linked by glycosidic (>C-O-

C<) bonds (Kacuráková, Capek et al. 2000). They are found in the middle lamella,

primary and secondary cell walls (Esau 1977; Raven, Evert et al. 1992). In infrared

spectra, they absorb strongly in the region associated with (C-O-C) and (C-OH) bonds

(1180-900 cm-1

). This frequency range will be referred to as the polysaccharide band.

Lignin is a complex three dimensional polymer comprised mainly of phenylpropanoid

units, with linkages and cross-linkages via carbon-carbon (C-C), ester (C-O-C) and a

variety of other linkages (Hammel, 1997). Lignin is found in the middle lamella, and in

the primary and secondary cell walls of xylem and fibres. As it is highly aromatic, it is

associated with the vibration of the C-C bonds of phenyl rings (1503 cm-1

), and ether

linkages associated with aromatic rings (1231 cm-1

). The band at 1503 cm-1

will be

referred to as the aromatic band, while the band at 1231 cm-1

will be referred to as the

ether linkage band. Together, these frequencies are referred to as lignin bands.

Plant oils and waxes are comprised of long-chain aliphatic “fatty” acids. Oils contain

triglycerols, which are in turn comprised of three fatty acids linked via ester bonds to

glycerol. Waxes are comprised of long-chain fatty acids linked to a long-chain alcohol

via an ester bond. Together with sterol and isoprenoid compounds, these compounds are

known as lipids. The carobonyl band at 1740-1730 cm-1

is associated with ester

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functional groups ((C=O)-O-C) from fatty acids, esters and lignin. It has been used here

to indicate the presence of lipids.

The spectral bands used in the analysis of Eucalyptus camaldulensis leaf tissue and

microbial signals are primarily based on the assignments used by Heraud et al. (Heraud,

Caine et al. 2007) in their analysis of Eucalyptus botryoides leaves. Assignment of

additional bands is based on Silverstien et al. (1974), Hori and Sugiyama (2003), Robert

et al., (2005), Seoudi et al.,(2005) and Bandekar and Krimm (1979) (see Table 7.1).

Table 7.1: Bands of interest in the FT-IR spectra obtained from decomposing leaf material in the range 1800-900 cm-

1. The column labelled “Wave Number” indicates the frequency limits between which each band was integrated,

“Band Assignment” indicates the molecular vibration responsible for absorbance in the stated frequency range (υ =

stretch; υs = symmetric stretch; υas = asymmetric stretch δ = deformation (bend)). The reference from which this

assignment was sourced is listed in the third column, and an indication of the probable source of these vibrations

within leaf material is indicated in the “notes” column.

Wave Number (cm-1) Band Assignment References Notes

1740-1730 ν(C=O) Heraud et al., 2007 Ester functional groups from fatty

acids, esters and lignin

1729-1713 ν(C=O) Silverstein et al., 1974 Aliphatic aldehydes and ketones

Amide I band

1700-1637

ν(C=O), ν s(C=C) Heraud et al., 2007 Mainly from proteins, also olefinic

and aromatic double bonds

1690-1655 ν(C=O) conjugated

to (C=C)

Silverstein et al., 1974 From quinones, aromatic aldehydes

and aromatic ketones

1620-1590 ν as(COO-) and

ν(C=C)

Heraud et al., 2007 From pectins and lignins,

respectively.

Amide II band

1543-1527

ν(C-H) and δ(N-H) Heraud et al., 2007 From proteins

Aromatic Band

1530-1486

ν(C=C) Heraud et al., 2007 Aromatic skeletal vibration from

lignins

1441-1396 ν s(COO-) Heraud et al., 2007 From pectins

1389-1259 ν s(NO2) Silverstein et al., 1974 Nitrates, nitramines, conjugated nitro

compounds, aromatic nitrates

1342-1266 ν(C-N) Silverstein et al., 1974 Aromatic amines

Amide III band

1330-1290

ν(C-H) and δ(N-H) Bandekar and Krimm,

1979

From proteins

1300-1255 ν as(NO2) Silverstein et al., 1974 Organic nitrates

Ether Linkage

1284-1185

ν(C-C), ν(C-O) Heraud et al., 2007 From lignins

~1261 δ(N-H) Seoudi et al., 2005 From N-acetyl side chain in chitin

1260-1180 νs(C-O-H) Silverstein et al., 1974 From polysaccharides, influenced by

hydrogen bonding

1250-1020 ν(C-N) Silverstein et al., 1974 Unconjugated C-N in aliphatic

amines

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Polysaccharides

1180-900

ν(C-O-C) Heraud et al., 2007 From polysaccharides

1180-1140 ν(C-O-C) Hori and Sugiyama , 2003 From β-1,4-glycosidic bonds from

microcrystalline cellulose

1085-1050 ν (C-O-H) Silverstein et al., 1974 Secondary alcohols groups from

sugars and phenols

1033 δ(C-O-C)

Hori and Sugiyama ,

2003; Robert et al, 2005

β-1,4-glucans from microcrystalline

cellulose

7.2.5 Data processing and image analysis

Spectral maps were constructed using Opus 6.5 software (Bruker Optik GmbH ,

Ettlingen, Germany). The Opus default integration method was used, whereby the peak

integration area is defined as the area under the curve between two selected wave

numbers, with a base line taken between the same two points. The wave number limits

for each band are shown in Table 7.1. Chemical maps were then constructed where x and

y co-ordinates are the spatial co-ordinates of a given sample, and the z co-ordinate is the

relative absorbance. Z-values were represented by a colour scale (Opus “rainbow” colour

scale) where lightest pink indicates the maximum and darkest blue the minimum relative

absorbance. Minimum absorbance was forced to zero excluding negative values and

maximum absorbance was set to a consistent value for maps of all samples at a given

frequency. High absorbance in a given band was correlated with structures and tissue

types within each leaf by overlaying IR absorbance contours on a visual image of each

sample. Images presented as absorbance contours overlaid on the bright field images are

used to highlight these correlations with plant structures, but are essentially the same as

full colour maps.

Spectra to be used in multivariate analysis were extracted from IR chemical images using

Opus 6.5 software. Firstly, maps indicating the distribution of absorbance in the amide I

band were created, and then spectra with a high absorbance were extracted from points in

the map.

Chemometric analysis of this data was carried out using “The Unscrambler” software

(CAMO Software AS, Oslo¸ Norway). Individual spectra with high absorbance in the

amide I band were sampled from Opus maps then imported into “The Unscrambler”.

These spectra were normalized by peak area, whereby the area under the curve was

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equalised for all spectra. The second derivative of the Amide I peak (1660-1630 cm-1

)

was then calculated using the Savitzky Golay function and 7 point smoothing. A

statistical examination of the derivatised data was carried out using Principal component

analysis (PCA), in order to determine whether differences in absorbance patterns in the

amide I band were significantly different between fresh E. camaldulensis leaves and

those that had undergone aqutic decomposition for 2 days. PCA employed leverage

correction, and the principle components were recalculated after removing outliers.

7.2.6 Light microscopy

Two sets of leaf sections, taken immediately adjacent to those used in FT-IR spectro-

microscopy, were stained using PAS and Crystal violet, respectively. These sections

were examined at 40×, 100× and 400× magnification using an Olympus BX 51compound

microscope equipped with an Olympus DP12 digital camera (Olympus Australia, Pty.

Ltd., Oakleigh, Victoria, Australia). Images were processed using Olysia software (Soft

Imaging System, Münster, Germany).

7.3 Results

Anatomical features of the vascular tissue in E. camaldulensis leaves are shown in Figure

7.3. Vascular traces are comprised of phloem fibres, phloem and xylem, and encased in a

bundle sheath. Cells comprising xylem and the fibre sheath cells are lignified, while

other vascular and non-vascular cells are not. In the mid-vein, vascular traces have the

distinctive appearance of a “smilie face”. Epidermal cells are impregnated with waxes

and oils known to inhibit fungal colonisation (Canhoto and Graça 1999). Anatomical

structures have been identified with reference to Esau (1977), Keating (1984), James and

Bell (1995), Ali et al. (2009) and Burley et al. (1971). Observations regarding the spatial

distribution of infra-red absorbance in various bands will refer to leaf structures as

identified here.

Very few fibre sheath cells are observed in the fresh leaf sample (Figure 7.3 (b)), as a

young leaf was selected to minimise the chance of fungal infection. This differs from

more mature leaves, such as the floodplain aged and partially decomposed leaves used in

this experiment. This is because the structure of the eucalyptus leaf vascular traces varies

with leaf age (England 2001). Young leaves may have very few fibres around the

vascular bundle, while in mature leaves a fibre sheath encircles the mid-vein close to the

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petiole (Figure 7.3, (a)), and fewer fibres are observed as the diameter of the leaf trace

decreases.

7.3.1 The chemical and structural components of fresh leaves

No fungal organisms were observed in sections of fresh leaf (Figure 7.4, a) or in fresh

leaf sections from the abiotic and biotic leaching treatments (Figure 7.4, b & c). Phloem

tissues were present in the fresh leaf between the xylem and the bundle sheath, but there

were very few fibre sheath cells. With abiotic leaching (Figure 7.4, b) few structural

changes were evident, but where leaching was accompanied by the activities of non-

fungal microorganisms from the water column (biotic)(Figure 7.4, c) the integrity of the

leaf began to break down. In this case, fissures began to develop on either side of the

fibre sheath and between the vascular bundle and the epidermis. There was also some

disintegration of the palisade mesophyll. As both leaching treatments were sealed after

water was added, it is likely that lack of oxygen inhibited fungal colonisation and the

breakdown observed in the biotic leaching treatment was due to anaerobic bacteria.

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Figure 7.3: Anatomical Structure of the mid-vein of Eucalyptus camaldulensis. Above we see a line drawing (a)

and an image of an 8µm cross section of the midvein of an E. camaldulensis leaf (b). The leaf has been stained with

periodic acid/Schiff’s (PAS) and is shown at 200x magnification. X = xylem; P = phloem; FS = fibre sheath; E =

epidermis and cuticle; BS = bundle sheath; PM = palisade mesophyll; SM = spongy mesophyll; C= collenchyma.

Lignin content in leaves is represented by absorbance at 1503 and 1231 cm-1

(see Table

7.1). In fresh leaves, the spatial distribution of these signals was tightly constrained in the

xylem (Figure 7.5, a (ii) & (iii)), but absorbance decreased and became more diffuse with

leaching and terrestrial aging (Figure 7.5, b, c & d (ii) and (iii)). Absorbance due to the

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presence of aldehydes and ketones was low in the fresh leaf sample (Figure 7.5, a (iv)),

and mainly associated with the bundle sheath.

FPA-FTIR data indicate that polysaccharides were present throughout the mid-vein of

fresh leaves, and in all tissue types (Figure 7.5, a,(v)). Images that also included a portion

of the mesophyll (Figure 7.6) indicate that most of the polysaccharides present in the

mesophyll of the fresh leaf (Figure 7.6, a) were lost during abiotic (Figure 7.6, b) or

biotic (Figure 7.6, c) leaching, and the polysaccharides that remained in the floodplain

aged leaf were confined to the vascular tissues (Figure 7.6, d).

The presence of proteins is indicated by absorbance in the amide bands (I, II, and III),

which indicate vibrations from C=O, NH2 and C-N, respectively, from within peptide

bonds. Absorbance in the amide I band was at its highest in the fresh leaf (Figure 7.5, a

(i)) and this absorbance was spatially correlated with the areas of the palisade mesophyll

adjacent to the vascular tissue. Relative absorbance in the amide I band was also high in

the phloem, and absorbance in the amide II and III bands in this area supports a

conclusion that this area contained protein. Areas of high absorbance in the amide II

band were also seen in the collenchyma, but this was not corroborated by amide I data

and may be due to the presence of lignin absorbing at similar frequencies. S-FTIR

mapping of a segment of the mid-vein showed that amide I absorbance had a patchy

distribution in the fresh leaf (Figure 7.7 (a)).

7.3.2 The chemical and structural components of terrestrial aged leaves

The terrestrially aged leaf (Figure 7.8, a & b (i)) had been colonised by fungi, which can

be seen between the xylem and fibre sheath in the vascular tissue. Phloem tissues were

no longer present. Large amounts of mesophyll tissue were also absent, and this area was

networked with fungal hyphae (Figure 7.8, a (i)). Reproductive structures were observed

close to, or protruding through, the epidermis (Figure 7.8, a (i) & c (ii)). Hyphae were

absent from oil vesicles.

In the terrestrially aged leaf section (Figure 7.5, d (i)) the area of high amide I absorbance

was smaller than in the fresh leaf, and was associated with fungal tissues rather than leaf

tissues. S-FTIR mapping of a segment of the mid-vein in a terrestrially aged leaf is

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shown in Figure 7.7, b (i). This finer scale image shows that amide I distribution was

correlated spatially with fungal tissues.

Figure 7.4: Compound microscope images leaf sections. Images are shown at 200× magnification and stained with

PAS. The fresh leaf (a), the leaf that was leached in autoclaved water (b), and the leaf that was leached in non-sterile

river water (c) show no sign of fungal colonisation. The leaf that was found on the floodplain (d) is extensively

colonised with fungal hyphae, which is observed as darkly staining tissues surrounding the xylem and fibre sheath, and

replacing the phloem and mesophyll. Fungal hyphae (H) are networked throughout the mesophyll region of the

leaf found on the floodplain.

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Figure 7.5: Broad scale images of the leaf mid-vein, acquired using the Focal Plane Array FT-IR micro-spectrometer

(x15 magnification). Here we show four micrometer sections through E. camaldulensis leaves that were (a) removed

from a living tree (fresh); (b) leached in autoclaves river water; (c) leached in untreated river water; and (d) found on

the surface of the floodplain (floodplain aged). Bright field images of the unstained leaf section are shown in the left-

most column, while images showing the spatial distribution of absorbance in the following bands are shown in

subsequent columns: (i) 1700-1637 cm-1; (ii) 1530-1486 cm-1; (iii) 1284-1185 cm-1; (iv) 1729-1713 cm-1; and (v)

1180-900 cm-1. Lightest pink indicates highest absorbance while darkest blue indicates lowest absorbance.

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Figure 7.6: Loss of polysaccharides on leaching. Here, the same leaf sections that were shown in Figure 7.4 (Bright

field) and Figure 7.5 are viewed with portions of the leaf mesophyll. FPA-FTIR spectral images show absorbance in

the polysaccharide band (1800-900 cm-1) of a fresh leaf (a), a leaf that has been leached abiotically (b) and biotically

(c) in the laboratory, and of a leaf found on the floodplain. Maximum absorbance is fixed at 120 au (lightest pink) and

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minimum absorbance is 0 au (darkest blue). The highly absorbing are to the left of the midvein in image (d) is a minor

vascular trace.

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The amide III band is associated with vibrations from proteins, but the vibrations from

the N-acetylated side-chains in chitin and organic nitrates may also absorb in this region

(see Table 7.1). In the terrestrially aged leaf (Figure 7.9, c (ii)) there was no absorbance

in this band associated with xylem, but areas of higher absorbance correlated spatially

with the fibre sheath cells, where microscopic observation suggests that fungi may have

been attacking the middle lamella.

Many fungal fruiting bodies were observed in both terrestrially aged and aquatically

decomposed samples. The structures tended to absorb in the amide I band (Figure 7.10,

a) with variable responses in the amide II and III bands (Figure 7.10, b & c). Spectra

indicated that aromatics (Figure 7.10, d), lipids and polysaccharides (data not shown)

were also present. Areas of higher absorbance values in the amide I band that are not

spatially correlated with higher absorbance in Amide II and III bands within the fruiting

body, but with aromatic bands suggest that protein-like substances are associated with

aromatic structures in fungal tissues.

While in the fresh leaf absorbance in the lignin bands was tightly correlated with cell

walls in the xylem, the signal became more diffuse and extended over a larger proportion

of the mid-vein region in the leached (Figure 7.5, b and c (ii) and (iii)) and floodplain

aged leaves (Figure 7.5, d, (ii) and (iii)).

All polysaccharides present in the terrestrially aged leaves (Figure 7.6, d) were associated

with leaf vascular tissues and epidermis, and with fungal tissues. High relative

absorbance in the polysaccharide band was concentrated in the vascular bundle of the

terrestrially aged leaf, as it was in the fresh leaf, but the exclusive association with xylem

was expanded to include phloem fibres and other cell types. FPA-FTIR and S-FTIR

images indicate a strong correlation between the spatial distribution of lignin and

polysaccharides and leaf vascular tissues, as might be expected given these molecules are

extensively cross-linked in cell walls (Figure 7.5, d (ii), (iii) and (v)). S-FTIR images

indicated that lignin and polysaccharides were present in xylem and fibre sheath cells,

although fibre sheath cells had higher absorbance in the polysaccharide region and lower

absorbance in the lignin regions, relative to xylem (Figure 7.7, b (ii), (iii) and (v)). Areas

of higher absorbance in both aromatic and polysaccharide bands were found immediately

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adjacent to fungal tissue (around the “eye” and at the base of the “smile”), with

absorbance then gradually declining with increasing distance.

Discrete molecules of aromatic aldehydes, such as veratryl aldehyde (3,4-

dimethoxybenzaldehyde), are formed as products of the break down of lignin by

oxidative enzymes (Tien and Kirk 1984), and their presence in dead leaves indicates

regions of microbial enzyme activity. In FTIR, aldehydes have strong absorbance in the

carbonyl band at 1729-1713 cm-1

. FPA-FTIR and S-FTIR images showing the

distribution and concentration of aldehydes in the mid vein of the terrestrially aged leaf

are shown in Figure 7.5 (d, (iv)) and Figure 7.7 (b, (iv)), respectively. These images

indicate that oxidative enzyme activity towards lignin was most active while the leaf was

on the floodplain, and that this activity was concentrated in areas close to fungal hyphae.

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Figure 7.7: S-FTIR spectral maps of sections through the leaf mid vein.

Fine scale maps of sections through the leaf mid-vein, acquired using an (S-FTIR) (x 36 magnification). Here we

show chemical maps extending from the upper epidermis towards the centre of the leaf mid-vein and including

tissues from the xylem. All maps are derived from the same 4µm E. camaldulensis leaf sections as FPA-FTIR

images and have a pixel size of 5µm x 5µm. Here we show (a) a fresh leaf, the segment is 80 x 330µm, and uses

5µm step between pixels; (b) a floodplain aged leaf, the segment is 80 x 212µm, and uses a 5 µm step between

pixels; and (c) a floodplain aged leaf that has been submerged in a floodplain wetland for 2 days, the segment is

40 x 90µm and uses a 2µm step between pixels. Bright field images of the unstained leaf section are shown in the

left-most column, while images showing the spatial distribution of absorbance in the following bands are shown

in subsequent columns: (i) 1700-1637 cm-1; (ii) 1530-1486 cm-1; (iii) 1284-1185 cm-1; (iv) 1729-1713 cm-1; and

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(v) 1180-900 cm-1. Lightest pink indicates highest absorbance while darkest blue indicates lowest absorbance.

The area of the sample shown in the infra-red images is indicated by the red matrix on the visible image.

Figure 7.8: Fungal features in wetland and river decomposed leaves.

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Fungal hyphae (H) are networked throughout the mesophyll in a leaf found on the floodplain (a (i)), and in a

leaf that had been decomposing in the river for 2 days (a (ii)). Fungal hyphae are shown to extend through the

space formerly occupied by the phloem in a leaf found on the floodplain (b (i)), and in a leaf that had been in the

river for 15 days (b (ii)). Leaf epidermis begins to break down after 60 days in the wetland (c (i)), and in the

river (c (ii). Fungal reproductive structures (R) can also be observed in a (i), b(i), and c(ii). Images are taken at

400x magnification using a compound microscope. Sections in a (i), b (i), c (i) and c(ii) are stained with PAS.

Sections in a (ii) and b (ii) are stained with gentian violet.

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Figure 7.9: Contour map images of absorbance in amide bands I, II and III.

Contour images are shown for a fresh leaf (60*225 µm, epidermis at base) (i), a leaf found on the floodplain

(60*120 µm, epidermis at base) (ii), a leaf that decomposed in the river for 15 days (60*150 µm, epidermis at

base) (iii) and a leaf that decomposed in the river for 107 days (60*120 µm, epidermis at top). Absorbance is

mapped for the amide I band (a), the amide II band (b), and the amide III (c). Each section extends from the

epidermis (E) through collenchyma (C) (i) or fungal hyphae (H) (ii-iv) to fibre sheath cells (FS) (absent in iv).

Phloem cells (P) (i) or more fungal tissues (H) then separate the fibre sheath cells from xylem (X) tissues.

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Figure 7.10: Fungal reproductive structure with contours indicating the distribution of absorbance

from amide bands and aromatics. This feature was located in the section of the leaf that had been

206

submerged in the river for 15 days (240*220 µm). Contour maps of absorbance in amide I (a), amide

II (b), amide III (c) and aromatic bands (d) are shown.

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7.3.3 The chemical and structural components of leaves partially decomposed

under aquatic conditions

After 2 days in the water the distribution of fungal tissue in aged leaves continued to be

centred on the leaf vascular traces, with hyphae extending into the remaining mesophyll.

The image seen in Figure 7.8, b (i) shows a vascular trace from the terrestrially aged leaf

section cut at an oblique angle. Fungal tissues (H) are located between xylem cells and

the fibre sheath, and these hyphae appear to be extending longitudinally along the trace.

Figure 7.8, b (ii) shows a longitudinal aspect of a vascular trace of the leaf that had

decomposed in the river for 15 days. Magenta coloured fungal hyphae (H) are seen

extending within the vascular trace between the striated xylem and the purple fibre sheath

cells. This pattern of colonisation suggests that fungal tissues have invaded the vascular

trace and are extending along them both internally and externally. At the same time,

hyphae extend into the mesophyll, breaking down the tissues and forming reproductive

structures (R) (Figure 7.8, a (i), b (i) & c (ii)).

Absorbance in the amide II band was poorly resolved in FPA-FTIR spectra. However, S-

FTIR spectral maps of leaves decomposing under aquatic conditions showed that

absorbance in this band was widespread over the leaf sections, similar to the distribution

of amide I absorbance. In the early stages of aquatic decay in the river (Figure 7.9 b (iii))

higher absorbance in the amide II band was correlated with fungal tissues. In the later

stages of decay there was no differentiation between fungal and leaf tissues (Figure 7.9, b

(iv)).

Absorbance in the amide III band is at times associated with fungal tissues, and at other

times associated with phloem fibresa or xylem. In leaves that had decayed in the river for

15 days high absorbance in the amide III band correlated with fungal tissues (Figure 7.9,

c (iii)). After 107 days in the river, areas of stronger absorbance were located within the

xylem and were not associated with fungal tissues (Figure 7.9, c (iv)). Leaves that had

decayed for 2-30 in the wetland showed higher amide III absorbance associated with the

fibre sheath and fungal tissues (data not shown).

After 15 days under aquatic conditions in the wetland (Figure 7.11, b) leaves exhibited a

loss of structural integrity, with a tendency to develop fissures on the periphery of the

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xylem. Most of the non-vascular tissue, except for the epidermal cells, had disappeared.

After 30 days of aquatic decomposition (Figure 7.11, c) xylem tissues tended to separate

from other tissues upon sectioning and mesophyll was absent. After 60 days there was

evidence of breakdown of the epidermal cell layer, and fungal attack of lignified tissues.

This trend continued until 107 days when the leaf consisted of the remnants of vascular

tissues and epidermal cells (Figure 7.11, e).

Leaves taken from aquatic conditions in the wetland had lower absorbance in the amide I

band than fresh or terrestrially aged leaves. However, this lower absorbance was more

widely distributed and extended over the entire mid-vein (Figure 7.12, a-e (i)). Sections

of leaves taken from the river had lower absorbance in this band than did fresh leaves,

but higher absorbance than floodplain aged leaves (Figure 7.13, a-e (i)). S-FTIR shows

areas of higher absorbance in river samples correlated spatially with fungal tissues

(Figure 7.9 a, (iii) & (iv)).

With decomposition, whether in the wetland or the river, the absorbance due to aromatics

became more diffuse, distributed over the area of the xylem but not tightly confined to

cell walls. FPA-FTIR images of aquatically decomposing leaves indicate similar patterns

of absorbance due to lignin and polysaccharides to those seen in the terrestrially aged leaf

(Figure 7.12 and Figure 7.13, a-e (ii), (iii) and (v)). Absorbance due to the presence of

aldehydes indicates that lignin was being decayed enzymatically during aquatic

decomposition (Figure 7.12, a-e, (iv) and Figure 7.13, a-e, (iv)). Temporal trends in this

process differed between the river and wetland (see below).

Polysaccharides were associated with leaf vascular tissues and fungal tissues in leaves

decaying under aquatic conditions (Figure 7.12, a-e, (v) and Figure 7.13, a-e, (v)). High

absorbance in the polysaccharide band correlated spatially with xylem and phloem fibres

in samples decomposing underwater for 2, 15 and 30 days. In later samples from the

wetland the phloem fibres were absent and absorbance in this band was lower. In later

samples taken from the river absorbance in this band was higher than in earlier stages of

aquatic decay. Some phloem fibres were present and absorbing is this band in the 60 days

river sample, and in addition to xylem, fungal tissues had high absorbance in this band in

the 107 day river sample.

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Carbonyl vibrations from lipids, waxes and oils are represented spectroscopically by

absorbance in the ester carbonyl band at 1740-1730 cm-1

. Leaf samples were saturated

with paraffin as part of the sample preparation process, but paraffin does not contain ester

groups. It is therefore unlikely that absorbance in this band is due to residual paraffin.

FPA-FTIR images in this wave band (not shown) revealed that lipids persisted in the leaf

cuticle and epidermis for at least 60 days after entering the aquatic environment.

Absorbance in this band was also observed within vascular tissues and associated with

fungal material. As lignin is also known to absorb in these frequencies, absorbance

associated with xylem was probably due to the presence of lignin.

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Figure 7.11: Compound microscope images of sections of terrestrially aged leaves that have partially

decomposed in a wetland. Images are shown at 200x magnification and stained with PAS. After 2

days in the wetland (a) fungal hyphae (H) can be observed around and within the vascular bundle

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and throughout the remaining mesophyll (M) tissue. After 15 days (b) fungal distribution is similar

to day 2, but fissures are developing in the mid-vein. After 30 days (c), 60 days (d), and 107 days (e)

leaf disintegration was continuing, and the fibre sheath had decayed. Xylem (X) and the epidermis

(E) remained intact.

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Figure 7.12: Broad scale images of the leaf mid-vein, acquired using the Focal Plane Array FT-IR micro-spectrometer

(x15 magnification). Here are shown four micrometer sections through floodplain aged E. camaldulensis leaves that

have been submerged in a floodplain wetland for (a) 2; (b) 15; (c) 30; (d) 60; and (e) 107 days. Visible light images of

the unstained leaf section are shown in the left-most column, while images showing the spatial distribution of

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absorbance in the following bands are shown in subsequent columns: (i) 1700-1637 cm-1; (ii) 1530-1486 cm-1; (iii)

1284-1185 cm-1; (iv) 1729-1713 cm-1; and (v) 1180-900 cm-1. Lightest pink indicates highest absorbance while

darkest blue indicates lowest absorbance.

Figure 7.13: FPA-FTIR images of the leaf midvein from leaves submerged in the Kiewa River.

Shown here are four micrometer sections through floodplain aged E. camaldulensis leaves that were submerged

in a floodplain wetland for (a) 2; (b) 15; (c) 30; (d) 60 and (f) 107 days. Visible light images of the unstained leaf

214

section are shown in the left-most column, while images showing the spatial distribution of absorbance in the

following bands are shown in subsequent columns: (i) 1700-1637 cm-1; (ii) 1530-1486 cm-1; (iii) 1284-1185 cm-1;

(iv) 1729-1713 cm-1; and (v) 1180-900 cm-1. Lightest pink indicates highest absorbance while darkest blue

indicates lowest absorbance.

215

Figure 7.14: Compound microscope images of sections of terrestrially aged leaves that have partially decomposed in a

river. Images are shown at 200x magnification and stained with PAS. After 2 days in the river (a) fungal hyphae can be

observed around and within the vascular bundle and throughout the mesophyll, some mesophyll tissue is still present.

After 15 days (b) fungal distribution is similar to day 2. Loss of integrity in is greater than in the wetland, possibly due

to sand blasting effects from suspended river sediments. Sand grains in the samples cause sections to disintegrate as the

microtome makes its cut. After 30 days (c), 60 days (d), and 107 days (e) leaf disintegration is continuing, and the fibre

sheath has decayed. Xylem and the epidermis remain intact and fungal reproductive structures become abundant.

216

7.3.4 Chemical and structural differences between leaves decomposing under

river and wetland conditions

Samples that had been submerged in the river showed a similar degree of fungal

colonisation to those submerged in the wetland, and fungal hyphae were distributed

throughout similar plant tissues (Figure 7.14). However, breakdown of the fibre sheath

cells was evident after 30 days in the river (Figure 7.14, c) but was not observed until

after 60 days in the wetland (Figure 7.11, d). A large fungal reproductive structure can be

seen on the lower side of the midvein in the leaf section from 107 days in the river

(Figure 7.14, e). Differences were observed in the structure of fungal hyphae and

reproductive structures between river and wetland samples, but no attempt was made to

identify or quantify the species present.

The magnitude of absorbance in the aromatic band (1503 cm-1

) (Figure 7.12, a-e, (ii)) did

not change appreciably over time for leaves decaying in the wetland, while leaves

decaying in the river showed increasing absorbance from lignin over time (Figure 7.13,

(ii)). Absorbance in the ether linkage band (1231 cm-1

), was associated with the xylem

and fibre sheath in submerged leaves. In the wetland (Figure 7.12, (iii)) absorbance was

high over the first 30 days, particularly at the margins of the xylem, and then declined. In

the river (Figure 7.13, (iii)) absorbance was moderate at day 2, declined at day 15, and

then continued to increase over the remainder of the experimental period. The areas of

highest absorbance were found at the margins of the xylem, where tissues were more

dense.

Absorbance due to the presence of aldehydes indicated that lignin was being decayed

enzymatically during the early stages of aquatic decomposition (up to day 30) in the

wetland (Figure 7.12, c-d (iv)). In the river, activity declined from day 2 to day 15

(Figure 7.13, b & c (iv)), then continued to increase reaching a maximum at 60 days

(Figure 7.13, d (iv)). This was consistent with changes in absorbance due to lignin. High

absorbance due to aldehydes can also be seen in the large fungal reproductive structure in

the lower portion of the river image for 107 days (Figure 7.13, e (iv)). Figure 7.7, c (iv)

shows absorbance due to aldehydes in a sample submerged for 2 days in the wetland

217

mapped using S-FTIR. This image shows that aldehydes were distributed around the

periphery of the fungal hyphae in the aquatically decomposing leaf.

7.3.5 Chemical gradients observed in decomposing leaf material using S-FTIR

The area around hyphae in the S-FTIR spectral map seen in Figure 7.7 (c) was examined

more closely in order to detect chemical changes due to the activity of fungal extra-

cellular enzymes. Eight spectra were extracted from a segment of the 2 day aquatically

decomposed sample that extended from the hyphal tissue (0 µm) into the xylem (19 µm).

These spectra are shown in Figure 7.15. These data indicate the concentration of

polysaccharides, ether linkages, aldehydes & ketones and lipids changes over a distance

of 19 µm in a way that is consistent with enzymatic break down of the leaf components.

Polysaccharide concentrations were lowest close to the hypha and increased with

increasing distance from fungal hyphae. Specifically, peaks at 1161, 1060 and 1033cm-1

are indicative of cellulose (Hori and Sugiyama 2003) and the height of these peaks

increased with distance from the hypha. This is pattern is consistent cellulose breakdown

by extracellular enzymes that cleave β-1,4-glycosidic linkages between pyranose rings.

Lignin is composed of phenyl rings connected by ether (and other) linkages. The band at

1530-1486 cm-1

, indicating the presence of phenyl rings, was not well resolved in spectra

correlating with fungal tissues, but resolution improved with increasing distance into

xylem tissues (blue spectra). Relative absorbance in this band decreased markedly

between 8 and 15 µm from the hypha, possibly indicating a transition between altered

and unaltered xylem. The ether linkage band (1284-1185 cm-1

) showed decreasing

absorbance over 10-15 µm extending away from the hypha, and increasing absorbance

over the remaining distance. Moreover, there was a shift in the peak maxima to higher

frequencies with increasing distance, indicating a change in the conformation of this

bond type, possibly due to more extensive hydrogen bonding.

The amide I and II bands (1700-1637 and 1543-1527 cm-1

), indicating protein, were

present over the first 1-6µm, although the peaks were poorly resolved. Over the latter 6-

19 µm, these peaks were absent, and a peak at 1590 cm-1

was observed. The peak at 1590

cm-1

is consistent with C=C stretching is aromatic rings from lignin (Michell 1992).

218

The carbonyl peak at 1740-1730 cm-1

was absent over the first 4-9 µm, corresponding

spatially with the hypha, but became well resolved in the latter 8-19 µm. This band is

primarily associated with fatty acids and esters but is also associated with lignin (Heraud,

Caine et al. 2007). The aldehyde and ketone peak (1729-1713 cm-1

) was only clearly

resolved in the 8-13 µm spectrum, which was measured from the interface between the

hypha and the xylem. This is an indication of lignin degradation in this narrow zone.

Figure 7.15: Spectra extracted from a segment of the S-FTIR map of the 2 day aquatically decomposed sample, shown

here at the left, with z-values representing absorbance in the amide I band. Each spectrum represents a pixel with an

area of 5 x 5 µm, which overlaps the adjacent pixel by 3 µm, to give a total transect length of 19 µm. Spectra are

presented in a colour similar to that shown in the S-FTIR map. Pink - orange spectra are derived from the fungal tissue,

blue spectra are derived from xylem and the green spectra are derived from the interface between these two tissue

types. Chemical gradients radiating away from fungal tissues can be observed for polysaccharides and lignin. Protein is

evident as amide peaks 1 and 2 in the first 2 spectra (pink and red). Carbonyl peaks indicating aldehydes & ketones and

lipids (and lignin) are also illustrated.

0.0E+00

2.0E-04

4.0E-04

6.0E-04

8.0E-04

1.0E-03

1.2E-03

1.4E-03

1.6E-03

1,8

00

1,7

68

1,7

36

1,7

04

1,6

71

1,6

39

1,6

07

1,5

75

1,5

43

1,5

11

1,4

79

1,4

46

1,4

14

1,3

82

1,3

50

1,3

18

1,2

86

1,2

54

1,2

21

1,1

89

1,1

57

1,1

25

1,0

93

1,0

61

1,0

29

99

6

96

4

93

2

90

0

Infra-red Frequency (cm-1)

Infr

a-re

d A

bso

rban

ce (

AU

)

0-5 µm 2-7 µm 4-9 µm 6-11 µm 8-13 µm 10-15 µm 12-17 µm 14-19 µm

Pro

tein

Lip

ids

Lig

nin

Lig

nin

Ald

eh

ydes

and

Ket

one

s

Polysaccharides

8-13µm

0-5µm

2-7µm

4-9µm

6-11µm

10-15µm

12-17µm

14-19µm

219

7.3.6 Differentiating between different protein structures using FT-IR

The second derivative of the amide I peak can be used to reveal structural features of

proteins (Byler and Susi 1986). In plant material the amide I peak has its apex at 1650

cm-1

(the α-helix position) (Heraud, Caine et al. 2007). In fungal tissues, the apex of this

peak is typically at 1648-1626 cm-1

(Szeghalmi et al., 2006) depending on the species.

The results shown in Figure 7.16 indicate that protein structures differed between the

fresh and partially decomposed leaves. In the fresh leaf, maxima were seen at 1671,

1665, 1655, 1645 and 1634 cm-1

, while in a leaf decayed in the water for 2 days maxima

were seen at 1671-1668, 1665, 1660, 1657, 1652-1650, 1644, 1641, 1637 and 1628 cm-1

.

Peaks from the spectra of partially decayed leaves had greater variation between

replicates than spectra from fresh leaves. PCA analysis of the derivatised spectra

indicated that amide I spectra from fresh leaves were substantially different from spectra

taken from partially decayed leaves (Figure 7.17). The loadings for each principal

component are shown in Table 7.2. This table indicates that the first three principal

components differ in terms of the absorbance frequency of the major protein structures.

Figure 7.16: The second derivative of three replicate spectra from fresh leaves, and three spectra

from leaves submerged in the wetland for 2 days. Only wave numbers corresponding to the amide I

peak are shown. Peak maxima represent different structural features of proteins such as extended

-1.20E-06

-1.00E-06

-8.00E-07

-6.00E-07

-4.00E-07

-2.00E-07

0.00E+00

2.00E-07

4.00E-07

6.00E-07

16

73

16

71

16

70

16

69

16

68

16

66

16

65

16

64

16

62

16

61

16

60

16

59

16

57

16

56

16

55

16

53

16

52

16

51

16

50

16

48

16

47

16

46

16

44

16

43

16

42

16

41

16

39

16

38

16

37

16

35

16

34

16

33

16

32

16

30

16

29

16

28

16

26

16

25

16

24

IR Frequency (cm-1)

Seco

nd

De

riva

tive

of

FT-I

R

Ab

sorb

ance

Fresh(a) Fresh(b) Fresh(c) 2 Day(a) 2 Day(b) 2 Day(c)

220

chains (low components) at 1624-1637 cm-1

, α-helices at 1654 cm-1

, unstructured components at 1645

cm-1

, and turns and bends at 1663-1694 cm-1

. Peak broadening and shifts to higher frequencies

indicate weaker hydrogen bonding in the molecule.

Figure 7.17: PCR analysis of extracted spectra. Spectra extracted from regions of the S-FTIR maps

that had strong absorbances in the amide I band were normalised and smoothed. The second

derivative of the absorbance within the amide I band was then taken, as this region is indicative of

protein structure. Principle components analysis indicated significant differences in protein

structure between the two sample groups.

Table 7.2: The 2nd

derivative peaks that gave positive loadings for principal components 1, 2 and 3.

The protein structure associated with these peaks, as reported by Byler and Susi (1986), is shown in

the column labelled “Protein structure”.

Frequency (cm-1

) Protein structure PC1 PC2 PC3

1623 low chain a 1623

1627 low chain a 1627

1629 low chain b 1629

1636 low chain c 1636

1637 low chain c 1637

1641 unordered 1641

1644 unordered 1644

1647 unordered 1647

1650 1650

1652 alpha-helices 1652

-2.00E-06

-1.50E-06

-1.00E-06

-5.00E-07

0.00E+00

5.00E-07

1.00E-06

1.50E-06

2.00E-06

2.50E-06

3.00E-06

-2.50E-

06

-2.00E-

06

-1.50E-

06

-1.00E-

06

-5.00E-

07

0.00E+0

0

5.00E-07 1.00E-06 1.50E-06

PC2

PC 1

Fresh Leaf

2 days Submerged

221

1653 alpha-helices 1653

1658 bend a 1658

1662 bend a 1662

1666 (aromatic aldehyde) 1666

1670 bend b 1670

1675 high chain 1675

7.3.7 Evidence for the formation of protein complexes

In the fresh and leached leaf samples there were no areas where amide I absorbance was

spatially correlated with absorbance due to aromatic rings and ether linkages (Figure 7.5,

a-c, (i), (ii) and (iii)). In the floodplain aged leaf, absorbance in these bands was

associated over the entire area of mid-vein xylem (Figure 7.5, d (i), (ii) and (iii)). In the

river samples there was initially a weak association between the spatial distribution of

amide I and aromatic absorbance, and but the strength of the association increased with

decomposition time (Figure 7.13, a-e (i) & (ii)). In the wetland samples these bands had a

strong spatial association throughout the aquatic decay period, even as amide I

absorbance decreased over time (Figure 7.12, a-e (i) & (ii)).

In S-FTIR images, absorbance in the amide I band was spatially correlated with fungal

hyphae (Figure 7.18, a, b, c and d (ii)) (although spatial resolution was not adequate to

discriminate all parts the hyphae from the background in (Figure 7.18, b (ii)), while

absorbance in the amide III band coincided spatially with fibre sheath cells in Figure

7.18, a & d (iii), but with fungal tissues in Figure 7.18, c (iii). Absorbance in the amide

III band coinciding with fibre sheath cells may be due to the presence of inorganic and

organic nitrates, nitramines, and conjugated nitro compounds which absorb at similar

frequencies to the N-H bonds from proteins normally associated with this wave band (see

Table 7.1). Low absorbance due to aromatics from these tissues (Figure 7.18, a & d (iv))

suggests these nitrogen compounds are not aromatic.

S-FTIR images show that high absorbance in the aromatic band was correlated with

xylem (Figure 7.18, a (iv)), immediately adjacent to an area of moderate absorbance in

the amide I band. A more detailed examination of a hypha (Figure 7.18, b (ii) & (iv)

again indicated that protein associated with the hypha was concentrated in the area

immediately adjacent to aromatic compounds. A similar pattern was observed in leaf

222

material that had been submerged in the river for 2 days (Figure 7.18, c & d, (ii) & (iv)).

In Figure 7.18, d (ii) only a small amount of protein was evident within the hypha while

adjacent areas of the hypha absorbed strongly in the aromatic band. Aromatics also

exhibited moderate absorbance in (lignified) fibre sheath cells.

223

Figure 7.18: Fine scale relationships between protein and lignin distribution

This figure shows a segment of the midvein (60*120 µm) (a) and a map focused on fungal tissue (26*26 µm) (b)

taken from the floodplain aged leaf. Similar maps are also shown for the leaf that was submerged in the Kiewa

224

River for 2 days ((c) (60*225 µm) and (d) (44*36 µm) respectively). Images shown include the bright field image

of the sample (i) indicating the area in which the FTIR map was acquired, and S-FTIR maps for amide I (ii),

amide III (iii) and aromatic bands (iv).

225

7.4 Discussion

7.4.1 Synchrotron infrared micro-spectroscopy

By combining Fourier transform infrared spectroscopy with high resolution microscopy,

fungal organisms within a substrate can be chemically characterized in a spatially explicit

context (Garidel and Boese 2007). However, the diameter of fungal hyphae is similar to

the wavelength of infra-red light (2.5-12.5 µm), and the mass of material to be analysed

is very low, and this can result in light scattering and weak signals (Kaminskyj, Jilkine et

al. 2008). The use of the synchrotron light source addresses these issues (Carr 2001;

Garidel and Boese 2007; Miller and Dumas 2006; Reffner, Martoglio et al. 1995),

making it possible for the biochemical components of individual hyphae to be identified

(Jilkine, Gough et al. 2008; Szeghalmi, Kaminskyj et al. 2007). FTIR microscopy has

been used to differentiate between fungal and bacterial infections in animals

(Erukhimovitch, Pavlov et al. 2005) and to detect phyto-pathogens (Erukhimovitch,

Tsror et al. 2005). In pure cultures, bacteria and fungi, and true fungi and Stramenopiles

can be distinguished spectrally (Erukhimovitch, Tsror et al. 2005). However, there have

been no reports where spectra from these organisms were able to be distinguished in a

complex natural matrix such as a decomposing leaf.

7.4.2 Fungal colonisation of E. camaldulensis leaves

Microscopic examination of leaf sections using a conventional bright field microscope

has shown that E. camaldulensis leaves are colonised by fungi while on the floodplain.

At this time, fungi attack both the mesophyll and the leaf vascular tissue, where they

specifically target phloem cells. Fungal hyphae probably invade minor veins at the leaf’s

periphery, where the fibre sheath is sparse or absent, and begin attacking phloem at that

point. Having penetrated the vascular network, hyphae appear to extend along the interior

of the vascular bundle and consume the phloem as they progress. At the same time,

fungal hyphae attack vascular tissues from the mesophyll, encircling the vascular bundle

and attacking cells in the bundle sheath and collenchyma.

On entering the aquatic phase, the aged leaf consists mainly of vascular tissues

sandwiched between the epidermal layers. The fibre sheath and xylem are attacked at this

time. The decay of fibre sheath cells proceeds more quickly than xylem degradation.

Xylem is attacked from its periphery, with outer cells being degraded while inner cells

remain unaltered. Xylem showed no significant structural changes prior to 60 days in the

226

wetland and 107 days in the river. The leaf epidermis also remained unaltered for at least

60 days after entering the water. It is likely that lipids present in the epidermal tissues

until that time prevented microbial attack of these cells.

7.4.3 Aquatic break down of lignin in E. camaldulensis leaves

Chemical maps obtained from FPA-FTIR and S-FTIR have shown that polysaccharides

are lost from the leaf mesophyll during the terrestrial phase as a result of leaching. This is

consistent with previous studies where chemical changes were measured in decomposing

leaves (Gallardo and Merino 1992; Serrano 1992; Suberkropp, Godshalk et al. 1976).

Light microscopy showed that fungal tissues were present throughout the mesophyll

tissue after terrestrial aging, while the tissues of the xylem, fibre sheath and epidermis are

largely intact but surrounded by fungal hyphae. This shows that non-lignified plant

tissues are consumed preferentially.

On entering the aquatic phase, the majority of polysaccharides available to micro-

organisms are those within the leaf vascular tissues, and these are extensively cross-

linked to lignin. Polysaccharides continue to be lost from the mesophyll, and

polysaccharide signals from this region of the leaf were increasingly associated with

fungal cell walls. Jilkine et al. (2008) found a strong peak at 990 cm-1

is associated with

trehalose in fungal cell walls, but this peak was not well resolved in the data from this

work.

High absorbance in the polysaccharide band from leaf vascular bundles suggests that

these tissues were rich in cellulose, and hemicellulose, while contributions from starches,

pectins and nucleic acids could not be ruled out (Heraud et al, 2007). The discrete wave

bands for these individual polysaccharides were mapped over the tissues, but the spatial

distribution of absorbance for each separate polysaccharide matched that of

polysaccharides generally, with minor differences in absorptivity.

As the majority of polysaccharides remaining in a leaf when it enters the aquatic phase

are cross-linked to lignin, lignin break down is a necessary first step before aquatic

organisms can obtain energy. While mineralisation of lignin is possible in aquatic

systems (Bergbauer, Moran et al. 1992; Moran, Benner et al. 1989), it is limited by

oxygen availability (Freeman, Ostle et al. 2004) and proceeds very slowly under anoxic

227

conditions (Benner, Maccubbin et al. 1984). Floodplain wetlands commonly suffer from

low dissolved oxygen (Baldwin and Mitchell 2000), particularly in warm weather (Ford,

Boon et al. 2002), and wetland sediments are generally anoxic (Baldwin and Mitchell

2000). Thus, it would be expected that lignin breakdown would proceed more slowly in

the wetland than in the river, or fail to proceed at all.

Both microscopic and FTIR data show that there is some degradation of lignified tissues

in the aquatic phase, and that degradation follows different temporal patterns in rivers

and wetlands. The spatial distribution of aldehydes, as illustrated in FPA-FTIR images,

indicates that this activity is highest in the floodplain aged leaf, but continues after

immersion in the wetland then declines to low levels after 60 days. This may be a

response to decreasing concentrations of dissolved oxygen as temperatures increased, or

depletion of exploitable substrate. The distribution of aldehydes in leaf samples

submerged in the river shows that lignin degradation increases over time and continues to

be active even after 107 days in the aquatic environment. While subject to the same

temperature regime as the wetland, flow in the river continually incorporates oxygen into

the water column, making it less likely to be limited by dissolved oxygen. It is clear that

enzyme mediated lignin breakdown is occurring in the submerged leaf detritus, but

fungal decay of lignified leaf material begins earlier in wetlands than in rivers.

The fungal enzymes responsible for the breakdown of lignin include laccase (phenol

oxidase) and lignin peroxidase. Laccase oxidizes phenolic moieties within the lignin

structure to firstly form phenolic radicals, then quinones. This reaction may proceed via a

number of different pathways, but generally C-C and C-O bonds are cleaved in side

chains, aromatic rings (Kawai et al, 1988), and between monomers (Harkin and Obst,

1974). Some organisms may only demethylate lignin side chains (Leonowicz et al,

2001). Lignin peroxidase requires the presence of hydrogen peroxide (generated by other

enzymes) for its activity. It catalyses a variety of C-C and C-O bond cleaving reactions,

including the oxidation of the alkyl side chains of lignin, cleavage of C-C bonds in the

side-chains of monomers, oxidation of veratryl alcohol (3,4-dimethoxybenzyl alcohol)

monomers to the related aldehydes and ketones, and hydroxylation of benzylic methylene

groups (Tien and Kirk 1984). The products of lignin peroxidase degradation of natural

lignin provide a substrate for laccase, and these enzymes commonly co-occur

(Leonowicz et al, 2001).

228

The end products of lignin breakdown reactions are largely ketones, acids and aldehydes,

including coniferyl aldehyde and sinapyl aldehyde. Vibrations from the C=O bond in

these compounds may absorb at frequencies within the amide I band. For example, the

carbonyl group in coniferyl aldehyde and sinapyl aldehyde absorb at 1661 cm-1

and 1655

cm-1

, respectively (Pomar et al, 2002). These monomers are extremely unstable and may

re-polymerise (Leonowicz et al, 1985), particularly in the presence of lignin peroxidase

(Sarkanen et al, 1991).

Earlier studies of leaves and wood decomposing in aquatic environments have revealed

increases in the absolute lignin content over the first 6-12 weeks of decomposition, with

corresponding increases in nitrogen and protein contents (Suberkropp, Godshalk et al.

1976; Weiland and Guyonnet 2003). The re-polymerisation of lignin monomers provides

a possible mechanism for the generation of “new” lignin. These new polymers may then

be deposited on pre-existing lignin. This would account for minimal changes in overall

spatial distribution of absorption in the aromatic band, while discrete correlation with a

rigid cell wall decreases with time. However it would not account for increases in the

mass of lignin in decomposing leaves.

Complexation of lignin breakdown products with enzyme proteins is a possible

mechanism for perceived increases in the absolute lignin content of decaying leaves. The

most probable location that these complexes would form is in areas where enzymes are

being secreted, such as the periphery of the xylem, where absorbance in the amide I band

is often most intense.

A well resolved peak at 1510-1515 cm-1

(aromatic skeletal vibrations) is a common

feature of the spectra of lignin from a variety of sources (Boeriu, Bravo et al. 2004;

Gosselink, Abächerli et al. 2004). The height of this peak (absorbance) in spectra from

xylem tissues decreased with distance from the hyphae. A significant reduction in the

intensity of this band between 8 and 10 µm from the hyphae is consistent with enzymatic

depolymerisation of lignins (Buta, Zadrazil et al. 1989). This band is poorly resolved in

spectra from fungal tissues, and this may be due to the presence of non-lignin aromatic

compounds such as those found in fungal melanins (Saiz-Jimenez, Ortega-Calvo et al.

1995). Changes in intensity and frequency of the band at 1284-1185 cm-1

(ether linkages

229

in lignin) indicates reduced absorbance due to νs(C-O-C) and increasing absorbance due

to νs(C-OH) (Michell 1992). This is consistent with lignin breakdown via enzymatic

cleavage of the linkages between monomers, such as occurs with the action of fungal

laccases and peroxidises (Leonowicz, Cho et al. 2001).

7.4.4 The formation of humic substances

Low level absorbance in the amide I band is likely to be due to either microbial extra-

cellular enzymes, or aromatic products of lignin break down (such as aromatic aldehydes

absorbing in the region 1710 - 1685 cm-1

(Silverstein, Bassler et al. 1974)). Earlier

studies (Bärlocher, Tibbo et al. 1989; Suberkropp, Godshalk et al. 1976) have shown that

fungal extracellular enzymes form complexes with plant derived tannins and

polyphenols. If this is occurring in submerged E. camaldulensis leaves, we should

observe increased spatial correlation between absorbance in the aromatic and amide

bands over time. As tannins and polyphenols are abundant in mature E. camaldulensis

leaves (Cork and Pahl 1996), and extra-cellular enzyme activity has been shown to occur

(see Chapter 4), it is probable that protein-soluble polyphenol complexes will form.

However, a large proportion of soluble polyphenols are lost via leaching during the

terrestrial phase (Serrano 1992), so concentrations of these complexes may be low. FPA-

FTIR and S-FTIR data suggest that such complexes may form in narrow zones at the

periphery of lignified plant structures.

Low level absorbance in the amide I band associated with lignin in wetland leaves, and

the Day 107 river leaf, may alternatively be due to carbonyl vibrations from lignin

decomposition products, such as quinones, other aromatic aldehydes and aromatic

ketones, which absorb at similar frequencies (Silverstein, Bassler et al. 1974). S-FTIR

shows that while amide peaks are present in spectra from fungal tissues, they are poorly

resolved in the region between hyphae and xylem (the region where enzymes are likely

to be active), and absent at distances of greater than 10 µm from the hypha. This suggests

that the distribution of protein-polyphenol complexes and lignin decomposition by-

products is patchy at small spatial scales.

The data show that amide III absorbance (1328-1290 cm-1

) correlates spatially with

amide I and II in the fresh leaf, a strong indication of protein. In decomposed leaves

amide III absorbance is associated with the fibre sheath and fungal tissues for the first

230

month. In the later stages of decay, this pattern continued in the river, while in the

wetland amide III absorbance became associated with xylem. Rice (1982) suggests that

nitrogen is associated with heterocyclic aromatic humic compounds in the later stages of

decay. It is likely that these humic compounds would include aromatic amines, which

show a strong C-N stretch absorption in 1342-1266 cm-1

(Silverstein, Bassler et al.

1974). Amide III absorbance associated with the fibre sheath is probably associated with

the production of aromatic amines as a result of fungal activity, but it is also consistent

with C-N ring vibrations of melanin in fungal cell walls (Saiz-Jimenez, Ortega-Calvo et

al. 1995). At the periphery of the xylem it is plausible that absorbance in this band is

associated with aromatic amines or protein complexes. However, where absorbance

peaks in amide I and II bands are absent, C-N bonds indicated by absorbance in the

amide III band are unlikely to be from protein.

The FPA-FTIR data show a weak yet constant spatial correlation between absorbance in

the aromatic band and absorbance in the amide I band in wetland leaf samples. In

samples from the river, high absorbances in these bands do not coincide spatially until 60

days, when rates of lignin degradation were higher. While this is consistent with the

formation of complexes between proteins and leaf polyphenols as proposed by

Suberkropp et al.(1976), a more controlled experiment with longer incubations times

would be desirable before drawing conclusions.

In both the wetland and the river, areas of highest absorbance in the polysaccharide band

were spatially correlated with lignified plant tissues, as indicated by absorbance in

aromatic and ether linkage bands. In the wetland, polysaccharide absorbance in these

regions decreased over time, while in the river, absorbance decreased between day 2 and

day 15, then increased for the remainder of the experimental period. This correlates with

patterns of lignin decay as revealed by absorbance from aldehydes, and suggests that C-

O-C bonds in polysaccharides are becoming disencumbered from crossed-linked lignin,

enabling greater vibration. For this reason, higher absorbance in the polysaccharide band

in observed in the river samples after lignin degradation has been initiated. We know

from chemical analysis of decomposing oak and hickory leaves in a stream that losses of

cellulose and hemicellulose closely reflect total weight loss, and that the rate of their loss

remains relatively constant (Suberkropp, Godshalk et al. 1976). We can therefore

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conclude that aquatic fungal communities are limited in their capacity to degrade and

consume polysaccharides in leaf vascular tissues by their ability to first degrade lignin.

The trends observed in the polysaccharide absorbance with increasing distance from

fungal hyphae, i.e. lower absorbance and broader peaks closer to fungal tissue, are

similar to those seen in the fungal decay of wood (Weiland and Guyonnet 2003). Lower

absorbance is associated with the depolymerisation of compounds such as cellulose,

hemicellulose and pectin, and peak broadening has been associated with changes in

intermolecular hydrogen bonding (Hsu 1997). These results show that polysaccharides

are being depolymerised, and are consistent with spectroscopic observations reported by

Gierlinger et al.(2008).

7.4.5 Protein and nitrogen enrichment

Both fresh and decomposed leaves contain protein, as indicated by absorbance in the

amide I band. The protein content of the fresh leaf was spatially correlated with

mesophyll cells, while in partially decomposed leaves protein is spatially correlated with

fungal tissues. The second derivative of the amide I peak was compared between these

two sources using PCA, and showed that these proteins were structurally different.

Further to this, derivative spectra from the fresh leaf were similar to each other, while

spectra in decomposing leaves varied significantly spatially and over time. It is likely that

this difference reflects the presence of a single dominant protein present in the fresh

leaf’s palisade mesophyll cells (ribulose bis-phosphate carboxylase; Heraud et al, 2005),

compared with a variety of different proteins associated with different species of fungi

(Szeghalmi, Kaminskyj et al. 2007), and hyphae of different ages and functions (Jilkine,

Gough et al. 2008; Kaminskyj, Jilkine et al. 2008). Thus, FPA-FTIR and S-FTIR spectral

mapping has revealed that leaf protein, from the mesophyll and phloem, is lost when

leaves age on the floodplain, and is replaced by protein with a markedly different

structure.

As we know that submerged leaves exhibit increased protein and nitrogen content when

colonised by aquatic fungi, it has been proposed that the source of this additional protein

is fungal biomass. The fungal mycelium is composed primarily of polysaccharides (80-

90%, Bartnicki-Garcia, 1968), but the cell wall has an outer layer that is primarily protein

(Klis, Mol et al. 2002). We can therefore expect fungal tissue to absorb strongly in the

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amide bands. The data presented here show that absorbance in the amide I band within

the decomposing E. camaldulensis leaf is spatially correlated with fungal tissues after

incubation in either the river or the wetland. However, we can not assume that all

nitrogen within the decomposing leaf is in the form of protein (Odum, Kirk et al. 1979).

Throughout aquatic leaf decomposition, low level absorbance in the amide I band is

spatially correlated with absorbance in the aromatic band and lignified plant tissues,

while higher level absorbance is found adjacent to the margins of the lignin. This is

consistent with the formation of protein – polyphenol complexes which later adhere to

the lignified tissues. However, given that absorbance at frequencies within the amide I

band is also characteristic of lignin breakdown products such as quinones and aromatic

aldehydes, we can not conclude from these data that protein complexes have formed. If

indeed the amide I band is indicating the presence of protein complexes, they are in very

low concentrations. Therefore, these results neither confirm nor contradict assertions by

Melillo et al. (1984), Odum et al.(1979) and Rice (1982) that a proportion of the leaf

protein is comprised of abiotic protein complexes.

Previous studies using FT-IR have shown that the amide II band may be poorly resolved

in mature hyphae, which have less protein than new growth, contain large vacuoles

(Szeghalmi, Kaminskyj et al. 2007) and may have melanised cell walls (Kaminskyj,

Jilkine et al. 2008; Martin, Haider et al. 1974; Zhong, Frases et al. 2008). This poor

resolution may be due to the presence of non-proteinaceous compounds in the leaf matrix

that have NH2 or N-H bending vibrations. Such compounds might include chitin,

chitosan, and primary and secondary amides, which absorb at similar frequencies to the

amide III band. The amide II peak was poorly resolved in the decomposed leaf sections.

As many of the hyphae observed via light microscopy were darkly coloured, and

probably had some chitin in the cell wall, poor resolution might be expected in spectra

spatially correlated with fungal hyphae. In areas of the leaf where fungal tissues were not

present, such as the xylem, poor resolution was probably due to nitrogenous

decomposition by-products within the leaf matrix.

Absorption in the amide III band results from (C-H) and (C-N) stretch vibrations and (N-

H) bending vibrations. This suggests Absorbance in this band may provide an indication

of the distribution of organic nitrogen within a decomposing leaf. In line with Rice’s

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(1982) suggested mechanism for nitrogen accumulation in decomposing plant material,

absorbance in this band was correlated with fungal protein in the early stages of aquatic

decay, but that the strength of this correlation decreased with time.

The results show that the protein and nitrogen in E. camaldulensis leaves are largely

confined to fungal tissues during the early phase of aquatic decomposition. As

decomposition progresses nitrogen and protein are increasingly associated with lignified

plant tissues. This tends to support the notion that, as the energy of the leaf substrate is

depleted, protein is entrained within abiotic complexes, and nitrogen is incorporated into

non-proteinaceous organic compounds. Fungal growth and metabolism provide a

possible mechanism for these transformations. For example, nitrogen is incorporated into

chitin and melanins. As the fungal tissues senesce, these cell wall components may be

further degraded to structures similar to the refractory heterocyclic compounds typical of

humic material. This suggests that aquatic saprobic fungi biologically synthesise the

precursors to humic substances in aquatic environments.

7.4.6 The ecological significance of these findings

The physical environment in rivers and wetlands differs markedly at scales that are

relevant to aquatic fungi. Even though the fungal community colonising submerged E.

camaldulensis leaves does not differ between these environments, it is now evident that

these communities utilise their substrate differently.

The dried and senesced leaves of E. camaldulensis represent a poor food source for

aquatic fauna (Bunn, De Deckker et al. 1986; Canhoto, Bärlocher et al. 2002; Canhoto

and Graça 1999; Glazebrook and Robertson 1999), and while terrestrial aging removes

many substances that may deter feeding, it results in a food source that is composed of

polysaccharides and little else. Of the polysaccharides remaining, most are cross-linked

to lignin, which limits bio-availability. So when these leaves enter the aquatic phase, they

are colonised by fungi with the enzymatic capacity to degrade lignified plant cell walls

(Leonowicz et al. 2001). These fungi, together with the substrate, then comprise a more

nutritious food resource to higher trophic levels (Bärlocher and Kendrick 1975; Chung

and Suberkropp 2009).

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Lignin decay commences immediately after leaf immersion in wetlands, but substantial

decay of lignin may be delayed for up to 2 months in rivers. This may be because leaves

submerged in the river accumulate a surface biofilm (Golladay and Sinsabaugh 1991),

composed of fungi, bacteria, algae and other organisms (Burns and Ryder 2001a; Schulze

and Walker 1997). This biofilm may generate bio-available carbon (Haack and McFeters

1982) that can be utilised more readily than lignified plant tissues. Under these

circumstances the breakdown of lignin would not be necessary until carbon generated

within the biofilm became limiting. In wetland environments where biofilm does not

accumulate on the leaf surface, the decomposer community would be required to initiate

lignin degradation at an earlier stage. Further experimental work would be required to

test this hypothesis.

Fungal ligninases are instrumental in the formation and transformation of aquatic humic

substances (Claus and Filip 1998). The fungal degradation of lignin leads to the

formation of refractory organic compounds that are unavailable to higher trophic levels

and become stored in wetland sediments. Fungal metabolic processes are also responsible

for sequestering nitrogen from the water column and incorporating it into proteins, chitin

and aromatic compounds. While these compounds may undergo transformations in food

webs and by other decomposer organisms, they represent the precursors of aquatic humic

substances.

We know that fungi tend to dominate the early stages of aquatic leaf decomposition in

terms of biomass and carbon mineralisation (Baldy, Gessner et al. 1995). However, it is

certain that bacteria also contribute to this process, and more so in floodplain wetlands

than in rivers (Baldy, Chauvet et al. 2002). Bacteria become dominant on submerged

leaves when particle size becomes small, but many of these species may not be capable

of degrading plant structural polymers (Suberkropp and Klug 1976). Bacteria are also the

dominant decomposer organisms where dissolved oxygen is absent, such as in wetland

sediments (Boon and Mitchell 1995; Boon, Virtue et al. 1996). While this work focuses

on fungi, we know that bacteria have the potential to make significant contributions to

protein enrichment and the break down of polysaccharides and lignin in submerged

leaves. It is important to quantify this contribution in future work.

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7.5 Conclusions

We have seen in earlier chapters that the fungal community on submerged E.

camaldulensis leaves differs from that found on other litter types, and has the potential to

accumulate significant amounts of biomass in both rivers and wetlands. As these leaves

represent an abundant carbon resource in Australian floodplain environments, fungal

breakdown of structural polymers and protein enrichment is a vital process connecting

riparian plant production to higher trophic levels. This work demonstrates that E.

camaldulensis leaves submerged in wetlands exhibit lignin degradation earlier than those

in rivers, and that this may be due to more limited bio-available carbon in wetland

benthic habitats.

Infrared mapping of leaves colonised by micro-organisms provides hitherto unseen

spatially explicit insights into decomposition mechanisms, and supports earlier theories

regarding nitrogen, protein and lignin enrichment of submerged leaves. Chemical

mapping using Synchrotron and Focal Plane Array FTIR micro-spectroscopy has

demonstrated that some microbial protein and other nitrogenous compounds may become

complexed with polyphenols, or incorporated into heterocyclic aromatic compounds

which are not bio-available. For this reason, C:N ratios may over-estimate the protein

content of the leaf.

The high spatial resolution of this instrument has enabled us to detect the transformation

of structural carbohydrates in plant vascular tissues into bio-available sugars as a result

of fungal enzyme activity. Furthermore, it has provided evidence supporting the theory

that lignin breakdown products released via oxidative enzyme activity form stable and

refractory complexes within the decomposing leaf, and humic precursor compounds

which may be stored in wetland sediments.

Given the limited replication in this exploratory research, further work is now required to

quantify the importance of these processes. For example, it would be desirable to isolate

the fungal species responsible for aquatic lignin degradation on submerged E.

camaldulensis leaves, and re-introduce them to sterilised leaves. This would allow the

contribution of fungi to leaf break down to be assessed independently to that of bacteria.

Spatially explicit chemical analysis of such leaves using S-FTIR would benefit from

concurrent mass spectroscopy or NMR analysis, to track the chemical changes produced

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by fungal colonisation. This may enable us to characterise “new” lignin polymers.

Further investigation of protein enrichment, carbohydrate breakdown, leaf biofilm

dynamics and humic substance formation in E. camaldulensis leaves by fungi using 15

N

and 13

C signatures may also be of benefit.

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Chapter 8

General Discussion

8.1 Carbon Cycling in Australian Floodplain Wetlands

Our knowledge of the role of fungi in wetland carbon cycles has previously relied upon

research performed outside Australia (Shearer, Descals et al. 2007). This research has

established an understanding of fungal metabolism, explored taxonomic diversity and

phylogenetic linkages between fungal groups, and investigated the ecology of aquatic

fungi. While it is possible to apply this knowledge in Australian settings, the uniqueness

of this country’s vegetation and climatic conditions demands that fungal communities in

Australian floodplain wetland systems be investigated directly.

The exchange of carbon and nutrients between the river channel and the floodplain

(including its wetlands) is discussed in the Flood Pulse Concept (Junk, Bayley et al.

1989), as is the adaptation of floodplain biota to periodic inundation. Fungi contribute to

this exchange by incorporating the carbon from plant detritus, produced on the floodplain

and in floodplain wetlands, into their own biomass. The propagules released into the

water column are then distributed by the river channels and fungal mycelia and

propagules may then be consumed by stream biota. Fungal activity may also facilitate

leaching from submerged detritus. In arid systems, fungi are instrumental in releasing

nutrients from stored biomass on the floodplain for use by plants, and their activities are

synchronised with moisture availability. This has been described as the Pulse-Reserve

paradigm (Collins, Sinsabaugh et al. 2008) and the flow of carbon through the system as

the “fungal loop”. These concepts are similar in approach, but deal with different

magnitudes of moisture pulses, and different spatial scales of carbon movement. In the

absence of a flood event, the movement of carbon stored on floodplains in arid

environments is quite restricted.

Carbon dynamics in Australian floodplain wetland systems have been investigated on a

number of fronts. These include the degradation of sediment carbon by bacteria, the

break-down of submerged plant material, detrital food chains and pelagic food webs. An

understanding of fungal processes and functions within this floodplain wetland

ecosystem can now be integrated into this body of knowledge.

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8.1.1 Carbon cycling in the sediments

In ephemeral floodplain wetlands a thin layer of oxic sediments overlies anoxic

sediments. The boundary between these layers varies in both space and time as the

wetlands flood and dry (Boon 2006). Rapid shifts to anoxic conditions are common

(Boon 2000). Under these conditions, sediment carbon is mainly metabolised by

methanogenic bacteria (Boon 2000), releasing methane and carbon dioxide firstly into

the water column and then to the atmosphere, and the rate of methane emission increases

with temperature (Boon, Mitchell et al. 1997). Aerobic decay of sediment carbon is

considered to be a secondary source of carbon dioxide emissions.

In Chapter 5, it was shown that wetland sediments contain little fungal biomass,

supporting the findings of Boon et al. (1996). Fungi do not appear to utilise sediment

carbon, but lay dormant ready to utilise available substrates should conditions become

suitable. Under the observed conditions, the impact of sediment fungi upon wetland

carbon cycles is likely to be small.

8.1.2 The decomposition of submerged and standing dead plant material

While autochthonous productions in wetlands due to the growth of macrophytes and

phytoplankton can be very high (Bott, Barnes et al. 1983; Briggs and Maher 1985;

Roberts and Ganf 1986), significant amounts of organic material are also derived from

allochthonous sources such as riparian vegetation (Briggs and Maher 1985; Campbell,

James et al. 1992a). This production is occasionally consumed by grazers but more

commonly decays in situ, either in the standing dead position or after entering the aquatic

phase (Buesing and Gessner 2006; Kuehn and Suberkropp 1998a). Where the water

column is oxygenated, decomposition may proceed quickly, but under anoxic conditions

decomposition is slow and organic matter may begin to accumulate in the sediments

(McLatchy and Reddy 1998).

In forested floodplain wetlands on Australian lowland rivers, the breakdown and

decomposition of allochthonous plant material (eucalyptus leaves) has been shown to

take around 4 months (Briggs and Maher 1983), with the latter half of this period being

dominated by fragmentation (Glazebrook 1995; Janssen and Walker 1999). I have shown

that mass loss in the earlier phase of aquatic plant litter decomposition is temporally

correlated with high fungal biomass (Chapters 4 and 6).

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During aerobic decomposition, complex carbohydrate polymers are cleaved by the

extracellular enzymes of micro-organisms into monomers such as glucose, which can

then be absorbed and metabolised (Chamier 1985). This thesis shows that material

submerged in Australian floodplain wetlands is colonised by fungi, and that the biomass,

structure and enzyme activity of the fungal community changes over time. Fungi have

been characterised as the most important group of micro-organisms responsible for the

aerobic decomposition of plant material in agricultural soils, forest soils, rivers and peat

lands (Baldy, Gessner et al. 1995; Cromack and Caldwell 1992; Moorhead and Reynolds

1992; Thormann 2006), and it is now clear that they also degrade plant material in

Australian floodplain wetlands.

8.1.3 Detrital food chains

Food chains represent the dominant pathway for carbon transformation in many

ecosystems. Bunn and Boon (1993) investigated food webs in Australian floodplain

wetlands and identified Cherax sp. as a major consumer of detrital carbon. They found

that consumer groups were consuming a relatively 13

C-depleted resource as well as

riparian and macrophyte vegetation. Methanotrophic bacteria and planktonic

Chlorophyta were suggested as candidates for the 13

C-depleted carbon source. Fungi

colonising submerged plant litter may also be candidates (Henn, Gleixner et al. 2002),

but tend to have δ13

C signatures similar to their substrates (Reid, Quinn et al. 2008). The

δ13

C signatures of Oomycetes colonising submerged plant detritus has not been

determined in Australian systems, so this group is another potentially13

C-depleted

resource.

The fungi that colonise submerged plant litter are an important food source to

invertebrates such as caddisfly larvae (Cargill, Cummins et al. 1985), and provide

elements of their diet essential for growth and reproduction (Albariño, Villanueva et al.

2008). Colonisation of submerged allochthonous litter by aquatic fungi and Oomycetes

also results in a temporary increase in the leaf C:N, improving their food value (Chapter

6). The absence of a terrestrial aging period in litter decomposition prior to inundation, as

may occur with summer flooding, may lead to lower levels of fungal colonisation and

consequently reduced food resources for emerging invertebrates. In rivers, the observed

increases in C:N in submerged leaves are larger (Chapter 6), as is the abundance and

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diversity of shredder invertebrates relative to floodplain wetlands (Boulton and Lloyd

1991).

8.1.4 Pelagic food webs and phytoplankton decomposition

Carbon dissolved in the water column is utilised mainly by planktonic bacteria (Findlay

and Sinsabaugh 1999). Large amounts of organic carbon are leached from E.

camaldulensis leaves when they are submerged, and up to 50 % of this is rapidly utilised

by planktonic bacteria (Baldwin 1999). Extracellular enzyme activity provides further

evidence that bacteria use much of the DOC in the water column of Australian floodplain

wetlands (Boon 1989; Boon 1990; Boon and Sorrell 1991; Scholz and Boon 1993a;

Scholz and Boon 1993b). In terms of carbon cycling, bacterial activity in these wetlands

has been found to be consistent with the microbial loop concept (Pomeroy and Wiebe

1988). It should be noted that only weak linkages between grazing food webs and the

microbial loop have been established (Thorp and Delong, 2002).

In the water column, the main fungal species are zoosporic fungi belonging to the

Oomycetes and Chytridiomycota, and their motile propagules (Voronin 2008). The

Chytrids are commonly parasitic on algae (Ellen Van Donk 1983; Kagami, Van Donk et

al. 2004; Lopez-Llorca and Hernandez 1996) and Kagami et al. (2007) described the

food chain whereby fungal parasites make algal carbon available to Daphnia as the

“Mycoloop”. The Oomycetes are opportunistic and may colonise either living or dead

organic material (Czeczuga, Godlewska et al. 2004; Czeczuga, Kozlowska et al. 2002;

Czeczuga, Mazalska et al. 2005), often infecting physiologically stressed fish. The fungal

community in the sediments may be important as a “seed bank”, storing encysted

propagules of zoosporic fungi until suitable substrates and conditions are present in the

wetland (Chapter 5). Thus a planktonic fungal community may be important to food

webs involving cladocerans, diatoms and zoosporic fungi, and may influence the

magnitude of algal detritus that accumulates in wetland sediments.

8.2 The Importance of Fungi in Australian Floodplain Wetlands

Gessner et al. (2007) outlined the conditions required to prove that fungi are important

decomposer organisms in a given environment. These conditions are as follows:

1. Fungi must be present;

2. Fungi must be able to grow and reproduce under the prevailing conditions;

3. Fungi must produce the enzymes necessary to degrade the tissues of local plants;

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4. Fungal activities should result in mass loss from organic matter; and

5. Fungi should compete successfully with other organisms in the system.

This study has shown that fungi are present on a variety of substrates in a floodplain

wetland in the Murray-Darling Basin (Chapters 3). Fungal propagules from wetland

water and sediments are able to colonise submerged E. camaldulensis leaves, grow,

produce enzymes that break down the leaf components, and induce losses of mass and

organic matter from the leaves (Chapter 4). Fungi are present in submerged wetland

sediments (Chapters 3 & 5), but biomass is extremely low and there is no evidence so far

that fungi either grow or utilise sediment carbon resources. On E. camaldulensis leaves,

fungi compete with aquatic Oomycetes for resources, and both groups accumulate high

biomass at some time during the leaf decomposition process. In order to prove that fungi

were important decomposers of plant detritus in these floodplain wetlands, it would be

necessary to demonstrate their ability to successfully reproduce.

The fungal communities in wetlands were observed to differ spatially within a wetland

(Chapter 3), but not between three different wetlands (Chapter 6). The spatial differences

observed in the pilot study may be due to stands of macrophytes and zones where

allochthonous litter is deposited and the associated substrate specific fungal

communities. In Chapter 6, only one substrate type was used and no spatial differences

were observed. However, Oomycete community structure was observed to differ between

wetlands on the same day (Chapter 6), indicating that the composition of these

communities is influenced by factors in addition to substrate type.

There is evidence to suggest that fungal community structure is influenced by moisture

content of drying wetland sediments (Chapters 3 and 5), and that these communities may

be primarily composed of chytrid and Oomycete propagules and fungi that parasitise

diatoms (Chapter 5). This community is unlikely to be active after extended periods of

inundation due to low oxygen availability, or when sediments are exposed and dry, due

to low moisture availability. The sediment moisture content and sediment inundation also

influence the fungal biomass and enzyme activity on E. camaldulensis leaves lying on

the sediment surface (Chapter 4).

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When leaves age on the floodplain, their chemical composition is altered when compared

with fresh leaves (Chapter 7), and these changes promote fungal colonisation (Chapter

4). However, the amount of organic matter lost from the submerged leaves does not

differ between these leaf types. High resolution FT-IR spectro-microscopy (Chapter 7)

has shown that polysaccharides and proteins in the leaf mesophyll are consumed rapidly,

and this is followed by the simultaneous degradation of polysaccharides and lignin in the

leaf’s vascular tissues. Lignin degradation begins sooner after inundation in wetlands

than in rivers, and oils and waxes in the leaf epidermis are degraded more slowly than

other leaf components.

8.3 Additional Observations

8.3.1 Fungal and Oomycete dynamics in rivers and wetlands

At sites where river flow was declining and wetlands were drying, Oomycetes biomass

was high when the C:N of leaf material was high and fungal biomass was low. This

suggests that the Oomycetes may be more important in improving the food quality of

submerged plant detritus under warm, low flow conditions. Thus, estimates of fungal

biomass based on ergosterol concentration only may underestimate the food value of

conditioned E. camaldulensis leaves under these conditions. As the Oomycetes

community structure differs between rivers and wetlands, and between sites, it is

probable that flow and water quality are driving these differences.

Based on the observations made throughout this thesis, I suggest that a pattern exists in

the way Oomycete and fungal organisms colonise leaf substrates in rivers and wetlands

(Figure 8.1). This pattern consists of three phases (roman numerals), which may vary in

duration. Leaves fall from a tree and may experience a period of decomposition on the

floodplain prior to entering the water. Then, in phase I, a peak in biomass occurs

immediately after immersion, comprised mainly of a few species of Oomycetes that

exhibit strong growth and decline quickly. In phase II, the Eumycota become established,

reach a peak in biomass and then decline. In this study, maximum biomass in this phase

was higher in rivers than in wetlands and endured over a longer period. Phase III occurs

after a disturbance, inducing stress. This disturbance may be high temperatures, low

flow, low oxygen availability, wetland drying or declining substrate quality. In this

phase, we see an increase in Oomycete biomass followed by a decline, while fungal

biomass remains stable but at low levels.

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This pattern in biomass accumulation can be related to the movement of the leaf substrate

through the wetland and floodplain environment, and the spatial distribution of various

types of fungal and Oomycete propagules throughout the environment. Whilst on the

tree, or during floodplain aging, leaves may attract the propagules of plant pathogens and

litter and soil fungi. There are many species of true fungi and Oomycetes in both these

groups. When the leaves enter the aquatic phase, they tend to float on the surface for

some time (personal observation). At this point, Oomycetes begin to grow, and

accumulate a large amount of biomass that also declines quickly. The leaf may also

encounter the propagules of aquatic and aero-aquatic hyphomycetes that float on the

surface of the water. The partially decomposed leaf then begins to sink through the water

column. Here, aquatic and aero-aquatic hyphomycetes begin to accumulate biomass, and

the leaf may encounter the propagules of zoosporic fungi. As the substrate quality and

the availability of oxygen declines, the fungal biomass also declines, until the leaf settles

on the sediment surface. If a disturbance such as wetland drying or periodic exposure due

to low river flows occurs, an increase in Oomycete biomass may be seen. The leaf may

encounter yeasts and dormant propagules in the sediments that are able to utilise the

remaining leaf resources should the wetland remain dry long enough for these fungi to

become established. The time the leaf spends in each of these phases is currently

unknown, but is likely to be extremely variable.

Members of the Oomycetes are able to secrete enzymes that degrade lipids (Beakes and

Ford 1983; Mozaffar and Weete 1993; Rand and Munden 1992), enabling them to

colonise the waxy surface of the Eucalyptus leaf, and infect living fish. This may explain

their abundance in phase I. Once this layer is breached, the Eumycota are able to colonise

the leaf mesophyll, and deter further presence of Oomycetes through competition for

resources or by producing chemical inhibitors (Fryar, Booth et al. 2005; Mille-Lindblom,

Fischer et al. 2006a). When high quality food resources in the leaf become diminished,

and dissolved oxygen levels are reduced, a different group of Oomycetes may then

compete successfully for use of the remaining resources. Increased lignin breakdown is

observed during phase III in streams.

The molecular methods used to compare fungal communities are sensitive, and reveal the

full diversity of the community. However, they are not able to distinguish between active

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fungal organisms and resistant propagules. In the case of mycorrhizal communities in a

pine forest, Taylor and Bruns (1999) found only 10% of the species comprising resistant

propagules were found actively colonising plant roots. Thus we should attempt to

discriminate between active and dormant fungal species before making inferences about

the impacts of changes in fungal community structure upon ecosystem functioning in

general, and carbon cycling specifically.

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Figure 8.1: A schematic diagram representing the temporal patterns observed when a leaf substrate

is colonised by fungi and Oomycetes (top) and an explanation for this temporal pattern in terms of

the spatial distribution of fungal propagules within the wetland and floodplain ecosystem (bottom).

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Time periods are indicated by roman numerals, indicating the biomass accumulating on the

substrate (top), and the likely position of the substrate when these organisms colonise and grow to

maximum biomass levels (bottom).

8.3.2 The decomposition of lignin by aquatic fungi

In Chapter 4 it was observed that oxidative enzyme activity on E. camaldulensis leaves

was higher under aquatic conditions that under floodplain conditions. This suggested that

lignin decomposition was occurring in submerged leaves. Hydrolytic enzyme activity

showed that polysaccharides were being degraded at the same time. These chemical

changes were examined in Chapter 7, and it was shown that E. camaldulensis leaves

submerged in wetlands exhibit lignin degradation earlier than those in rivers. This may

be due to more limited bio-available carbon in wetland benthic habitats (Chapter 7).

Chemical mapping using Synchrotron- and FPA-FTIR micro-spectroscopy revealed

hitherto-unseen spatially-explicit insights into decomposition mechanisms in leaves

colonised by micro-organisms, and provided evidence to support earlier theories that

lignin breakdown products released via oxidative enzyme activity form stable and

refractory complexes within the decomposing leaf. Microbial protein and other

nitrogenous compounds appear to become complexed with polyphenols, or incorporated

into heterocyclic aromatic compounds which are not bio-available. Energy for lignin

degradation may be derived from the enzymatic break-down of bio-available sugars,

which occurs simultaneously with lignin degradation.

8.3.3 Issues of scale

During an ecosystem disturbance such as flooding or drying it is highly probably that the

biomass and diversity of the fungal community will be altered, but the effect on different

species and individual organisms will vary between physical locations and with

disturbances of different types and magnitudes (Morris, Friese et al. 2007). However, the

spatial scale over which such a disturbance occurs differs from the scale at which the

response of fungal communities was measured in this study.

Fungal communities are intrinsically patchy. Within a plant substrate such as a leaf, there

is variability in the distribution of fungal species, and patchy distribution of plant detritus

within a wetland leads to patchy distribution of fungal communities on the wetland scale.

In addition, wetland ecosystems occur as patches within the floodplain landscape. Thus,

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fungal ecology in wetlands lends itself to analysis in terms of patch dynamics models

(Pattee and Chergui 1995; Pickett and White 1985; Townsend 1989). Under this

paradigm, at the landscape scale major flood events may disperse fungal propagules

allowing re-colonisation of disturbed sites. Ecosystem scale disturbances such as wetland

drying would influence substrate availability, leading to changes in the biomass and

structure of fungal communities on patch (e.g. collection of submerged plant debris, reed

stand) and community (individual leaves) scales. At the same time, altered fungal

community structure and function may influence change at patch and ecosystem levels.

For example, increases in the abundance of opportunistic pathogens may limit

phytoplankton productivity (Kagami, de Bruin et al. 2007), or loss of saprotrophic

species may impact on nutrient availability and alter plant productivity (Hunt and Wall

2002).

Flooding and drying in floodplain wetlands are predictable disturbances, and fungal

communities in these ecosystems will have developed resilience to these changes

(Morris, Friese et al. 2007). That is, they have the capacity to return to a pre-disturbance

state. However, river regulation and climate change have led to changes in the temporal

pattern of these disturbances, reducing their predictability, and perhaps impacting upon

the resilience of the fungal community.

The experimental findings in this thesis have investigated fungal dynamics at the

community scale, and the inherent patchiness has often resulted in large variations about

mean results. Changes in the structure and functioning of the fungal communities

examined are likely to influence processes within the wetland ecosystem, but

extrapolation of these findings to the ecosystem scale should be applied with extreme

caution. A larger number of replicates would be needed at the patch and ecosystem scales

before confident predictions could be made as to the effect of changes in the fungal

community on the functioning of floodplain wetlands in general.

8.4 Knowledge Gaps

The work of Thomas et al. (1992) showed that fungal community structure differs

between leaf substrates from different plant species. Evidence to suggest that this may

also be true in wetlands was presented in Chapter 3. However, further work is required to

show that fungal communities differ between the standing dead blades of different

248

species of aquatic macrophytes, and the submerged detritus of other wetland plants. The

identity of fungal species parasitising phytoplankton and the role of fungi in

decomposing dead algal cells in sediments also remain entirely unexplored in Australian

wetlands.

The response of the fungal communities in Australian floodplain wetlands to

environmental variables has not been investigated. These include their response to

changes in temperature and season, and changes in water quality such as increased

concentration of dissolved nutrients, salts and potentially toxic substances, and decreased

oxygen concentrations. In particular, it would be an advantage to know whether the

fungal communities in floodplain wetlands exhibit resilience to disturbances such as

increased salinity, acidification and black water events.

It has been shown that leaf conditioning results in increased nitrogen content (Melillo,

Aber et al. 1982; Suberkropp, Godshalk et al. 1976; Thornton 1965 and Chapter 6).

However, we do not know which organisms are responsible for this process in wetlands,

or the origin of the additional nitrogen. Research in river systems in the USA and Canada

has shown that aquatic hyphomycetes are able to utilise dissolved nitrogen from the

water column (Sridhar and Bärlocher 2000; Suberkropp and Chauvet 1995), but this may

vary with the concentration of nitrogen both in the water column and in the substrate.

Laboratory experiments under flowing and standing water conditions might utilise 15

N

and 13

C enriched water to track the sources and sinks of nitrogen in decomposing plant

detritus.

It would also be an advantage to determine what proportions of carbon loss from

submerged leaves can be attributed to leaching and fungal production. Personal

observation suggests that fresh E. camaldulensis leaves submerged in sterile water

undergo virtually no leaching, and that some microbial activity is required to breach the

waxy leaf cuticle and initiate the leaching processes. This needs to be confirmed

experimentally.

While evidence from rivers (Chessman 1986; Davis and Winterbourne 1977) and

laboratory experiments (Albariño, Villanueva et al. 2008; Bärlocher and Kendrick 1975;

Graça, Maltby et al. 1993; Graça, Pozo et al. 2002) has shown that invertebrate shredders

249

consume aquatically conditioned leaves, it is not known whether shredders consume

conditioned eucalypt leaves submerged in wetlands. Given that these leaves do not

improve in food quality to the same extent as those in rivers, and that shredders are in

low abundance in these systems, it is possible that these leaves are not consumed by

shredders. In wetlands, it is possible that scraper organisms such as aquatic snails are

consuming the fungal biomass from the surface of submerged leaves.

8.5 Future Directions

In this thesis, T-RFLP has been used to compare the community structure between

samples of substrates. This method does not endeavour to identify the species involved.

However, without knowledge of the species comprising the fungal community it is

difficult to determine how environmental disturbances might influence the structure and

function of the fungal community, or how changes in the fungal community might

impact upon the wetland ecosystem. Therefore, future investigations into fungal ecology

in wetlands should attempt to identify species colonising plant, phytoplankton and

sediment substrates. This might be done using conventional culturing techniques to

isolate and grow wetland fungi before describing their morphology and sequencing the

DNA, or utilise the emerging metagenomics technologies that enable us to derive DNA

sequences (usually 18S) of all organisms present within a sample.

The extraction of ergosterol from fungal biomass provides an efficient method for

estimating fungal biomass on plant substrates, given some understanding of the

composition of the fungal community. However, this method does not provide an

estimate of Oomycetes biomass, and there is no similar method currently available that

can do so with the ease and speed of the ergosterol method. As the cell wall sterols in the

Oomycetes are also found in other aquatic organisms, molecular methods such as real-

time quantitative PCR may provide the best options for developing a useful Oomycete

biomass estimation method.

Time constraints did not allow concurrent measurement of extra-cellular enzyme activity

and substrate chemical composition in river and wetland conditioned leaves (Chapter 6

and 7). The simultaneous measurement of these variables would have enabled an

exploration of relationships between the metabolic processes of fungi and the breakdown

of components of the leaf substrate. Samples have been retained and there is potential for

250

this work to be conducted in the future. Spatially explicit chemical analysis of partially

decomposed leaves using S-FTIR would be further complemented by concurrent mass

spectroscopy or NMR analysis, to track the chemical changes produced by fungal

colonisation. This may enable the characterisation of “new” lignin polymers.

Further work is required to quantify the rate of breakdown of lignin, polysaccharides,

lipids and proteins in decomposing leaves, and to link this to submerged detritus biomass

estimates. This would allow conclusions to be drawn regarding the importance of these

processes to wetland carbon cycling. In addition, the fungal species responsible for

aquatic lignin degradation on submerged E. camaldulensis leaves should be isolated and

identified, and the relative contribution of these organisms and bacteria should be

determined.

8.6 Concluding Remarks

In floodplain wetlands, fungi are important in the breakdown of plant structural

polymers, and in enriching detritus with protein. In this role, they form a vital link

between plant production and higher trophic levels, and may be instrumental in the

formation of aquatic humic substances. As a field of enquiry, the study of the ecology of

aquatic fungi in floodplain wetlands is in its infancy. There is also ample opportunity for

exploration of fungal taxonomy and ecophysiology. A range of emerging spectroscopic,

molecular and isotopic techniques could possibly be applied to explore these areas to

improve our understanding of wetland ecosystem nutrient cycles and biotic interactions.

251

Appendix 1: Optimising Noise Reduction

The affect of different peak area threshold values on data richness is illustrated in Figure

8.2. The optimum threshold value is indicated by the data point immediately following

an initial change in slope of the data series. The graph indicates that optimum noise

reduction lies between 0.1 and 0.5% for most samples. Therefore, data analysis was

performed using a noise reduction of 0.25%.

Figure 8.2: This graph illustrates the number of data points remaining for each T-RFLP

sample after noise was removed. The percentage of noise reduction is shown on the x-

axis, while the number of taxonomic units represented is shown on the y-axis.

Noise reduction

0.00

10.00

20.00

30.00

40.00

50.00

60.00

70.00

80.00

90.00

0.01 0.1 0.25 0.5 1 3%

% sample data removed

To

tal N

o. F

rag

me

nts

1_Hin1 2_Hin1 3_Hin1



10_Hin1 11_Hin1 12_Hin1

13_Hin1 14_Hin1 15_Hin1

16_Hin1 17_Hin1 18_Hin1

19_Hin1 20_Hin1 21_Hin1

22_Hin1 23_Hin1 24_Hin1

27_Hin1 28_Hin1 29_Hin1

30_Hin1 31_Hin1 32_Hin1

33_Hin1 34_Hin1

252

References

Abbott I, Vanheurck P, Burbidge T, Williams M (1993) Damage caused by insects and

fungi to eucalypt foliage - Spatial and temporal patterns in Mediterranean forest of

Western-Australia. Forest Ecology and Management 58(1-2), 85-110.

Abdel-Raheem A, Shearer CA (2002) Extracellular enzyme production by freshwater

ascomycetes. Fungal Diversity 11, 1-19.

Abdel-Raheem AM, Ali EH (2004) Lignocellulolytic enzyme production by aquatic

hyphomycetes species isolated from the Nile's delta region. Mycopathologia 157(3), 277-

286.

Abdullah SK, Taj-Aldeen SJ (1989) Extracellular enzymatic activity of aquatic and aero-

aquatic conidial fungi. Hydrobiologia 174, 217-223.

Adams DD, van Eck GTM (1988) Biogeochemical cycling of organic carbon in the

sediments of the Grote Rug reservoir. Archiv für Hydrobiologie 31, 319-330.

Adl SM, Simpson AGB, et al. (2005) The new higher level classification of eukaryotes

with emphasis on the taxonomy of protists. The Journal of Eukaryotic Microbiology

52(5), 399-451.

Aguiar A, de Souza-Cruz PB, Ferraz A (2006) Oxalic acid, Fe3+-reduction activity and

oxidative enzymes detected in culture extracts recovered from Pinus taeda wood chips

biotreated by Ceriporiopsis subvermispora. Enzyme and Microbial Technology 38(7),

873-878.

Aiken GR, McKnight DM, Wershaw RL, MacCarthy P (1985) An introduction to humic

substances in soil, sediment and water. In 'Humic substances in soil, sediment and water:

Geochemistry, isolation and characterisation.' pp. 1-9. (John Wiley and Sons: New

York)

Al-Rekabi SAW, Al-Jubori SS, Shneshil SS (2005) A study of the pathogenicity of two

aquatic fungi on carp eggs. Egyptian Journal of Aquatic Research 31, 149-156.

Albariño R, Villanueva VD, Canhoto C (2008) The effect of sunlight on leaf litter quality

reduces growth of the shredder Klapopteryx kuscheli. Freshwater Biology 53(9), 1881-

1889.

Ali I, Abbas SQ, Mameed M, Naz N, Zafar S, Kanwal S (2009) Leaf anatomical

adaptations in some exotic species of Eucalyptus L'Her. (Myrtaceae). Pakistan Journal of

Botany 41(6), 2717-2727.

Alverez S, Rico E, Guerrero MC, Montes C (2001) Decomposition of Juncus maritimus

in two shallow lakes of Donana National Park. International Review of Hydrobiology

86(4-5), 541-554. [In English]

253

American Public Health Association, American Water Works Association, Water

Pollution Control Federation, Franson MAH (Eds) (1989) 'Standard methods for the

examination of water and wastewater (17th ed. edn).' (American Public Health

Association: Washington, D.C., USA)

Ander P (1994) The cellobiose-oxidizing enzymes CBQ and CbO as related to lignin and

cellulose degradation: a review. FEMS Microbiology Reviews 13(2-3), 297-311.

Andjic V, Hardy GES, Cortinas MN, Wingfield MJ, Burgess TI (2007) Multiple gene

genealogies reveal important relationships between species of Phaeophleospora infecting

Eucalyptus leaves. FEMS Microbiology Letters 268(1), 22-33.

Anesio AM, Tranvik LJ, Granéli W (1999) Production of inorganic carbon from aquatic

macrophytes by solar radiation. Ecology 80(6), 1852-1859.

Aro N, Pakula T, Penttila M (2005) Transcriptional regulation of plant cell wall

degradation by filamentous fungi. FEMS Microbiology Reviews 29(4), 719-739.

Arteau M, Labrie S, Roy D (2010) Terminal-restriction fragment length polymorphism

and automated ribosomal intergenic spacer analysis profiling of fungal communities in

Camembert cheese. International Dairy Journal 20(8), 545-554.

Arthington AH, Balcombe SR, Wilson GA, Thoms MC, Marshall J (2005) Spatial and

temporal variation in fish-assemblage structure in isolated waterholes during the 2001

dry season of an arid-zone floodplain river, Cooper Creek, Australia. Marine and

Freshwater Research 56(1), 25-35.

Artigas Alejo J (2008) The role of fungi and bacteria on the organic matter

decomposition process in streams: Interaction and relevance in biofilms. PhD Thesis,

Universitat de Girona, Spain

Aseada T, Nam LH, Hietz P, Tanaka N, Karunaratne S (2002) Seasonal fluctuations in

live and dead biomass of Phragmites australis as described by a growth and

decomposition model: implications of duration of aerobic conditions for litter

mineralisation and sedimentation. Aquatic Botany 73, 223-239.

Ayers AR, Ayers SB, Eriksson KE (1978) Cellobiose oxidase, purification and partial

characterization of a hemoprotein from Sporotrichum pulverulentum. European Journal

of Biochemistry 90(1), 171-181.

Azam F, Cho BC (1987) Bacterial utilisation of organic matter in the sea. In 'Ecology of

Microbial Communities.' (Eds M Fletcher, TRG Gray and JG Jones) pp. 261-281.

(Cambridge University Press: Cambridge)

Azam F, Fenchel T, Field JG, Gray JS, Meyer-Reil LA, Thingstad TF (1983) The

ecological role of water column microbes in the sea. Marine Ecology - Progress Series

10, 257-263.

Bacon P, Stone C, Binns DL, Leslie D, Edwards DW (1993) Relationships between

water availability and Eucalyptus camaldulensis growth in a riparian forest. Journal of

Hydrology 150, 541-561.

254

Bailey PCE, Watkins SC, Morris KL, Boon PI (2003) Do Melaleuca ericifolia SM.

leaves suppress organic matter decay in freshwater wetlands? Archiv für Hydrobiologie

156(2), 225-240. [In English]

Balcombe SR, Closs GP, Suter PJ (2007) Density and distribution of epiphytic

invertebrates on emergent macrophytes in a floodplain billabong. River Research and

Applications 23(8), 843-857.

Baldrian P (2006) Fungal laccases - occurrence and properties. FEMS Microbial Review

30(2), 215-242.

Baldrian P, Valaskova V (2008) Degradation of cellulose by basidiomycetous fungi.

FEMS Microbiology Reviews 32(3), 501-521.

Baldwin DS (1999) Dissolved organic matter and phosphorus leached from fresh and

'terrestrially' aged river red gum leaves: implications for assessing river-floodplain

interactions. Freshwater Biology 41, 675-685. [In English]

Baldwin DS, Hall KC, Rees GN, Richardson AJ (2007) Development of a protocol for

recognizing sulfidic sediments (potential acid sulfate soils) in freshwater wetlands.

Ecological Management & Restoration 8(1), 56-60.

Baldwin DS, Humphries P (2001) Living with less: The role of drought in aquatic

ecosystems. Wetlands Alive 5(2), 8.

Baldwin DS, Mitchell AM (2000) The effects of drying and re-flooding on the sediment

and soil nutrient dynamics of lowland river-floodplain systems: a synthesis. Regulated

Rivers: Research and Management 16(5), 457-467.

Baldy V, Chauvet E, Charcosset J-Y, Gessner MO (2002) Microbial dynamics associated

with leaves decomposing in the mainstem and floodplain pond of a large river. Aquatic

Microbial Ecology 28(1), 25-36.

Baldy V, Gessner MO, Chauvet E (1995) Bacteria, fungi and the breakdown of leaf litter

in a large river. Oikos 74, 93-102.

Baldy V, Gobert V, Guerold F, Chauvet E, Lambrigot D, Charcosset JY (2007) Leaf

litter breakdown budgets in streams of various trophic status: effects of dissolved

inorganic nutrients on microorganisms and invertebrates. Freshwater Biology 52(7),

1322-1335.

Bandekar J, Krimm S (1979) Vibrational analysis of peptides, polypeptides, and proteins:

Characteristic amide bands of β-turns Proceedings of the National Academy of Sciences

of the United States of America 76 (2), 774-777.

Bärlocher F (1981) Fungi on the food and in the faeces of Gammarus pulex.

Transactions of the British Mycological Society 76, 160-165.

Bärlocher F (1982) The contribution of fungal enzymes to the digestion of leaves by

Gammarus fossarum Koch (Amphipoda). Oecologia 52, 1-4.

255

Bärlocher F (1985) The role of fungi in the nutrition of stream invertebrates. Botanical

Journal of the Linnean Society 91, 83-94.

Bärlocher F (1990) Factors that delay colonization of fresh alder leaves by aquatic

hyphomycetes Archiv für Hydrobiologie 119(3), 249-255.

Bärlocher F (1992) 'The ecology of aquatic hyphomycetes.' (Springer-Verlag) 225

Bärlocher F (2005) Leaching. In 'Methods to study litter decomposition: A practical

guide.' (Eds MAS Graca, F Bärlocher and MO Gessner) pp. 33-42. (Springer: Dordrecht,

The Netherlands)

Bärlocher F, Canhoto C, Graça MAS (1995) Fungal colonization of alder and eucalypt

leaves in two streams in central Portugal. Archiv für Hydrobiologie 133(4), 457-470.

Bärlocher F, Corkum M (2003) Nutrient enrichment overwhelms diversity effects in leaf

decomposition by stream fungi. Oikos 101(2), 247-252.

Bärlocher F, Graça MAS (2002) Exotic riparian vegetation lowers fungal diversity but

not leaf decomposition in Portuguese streams. Freshwater Biology 47(6), 1123-1135.

Bärlocher F, Kendrick B (1974) Dynamics of the fungal population on leaves in a stream.

The Journal of Ecology 62(3), 761-791.

Bärlocher F, Kendrick B (1975) Assimilation efficiency of Gammarus pseudolimnaeus

(Amphipoda) feeding on fungal mycelium or autumn-shed leaves Oikos 26(1), 55-59.

Bärlocher F, Mackay RJ, Wiggins GB (1978) Detritus processing in a temporary vernal

pool in southern Ontario. Archiv für Hydrobiologie 81(3), 269-295.

Bärlocher F, Oertli JJ (1978) Inhibitors of aquatic hyphomycetes in dead conifer needles.

Mycologia 70(5), 964-974.

Bärlocher F, Rosset J (1981) Aquatic hyphomycete spora of two Black Forest and two

Swiss Jura streams. Transactions of the British Mycological Society 76(3), 479-483.

Bärlocher F, Seena S, Wilson KP, Williams DD (2008) Raised water temperature lowers

diversity of hyporheic aquatic hyphomycetes. Freshwater Biology 53(2), 368-379.

Bärlocher F, Tibbo P, Christie S (1989) Formation of phenol-protein complexes and their

use by two stream invertebrates. Hydrobiologia 173(3), 243-249.

Battle JM, Golladay SW (2001) Hydroperiod influence on breakdown of leaf litter in

cypress-gum wetlands. The American Midland Naturalist 146, 128–145

Batzer DP, Wissinger SA (1996) Ecology of insect communities in nontidal wetlands.

Annual Review of Entomology 41(1), 75-100.

Bayley PB (1991) The flood pulse advantage and the restoration of river-floodplain

systems. Regulated Rivers: Research and Management 6(2), 75-86.

256

Beakes G, Ford H (1983) Esterase isoenzyme variation in the genus Saprolegnia, with

particular reference to the fish-pathogenic S. diclina-parasitica Complex. Journal of

General Microbiology 129(8), 2605-2619.

Bengtsson G (1992) Interactions between fungi, bacteria and beech leaves in a stream

microcosm. Oecologia 89, 542-549.

Benner R, Maccubbin AE, Hodson RE (1984) Anaerobic biodegradation of the lignin

and polysaccharide components of lignocellulose and synthetic lignin by sediment

microflora. Applied and Environmental Microbiology 47, 998-1004.

Benner R, Opsahl S (2001) Molecular indicators of the sources and transformations of

dissolved organic matter in the Mississippi River plume. Organic Geochemistry 32, 597-

611. [In English]

Bent Tolstrup C, Per Åkesson M (1982) Loss-on-ignition and carbon content in a beech

forest soil profile. Ecography 5(4), 376-380.

Bergbauer M, Moran MA, Hodson RE (1992) Decomposition of lignocellulose from a

freshwater macrophyte by aero-aquatic fungi. Microbial Ecology 23(2), 159-167.

Bertoni MD, Cabral D (1988) Phyllosphere of Eucalyptus viminalis. II: Distribution of

endophytes. Nova Hedwigia 46(3-4), 491-502. [In Anglais]

Bertrand C, Coute A, Cazaubon A (2004) Fungal parasitism of the diatom Asterionella

formosa Hassall (Bacillariophyceae) by Chytridiomycota. Annales De Limnologie-

International Journal of Limnology 40(1), 63-69.

Bettucci L, Saravay M (1993) Endophytic fungi of Eucalyptus globulus : A preliminary

study. Mycological Research 97(6), 679-682.

Birk EM (1979) Disappearance of overstorey and understorey litter in an open eucalypt

forest. Australian Journal of Ecology 4, 207-222.

Boddy L (1993) Saprotrophic cord-forming fungi: warfare strategies and other ecological

aspects. Mycological Research 97(6), 641-655.

Boeriu CG, Bravo D, Gosselink RJA, van Dam JEG (2004) Characterisation of structure-

dependent functional properties of lignin with infrared spectroscopy. Industrial Crops

and Products 20(2), 205-218.

Bolton RG, Boddy L (1993) Characterization of the spatial aspects of foraging mycelial

cord systems using fractal geometry. Mycological Research 97(6), 762-768.

Boon P (1991) Bacterial assemblages in rivers and billabongs of Southeastern Australia.


Boon PI (1989) Organic matter degradation and nutrient regeneration in Australian

freshwaters: I. Methods for exoenzyme assays in turbid aquatic environments. Archiv für

Hydrobiologie 115(3), 339-359.

257

Boon PI (1990) Organic matter degradation and nutrient regeneration in Australian

freshwaters: II. Spatial and temporal variation and relation with environmental

conditions. Archiv für Hydrobiologie 117(4), 405-436. [In English]

Boon PI (2000) Carbon cycling in Australian wetlands: the importance of methane.

Verhandlungen der Internationalen Vereinigung für Theoretische und Angewandte

Limnologie 27, 37-50.

Boon PI (2006) Biogeochemistry and bacterial ecology of hydrologically dynamic

wetlands. In 'Ecology of Freshwater and Estuarine Wetlands.' (Eds DP Batzer and RR

Sharitz) pp. 115-176. (University of California Press: Berkeley)

Boon PI, Johnstone L (1997) Organic matter decay in coastal wetlands: an inhibitory role

for essential oil from Melaleuca alternifolia leaves? Archiv für Hydrobiologie 134, 433-

449.

Boon PI, Mitchell A, Lee K (1997) Effects of wetting and drying on methane emissions

from ephemeral floodplain wetlands in south-eastern Australia. Hydrobiologia 357(1),

73-87.

Boon PI, Mitchell AM (1995) Methanogenesis in the sediments of an Australian

freshwater wetland: comparison with aerobic decay, and factors affecting

methanogenesis. FEMS Microbiology and Ecology 18(175-190.).

Boon PI, Sorrell BK (1991) Biogeochemistry of billabong sediments. I. The effect of

macrophytes. Freshwater Biology 26(2), 209-226.

Boon PI, Virtue P, Nichols PD (1996) Microbial consortia in wetland sediments: A

biomarker analysis of the effects of hydrological regime, vegetation and season on

benthic microbes. Marine and Freshwater Research 47, 27-41.

Borneman J, Hartin RJ (2000) PCR primers that amplify fungal rRNA genes from

environmental samples. Applied and Environmental Microbiology 66(10), 4356-4360.

Bornette G, Amoros C, Lamouroux N (1998) Aquatic plant diversity in riverine

wetlands: the role of connectivity. Freshwater Biology 39(2), 267-283.

Boswell GP, Jacobs H, Davidson FA, Gadd GM, Ritz K (2002) Functional consequences

of nutrient translocation in mycelial fungi. Journal of Theoretical Biology 217(4), 459-

477.

Bott TL, Barnes JR, Minshall GW (1983). In 'Primary productivity in streams.' pp. 29-

53. (Plenum Press: New York)

Boulton A, Lake P (1992) The ecology of two intermittent streams in Victoria, Australia.

Freshwater Biology 27(1), 123-138.

Boulton AJ, Boon PI (1991) A review of methodology used to measure leaf litter

decomposition in lotic environments: time to turn over an old leaf? Australian Journal of

Marine and Freshwater Research 42, 1-43.

258

Boulton AJ, Lloyd L (1991) Macroinvertebrate assemblages in floodplain habitats of the

lower River Murray, South Australia. Regulated Rivers: Research and Management 6,

183-201.

Boundy I Histological Methods. In 'DEV2011: Early human development from cells to

tissues. ' pp. 6. (Monash University, Department of Anatomy and Developmental

Biology)

Bourbonnais R, Paice MG (1990) Oxidation on non-phenolic substrates: An expanded

role for laccase in lignin biodegradation. FEBS Letters 267(1), 99-102.

Boyd CE, Goodyear CP (1971) Nutritive quality of food in ecological systems. Archiv

für Hydrobiologie 69, 256-270.

Bren LJ (1988) Effects of river regulation on flooding of a riparian red gum forest on the

River Murray, Australia. Regulated Rivers: Research and Management 2, 65-77.

Briggs S, Maher M, Tongway D (1993) Dissolved and particulate organic carbon in two

wetlands in southwestern New South Wales, Australia. Hydrobiologia 264(1), 13-19.

Briggs SV, Maher MT (1983) Litter fall and leaf decomposition in a river red gum

(Eucalyptus camaldulensis) swamp. Australian Journal of Botany 31, 307-316.

Briggs SV, Maher MT (1985) Limnological studies of waterfowl habitat in south-western

New South Wales. II. Aquatic macrophyte productivity. Australian Journal of Marine

and Freshwater Research 36, 707-15. [In English]

Briggs SV, Maher MT, Tongway DJ (1985) Dry matter and nutrient loss from

decomposing Vallisneria spiralis L. Aquatic Botany 22(3-4), 387-392.

Brinson MM (1977) Decomposition and Nutrient Exchange of Litter in an Alluvial

Swamp Forest. Ecology 58(3), 601-609.

Brinson MM, Lugo AE, Brown S (1981) Primary productivity, decomposition and

consumer activity in freshwater wetlands. Annual Review of Ecology and Systematics 12,

123-161.

Brock MA (1994) Aquatic vegetation in inland wetlands. In 'Australian vegetation.' 2nd

edn. Ed. RH Groves) pp. 437-466. (The Press Syndicate of the University of Cambridge:

Cambridge, UK)

Brune A, Frenzel P, Cypionka H (2000) Life at the oxic-anoxic interface: microbial

activities and adaptations. FEMS Microbiology Reviews 24(5), 691-710.

Bruning K (1991) Effects of temperature and light on the population dynamics of the

Asterionella-Rhizophydium association. Journal of Plankton Research 13(4), 707-719.

Buchan A, Newell SY, Butler M, Biers EJ, Hollibaugh JT, Moran MA (2003) Dynamics

of bacterial and fungal communities on decaying salt marsh grass. Applied and

Environmental Microbiology 69(11), 6676–6687.

259

Buesing N, Gessner MO (2006) Benthic bacterial and fungal productivity and carbon

turnover in a freshwater marsh. Applied and Environmental Microbiology 72(1), 596-

605.

Bunn SE (1988a) Processing of leaf litter in a northern jarrah forest stream, Western

Australia: I. Seasonal differences. Hydrobiologia 162, 201-210.

Bunn SE (1988b) Processing of leaf litter in two northern jarrah forest streams, Western

Australia: II. The role of macroinvertebrates and the influence of soluble polyphenols

and inorganic sediment. Hydrobiologia 162, 211-223. [In English]

Bunn SE, Boon PI (1993) What sources of organic carbon drive food webs in billabongs?

A study based on stable isotope analysis. Oecologia 96(1), 85-94.

Bunn SE, De Deckker P, Williams WD (1986) Origin and fate of organic matter in

Australian upland streams. In 'Limnology in Australia.' (Eds P De Deckker and WD

Williams) pp. 277-291. (Dr W. Junk Publishers, CSIRO: Dordrect, Melbourne)

Burford MA, Cook AJ, Fellows CS, Balcombe SR, Bunn SE (2008) Sources of carbon

fuelling production in an arid floodplain river. Marine and Freshwater Research 59(3),

224-234.

Burgaud G, Arzur D, Durand L, Cambon-Bonavita M-A, Barbier G (2010) Marine

culturable yeasts in deep-sea hydrothermal vents: species richness and association with

fauna. FEMS Microbiology Ecology 73(1), 121-133.

Burley J, Wood PJ, Hans AS (1971) Variation in leaf characteristics among provenances

of Eucalyptus camaldulensis Dehn. grown in Zambia. Australian Journal of Botany 19,

237-49.

Burns A, Ryder DS (2001a) Potential for biofilms as biological indicators in Australian

riverine systems. Ecological Management & Restoration 2(1), 53-64.

Burns A, Ryder DS (2001b) Response of bacterial extracellular enzymes to inundation of

floodplain sediments. Freshwater Biology 46(10), 1299-1307.

Buta JG, Zadrazil F, Galletti GC (1989) FT-IR determination of lignin degradation in

wheat straw by white rot fungus Stropharia rugosoannulata with different oxygen

concentrations. Journal of Agricultural and Food Chemistry 37(5), 1382-1384.

Butler MJ, Day AW (1998) Fungal melanins: a review. Canadian Journal of

Microbiology 44(12), 1115–1136.

Byler DM, Susi H (1986) Examination of the secondary structure of proteins by

deconvolved FTIR spectra. Biopolymers 25(3), 469-487.

Cabral D (1985) Phyllosphere of Eucalyptus viminalis : Dynamics of fungal populations.

Transactions of the British Mycological Society 85(3), 501-511.

260

Calbet A, Landry MR (1999) Mesozooplankton influences on the microbial food web:

Direct and indirect trophic interactions in the oligotrophic open ocean. Limnology and

Oceanography 44(6), 1370-1380

Campbell IC, Fuchshuber L (1995) Polyphenols, condensed tannins, and processing rates

of tropical and temperate leaves in an Australian stream. Journal of the North American

Benthological Society 14(1), 174-182.

Campbell IC, James KR, Hart BT, Devereaux A (1992a) Allochthonous coarse

particulate organic material in forest and pasture reaches of two south-eastern Australian

streams. I. Litter accession. Freshwater Biology 27, 341-352.

Campbell IC, James KR, Hart BT, Devereaux A (1992b) Allochthonous coarse

particulate organic material in forest and pasture reaches of two south-eastern Australian

streams. II. Litter processing. Freshwater Biology 27, 353-365.

Canhoto C, Bärlocher F, Graça MAS (2002) The effects of Eucalyptus globulus oils on

fungal enzymatic activity. Archiv für Hydrobiologie 154(1), 121-132.

Canhoto C, Graça MAS (1995) Food value of introduced eucalypt leaves for a

Mediterranean stream detritivore: Tipula lateralis. Freshwater Biology 34(2), 209-214.

Canhoto C, Graça MAS (1996) Decomposition of Eucalyptus globulus leaves and three

native leaf species (Alnus glutinosa, Castanea sativa and Quercus faginea) in a

Portuguese low order stream. Hydrobiologia 333(2), 79-85.

Canhoto C, Graça MAS (1999) Leaf barriers to fungal colonization and shredders

(Tipula lateralis) consumption of decomposing Eucalyptus globulus. Microbial Ecology.

37(3), 163-172.

Cappenberg TE (1988) Quantification of aerobic and anaerobic carbon mineralisation at

the sediment-water interface. Ergebnisse der Limnologie. 31, 307-317.

Cargill AS, Cummins KW, Hanson BJ, Lowry RR (1985) The role of lipids, fungi, and

temperature in the nutrition of a shredder caddisfly, Clistoronia magnifica. Freshwater

Invertebrate Biology 4(2), 64-78.

Carr GL (2001) Resolution limit for infrared microspectroscopy explored with

synchrotron radiation. Rev. Sci. Instr. 72(3), 1613-1619.

Chamier A-C (1985) Cell-wall degrading enzymes of aquatic hyphomycetes: a review.

Botanical Journal of the Linnean Society 91, 67-81.

Chamier A-C, Dixon PA (1982a) Pectinases in leaf degradation by aquatic

hyphomycetes. I. The field study: the colonisation pattern of aquatic hyphomycetes on

leaf packs in a Surrey stream. Oecologia 52, 109-115.

Chamier A-C, Dixon PA (1982b) Pectinases in leaf degradation by aquatic

hyphomycetes: the enzymes and leaf maceration. Journal of General Microbiology 128,

2469-2483.

261

Chamier A-C, Dixon PA, Archer SA (1984) The spatial distribution of fungi on

decomposing alder leaves in a freshwater stream. Oecologia 64(1), 92-103.

Charcosset J-Y, Chauvet E (2001) Effect of culture conditions on ergosterol as an

indicator of biomass in the Aquatic Hyphomycetes. Applied and Environmental

Microbiology 67(5), 2051-2055.

Chauvet E (1987) Changes in the chemical composition of alder, poplar and willow

leaves during decomposition in a river. Hydrobiologia 148(1), 35-44.

Chauvet E, Fabre E, Elosegul A, Pozo J (1997) The impact of eucalypt on the leaf-

associated aquatic hyphomycetes in Spanish streams. Canadian Journal of Botany 75,

880-887.

Chergui H, Pattee E (1993) Flow and retention of particulate organic matter in riparian

fluvial habitats under different climates. Hydrobiologia 251, 137-142.

Chessman B (1986) Dietary studies of aquatic insects from two Victorian rivers. .

Australian Journal of Marine and Freshwater Research 37(2), 129-146.

Choudhury D (1988) Herbivore induced changes in leaf-litter resource quality: A

neglected aspect of herbivory in ecosystem nutrient dynamics. Oikos 51(3), 389-393.

Chróst RJ (1990) Microbial ectoenzymes in aquatic environments. In 'Aquatic microbial

ecology: Biochemical and molecular approaches.' First edn. (Eds J Overbeck and RJ

Chrost) pp. 47-78. (Springer-Verlag: New York)

Chung N, Suberkropp K (2009) Contribution of fungal biomass to the growth of the

shredder, Pycnopsyche gentilis (Trichoptera: Limnephilidae). Freshwater Biology

54(11), 2212-2224.

Chynoweth DP (1996) Environmental impact of biomethanogenesis. Environmental

Monitoring and Assessment 42(1), 3-18.

Clarke KR, Somerfield PJ, Airoldi L, Warwick RM (2006) Exploring interactions by

second-stage community analyses. Journal of Experimental Marine Biology and Ecology

338(2), 179-192.

Clarke KR, Warwick RM (2001) 'Change in marine communities: An approach to

statistical analysis and interpretation.' 2nd edn. (Primer-E Plymouth)

Claus H, Filip Z (1998) Degradation and transformation of aquatic humic substances by

laccase-producing fungi Cladosporium cladosporioides and Polyporus versicolor. Acta

Hydrochimica et Hydrobiologica 26(3), 180-185.

Cochrane VW (1956) The anaerobic dissimilation of glucose by Fusarium lini.

Mycologia 48(1), 1-12.

Cole J, Findlay S, Pace ML (1988) Bacterial production in fresh and saltwater

ecosystems: a cross-system overview. Marine Ecology - Progress Series 43, 1-10.

262

Collado J, Platas G, Paulus B, Bills GF (2007) High-throughput culturing of fungi from

plant litter by a dilution-to-extinction technique. FEMS Microbiology Ecology 60(3),

521-533.

Collins SL, Sinsabaugh RL, Crenshaw C, Green L, Porras-Alfaro A, Stursova M, Zeglin

LH (2008) Pulse dynamics and microbial processes in aridland ecosystems. Journal of

Ecology 96(3), 413-420.

Conant RT, Drijber RA, Haddix ML, Parton WJ, Paul EA, Plante AF, Six J, Steinweg

JM (2008) Sensitivity of organic matter decomposition to warming varies with its

quality. Global Change Biology 14(4), 868-877.

Connell JH (1978) Diversity in tropical rain forests and coral reefs. Science 199(4335),

1302-1310.

Convention on Wetlands (Ramsar I, 1971) Classification System for Wetland Type. In

'Key Documents of the Ramsar Convention. ')

Cooke RC, Whipps JM (1993) 'Ecophysiology of fungi.' First edn. (Blackwell Scientific

Publications: Oxford) 37-41

Cork SJ, Pahl L (1996) The possible influence of nutritional factors on diet and habitat

selection by the ring tail possum (Pseudocheirus peregrinus). In 'Possums and Gliders.'

2nd edn. (Eds A Smith and I Hume) pp. 267-276. (Surrey Beatty and Sons Pty Ltd and

The Australian Mammal Society)

Cornut J, Elger A, Lambrigot D, Marmonier P, Chauvet E (2010) Early stages of leaf

decomposition are mediated by aquatic fungi in the hyporheic zone of woodland streams.

Freshwater Biology, doi: 10.1111/j.1365-2427.2010.02483.x.

Coûteaux M-M, Bottner P, Berg B (1995) Litter decomposition, climate and litter

quality. Tree 10(2), 63-66.

Cowling EB, Merrill W (1966) Nitrogen in wood and its role in wood deterioration.

Canadian Journal of Botany 44(11), 1539-1554

Cowling SW, Waid JS (1963) Aquatic hyphomycetes in Australia. Australian Journal of

Science 26, 122-123.

Cromack KJ, Caldwell BA (1992) The role of fungi in litter decomposition and nutrient

cycling. In 'The Fungal Community: It's organisation and role in the ecosystem.' Second

edn. (Eds GC Carroll and DT Wicklow) pp. 653-668. (MarcelDekker, Inc: New York)

Cullen D, Kersten PJ (1992) Fungal enzymes for ligno-cellulose degradation. In 'Applied

Molecular Genetics of Filamentous Fungi.' (Eds JR Kinghorn and G Turner) pp. 100-

131. (Blackie: London)

Cummins KW, Klug MJ (1979) Feeding ecology of stream invertebrates. Annual Review

of Ecology and Systematics 10(1), 147-172.

263

Czeczuga B, Godlewska A, Kiziewicz B (2004) Aquatic fungi growing on feathers of

wild and domestic bird species in limnologically different water bodies. Polish Journal

of Environmental Studies 13(1), 21-31.

Czeczuga B, Kozlowska M, Godlewska A (2002) Zoosporic aquatic fungi growing on

dead specimens of 29 freshwater crustacean species. Limnologica 32(2), 180-193.

Czeczuga B, Mazalska B, Godlewska A, Muszynska E (2005) Aquatic fungi growing on

dead fragments of submerged plants. Limnologica 35(4), 283-297.

Czeczuga B, Muszyńska E, Godlewska A, Mazalska B (2007) Aquatic fungi and

straminipilous organisms on decomposing fragments of wetland plants. Mycologia

Balcanica 4 (1-2), 31-44.

Das M, Royer TV, Leff LG (2007) Diversity of fungi, bacteria, and actinomycetes on

leaves decomposing in a stream. Applied and Environmental Microbiology 73(3), 756-

767.

Davies DR, Theodorou MK, Brooks AE, Trinci APJ (1993) Influence of drying on the

survival of anaerobic fungi in rumen digesta and faeces of cattle. FEMS Microbiology

Letters 106, 59-64.

Davis SF, Winterbourne MJ (1977) Breakdown and colonisation of Nothofagus leaves in

a New Zealand stream. Oikos 28, 250-255.

Day FP (1983) Effects of flooding on leaf litter decomposition in microcosms. Oecologia

56(2), 180-184.

de la Cruz AA, Gabriel BC (1974) Caloric, elemental, and nutritive changes in

decomposing Juncus roemerianus leaves. Ecology 55(4), 882-886.

de Vries RP, Visser J (2001) Aspergillus enzymes involved in degradation of plant cell

wall polysaccharides. Microbiology and Molecular Biology Reviews 65(4), 497-522.

Deobald LA, Crawford DL (1992) Lignocellulose biodegradation. In 'The fungal

community: Its organisation and role in the ecosystem.' (Eds GC Carroll and DT

Wicklow) pp. 730-737. (Marcel Dekker, Inc.: New York)

Dick MW (1962) The occurrence and distribution of Saprolegniaceae in certain soils of

South-East England. II. Distribution within defined areas Journal of Ecology 50(1), 119-

127.

Dick MW (1963) The occurrence and distribution of Saprolegniaceae in certain soils of

South-East England: III. Distribution in relation to pH and water content

The Journal of Ecology 51(1), 75-81.

Dick MW (1969) Morphology and taxonomy of the Oomycetes, with special reference to

Saprolegniaceae, Leptomitaceae and Pythiaceae I. Sexual reproduction. New Phytologist

68(3), 751-775.

264

Dick MW (1972) Morphology and taxonomy of the Oomycetes, with special reference to

Saprolegniaceae, Leptomitaceae, and Pythiaceae. II. Cytogenetic systems. New

Phytologist 71(6), 1151-1159.

Dick MW (1976a) The ecology of aquatic phycomycetes. In 'Recent advances in aquatic

mycology.' Ed. EBG Jones) pp. 513-542. (Elek Science: London, England.)

Dick MW (1976b) The ecology of the aquatic phycomyces. In 'Recent advances in

aquatic mycology.' Ed. EBG Jones). (Elek Science: London)

Díez-Navajas AM, Greif C, Poutaraud A, Merdinoglu D (2007) Two simplified

fluorescent staining techniques to observe infection structures of the oomycete

Plasmopara viticola in grapevine leaf tissues. Micron 38(6), 680-683.

Dix NJ, Webster J (1995) 'Fungal ecology.' First edn. (Chapman and Hall: London) 145-

171

Don RH, Cox PT, Wainwright BJ, Baker K, Mattick JS (1991) 'Touchdown' PCR to

circumvent spurious priming during gene amplification. Nucleic Acids Research 19(14),

4008.

Dorado J, van Beek TA, Claassen FW, Sierra-Alvarez R (2001) Degradation of lipophilic

wood extractive constituents in Pinus sylvestris by the white-rot fungi Bjerkandera sp.

and Trametes versicolor. Wood Science and Technology 35(1), 117-125.

Edwards PB (1982) Do waxes on juvenile eucalyptus leaves provide protection from

grazing insects? Australian Journal of Ecology 7(4), 347-352.

Eicker A (1973) The mycoflora of Eucalyptus maculata leaf litter. Soil Biology and

Biochemistry 5, 441-448.

Ellen Van Donk JR (1983) The effect of fungal parasitism on the succession of diatoms

in Lake Maarsseveen I (The Netherlands). Freshwater Biology 13(3), 241-251.

Ellis I, Meredith S (2004) Aquatic fauna survey of wetlands 351 and 491 near

Wentworth, South-West NSW. Murray-Darling Freshwater Research Centre,

NSW Murray Wetlands Working Group Inc.,, No. 01/2005.

Elton CS (1927) 'Animal ecology.' (Sidgwick & Jackson, LTD.: London) 207

Elwood JW, Newbold JD, Trimble AF, Stark RW (1981) The limiting role of phosphorus

in a woodland stream ecosystem: effects of P enrichment on leaf decomposition and

primary producers. Ecology 62(1), 146-158.

Emerson R, Natvig DO (1981) Adaption of fungi to stagnant waters. In 'The Fungal

Community: Its Organisation and Role in the Ecosystem. Vol. 2.' (Eds DT Wicklow and

GC Carroll) pp. 109-128. (Marcel Dekker: New York)

England JR (2001) Changes in the morphology and anatomy of leaves and sapwood of

Eucalyptus regnans F. Muell. with age., University of Melbourne, Melbourne

265

Enriquez S, Duarte CM, Sand-Jensen K (1993) Patterns in decomposition rates among

photosynthetic organisms: the importance of detritus C:N:P content. Oecologia 94, 457-

471.

Erickson K-EL, Blanchette RA, Ander P (1990a) The biodegradation of cellulose. In

'Microbial and Enzymatic Degradation of Wood and Wood Components.' First edn. pp.

99-113. (Springer-Verlag: Berlin)

Erickson K-EL, Blanchette RA, Ander P (1990b) 'Microbial and enzymatic degradation

of wood and wood components.' First edn. (Springer-Verlag: Berlin) 407

Erickson K-EL, Blanchette RA, Ander P (1990c) Morphological aspects of wood

degradation by fungi and bacteria. In 'Microbial and Enzymatic Degradation of Wood

and Wood Components.' First edn. pp. 1-21. (Springer-Verlag: Berlin)

Erukhimovitch V, Pavlov V, Talyshinsky M, Souprun Y, Huleihel M (2005) FTIR

microscopy as a method for identification of bacterial and fungal infections. Journal of

Pharmaceutical and Biomedical Analysis 37(5), 1105-1108.

Erukhimovitch V, Tsror L, Hazanovsky M, Talyshinsky M, Mukmanov I, Souprun Y,

Huleihel M (2005) Identification of fungal phyto-pathogens by Fourier-transform

infrared (FTIR) microscopy. Journal of Agricultural Technology 1(1), 145-152.

Esau K (1977) 'Anatomy of seed plants.' 2nd edn. (John Wiley and Sons: Brisbane)

Esteves FA, Barbieri R (1983) Dry weight and chemical changes during decomposition

of tropical macrophytes in Lobo Reservoir - São Paulo, Brazil. Aquatic Botany 16(3),

285-295.

Evans WC, Fuchs G (1988) Anaerobic degradation of aromatic compounds. Annual

Review of Microbiology 42 289-317.

Evert RF, Esau K (2006) 'Esau's Plant anatomy: meristems, cells, and tissues of the plant

body : their structure, function, and development.' (John Wiley and Sons: Hoboken, New

Jersey) 601

Fackler K, Schwanninger M, Gradinger C, Hinterstoisser B, Messner K (2007)

Qualitative and quantitative changes of beech wood degraded by wood-rotting

basidiomycetes monitored by Fourier transform infrared spectroscopic methods and

multivariate data analysis. FEMS Microbiology Letters 271 (2), 162-169.

Ferreira V, Gulis V, Graça M (2006) Whole-stream nitrate addition affects litter

decomposition and associated fungi but not invertebrates. Oecologia 149(4), 718-729.

Field JI, Webster J (1983) Anaerobic survival of aquatic fungi. Transactions of the

British Mycological Society 81(2), 365-369.

Filip Z, Claus H (2002) Formation, Utilization, and Transformation of Some Refractory

Organic Substanes by Aquatic Microorganisms. In 'Refractory Organic Substances in the

Environment.' First edn. (Eds FH Frimmel, G Abbt-Braun, KG Heumann, B Hock, H-D

266

Ludemann and M Spiteller) pp. 311-320. (Wiley-VCH: Weinheim (Federal Republic of

Germany))

Findlay S, Howe K, Austin HK (1990) Comparison of detritus dynamics in two tidal

freshwater wetlands. Ecology 71(1), 288-295.

Findlay S, Sinsabaugh RL (1999) Unravelling the sources and bioavailability of

dissolved organic matter in lotic aquatic ecosystems. Marine and Freshwater Research

50, 781-90.

Findlay SEG, Arsuffi TL (1989) Microbial growth and detritus transformations during

decomposition of leaf litter in a stream. Freshwater Biology 21(2), 261-269.

Fischer H, Bergfur J, Goedkoop W, Tranvik LJ (2009) Microbial leaf degraders in boreal

streams: bringing together stochastic and deterministic regulators of community

composition. Freshwater Biology 54(11), 2276-2289.

Fischer H, Mille-Lindblom C, Zwimmann E, Tranvik LJ (2006) Contribution of fungi

and bacteria to the formation of dissolved organic carbon from decaying common reed

(Phragmites australis). Archiv für Hydrobiologie 166(1), 79-97. [In Anglais]

Fisher JP, Davey RA, Webster J (1983) Degradation of lignin by aquatic and aeroaquatic

hyphomycetes. Trans Br Mycol Soc 80(1), 166-168.

Fisher PJ, Webster J (1979) Effect of oxygen and carbon dioxide on growth of four aero-

aquatic hyphomycetes. Transactions of the British Mycological Society 72(1), 57-61.

Ford PW, Boon PI, Lee K (2002) Methane and oxygen dynamics in a shallow floodplain

lake: the significance of periodic stratification. Hydrobiologia 485(1 - 3), 97-110.

Ford TE (1993) 'Aquatic microbiology: An ecological approach.' First edn. (Blackwell

Scientific Publications: Boston)

Francis C, Sheldon F (2002) River red gum (Eucalyptus camaldulensis Dehnh) organic

matter as a carbon source in the lower Darling River, Australia. Hydrobiologia 481, 113-

124.

Frankland JC (1982) Biomass and nutrient cycling by decomposer basidiomycetes. In

'Decomposer basidiomycetes: their biology and ecology.' (Eds JC Frankland, JN Hedger

and MJ Swift) pp. 242-261. (Cambridge University Press: New York)

Frankland JC, Lindley DK, Swift MJ (1978) A comparison of two methods for the

estimation of mycelial biomass in leaf litter. Soil Biology & Biochemistry 10, 323-333.

Frazier P, Page K (2006) The effect of river regulation on floodplain wetland inundation,

Murrumbidgee River, Australia. Marine and Freshwater Research 57, 133-141.

Freeman C, Ostle N, Fenner N, Kang H (2004) A regulatory role for phenol oxidase

during decomposition in peatlands. Soil Biology & Biochemistry 36, 1663-1667.

267

Freeman C, Ostle N, Kang H (2001) An enzymic 'latch' on a global carbon store. Nature

409(6817), 149-149.

Fryar SC, Booth W, Davies J, Hodgkiss IJ, Hyde KD (2005) Evidence of in situ

competition between fungi in freshwater. Fungal Diversity 18, 59-71.

Gallardo A, Merino J (1992) Nitrogen immobilization in leaf litter at two Mediterranean

ecosystems of SW Spain. Biogeochemistry 15(3), 213-228.

Gamauf C, Metz B, Seiboth B (2007) Degradation of plant cell wall polymers by fungi.

In 'Environmental and Microbial Relationships. Vol. 4.' (Eds CP Kubicek and IS

Druzhinina) pp. 325-340. (Springer Berlin Heidelberg)

Gams W (2007) Biodiversity of soil-inhabiting fungi. Biodiversity and Conservation 16,

69-72.

Garidel P, Boese M (2007) Mid Infrared Microspectroscopic Mapping and Imaging: A

Bio-Analytical Tool for Spatially and Chemically Resolved Tissue Characterisation and

Evaluation of Drug Permeation Within Tissues. Microscopy Research and Technique 70,

336-349.

Gehrke PC, Revell MB, Philbey AW (1993) Effects of river red gum, Eucalyptus

camaldulensis, litter on golden perch, Macquaria ambigua. Journal of Fish Biology 43,

265-279.

Gell PA, Sluiter IR, Fluin J (2002) Seasonal and interannual variations in diatom

assemblages in Murray River connected wetlands in north-west Victoria, Australia.

Marine and Freshwater Research 53(6), 981-992.

Gessner MO (2001) Mass loss, fungal colonisation and nutrient dynamics of Phragmites

australis leaves during senescence and early aerial decay. Aquatic Botany 69, 325-339.

[In English]

Gessner MO (2005) Ergosterol as a measure of fungal biomass. In 'Methods to Study

Litter Decomposition: A Practical Guide.' First edn. (Eds MAS Graca, F Bärlocher and

MO Gessner) pp. 329. (Springer: Dordrecht,, The Netherlands)

Gessner MO, Bauchrowitz MA, Escautier M (1991) Extraction and quantification of

ergosterol as a measure of fungal biomass in leaf litter. Microbial Ecology 22, 285-291.

Gessner MO, Chauvet E (1993) Ergosterol-to-biomass conversion factors for aquatic

hyphomycetes. Applied and Environmental Microbiology 59(2), 502-507.

Gessner MO, Chauvet E (1994) Importance of stream microfungi in controlling

breakdown rates of leaf litter. Ecology 75(6), 1807-1817.

Gessner MO, Gulis V, Kuehn KA, Chauvet E, Suberkropp K (2007) Fungal decomposers

of plant litter in aquatic ecosystems. In 'The Mycota IV: Environmental and microbial

relationships. Vol. IV.' Second edn. (Eds CP Kubicek and IS Druzhinina) pp. 301-324.

(Springer-Verlag: Berlin)

268

Gessner MO, Robinson CT, Ward JV (1998) Leaf breakdown in streams of an alpine

glacial floodplain: dynamics of fungi and nutrients. Journal of the North American

Benthological Society 17, 403-419.

Gessner MO, Schieferstein B, Muller U, Barkmann S, Lenfers U (1996) A partial budget

of the primary organic carbon flows in the littoral zone of a hardwater lake. Aquatic

Botany 55, 93-105.

Gessner MO, Schwoerbel JK (1989) Leaching kinetics of fresh leaf-litter with

implications for the current concept of leaf-processing in streams. Archiv für

Hydrobiologie 115(1), 81-90.

Gessner MO, Van Ryckegem G (2003) 'Water fungi as decomposers in freshwater

ecosystems.' (John Wiley & Sons: New York)

Gibb E, Walsh JH (1980) Effect of nutritional factors and carbondioxide on growth of

Fusarium moniliforme and other fungi in reduced oxygen concentrations. Transactions of

the British Mycological Society 74(111-118.).

Gierlinger N, Goswami L, Schmidt M, Burgert I, Coutand C, Rogge T, Schwanninger M

(2008) In situ FT-IR microscopic study on enzymatic treatment of poplar wood cross-

sections. Biomacromolecules 9(8), 2194-2201.

Gilles M, Zhao J, An M, Agboola S (2010) Chemical composition and antimicrobial

properties of essential oils of three Australian Eucalyptus species. Food Chemistry

119(2), 731-737.

Glazebrook HS (1995) The effect of floods on leaf litter breakdown rates and nutrient

dynamics in a river red gum (Eucalyptus camaldulensis Dehnh.) forest. Charles Sturt

University, Faculty of Science and Agriculture, School of Environmental and

Information Sciences.,

Glazebrook HS, Robertson AI (1999) The effect of flooding and flood timing on leaf

litter breakdown rates and nutrient dynamics in a river red gum (Eucalyptus

camaldulensis) forest. Australian Journal of Ecology 24, 625-635.

Gleason FH, Kagami M, Lefevre E, Sime-Ngando T (2008) The ecology of chytrids in

aquatic ecosystems: roles in food web dynamics. Fungal Biology Reviews 22, 17-25.

Glen M, Tommerup IC, Bougher NL, O'Brien PA (2001) Specificity, sensitivity and

discrimination of primers for PCR-RFLP of larger basidiomycetes and their applicability

to identification of ectomycorrhizal fungi in Eucalyptus forests and plantations.

Mycological Research 105(2), 138-149.

Godshalk GL, Wetzel RG (1978a) Decomposition of aquatic angiosperms. I. Dissolved

components. Aquatic Botany 5, 281-300.

Godshalk GL, Wetzel RG (1978b) Decomposition of aquatic angiosperms. II. Particulate

components. Aquatic Botany 5, 301-327.

269

Golladay SW, Sinsabaugh RL (1991) Biofilm development on leaf and wood surfaces in

a boreal river. Freshwater Biology 25(3), 437-450.

Goncalves JF, Graca MAS, Callisto M (2007) Litter decomposition in a Cerrado

savannah stream is retarded by leaf toughness, low dissolved nutrients and a low density

of shredders. Freshwater Biology 52(8), 1440-1451.

Goonan PM, Beer JA, Thompson TB, Suter PJ (1992) Wetlands of the River Murray

floodplain, South Australia. 1. Preliminary survey of the biota and physio-chemistry of

ten wetlands from Chowilla to Mannum. Transactions of the Royal Society of South

Australia 116(3), 81-94.

Goos RD (1987) Fungi with a twist: The helicosporous hyphomycetes. Mycologia 79(1),

1-22.

Gosselink RJA, Abächerli A, Semke H, Malherbe R, Käuper P, Nadif A, van Dam JEG

(2004) Analytical protocols for characterisation of sulphur-free lignin. Industrial Crops

and Products 19(3), 271-281.

Graça MAS, Maltby L, Calow P (1993) Importance of fungi in the diet of Gammarus

pulex and Asellus aquaticus. II. Effects on growth, reproduction and physiology.

Oecologia 96(3), 304-309.

Graça MAS, Pozo J, Canhoto C, Elosegi A (2002) Effects of Eucalyptus plantations on

detritus, decomposers, and detritivores in streams. The Scientific World 2, 1173-1185.

Grant TR, Temple-Smith PD (1998) Field biology of the platypus (Ornithorhynchus

anatinus): historical and current perspectives. Philosophical Transactions of the Royal

Society B: Biological Sciences 353(1372), 1081–1091.

Green SJ, Freeman S, Hadar Y, Minz D (2004) Molecular tools for isolate and

community studies of Pyrenomycete fungi. Mycologia 96(3), 439-451.

Gribben D, Rees G, Croome R (2003) Development of substantial populations of

anoxygenic phototrophic bacteria and aerobic phototrophs in Normans Lagoon, a

"billabong" adjacent to the Murray River, South-eastern Australia. Lake & Reservoirs,

Research & Management 8, 95-104.

Gribben DL (2000) Norman's Lagoon: an examination of its phytoplankton and

photosynthetic bacterial populations, and the physico-chemical factors influencing them.,

La Trobe University, Wodonga

Griffin DM (1972) 'Ecology of Soil Fungi.' (Chapman and Hall: London)

Grime JP (1973) Competitive exclusion in herbaceous vegetation. Nature 242(5396),

344-347

Guglielmo F, Bergemann SE, Gonthier P, Nicolotti G, Garbelotto M (2007) A multiplex

PCR-based method for the detection and early identification of wood rotting fungi in

standing trees. Journal of Applied Microbiology 103(5), 1490–1507.

270

Gulis V (2001) Are there any substrate preferences in aquatic hyphomycetes?


Gulis V, Ferreira V, Graça MAS (2006) Stimulation of leaf litter decomposition and

associated fungi and invertebrates by moderate eutrophication: implications for stream

assessment. Freshwater Biology 51(9), 1655 - 1669.

Gulis V, Kuehn KA, Suberkropp K (2009) Fungi. In 'The Encyclopedia of Inland Waters

'. Ed. GE Likens). (Elsevier )

Gulis V, Stephanovich AI (1999) Antibiotic effects of some aquatic hyphomycetes.


Gulis V, Suberkropp K (2003) Effect of inorganic nutrients on relative contributions of

fungi and bacteria to carbon flow from submerged decomposing leaf litter. Microbial

Ecology 45(1), 11-19.

Gulis V, Suberkropp K (2004) Effects of whole-stream nutrient enrichment on the

concentration and abundance of aquatic hyphomycete conidia in transport. Mycologia

96(1), 57-65.

Guo LB, Sims REH (2001) Effects of light, temperature, water and meatworks effluent

irrigation on eucalypt leaf litter decomposition under controlled environmental

conditions. Applied Soil Ecology 17, 229-237.

Gutknecht J, Goodman R, Balser T (2006) Linking soil process and microbial ecology in

freshwater wetland ecosystems. Plant and Soil 289(1), 17-34.

Haack TK, McFeters GA (1982) Nutritional relationships among microorganisms in an

epilithic biofilm community. Microbial Ecology 8(2), 115-126.

Hall KC, Baldwin DS, Rees GN, Richardson AJ (2006) Distribution of inland wetlands

with sulfidic sediments in the Murray-Darling Basin, Australia. Science of The Total

Environment 370(1), 235-244.

Hallett IC, Dick MW (1981) Seasonal and diurnal fluctuations of oomycete propagule

numbers in the free water of a freshwater lake. Journal of Ecology 69, 671-692.

Hallett IC, Dick MW (1986) Fine structure of zoospore cyst ornamentation in the

Saprolegniaceae and Pythiaceae. Transactions of the British Mycological Society 86(3),

457-463.

Hammel KE (1997) Fungal degradation of lignin. In 'Driven by nature: Plant litter quality

and decomposition.' (Eds G Cadisch and KE Giller) pp. 33-45. (CAB International)

Hanssen JF, Thingstad TF, Goksøyr J (1974) Evaluation of hyphal lengths and fungal

biomass in soil by a membrane filter technique. Oikos 25(1), 102-107.

Hart BT, Bailey PCE, Edwards R, Hortle K, James KR, McMahon A, Meredith C,

Swadling K (1991) A review of the salt sensitivity of the Australian freshwater biota.

Hydrobiologia 210(105-144).

271

Hart BT, McGregor RJ (1980) Limnological survey of eight billabongs in the Magela

Creek catchment, Northern Territory. Australian Journal of Marine and Freshwater

Research 31, 611-626.

Harvey PJ, Schoemaker HE, Palmer JM (1986) Veratryl alcohol as a mediator and the

role of radical cations in lignin biodegradation by Phanerochaete chrysosporium. FEBS

Letters 195(1,2), 242-246.

Hawksworth DL, Rossman AY (1997) Where are al the undescribed Fungi?

Phytopathology 87, 888-891.

Headley JV, Peru KM, Verma B, Robarts RD (2002) Mass spectrometric determination

of ergosterol in a prairie natural wetland. Journal of Chromatography A 958(1-2), 149-

156.

Healey M, Thompson D, Robertson A (1997) Amphibian communities associated with

billabong habitats on the Murrumbidgee floodplain, Australia. Australian Journal of

Ecology 22(3), 270-278.

Hendel B, Marxsen J (2000) Extracellular enzyme activity associated with degradation of

beech wood in a central European stream. International Review of Hydrobiology 85(1),

95-105.

Hendel B, Sinsabaugh RL, Marxsen J (2005) Lignin-degrading enzymes: Phenoloxidase

and peroxidase. In 'Methods to study litter decomposition: A practical guide.' First edn.

(Eds MAS Graca, F Barlocher and MO Gessner) pp. 267-277. (Springer: Dordrecht, The

Netherlands)

Henn MR, Chapela IH (2000) Differential C isotope discrimination by fungi during

decomposition of C3- and C4- derived sucrose. Applied and Environmental

Microbiology 66(10), 4180-4186.

Henn MR, Gleixner G, Chapela IH (2002) Growth-dependent stable carbon isotope

fractionation by Basidiomycete fungi: δ13C pattern and physiological process. Applied

and Environmental Microbiology 68(10), 4956–4964.

Heraud P, Caine S, Sanson G, Gleadow R, Wood BR, McNaughton D (2007) Focal plane

array infrared imaging: a new way to analyse leaf tissue. New Phytologist 173(1), 216-

225.

Hessen DO (1998) Food webs and carbon cycling in humic lakes. In 'Aquatic Humic

Substances: Ecology and Biogeochemistry. Vol. 133.' (Eds DO Hessen and LJ Tranvik)

pp. 285-315. (Springer: Berlin)

Hieber M, Gessner MO (2002) Contribution of stream detritivores, fungi, and bacteria to

leaf breakdown based on biomass estimates. Ecology 83(4), 1026-1038.

Hillman T (1995) Billabongs, floodplains and the health of rivers. Water 22(2), 16-19.

272

Hillman TJ (1986) Billabongs. In 'Limnology in Australia.' (Eds P de Deckker and WD

Williams) pp. 457-470. (CSIRO/Junk: Melbourne/Dordrecht)

Hillman TJ, Quinn GP (2002) Temporal changes in macroinvertebrate assemblages

following experimental flooding in permanent and temporary wetlands in an Australian

floodplain forest. Regulated Rivers: Research and Management 18(2), 137-154.

Hillman TJ, Shiel RJ (1991) Macro- and microinvertebrates in Australian billabongs.


Limnologie 24(3), 1581-1587.

Hladyz S, Gessner MO, Giller PS, Pozo J, Woodward G (2009) Resource quality and

stoichiometric constraints on stream ecosystem functioning. Freshwater Biology 54(5),

957-970.

Holt DM, Jones EBG (1983) Bacterial degradation of lignified wood cell walls in

anaerobic aquatic habitats. Applied and Environmental Microbiology 46(3), 722-727.

Hood E, W., Williams MW, McKnight DM (2005) Sources of dissolved organic matter

(DOM) in a Rocky Mountain stream using chemical fractionation and stable isotopes.

Biogeochemistry 74, 231-255.

Hori R, Sugiyama J (2003) A combined FT-IR microscopy and principal component

analysis on softwood cell walls. Carbohydrate Polymers 52(4), 449-453.

Houghton RA (1995) Land-use change and the carbon cycle. Global Change Biology

1(4), 275-287.

Howitt JA (2003) Photochemical degradation of aquatic dissolved organic matter: the

role of suspended iron oxides. PhD Thesis, Monash University, Melbourne

Howitt JA, Baldwin DS, Rees GN, Hart BT (2004) Facilitated heterogeneous

photodegradation of dissolved organic matter by particulate iron. Environmental

Chemistry 1(3), 197-205.

Howitt JA, Baldwin DS, Rees GN, Williams JL (2007) Modelling blackwater: Predicting

water quality during flooding of lowland river forests. Ecological Modelling 203 (3-4),

229-242.

Hsu C-PS (1997) Infrared Spectroscopy. In 'Handbook of instrumental techniques for

analytical chemistry.' Ed. FA SETTLE) pp. 247-283. (Prentice Hall: Upper Saddle River,

NJ)

Hulthe G, Hulth S, Hall POJ (1998) Effect of oxygen on degradation rate of refractory

and labile organic matter in continental margin sediments. Geochimica et Cosmochimica

Acta 62(8), 1319-1328.

Hulvey JP, Padgett DE, Bailey JC (2007) Species boundaries within Saprolegnia

(Saprolegniales, Oomycota) based on morphological and DNA sequence data. Mycologia

99(3), 421-429.

273

Hunt HW, Wall DH (2002) Modelling the effects of loss of soil biodiversity on

ecosystem function. Global Change Biology 8(1), 33-50.

Hutson BR (1985) Rates of litterfall and organic matter turnover at three South

Australian indigenous forest sites. Australian Journal of Ecology 10, 351-359.

Hutson BR, Veitch LG (1985) Relationships between litterfall rate, litter mass and

decomposition rate in Eucalyptus forests in southeastern Australia. Australian Journal of

Ecology 10, 443-450. [In English]

Hyde KD, Goh TK (1998) Fungi on submerged wood in Lake Barrine, north Queensland,

Australia. Mycological Research 102(6), 739-749.

Hyde KD, Lee SY (1995) Ecology of mangrove fungi and their role in nutrient cycling:

what gaps occur in our knowledge? Hydrobiologia 295(1), 107-118.

Ingold CT (1942) Aquatic hyphomycetes of decaying alder leaves. Transactions of the

British Mycological Society 25, 339-417.

Ingold CT (1961) Another aquatic spore-type with clamp connexions. Transactions of

the British Mycological Society 44(1), 27-30, IN2.

Ingold CT (1976) The morphology and biology of freshwater fungi excluding

phycomycetes. In 'Recent Advances in Aquatic Mycology.' Ed. EBG Jones). (Elek

Science: London)

Inkley MD, Wissinger SA, Baros BL (2008) Effects of drying regime on microbial

colonization and shredder preference in seasonal woodland wetlands. Freshwater

Biology 53(3), 435-445.

Iqbal SH (1976) Effect of pH on sporulation of freshwater hyphomycetes. Biologia

(Lahore) 22, 143-153.

Iqbal SH, Webster J (1973) The trapping of aquatic hyphomycetes spores by air bubbles.


Isaac S (1997) Fungi naturally form many diverse biochemical products, some of which

are now commercially important; how and why do they do this? Mycologist 11(4), 182-

183.

Jaatinen K, Fritze H, Laine J, Laiho R (2007) Effects of short- and long-term water-level

drawdown on the populations and activity of aerobic decomposers in a boreal peatland.

Global Change Biology 13, 1-20.

Jackson CR, Foreman CM, Sinsabaugh RL (1995) Microbial enzyme activities as

indicators of organic matter processing rates in a Lake Erie coastal wetland. Freshwater

Biology 34(2), 329-342.

Jackson EF, Jackson CR (2008) Viruses in wetland ecosystems. Freshwater Biology

53(6), 1214-1227.

274

Jackson MA, Bothast RJ (1990) Carbon concentration and carbon-to-nitrogen ratio

influence submerged-culture conidiation by the potential bioherbicide Colletotrichum

truncatum NRRL 13737. Applied and Environmental Microbiology 56(11), 3435-3438.

James KR, Hart BT, Bailey PCE, Blinn DW (2009) Impact of secondary salinisation on

freshwater ecosystems: effect of experimentally increased salinity on an intermittent

floodplain wetland. Marine and Freshwater Research 60(3), 246-258.

James SA, Bell DT (1995) Morphology and anatomy of leaves of Eucalyptus

camaldulensis clones: Variation between geographically separated locations. Australian

Journal of Botany 43(4), 415-433.

Janssen MA, Walker KF (1999) Processing of riparian and wetland plant litter in the

River Murray, South Australia. Hydrobiologia 411, 53-64.

Jeffries TW, Choi S, Kirk TK (1981) Nutritional regulation of lignin degradation by

Phanerochaete chrysosporium. Applied and Environmental Microbiology 42(2), 290-

296.

Jenkins KM, Boulton AJ (2007) Detecting impacts and setting restoration targets in arid-

zone rivers: aquatic micro-invertebrate responses to reduced floodplain inundation.

Journal of Applied Ecology 44(4), 823-832.

Jennings DH, Lysek G (1996) 'Fungal biology: Understanding the fungal lifestyle.' 1 edn.

(BIOS Scientific Publishers Limited: Oxford, UK) 156

Jilkine K, Gough KM, Julian R, Kaminskyj SGW (2008) A sensitive method for

examining whole-cell biochemical composition in single cells of filamentous fungi using

synchrotron FTIR spectromicroscopy. Journal of Inorganic Biochemistry 102(3), 540-

546.

Jobard M, Rasconi S, Sime-Ngando T (2010) Diversity and functions of microscopic

fungi: a missing component in pelagic food webs. Aquatic Sciences - Research Across

Boundaries 72(3), 255-268.

Johnson TWJ, Seymour RL, Padgett DE (2002) 'The biology and systematics of the

Saprolegniaceae.' (Ilumina: Educational Resources for Science and Mathematics

Published online)

Jönsson LJ, Palmqvist E, Nilvebrant NO, Hahn-Hägerdal B (1998) Detoxification of

wood hydrolysates with laccase and peroxidase from the white-rot fungus Trametes

versicolor. Applied Microbiology and Biotechnology 49(6), 691-697.

Junk W, Bayley P, Sparks R (1989) The flood pulse concept in river-floodplain system.

Canadian special publication of fisheries and aquatic sciences 106, 110-127.

Juntheikki M-R, Julkunen-Tiitto R (2000) Inhibition of β-glucosidase and esterase by

tannins from Betula, Salix, and Pinus Species. Journal of Chemical Ecology 26(5), 1151-

1165.

275

Kacuráková M, Capek P, Sasinková V, Wellner N, Ebringerová A (2000) FT-IR study of

plant cell wall model compounds: pectic polysaccharides and hemicelluloses.

Carbohydrate Polymers 43(2), 195-203.

Kagami M, de Bruin A, Ibelings B, Van Donk E (2007) Parasitic chytrids: their effects

on phytoplankton communities and food-web dynamics. Hydrobiologia 578(1), 113-129.

Kagami M, Van Donk E, de Bruin A, Rijkeboer M, Ibelings BW (2004) Daphnia can

protect diatoms from fungal parasitism. Limnology and Oceanography 49(3), 680-685.

Kaminskyj SGW, Jilkine K, Szeghalmi A, Gough KM (2008) High spatial resolution

analysis of fungal cell biochemistry - bridging the analytical gap using synchrotron FTIR

spectromicroscopy. FEMS Microbiology Letters 284(1), 1-8.

Kapich AN, Steffen KT, Hofrichter M, Hatakka A (2005) Involvement of lipid

peroxidation in the degradation of a non-phenolic lignin model compound by manganese

peroxidase of the litter-decomposing fungus Stropharia coronilla. Biochemical and

Biophysical Research Communications 330(2), 371-377.

Kaplan LA, Newbold JD (2003) The role of monomers in stream ecosystem metabolism.

In 'Aquatic ecosystems: interactivity of dissolved organic matter.' (Eds S Findlay and RL

Sinsabaugh) pp. 121-137. (Elsevier Science (USA): San Diego, California)

Kasuga T, Taylor JW, White TJ (1999) Phylogenetic relationships of varieties and

geographical groups of the human pathogenic fungus Histoplasma capsulatum Darling.

Journal of Clinical Microbiology 37(3), 653-663.

Kaushik NK, Hynes HBN (1971) The fate of dead leaves that fall into streams. Archiv

für Hydrobiologie 68, 645-515.

Keating RC (1984) Leaf histology and its contribution to relationships in the Myrtales.

Annals of the Missouri Botanical Garden 71(3), 801-823.

Keishi T, Yasoo I, Tagiru O, Kunihiko K, Keiko S (2006) Phylogenetic analysis of

manganese-oxidizing fungi isolated from manganese-rich aquatic environments in

Hokkaido, Japan. Limnology 7(3), 219-223.

Kennett R, Roe J, Hodges K, Georges A (2009) Chelodina longicollis (Shaw 1974) -

Eastern long-necked turtle, common long-necked turtle, common snake-necked turtle. In

'Conservation biology of Australian freshwater turtles and tortoises: A complilation

project of the IUCN/SSC Toirtoise and freshwater turtle specialist group.' (Eds AGJ

Rodin, PCH Pritchard, PPv Dijk, RA Saumure, KA Buhlmann, JB Iverson and RA

Mittermeier) pp. 031.1-031.8. (Chelonian Research Foundation)

Khallil AM, El-Hissy FT, Ali EH (1995) Seasonal fluctuations of aquatic fungi recovered

from Egyptian soil (Delta Region). Journal of Basic Microbiology 35(2), 93-102.

Khan AG (1993) Occurrence and importance of mycorrhizae in aquatic trees of New

South Wales, Australia. Mycorrhiza 3(1), 31-38.

276

Kiesecker JM, Blaustein AR, Miller CL (2001) Transfer of a Pathogen from Fish to

Amphibians. Conservation Biology 15(4), 1064-1070.

King GM (1986) Characteristics of -glucosidase activity in intertidal marine sediments.

Applied and Environmental Microbiology 51, 373-380.

Kingsford RT, Curtin AL, Porter J (1999) Water flows on Cooper Creek in arid Australia

determine 'boom' and 'bust' periods for waterbirds. Biological Conservation 88(2), 231-

248.

Kingsford RT, Norman FI (2002) Australian waterbirds - products of the continent's

ecology. Emu 102, 47-69.

Kiørboe T (1993) Turbulence, phytoplankton cell size, and the structure of pelagic food

webs Advances in Marine Biology 29, 1-61.

Kirk TK, Farrell RL (1987) Enzymatic combustion: the microbial degradation of lignin.

Annual Review of Microbiology 41, 465-505.

Kirschbaum MUF (1995) The temperature dependence of soil organic matter

decomposition, and the effect of global warming on soil organic C storage. Soil Biology

& Biochemistry 27(6), 753-760.

Kirschbaum MUF (2006) The temperature dependence of organic-matter decomposition-

-still a topic of debate. Soil Biology and Biochemistry 38(9), 2510-2518.

Kjoller A, Struwe S (1980) Microfungi of decomposing red alder leaves and thier

substrate utilisation. Soil Biology & Biochemistry 12, 425-431. [In English]

Kjoller A, Struwe S (1992) Functional groups of microfungi in decomposition. In 'The

Fungal Community: Its organisation and Role in the Ecosystem. Vol. 9.' Second edn.

(Eds GC Carroll and DT Wicklow) pp. 619-630. (Marcel Dekker, Inc.: New York)

Klis FM, Mol P, Hellingwerf K, Brul S (2002) Dynamics of cell wall structure in

Saccharomyces cerevisiae. FEMS Microbiology Reviews 26(3), 239-256.

Kominkova D, Kuehn KA, Buesing N, Steiner D, Gessner MO (2000) Microbial

biomass, growth, and respiration associated with submerged litter of Phragmites

australis decomposing in a littoral reed stand of a large lake. Aquatic Microbial Ecology

22, 271-282. [In English]

Kominoski JS, Hoellein TJ, Kelly JJ, Pringle CM (2009) Does mixing litter of different

qualities alter stream microbial diversity and functioning on individual litter species?

Oikos 118(3), 457-463.

Konen ME, Jacobs PM, Burras CL, Talaga BJ, Mason JA (2002) Equations for

Predicting Soil Organic Carbon Using Loss-on-Ignition for North Central U.S. Soils. Soil

Science Society of America Journal 66(6), 1878-1881.

277

Kuehn KA, Lemke MJ, Suberkropp K, Wetzel RG (2000) Microbial biomass and

production associated with decaying leaf litter of emergent macrophyte Juncus effusus.

Limnology and Oceanography 45(4), 862-870. [In English]

Kuehn KA, Suberkropp K (1998a) Decomposition of standing litter of the freshwater

macrophyte Juncus effusus. Freshwater Biology 40, 717-727. [In English]

Kuehn KA, Suberkropp K (1998b) Diel fluctuations in microbial actvity associated with

standing dead leaf litter of the emergent macrophyte Juncus effusus. Aquatic Microbial

Ecology 14, 171-182.

Kumar M, Shukla PK (2005) Use of PCR targeting of Internal Transcribed Spacer

regions and single-stranded conformation polymorphism analysis of sequence variation

in different regions of rRNA genes in fungi for rapid diagnosis of mycotic keratitis.

Journal of Clinical Microbiology 43(2), 662-668.

Lake PS (2000) Disturbance, patchiness, and diversity in streams. Journal of the North

American Benthological Society 19(4), 573-592.

Lamour KH, Win J, Kamoun S (2007) Oomycete genomics: new insights and future

directions. FEMS Microbiology Letters 274(1), 1-8.

Landeweert R, Veenman C, Kuyper TW, Fritze H, Wernars K, Smit E (2003)

Quantification of ectomycorrhizal mycelium in soil by real-time PCR compared to

conventional quantification techniques. FEMS Microbiology Ecology 45(3), 283-292.

Langhans S, Tockner K (2006) The role of timing, duration, and frequency of inundation

in controlling leaf litter decomposition in a river-floodplain ecosystem (Tagliamento,

northeastern Italy). Oecologia 147(3), 501-509.

Latijnhouwers M, de Wit PJGM, Govers F (2003) Oomycetes and fungi: similar

weaponry to attack plants. Trends in Microbiology 11(10), 462-469.

Laudon H, Kohler S, Buffam I (2004) Seasonal TOC export from seven boreal

catchments in northern Sweden. Aquatic Sciences 66, 223-230.

Lawley B, Ripley S, Bridge P, Convey P (2004) Molecular Analysis of Geographic

Patterns of Eukaryotic Diversity in Antarctic Soils. Applied and Environmental

Microbiology 70(10), 5963-5972.

Leahy PJ, Tibby J, Kershaw AP, Heijnis H, Kershaw JS (2005) The impact of European

settlement on Bolin Billabong, a Yarra River floodplain lake, Melbourne, Australia.

River Research and Applications 21(2-3), 131-149.

Leclerc MC, Guillot J, Deville M (2000) Taxonomic and phylogenetic analysis of

Saprolegniaceae (Oomycetes) inferred from LSU rDNA and ITS sequence comparisons.

Antonie van Leeuwenhoek 77(4), 369-377.

Lee SB, Taylor JW (1990) Isolation of DNA from fungal mycelia and spores. In 'PCR

Protocols: A guide to methods and applications.' pp. 282-287. (Academic Press, Inc.:

San Diego)

278

Lefevre E, Bardot C, Noel C, Carrias J-F, Viscogliosi E, Amblard C, Sime-Ngando T

(2007) Unveiling fungal zooflagellates as members of freshwater picoeukaryotes:

evidence from a molecular diversity study in a deep meromictic lake. Environmental


Lehninger AL, Nelson DL, Cox MM (1993) The citric acid cycle. In 'Principles of

biochemistry.' pp. 446-478. (Worth Publishers: New York)

Lenz J (1992) Microbial loop, microbial food web and classical food chain: Thier

significance in pelagic marine ecosystems. Ergebnisse der Limnologie. 37, 265-278.

Leonowicz A, Cho N, Luterek J, Wilkolazka A, Wojtas-Wasilewska M, Matuszewska A,

Hofrichter M, Wesenberg D, Rogalski J (2001) Fungal laccase: properties and activity on

lignin. Journal of Basic Microbiology 41(3-4), 185-227.

Liakos D (1986) A comparative study of decomposition rates of leaves of the River Red

Gum (Eucalyptus camaldulensis Dehnh.) and the Weeping Willow (Salix babylonica L.)

in the River Murray. University of Adelaide., Adelaide

Libkind D, Brizzio S, Ruffini A, Gadanho M, van Broock M, Paulo Sampaio J (2003)

Molecular characterization of carotenogenic yeasts from aquatic environments in

Patagonia, Argentina. Antonie van Leeuwenhoek 84(4), 313-322.

Lindroth RL, Kopper BJ, Parsons WFJ, Bockheim JG, Karnosky DF, Hendrey GR,

Pregitzer KS, Isebrands JG, Sober J (2001) Consequences of elevated carbon dioxide and

ozone for foliar chemical composition and dynamics in trembling aspen (Populus

tremuloides) and paper birch (Betula papyrifera). Environmental Pollution 115(3), 395-

404.

Ljungdahl LG, Eriksson K-E (1985) Ecology of microbial cellulose degradation.

Advances in Microbial Ecology 8, 237-299.

Lloyd L, Atkins B, Boon PI, Jacobs T, Roberts J (1992) Natural processes within

Murray-Darling floodplain wetlands. In 'Murray-Darling Basin floodplain wetlands

management. ' (Eds T Sharley and C Huggan). (Murray-Darling Basin Commission:

Albury, NSW)

Lopez-Llorca LV, Hernandez P (1996) Infection of the green alga Oocystis lacustris

Chod. with the Chytrid fungus Diplochytridium deltanum (Masters) Karling. An SEM

study. Micron 27(5), 355-358.

Lopez-Real JM, Swift MJ (1977) Formation of pseudosclerotia ('zone lines') in wood

decayed by Armillaria mellea and Stereum hirsutum: III. Formation in relation to

composition of gaseous atmosphere in wood. Transactions of the British Mycological

Society 68(3), 321-325.

Lord JC, Roberts DW (1985) Effects of salinity, pH, organic solutes, anaerobic

conditions, and the presence of other microbes on production and survival of Lagenidium

giganteum (Oomycetes: Lagenidiales) zoospores. Journal of Invertebrate Pathology

45(3), 331-338.

279

Love P (2005) Decomposition of fire-affected Eucalyptus viminalis leaves in a

freshwater ecosystem. Honours Thesis, University of Canberra, Canberra, Australia

Lund A (1934) Studies on Danish freshwater phycomycetes and notes on their

occurrence particularly relative to the hydrogen ion concentration of the water. Det

Kongelige Danske Videnskabernes Selskabs Naturvidenskabelige og Mathematiske

Afhandlinger 9(6), 1-97.

Luo Q, Krumholz LR, Najar FZ, Peacock AD, Roe BA, White DC, Elshahed MS (2005)

Diversity of the microeukaryotic community in sulfide-rich Zodletone Spring

(Oklahoma). Applied and Environmental Microbiology 71(10), 6175-6184.

Macauley BJ, Thrower LB (1966) Succession of fungi in leaf litter of Eucalyptus

regnans. Transactions of the British Mycological Society 49, 509-20.

Mackensen J, Bauhus J, Webber E (2003) Decomposition rates of coarse woody debris -

A review with particular emphasis on Australian tree species. Australian Journal of

Botany 51(1), 27-37.

MacNally R, Parkinson A, Horrocks G, Young M (2002) Current loads of coarse woody

debris on Southeastern Australian floodplains: Evaluation of change and implications for

restoration. Restoration Ecology 10, 627-635.

Magoulick DD, Kobza RM (2003) The role of refugia for fishes during drought: a review

and synthesis. Freshwater Biology 48(7), 1186-1198.

Maheshwari BL, Walker KF, McMahon TA (1995) Effects of regulation on the flow

regime of the River Murray, Australia. Regulated Rivers: Research and Management 10,

15-38.

Manahan SE (1994) 'Environmental Chemistry.' 6th edn. (Lewis Publishers: London)

Mann CJ, Wetzel RG (1995) Dissolved organic carbon and its utilization in a riverine

wetland ecosystem. Biogeochemistry 31(2), 99-120.

Marchant R (1982) Seasonal variation in the macroinvertebrate fauna of billabongs along

Magela Creek, Northern Territory. Australian Journal of Marine and Freshwater

Research 33(2), 329-342.

Margules and Partners, P and J Smith Ecological Consultants, Murray-Darling Basin

Commission (Australia), Victorian Department of Conservation Forests and Lands

(1990) Riparian vegetation of the River Murray / report prepared by Margules and

Partners Pty. Ltd., P. and J. Smith Ecological Consultants and Department of

Conservation, Forests and Lands Murray-Darling Basin Commission, Canberra, A.C.T.

Margules and Partners Pty Ltd, P and J Smith Ecological Consultants, Department of

Conservation Forests and Lands (1990) Riparian vegetation of the River Murray: A

summary. Murray-Darling Basin Commission (Australia), Canberra.

280

Marschner B, Kalbitz K (2002) Controls on the availability and biodegradability of

dissolved organic matter in soils. Geoderma 113, 211-235.

Martin JP, Haider K, Saiz-Jimenez C (1974) Sodium amalgam reductive degradation of

fungal and model phenolic polymers, soil humic acids, and simple phenolic compounds.

Soil Science Society of America Journal 38(5), 760-765.

Martin KJ, Rygiewicz PT (2005) Fungal-specific PCR primers developed for analysis of

the ITS region of environmental DNA extracts. BMC Microbiology 5, 28-38.

Mayor JR, Schuur EAG, Henkel TW (2009) Elucidating the nutritional dynamics of

fungi using stable isotopes. Ecology Letters 12(2), 171-183.

McCarthy B, Conallin A, D'Santos P, Baldwin D (2006) Acidification, salinization and

fish kills at an inland wetland in south-eastern Australia following partial drying.

Ecological Management & Restoration 7(3), 218-223.

McLatchy GP, Reddy KR (1998) Regulation of organic matter decomposition and

nutrient release in a wetland soil. Journal of Environmental Quality 27(5), 1268-1274.

McMahon R, Hunter RD, Russell-Hunter WD (1974) Variation in aufwuchs at six

freshwater habitats in terms of carbon biomass and of carbon: Nitrogen ratio.

Hydrobiologia 45(4), 391-404.

McNeil DG, Closs GP (2007) Behavioural responses of a south-east Australian

floodplain fish community to gradual hypoxia. Freshwater Biology 52(3), 412-420.

McVeigh HP, Munro J, Embley TM (1996) Molecular evidence for the presence of novel

actinomycete lineages in a temperate forest soil. Journal of Industrial Microbiology and

Biotechnology 17(3), 197-204.

Medeiros AO, Pascoal C, Graca MAS (2009a) Diversity and activity of aquatic fungi

under low oxygen conditions. Freshwater Biology 54, 142-149.

Medeiros AO, Pascoal C, GraÇA MAS (2009b) Diversity and activity of aquatic fungi

under low oxygen conditions. Freshwater Biology 54(1), 142-149.

Melillo J, Aber J, Linkins A, Ricca A, Fry B, Nadelhoffer K (1989) Carbon and nitrogen

dynamics along the decay continuum: Plant litter to soil organic matter. Plant and Soil

115(2), 189-198.

Melillo JM, Aber JD, Muratore JF (1982) Nitrogen and lignin control of hardwood leaf

litter decomposition dynamics. Ecology 63(3), 621-626.

Melillo JM, McGuire AD, Kicklighter DW, Berrien Moore III, Vorosmarty CJ, Schloss

AL (1993) Global climate change and terrestrial net primary production. Nature 363,

234-240.

Melillo JM, Naiman RJ, Aber JD, Linkins AE (1984) Factors controlling mass loss and

nitrogen dynamics of plant litter decaying in northern streams. Bulletin of Marine

Science 35, 341-356.

281

Menge BA, Sutherland JP (1987) Community regulation: Variation in disturbance,

competition, and predation in relation to environmental stress and recruitment. The

American Naturalist 130(5), 730-757.

Merritt RW, Cummins KW (1978) 'An introduction to the aquatic insects of North

America.' (Kendall/Hunt Publishing Company: Dubuque, Iowa) 441

Meyer JL (1980) Dynamics of phosphorus and organic matter during leaf decomposition

in a forest stream. Oikos 34(1), 44-53.

Meyer JL (1994) The microbial loop in flowing waters. Microbial Ecology 28(2), 195-

199.

Meyer SLF, Carta LK, Rehner SA (2005) Morphological variability and molecular

phylogeny of the nematophagous fungus Monacrosporium drechsleri. Mycologia 97(2),

405-415.

Michell AJ (1992) FTIR spectroscopic studies of the reactions of wood and of lignin

model compounds with inorganic agents. Wood Science and Technology 27(1), 69-80.

Miles CJ, Brezonik PL (1981) Oxygen consumption in humic-colored waters by a

photochemical ferrous-ferric catalytic cycle. Environmental Science & Technology 15(9),

1089-1095.

Mille-Lindblom C, Fischer H, J. Tranvik L (2006a) Antagonism between bacteria and

fungi: substrate competition and a possible tradeoff between fungal growth and tolerance

towards bacteria. Oikos 113(2), 233.

Mille-Lindblom C, Fischer H, Tranvik LJ (2006b) Litter-associated bacteria and fungi - a

comparison of biomass and communities across lakes and plant species. Freshwater

Biology 51(4), 730-741.

Mille-Lindblom C, Tranvik LJ (2003) Antagonism between bacteria and fungi on

decomposing aquatic plant litter. Microbial Ecology 45(2), 173-182.

Mille-Lindblom C, Wachenfeldt Ev, Tranvik LJ (2004) Ergosterol as a measure of living

fungal biomass: persistence in environmental samples after fungal death. Journal of

Microbiological Methods 59, 253-262. [In English]

Miller LM, Dumas P (2006) Chemical imaging of biological tissue with synchrotron

infrared light. Biochimica et Biophysica Acta 1758(7), 846-57.

Mitchell D (1992) Floodplain wetlands of the Murray-Darling Basin: Management issues

and challenges. In 'Murray-Darling Basin Floodplain Wetlands Management. ' (Murray-

Darling Basin Commission: Albury, NSW)

Mitsch WJ, Gosselink JG (1993a) Biogeochemistry of wetlands. In 'Wetlands.' Second

edn. pp. 114-163. (Van Nostrand Reinhold: New York)

282

Mitsch WJ, Gosselink JG (1993b) Riparian wetlands. In 'Wetlands.' Second edn. (Van

Nostrand Reinhold: New York)

Mitsch WJ, Gosselink JG (Eds) (2000) 'Wetlands (Third edn).' (John Wiley and Sons

Inc.: New York)

Miyata N, Tani Y, Iwahori K, Soma M (2004) Enzymatic formation of manganese oxides

by an Acremonium-like hyphomycete fungus, strain KR21-2. FEMS Microbiology

Ecology 47(1), 101-109.

Mo Bio Laboratories I (2003) UltraCleanTM

Soil DNA Iswolation Kit Instruction

Manual. Mo Bio Laboratories, Inc., No. Version 10292003, Solana Beach, California,

USA.

Moller J, Miller M, Kjoller A (1999) Fungal-bacterial interaction on beech leaves:

influence on decomposition and dissolved organic carbon quality. Soil Biology and

Biochemistry 31(3), 367-374.

Money NP (1998) Why oomycetes have not stopped being fungi. Mycol Res 102(6), 767-

768.

Moorhead DL, Reynolds JF (1992) Modelling contributions of decomposer fungi in

nutrient cycling. In 'The Fungal Community: Its organisation and Role in the Ecosystem.

Vol. 9.' Second edn. (Eds GC Carroll and DT Wicklow) pp. 691-714. (Marcel Dekker,

Inc.: New York)

Moorhead DL, Sinsabaugh RL (2006) A theoretical model of litter decay and microbial

interaction. Ecological Monographs 76(2), 151-174.

Moran MA, Benner R, Hodson RE (1989) Kinetics of microbial degradation of vascular

plant material in two wetland ecosystems. Oecologia 79(2), 158-167.

Moran MA, Zepp RG (1997) Role of photoreactions in the formation of biologically

labile compounds from dissolved organic matter. Limnology and Oceanography 42(6),

1307-1316.

Morris SJ, Friese CF, Allen MF (2007) Disturbance in natural ecosystems: Scaling from

fungal diversity to ecosystem functioning. In 'The Mycota IV: Environmental and

Microbial Relationships. Vol. IV.' Second edn. (Eds CP Kubicek and IS Druzhinina) pp.

350. (Springer-Verlag: Berlin)

Mozaffar Z, Weete J (1993) Purification and properties of an extracellular lipase from

Pythium ultimum. Lipids 28(5), 377-382.

Mulholland PJ (1981) Organic carbon flow in a swamp-stream ecosystem. Ecological

Monographs 51(3), 307-322.

Mulholland PJ, Kuenzler EJ (1979) Organic carbon export from upland and forested

wetland watersheds. Limnology and Oceanography 24(5), 960-966.

283

Nagy LA, MaCauley BJ (1982) Eucalyptus leaf-litter decomposition: Effects of relative

humidity and substrate moisture content. Soil Biology & Biochemistry 14, 233-236.

Nawawi A (1985) Basidiomycetes with branched, water-borne conidia. Botanical

Journal of the Linnean Society 91(1-2), 51-60.

Nechwatal J, Wielgoss A, Mendgen K (2008) Diversity, host, and habitat specificity of

Oomycete communities in declining reed stands (Phragmites australis) of a large

freshwater lake. Mycological Research 112(6), 689-696.

Neckles HA, Neill C (1994) Hydrologic control of litter decomposition in seasonally

flooded prairie marshes. Hydrobiologia 286, 155-165.

Neher D (1999) Soil community composition and ecosystem processes: Comparing

agricultural ecosystems with natural ecosystems. Agroforestry Systems 45(1), 159-185.

Newell S, Hicks R (1982) Direct-count estimates of fungal and bacterial biovolume in

dead leaves of smooth cordgrass (Spartina alterniflora Loisel.). Estuaries and Coasts

5(4), 246-260.

Newell SY (1992) Estimating fungal biomass and productivity in decomposing litter. In

'The Fungal Community: It's organisation and role in the Ecosystem.' (Eds GC Carroll

and DT Wicklow) pp. 521-561. (Marcel Dekker, Inc.: New York)

Newell SY (1995) Minimizing ergosterol loss during preanalytical handling and shipping

of samples of plant litter. Applied and Environmental Microbiology 61(7), 2794-2797.

Newell SY (2001) Fungal biomass and productivity in standing-decaying leaves of black

needlerush (Juncus roemerianus). Marine and Freshwater Research 52(2), 249-255.

Newell SY, Bärlocher F (1993) Removal of fungal and total organic matter from

decaying cordgrass leaves by shredder snails. Journal of Experimental Marine Biology

and Ecology 171(1), 39-49.

Newell SY, Moran MA, Wicks R, Hodson RE (1995) Productivities of microbial

decomposers during early stages of decomposition of leaves of a freshwater sedge.

Freshwater Biology 34, 135-148.

Newell SY, Statzell-Tallman A (1982) Factors for conversion of fungal biovolume

values to biomass, carbon and nitrogen: Variation with mycelial ages, growth conditions,

and strains of fungi from a salt marsh Oikos 39(2), 261-268

Newman RM (1991) Herbivory and detritivory on freshwater macrophytes by

invertebrates: A review. Journal of the North American Benthological Society 10(2), 89-

114.

Nichols-Orians C (1991) Differential effects of condensed and hydrolyzable tannin on

polyphenol oxidase activity of attine symbiotic fungus. Journal of Chemical Ecology

17(9), 1811 - 1819

284

Nielsen DL, Brock MA, Rees G, Baldwin DS (2003) Effects of increasing salinity on

freshwater ecosystems in Australia Australian Journal of Botany 51, 655-665.

Nielsen DL, Hillman TJ, Smith FJ, Shiel RJ (2002) The influence of seasonality and

duration of flooding on zooplankton in experimental billabongs. River Research and

Applications 18(3), 227-237.

Nikolcheva L, Bärlocher F (2004) Taxon-specific fungal primers reveal unexpectedly

high diversity during leaf decomposition in a stream. Mycological Progress 3(1), 41-49.

Nikolcheva LG, Bärlocher F (2005) Seasonal and substrate preferences of fungi

colonizing leaves in streams: traditional versus molecular evidence. Environmental


Nikolcheva LG, Cockshutt AM, Bärlocher F (2003) Determining diversity of freshwater

fungi on decaying leaves: comparison of traditional and molecular approaches. Applied

and Environmental Microbiology 69, 2548-2554.

Ning NSP, Nielsen DL, Hillman TJ, Suter PJ (2008) Evaluation of a new technique for

characterizing resting stage zooplankton assemblages in riverine slackwater habitats and

floodplain wetlands. Journal Of Plankton Research 30(4), 415-422.

Nout MJR, Bonants-Van Laarhoven TMG, Jongh Pd, Koster PGd (1987) Ergosterol

content of Rhizopus oligosporus NRRL 5905 grown in liquid and solid substrates.

Applied Microbiology and Biotechnology 26, 456-461.

Noy-Meir I (1973) Desert ecosystems, environments and producers. Annual Review of

Ecology and Systematics 4, 25-51.

O'Connell AM, Grove TS (1987) Seasonal variation in C2H2 reduction (N2-fixation) in

the litter layer of eucalypt forests of South-Western Australia. Soil Biology and

Biochemistry 19(2), 135-142.

O'Connell M (1998) Dissolved organic carbon from components of floodplain litter:

release and bioavailability. Honours Thesis, Charles Sturt university,

O'Connell M, Baldwin DS, Robertson AI, Rees G (2000) Release and bioavailability of

dissolved organic matter from floodplain litter: influence of origin and oxygen levels.

Freshwater Biology 45, 333-342.

O'Connor NA (1992) Quantification of submerged wood in a lowland Australian stream

system. Freshwater Biology 27, 387-395.

Oda Y, Saito K, Yamauchi H, Mori M (2002) Lactic acid fermentation of potato pulp by

the Fungus Rhizopus oryzae. Current Microbiology 45(1), 1-4.

Odum WE, Kirk PW, Zieman JC (1979) Non-protein nitrogen compounds associated

with particles of vascular plant detritus Oikos 32(3), 363-367

Ogle K, Reynolds JF (2004) Plant responses to precipitation in desert ecosystems:

integrating functional types, pulses, thresholds and delays. Oecologia 141, 282-294.

285

Ohmart CP, Edwards PB (1991) Insect herbivory on Eucalyptus. Annual Review of

Entomology 36, 637-657.

Olson FCW (1950) Quantitative estimates of filamentous algae. Transactions of the

American Microscopy Society 59, 272-279.

Orsborne JL (1987) Effects of temperature, relative humidity and substratum moisture

content on the decomposition and colonisation of eucalypt leaf litter by fungi. PhD

Thesis, La Trobe University, Bundoora

Pace ML, Cole JJ, Carpenter SR, Kitchell JF, Hodgson JR, Van de Bogert MC, Bade DL,

Kritzberg ES, Bastviken D (2004) Whole-lake carbon-13 additions reveal terrestrial

support of aquatic food webs. Nature 427, 240-243.

Padgett DE, Kendrick AS, Hearth JH, Wm DW (1988) Influence of salinity, temperature,

and nutrient availability on the respiration of saprolegniaceous fungi (Oomycetes).

Ecography 11(2), 119-126.

Parish F, Sirin A, Charman D, Joosten H, Minayeva T, Silvius M, Stringer L (2008)

Assessment on Peatlands, Biodiversity and Climate Change: Main Report. Global

Environment Centre, Kuala Lumpur and Wetlands International, Wageningen.

Park HC, Sorenson WG, Davis RJ (1978) Aquatic Oomycetes in farm ponds in Bryan

County, Oklahoma. Proceedings of the Oklahoma Academy of Science 58, 48-53.

Parkinson SM, Wainwright M, Killham K (1989) Observations on oligotrophic growth of

fungi on silica gel. Mycological Research 93(4), 529-534.

Pasanen A-L, Yli-Pietila K, Pasanen P, Kalliokoski P, Tarhanen J (1999) Ergosterol

content in various fungal species and biocontaminated building materials. Applied and

Environmental Microbiology 65(1), 138-142.

Pascoal C, Cassio F (2004) Contribution of fungi and bacteria to leaf litter decomposition

in a polluted river. Applied and Environmental Microbiology 70(9), 5266-5273.

Pascoal C, Cássio F, Marcotegui A, Sanz B, Gomes P (2005) Role of fungi, bacteria, and

invertebrates in leaf litter breakdown in a polluted river. Journal of the North American

Benthological Society. 24(4), 784-797.

Pattee E, Chergui H (1995) The application of habitat templets and traits to

Hyphomycete fungi in a mid-European river system. Freshwater Biology 33, 525-539.

Paustian K, Schnurer J (1987) Fungal growth response to carbon and nitrogen limitation:

A theoretical model. Soil Biology & Biochemistry 19(5), 613-620.

Pereira JS, Kozlowski TT (1976) Leaf anatomy and water relations of Eucalyptus

camaldulensis and E. globulus seedlings. Canadian Journal of Botany 54(24), 2868-

2880.

286

Pettitt TR, Wakeham AJ, Wainwright MF, White JG (2002) Comparison of serological,

culture, and bait methods for detection of Pythium and Phytophthora zoospores in water.

Plant Pathology 51(6), 720-727.

Pickering AD, Willoughby LG, McGrory CB (1979) Fine structure of secondary

zoospore cyst cases of Saprolegnia isolates from infected fish. Transactions of the

British Mycological Society 72(3), 427-436.

Pickett S, White P (Eds) (1985) 'The ecology of natural disturbance and patch dynamics.'

(Academic Press: New York, NY, USA)

Pidgeon RW, Cairns SC (1981) Decomposition and colonisation by invertebrates of

native and exotic leaf material in a small stream in New England (Australia).

Hydrobiologia 77, 113-127.

Pieczynska E (1993) Detritus and nutrient dynamics in the shore zone of lakes: a review.

Hydrobiologia 251, 49-58. [In English]

Pind A, Freeman C, Lock MA (1994) Enzymic degradation of phenolic materials in

peatlands — measurement of phenol oxidase activity. Plant and Soil 159(2), 227-231.

Pomeroy L, Wiebe W (1988) Energetics of microbial food webs. Hydrobiologia 159(1),

7-18.

Porter D, Farnham WF (1986) Mycaureola dilseae, a marine Basidiomycete parasite of

the red alga, Dilsea carnosa. Transactions of the British Mycological Society 87(4), 575-

582.

Pozo J, Basaguren A, Elosegui A, Molinero J, Fabre E, Chauvet E (1998) Afforestation

with Eucalyptus globulus and leaf litter decomposition in streams of northern Spain.

Hydrobiologia 373/374, 101-109.

Premdas PD, Kendrick B (1991) Seasonal sporulation of some aero-aquatic fungi. Archiv

für Hydrobiologie 122(4), 479-482.

Presland AJ (1982) Litter production and decomposition from an overstorey of

Eucalyptus spp. on two catchments in the New England region of New South Wales.

Austral Ecology 7(2), 171-180.

Pressey RL (1986) 'Wetlands of the River Murray below Lake Hume : a summary.'

(River Murray Commission: Canberra) 13

Pressey RL (1990) Wetlands. In 'The Murray.' (Eds N Mackay and D Eastburn).

(Murray-Darling Basin Commission: Canberra, Australia)

Price IP, Talbot PHB (1966) An aquatic hyphomycete in a lignicolous habitat. Australian

Journal of Botany 14, 19-23.

Qiu S, McComb AJ, Bell RW, Davis JA (2003) Nutrient response to soil and litter

metabolic activity in a transect across a seasonal wetland. Marine and Freshwater

Research 54(3), 243-252.

287

Qiu S, McComb AJ, Bell RW, Davis JA (2005) Response of soil microbial activity to

temperature, moisture, and litter leaching on a wetland transect during seasonal refilling.

Wetlands Ecology and Management 13(1), 43-54.

Quinn GP, Hillman TJ, Cook R (2000) The response of macroinvertebrates to inundation

in floodplain wetlands: A possible effect of river regulation? Regulated Rivers: Research

and Management 16, 469-477.

Rand TG, Munden D (1992) Enzyme involvement in the invasion of brook char,

Salvelinus fontinalis (Mitchill), eggs by Saprolegnia diclina (Oomycotina:

Saprolegniaceae). Journal of fish diseases 15 (1), 91

Ravelet C, Grosset C, Alary J (2001) Quantitation of ergosterol in river sediment by

liquid chromatography. Journal of Chromatographic Science 39(6), 239-242.

Raven PH, Evert RF, Eichhorn SE (1992) 'Biology of Plants.' Fifth edn. (Worth

Publishers: New York) 791

Reddy KR, DeLaune RD (2008) 'Biogeochemistry of wetlands: science and applications.'

(Taylor & Francis: Boca Raton)

Reddy KR, Patrick WH (1975) Effect of alternate aerobic and anaerobic conditions on

redox potential, organic matter decomposition and nitrogen loss in a flooded soil. Soil

Biology and Biochemistry 7(2), 87-94.

Rees GN, Baldwin DS, Watson GO, Perryman S, Nielsen DL (2004) Ordination and

significance testing of microbial community composition derived from terminal

restriction fragment length polymorphisms: application of multivariate statistics. Antonie

van Leeuwenhoek 86(4), 339-347.

Reffner JA, Martoglio PA, Williams GP (1995) FTIR microscope analysis with

synchrotron radiation: The microscope optics and system performance. Rev. Sci. Instr.

66.

Reid DJ, Lake PS, Quinn GP, Reich P (2008) Association of reduced riparian vegetation

cover in agricultural landscapes with coarse detritus dynamics in lowland streams.

Marine and Freshwater Research 59(11), 998-1014.

Reid DJ, Quinn GP, Lake PS, Reich P (2008) Terrestrial detritus supports the food webs

in lowland intermittent streams of south-eastern Australia: a stable isotope study.


Reid M, Brooks J (2000) Detecting effects of environmental water allocations in

wetlands of the Murray–Darling Basin, Australia. Regulated Rivers: Research and

Management 16(5), 479-496.

Reid M, Ogden R (2009) Factors affecting diatom distribution in floodplain lakes of the

southeast Murray Basin, Australia and implications for palaeolimnological studies.

Journal of Paleolimnology 41(3), 453-470.

288

Reid M, Quinn G (2004) Hydrologic regime and macrophyte assemblages in temporary

floodplain wetlands: Implications for detecting responses to environmental water

allocations. Wetlands 24(3), 586-599.

Reith F, Drake HL, Kusel K (2002) Anaerobic activities of bacteria and fungi in

moderately acidic conifer and deciduous leaf litter. FEMS Microbiology Ecology 41(1),

27-35.

Reynolds JF, Kemp PR, Ogle K, Fernandez RJ (2004) Modifying the 'pulse-reserve'

paradigm for deserts of North America: precipitation pulses, soil water, and plant

responses. Oecologia 141, 194-210.

Rice DL (1982) The detritus nitrogen problem: New observations and perspectives from

organic geochemistry. Marine Ecology - Progress Series 9, 153-162.

Rinklebe J, Langer U (2006) Microbial diversity in three floodplain soils at the Elbe

River (Germany). Soil Biology and Biochemistry 38(8), 2144-2151.

Ritz K (2007) The plate debate: Cultivable communities have no utility in contemporary

environmental microbial ecology. FEMS Microbiology Ecology 60(3), 358-362.

Roache MC, Bailey PC, Boon PI (2006) Effects of salinity on the decay of the freshwater

macrophyte, Triglochin procerum. Aquatic Botany 84(1), 45-52.

Robert P, Marquis M, Barron C, Guillon F, Saulnier L (2005) FT-IR investigation of cell

wall polysaccharides from cereal grains. Arabinoxylan infrared assignment. Journal of

Agricultural and Food Chemistry 53(18), 7014-7018.

Roberts J (2006) Field Assessment. Contribution to Stage 1 Report on the project

“Identification of wetlands for potential rehabilitation and water savings within the River

Murray system, NSW”. . Report to NSW Murray Wetlands Working Group., No. JR

07/2006, Canberra.

Roberts J, Ganf GG (1986) Annual production of Typha orientalis Presl. in inland

Australia. Australian Journal of Marine and Freshwater Research 37, 659-668.

Roberts L (1899) Experimental note on the ferments of the ringworm fungi. British

Medical Journal 7 January 1899, 13-14.

Roberts RE (1963) A study of the distribution of certain members of the Saprolegniales.


Robertson AI, Bacon P, Heagney G (2001) The responses of floodplain primary

production to flood frequency and timing. Journal of Applied Ecology 38, 126-136.

Robertson AI, Bunn SE, Boon PI, Walker KF (1999) Sources, sinks and transformations

of organic carbon in Australian floodplain rivers. Marine and Freshwater Research 50,

813-829.

289

Robertson AI, Healey, King (1997) Experimental manipulations of the biomass of

introduced carp (Cyprinus carpio) in billabongs. II. Impacts on benthic properties and

processes. Marine and Freshwater Research 48, 445-54.

Robertson AI, Klomp N, Lunt ID (1997) 'Land-water linkages in floodplain river

systems: the influence of domestic stock.' (Elsevier Science: Oxford)

Robinson CH, Borisova OB, Callaghan TV, Lee JA (1996) Fungal hyphal length in litter

of Dryas octopetala in a high-Arctic polar semi-desert, Svalbard. Polar Biology 16(1),

71-74.

Rodriguez RJ, Cullen D, Kurtzman CP, Khachatourians GG, Hegedus DD (2004)

Molecular methods for discriminating taxa, monitoring species, and assessing fungal

diversity. In 'Biodiversity of Fungi: Inventroy and monitoring methods.' pp. 77-102.

(Elsevier Academic Press: London)

Romani AM, Fischer H, Mille-Lindblom C, Tranvik LJ (2006) Interactions of bacteria

and fungi on decomposing litter: Differential extracellular enzyme activities. Ecology

87(10), 2559-2569.

Rosset J, Bärlocher F, Oertli J (1982) Decomposition of conifer needles and deciduous

leaves in two Black Forest and two Swiss Jura streams. Internationale Revue der

Gesamten Hydrobiologie und Hydrographie 67(5), 695-711.

Ryder DS, Horwitz P (1995) Seasonal water regimes and leaf litter processing in a

wetland on the Swan Coastal Plain, Western Australia. Marine and Freshwater Research

46(7), 1077-1084.

Sabetta L, Costantini ML, Maggi O, Persiani AM, Rossi L (2000) Interactions between

detritivores and microfungi during the leaf detritus decomposition in a volcanic lake

(Lake Vico - central Italy). Hydrobiologia 439(1-3), 49-60.

Safe S (1973) The effect of the environment on the free and hydrosoluble sterols of

Mucor rouxii. Biochimica et Biophysica Acta 326, 471-475.

Sait L, Galic M, Strugnell RA, Janssen PH (2003) Secretory antibodies do not affect the

composition of the bacterial microbiota in the terminal ileum of 10-week-old mice.

Applied and Environmental Microbiology 69(4), 2100-2109.

Saiz-Jimenez C, Ortega-Calvo JJ, de Leeuw JW (1995) The chemical structure of fungal

melanins and their possible contribution to black stains in stone monuments. Science of

The Total Environment 167(1-3), 305-314.

Sampaio A, Cortes R, Leão C (2004) Yeast and macroinvertebrate communities

associated with leaf litter decomposition in a second order stream. International Review

of Hydrobiology 89(5-6), 453-466.

Sanders PF, Anderson JM (1979) Colonization of wood blocks by aquatic hyphomycetes.


290

Sanders RW, Porter KG, Bennett SJ, DeBiase AE (1989) Seasonal patterns of

bacterivory by flagellates, ciliates, rotifers, and cladocerans in a freshwater planktonic

community. Limnology and Oceanography 34(4), 673-687.

Sati SC, Belwal M (2005) Aquatic hyphomycetes as endophytes of riparian plant roots.

Mycologia 97(1), 45-49.

Scheffer M (1998) Vegetation. In 'Ecology of shallow lakes. Vol. 22.' pp. 210-286.

(Kluwer Academic Publishers: Dordrecht, The Netherlands)

Schlesinger WH, Melack JM (1981) Transport of organic carbon in the world's rivers.

Tellus 33, 172-187.

Schoenlein-Crusius IH, Pires-Zottarelli CLA, Milanez AI, Humphreys RD (1999)

Interaction between the mineral content and the occurrence number of aquatic fungi in

leaves submerged in a stream in the Atlantic rainforest, São Paulo, Brazil. Revista

Brasileira de Botânica 22, 133-139.

Scholz O, Boon PI (1993a) Biofilm development and extracellular enzyme activities on

wood in billabongs of south-eastern Australia. Freshwater Biology 30, 359-368.

Scholz O, Boon PI (1993b) Biofilms on submerged river red gum (Eucalyptus

camaldulensis Dehnh. Myrtaceae) wood in billabongs: an analysis of bacterial

assemblages using phospholipid profiles. Hydrobiologia 259, 169-178.

Schulze DJ, Walker KF (1997) Riparian eucalypts and willows and their significance for

aquatic invertebrates in the River Murray, South Australia. Regulated Rivers: Research

and Management 13, 557-577.

Sedell JR, Dahm CN (1990) Spatial and temporal scales of dissolved organic carbon in

streams and rivers. In 'Organic Acids in Aquatic Ecosystems.' (Eds EM Perdue and ET

Gjessing) pp. 261-279. (John Wiley and Sons Ltd.: Chichester)

Seo DC, DeLaune RD (2010) Effect of redox conditions on bacterial and fungal biomass

and carbon dioxide production in Louisiana coastal swamp forest sediment. Science of

The Total Environment 408(17), 3623-3631.

Seoudi R, Nada AMA, Elmongy SA, Hamed SS (2005) Fourier transform infrared

spectroscopic and AC electrical conductivity studies of chitin and its derivatives. Journal

of Applied Polymer Science 98(2), 936-943.

Serrano L (1992) Leaching from vegetation of soluble polyphenolic compounds, and

their abundance in temporary ponds in the Doñana National Park (SW Spain).

Hydrobiologia 229, 43-50.

Serrano L, Boon PI (1991) Effect of polyphenolic compounds on alkaline phosphatase

activity: its implication for phosphorus regeneration in Australian waters. Archiv für

Hydrobiologie 123, 1-19.

Shearer CA (1992) The role of woody debris. In 'The Ecology of Aquatic

Hyphomycetes.' Ed. F Barlocher) pp. 77-98. (Springer-Verlag: Berlin)

291

Shearer CA, Descals E, et al. (2007) Fungal biodiversity in aquatic habitats. Biodiversity

and Conservation 16, 49-67.

Shearer CA, von Bodman SB (1983) Patterns of occurrence of ascomycetes associated

with decomposing twigs in a midwestern stream. Mycologia 75(May/June), 518-530.

Shearer CA, Webster J (1985) Aquatic hyphomycete communities in the River Teign I.

Longitudinal distribution patterns. Transactions of the British Mycological Society 84(3),

489-501.

Sheldon F, Boulton AJ, Puckridge JT (2002) Conservation value of variable

connectivity: aquatic invertebrate assemblages of channel and floodplain habitats of a

central Australian arid-zone river, Cooper Creek Biological Conservancy 103(1), 13-31.

Sherr E, Sherr B (2002) Significance of predation by protists in aquatic microbial food

webs. Antonie van Leeuwenhoek 81(1), 293-308.

Sherr EB, Sherr BF (1991) Planktonic microbes: Tiny cells at the base of the ocean's

food webs. Trends in Ecology & Evolution 6(2), 50-54.

Shiel RJ (1983) Rotifer communities of billabongs in northern and south-eastern

Australia. Hydrobiologia 104(1), 41-47.

Shilla D, Aseada T, Fujino T, Sanderson B (2006) Decomposition of dominant

submerged macrophytes: implications for nutrient release in Myall Lake, NSW,

Australia. Wetlands Ecology and Management 14, 427-433.

Shimada M, Akamtsu Y, Tokimatsu T, Mii K, Hattori T (1997) Possible biochemical

roles of oxalic acid as a low molecular weight compound involved in brown-rot and

white-rot wood decays. Journal of Biotechnology 53(2-3), 103-113.

Shine R (1987) Intraspecific variation in thermoregulation, movements and habitat use

by Australian blacksnakes, Pseudechis porphyriacus (Elapidae). Journal of Herpetology

21(3), 165-177.

Silvar C, Diaz J, Merino F (2005 ) Real-time polymerase chain reaction quantification of

Phytophthora capsici in different pepper genotype Phytopathology 95(12), 1423-1429.

[In English ]

Silverstein RM, Bassler GC, Morrill TC (1974) 'Spectrometric identification of organic

compounds.' Third edn. (John Wiley & Sons: Brisbane) 340

Singer R (1988) The role of fungi in periodically inundated Amazonian forests. Vegetatio

78, 27-30.

Singh BK, Nazaries L, Munro S, Anderson IC, Campbell CD (2006) Use of multiplex

terminal restriction fragment length polymorphism for rapid and simultaneous analysis of

different components of the soil microbial community. Applied and Environmental

Microbiology 72(11), 7278-7285.

292

Singhal RN, Jeet S, Davies RW (1987) Experimental transmission of Saprolegnia and

Achlya to fish. Aquaculture 64(1), 1-7.

Sinsabaugh RL, Gallo ME, Lauber C, Waldrop MP, Zak DR (2005) Extracellular

enzyme activities and soil organic matter dynamics for northern hardwood forests

receiving simulated nitrogen deposition. Biogeochemistry 75(2), 201-215.

Sinsabaugh RL, Moorhead DL, Linkins AE (1994) The enzymic basis of plant litter

decomposition: emergence of an ecological process. Applied Soil Ecology 1, 97-111.

Smith D (2010) Environmental data for Quat Quatta East lagoon, 2008-2009. In. '

Environmental data for Quat Quatta East lagoon, 28/10/2008-22/4/2009 edn. Ed. J

Williams): Albury-Wodonga, Victoria)

Smith WK, Gao W, Steltzer H, Wallenstein MD, Tree R (2009) Moisture availability

influences the effect of ultraviolet-B radiation on leaf litter decomposition. Global

Change Biology 16(1), 484-495.

Söderhäll K, Dick MW, Clark G, Fürst M, Constantinescu O (1991) Isolation of

Saprolegnia parasitica from the crayfish Astacus leptodactylus. Aquaculture 92, 121-

125.

Sorrell BK, Boon PI (1992a) Biogeochemistry of billabong sediments. II. Seasonal

variations in methane production. Freshwater Biology 27(3), 435-445.

Sorrell BK, Boon PI (1992b) Biogeochemistry of billabong sediments. II. Seasonal

variations in methane production. Freshwater Biology 27(3), 435-445.

Sridhar KR, Bärlocher F (1992) Endophytic aquatic hyphomycetes of roots of spruce,

birch and maple. Mycological Research 96(4), 305-308.

Sridhar KR, Bärlocher F (2000) Initial colonization, nutrient supply, and fungal activity

on leaves decaying in streams. Applied and Environmental Microbiology 66(3), 1114-

1119.

Sridhar KR, Duarte S, Cássio F, Pascoal C (2009) The role of early fungal colonizers in

leaf-litter decomposition in Portuguese streams impacted by agricultural runoff.

International Review of Hydrobiology 94(4), 399-409.

Stelzer RS, Heffernan J, Likens GE (2003) The influence of dissolved nutrients and

particulate organic matter quality on microbial respiration and biomass in a forest stream.

In. Vol. 48'. pp. 1925-1937)

Stone C, Bacon P (1994) Relationships among moisture stress, insect herbivory, foliar

cineole content and the growth of river red gum Eucalyptus camaldulensis. Journal of

Applied Ecology 31, 604-612.

Stone C, Bacon PE (1995) Leaf dynamics and insect herbivory in a Eucalyptus

camaldulensis forest under moisture stress. Australian Journal of Ecology 20(4), 473-

481.

293

Suberkropp K (1984) Effect of temperature on seasonal occurrence of aquatic

hyphomycetes. Transactions of the British Mycological Society 82(1), 53-62.

Suberkropp K (1991a) Aquatic hyphomycete communities. In 'The fungal community.'

(Eds GC Carroll and DT Wicklow). (Marcel Dekker: New York)

Suberkropp K (1991b) Relationships between growth and sporulation of aquatic

hyphomycetes on decomposing leaf litter. Mycological Research 95(7), 843-850.

Suberkropp K (1992) Interactions with Invertebrates. In 'The Ecology of Aquatic

Hyphomycetes. Vol. 94.' Ed. F Barlocher) pp. 118-. (Springer-Verlag: Berlin)

Suberkropp K (1995) The influence of nutrients on fungal growth, productivity, and

sporulation during leaf breakdown in streams Canadian Journal of Botany 1(1).

Suberkropp K (1997) Annual production of leaf-decaying fungi in a woodland stream.


Suberkropp K (1998) Effect of dissolved nutrients on two aquatic hyphomycetes growing

on leaf litter. Mycological Research 102(8), 998-1002.

Suberkropp K (2001) Fungal growth, production and sporulation during leaf

decomposition in two streams. Applied and Environmental Microbiology 67(11), 5063-

5068.

Suberkropp K, Arsuffi TL, Anderson JP (1983) Comparison of degradative ability,

enzymatic activity, and palatability of aquatic hyphomycetes grown on leaf litter. Applied

and Environmental Microbiology 46(1), 237-244. [In English]

Suberkropp K, Chauvet E (1995) Regulation of leaf breakdown by fungi in streams:

Influences of water chemistry. Ecology 76(5), 1433-1445.

Suberkropp K, Gessner MO, Chauvet E (1993) Comparison of ATP and ergosterol as

indicators of fungal biomass associated with decomposing leaves in streams. Applied and

Environmental Microbiology 59(10), 3367-3372.

Suberkropp K, Godshalk GL, Klug MJ (1976) Changes in the chemical composition of

leaves during processing in a woodland stream. Ecology 57, 720-727. [In English]

Suberkropp K, Klug MJ (1976) Fungi and bacteria associated with leaves during

processing in a woodland stream. Ecology 57( 4), 707-719.

Suberkropp K, Klug MJ (1980) The maceration of deciduous leaf litter by aquatic

hyphomycetes. Canadian Journal of Botany 58, 1025-1031.

Suter PJ, Hawking JH (2002) Aquatic macroinvertebrates of the Murray River. Victorian

Naturalist 119, 186-200.

Suter SGA (2009) Aquatic fungi in an alpine stream of south-eastern Australia Honours

Thesis, La Trobe University, Albury-Wodonga

294

Swart HJ (1986) A preliminary survey of aquatic hyphomycetes in Victoria, Australia.

Transactions of the British Mycological Society 86(3), 497-501.

Swift MJ, Heal OW, Anderson JM (1979) The influence of the physico-chemical

environment on decomposition process. In 'Decomposition in terrestrial ecosystems.' pp.

220-266. (Blackwell Scientific Publications: Melbourne)

Szeghalmi A, Kaminskyj SGW, Gough KM (2007) A synchrotron FTIR

microspectroscopy investigation of fungal hyphae grown under optimal and stressed

conditions. Analytical and Bioanalytical Chemistry 387(5), 1779-1789.

Tan L-W, Shiel RJ (1993) Responses of billabong rotifer communities to inundation.

Hydrobiologia 255-256(1), 361-369.

Taylor BR, Parkinson D, Parsons WFJ (1989) Nitrogen and lignin content as predictors

of litter decay rates: A microcosm test. Ecology 70(1), 97-104.

Taylor DL, Bruns TD (1999) Community structure of ectomycorrhizal fungi in a Pinus

muricata forest: minimal overlap between the mature forest and resistant propagule

communities. Molecular Ecology 8(11), 1837-1850.

Terzi V, Morcia C, Faccioli P, Vale G, Tacconi G, Malnati M (2007) In vitro antifungal

activity of the tea tree (Melaleuca alternifolia) essential oil and its major components

against plant pathogens. Letters in Applied Microbiology 44 (6), 613-618.

Theodorou MK, Lowe SE, Trinci APJ (1992) Anaerobic fungi and the rumen ecosystem.

In 'The Fungal Community: Its Organization and Role in the Ecocsystem.' 2nd edn. (Eds

GC Carroll and DT Wicklow) pp. 43-72. (Marcel Dekker, Inc.: New York)

Thomas K (1992) The ecology of aquatic hyphomycetes in an Australian upland stream.

PhD Thesis, The Australian National University, Canberra

Thomas K (1996) Freshwater fungi. In 'Fungi of Australia. Vol. 1B - Introduction to

fungi in the environment.' (Eds K Mallet and C Grgurinovic) pp. 1-37. (CSIRO

PUBLISHING,

The Australian Biological Resources Study,: Canberra)

Thomas K, Chilvers GA, Norris RH (1989) Seasonal occurrence of conidia of aquatic

hyphomycetes (Fungi) in Lees Creek, Australian Capital Territory. Australian Journal of

Marine and Freshwater Research 40, 11-23.

Thomas K, Chilvers GA, Norris RH (1992) Aquatic hyphomycetes from different

substrates: Substrate preference and seasonal occurrence. Australian Journal of Marine

and Freshwater Research 43, 491-509.

Thompson IA, Huber DM, Guest CA, Schulze DG (2005) Fungal manganese oxidation

in a reduced soil. Environmental Microbiology 7(9), 1480-1487. [In English]

Thormann M, N., Currah R, S., Bayley S, E. (2003) Succession of microfungal

assemblages in decomposing peatland plants. Plant and Soil 250(2), 323-333.

295

Thormann MN (2006) Diversity and function of fungi in peatlands: A carbon cycling

perspective. Canadian Journal of Soil Science 86, 281-293.

Thornton DR (1963) The physiology and nutrition of some aquatic hyphomycetes.

Journal of General Microbiology 33, 23-31.

Thornton DR (1965) Amino acid analysis of fresh leaf litter and the nitrogen nutrition of

some aquatic hyphomycetes. Canadian Journal of Microbiology 11, 657-662.

Thorp JH, Delong MD (2002) Dominance of autochthonous autotrophic carbon in food

webs of heterotrophic rivers. Oikos 96(3), 543-550.

Thrane U, Anderson B, Frsvad JC, Smedsgaard J (2007) The exo-metabolome in

filamentous fungi. In 'Metabolomics. Vol. 18.' (Eds J Nielsen and MC Jewett) pp. 235-

252. (Springer-Verlag: Berlin, Heidelberg)

Thurman EM (1985) 'Organic geochemistry of natural waters.' (Martinus Nijhoff / Dr W.

Junk: Dordrecht, Netherlands) 497

Tien M, Kirk TK (1984) Lignin-degrading enzyme from Phanerochaete chrysosporium:

Purification, characterization, and catalytic properties of a unique H2O2-Requiring

Oxygenase. Proceedings of the National Academy of Sciences of the United States of

America 81(8), 2280-2284.

Toberman H, Freeman C, Evans C, Fenner N, Artz RRE (2008) Summer drought

decreases soil fungal diversity and associated phenol oxidase activity in upland Calluna

heathland soil. FEMS Microbiology Ecology 66(2), 426-436.

Townsend CR (1989) The patch dynamics concept of stream community ecology.

Journal of the North American Benthological Society 8(1), 36-50.

Tranvik LJ, van Hees PAW, Lundstrom US (2004) Abundance and functions of natural

organic matter species in soil and water. Aquatic Sciences 66(2), 149-150.

Trinci APJ, Davies DR, Gull K, Lawrence MI, Nielsen BB, Rickers A, Theodorou MK

(1994) Anaerobic fungi in herbivorous animals. Mycological Research 98, 129-152.

Trinder CJ, Johnson D, Artz RRE (2008) Interactions among fungal community

structure, litter decomposition and depth of water table in a cutover peatland. FEMS

Microbiology Ecology 64(3), 433 - 448.

Triska FJ, Sedell JR (1976) Decomposition of four species of leaf litter in response to

nitrate manipulation. Ecology 57(4), 783-792.

Valikhanov BH (1988) Cellulose and ligninolytic properties of freshwater fungi.

Ukrayins'Kyi Botanichnyi Zhurnal 45(6), 46-50.

Van Laere A (1994) Intermediary metabolism. In 'The Growing Fungus.' pp. 211-238.

(Springer Netherlands )

296

Van Ryckegem G, Gessner M, Verbeken A (2007) Fungi on leaf blades of Phragmites

australis in a brackish tidal marsh: Diversity, succession, and leaf decomposition.


Van Veen JA, Paul EA (1979) Conversion of biovolume measurements of soil

organisms, grown under various moisture tensions, to biomass and their nutrient content.

Applied and Environmental Microbiology 37(4), 686-692.

Van Wensem J, Van Straalen NM, Kooijman SALM (1997) Carbon and nitrogen fluxes

in decomposing leaf litter with microbial-detritivore interactions: model simulations

compared to microcosm ecotoxicity tests. Ecological Modelling 96, 175-189.

van West P (2006) Saprolegnia parasitica, an oomycete pathogen with a fishy appetite:

new challenges for an old problem. Mycologist 20(3), 99-104.

Vannote RL, Minshall GW, Cummins KW, Sedell JR, Cushing CE (1980) The river

continuum concept. Canadian Journal of Aquatic Sciences 37, 130-137.

Verma B, Robarts RD, Headley JV (2003) Seasonal changes in fungal production and

biomass on standing dead Scirpus lacustris litter in a northern prairie wetland. Applied

and Environmental Microbiology 69(2), 1043-1050. [In English]

Victorian State Government, EPA Victoria, Murray-Darling Basin Commission

(Australia), Waterwatch Victoria, Victorian Catchment Management Council, National

Landcare Program (2010) Victorian Water Resources Data Warehouse. In. Vol. 2010'.)

Voelker BM, Morel FoMM, Sulzberger B (1997) Iron redox cycling in surface waters:

Effects of humic substances and light. Environmental Science & Technology 31(4),

1004-1011.

Voglmayr H (1997) Helicodendron praetermissum sp. nov. and Spirosphaera carici-

graminis sp. nov., aero-aquatic fungi on monocotyledonous debris. Canadian Journal of

Botany 75, 1772-1777.

Volk CJ, Volk CB, Kaplan LA (1997) Chemical composition of biodegradable dissolved

organic matter in streamwater. Limnology and Oceanography 42(1), 39-44. [In English]

Voronin L (2008) Zoosporic fungi in freshwater ecosystems. Inland Water Biology 1(4),

341-346.

Wainwright M (1992) The Impact of Fungi on Environmental Biogeochemistry. In 'The

fungal community: Its organization and role in the ecosystem.' Second edn. (Eds GC

Carroll and DT Wicklow) pp. 601-618. (Marcel Dekker, Inc: New York)

Walker J (1996) Key to orders. In 'Fungi of Australia Vol. 1A.' pp. 67-120. (Australian

Biological Resources Study: Canberra)

Wang FL, Bettany JR (1995) Carbon and nitrogen losses from undisturbed soil columns

under short-term flooding conditions. Canadian Journal of Soil Science 75, 333-341.

297

Warren RAJ (1996) Microbial hydrolysis of polysaccharides. Annual Review of


Watkins SC, Quinn GP, Gawne B (2010) Changes in organic-matter dynamics and

physicochemistry, associated with riparian vegetation loss and river regulation in

floodplain wetlands of the Murray River, Australia. Marine and Freshwater Research

61(10), 1207-1217.

Webster J (1977) Seasonal observations on aquatic hyphomycetes on oak leaves on the

ground. Transactions of the British Mycological Society 68, 108-111.

Webster JR, Benfield EF (1986) Vascular plant breakdown in freshwater ecosystems.

Annual Review of Ecology and Systematics 17, 567-94. [In English]

Weiland JJ, Guyonnet R (2003) Study of chemical modifications and fungi degradation

of thermally modified wood using DRIFT spectroscopy. Holz als Roh- und Werkstoff

61(3), 216-220.

Weinstein MP, Kreeger DA, Newell SY, Porter D (2002) Microbial Secondary

Production from Salt Marsh-Grass Shoots, and Its Known and Potential Fates. In

'Concepts and Controversies in Tidal Marsh Ecology.' pp. 159-185. (Springer

Netherlands)

Westermann P (1993) Wetland and swamp microbiology. In 'Aquatic Microbiology: An

ecological approach.' First edn. Ed. TE Ford) pp. 215-238. (Blackwell Scientific

Publications: Boston)

Wetzel R (1992) Gradient-dominated ecosystems: sources and regulatory functions of

dissolved organic matter in freshwater ecosystems. Hydrobiologia 229(1), 181-198.

Wetzel RG (1990) Land-water interfaces: Metabolic and limnological regulators.


Limnologie 24, 6-24.

Wetzel RG, Howe MJ (1999) High production in a herbaceous perrenial plant achieved

by continuous growth and synchronised population dynamics. Aquatic Botany 64, 111-

129.

White TJ, Bruns T, Lee SB, Taylor JW (1990) Amplification and direct sequencing of

fungal ribosomal RNA genes for phylogenetics. In 'PCR Protocols: A guide to methods

and applications.' First edn. pp. 315-322. (Academic Press Inc.: San Diego)

Whiting GJ, Chanton JP (1993) Primary production control of methane emission from

wetlands. Nature 364(6440), 794-795.

Whittaker R, Likens G (1973a) Primary production: The biosphere and man. Human

Ecology 1(4), 357-369.

Whittaker RH, Likens GE (1973b) Carbon in the biota. Brookhaven Symposium on

Biology 30, 281-302.

298

Wicklow DT (1981) Interference competition and the organisation of fungal

communities. In 'The fungal community.' (Eds DT Wicklow and GC Carroll) pp. 731-

742. (Marcel Dekker: New York)

Williams CJ, Shingara EA, Yavitt JB (2000) Phenol oxidase activity in peatlands in New

York state: Response to summer drought and peat type. Wetlands 20(2), 416–421.

Williams DD (2006) 'The biology of temporary waters.' (Oxford University Press:

Oxford)

Willoughby LG, Archer JF (1973) The fungal spora of a freshwater stream and its

colonization pattern on wood. Freshwater Biology 3(3), 219-239.

Willoughby LG, Redhead K (1973) Observations on the utilization of soluble nitrogen by

aquatic fungi in nature. Transactions of the British Mycological Society 60, 598-601.

Wong MKM, Goh T-K, Hodgkiss IJ, Hyde KD, Ranghoo VM, Tsui CKM, Ho W-H,

Wong WSW, Yuen T-K (1998) Role of fungi in freshwater ecosystems. Biodiversity and

Conservation 7(9), 1187-1206.

Wood TG (1974) Field investigations on the decomposition of leaves of Eucalyptus

delegatensis in relation to environmental factors. Pedobiologia 14, 343-371.

Woollard P, Vestjens WJM, Maclean L (1978) The ecology of the eastern water rat

Hydromys chrysogaster at Griffith, N.S.W.: Food and feeding habits. Australian Wildlife

Research 5(1), 59-73.

Wurzbacher CM, Bärlocher F, Grossart H-P (2010) Fungi in lake ecosystems. Aquatic


Zepp RG, Callaghan TV, Erickson DJ (1998) Effects of enhanced solar ultraviolet

radiation on biogeochemical cycles. Journal of Photochemistry and Photobiology B:

Biology 46, 69-82.

Zepp RG, Faust BC, Hoigne J (1992) Hydroxyl radical formation in aqueous reactions

(pH 3-8) of iron(II) with hydrogen peroxide: the photo-Fenton reaction. Environmental

Science & Technology 26(2), 313-319.

Zhong J, Frases S, Wang H, Casadevall A, Stark RE (2008) Following fungal melanin

biosynthesis with solid-state NMR: biopolymer molecular structures and possible

connections to cell-wall polysaccharides. Biochemistry 47 (16), 4701-10.

Zimmer M, Oliveira R, Rodrigues E, Graça MAS (2005) Degradation of leaf litter

phenolics by aquatic and terrestrial isopods. Journal of Chemical Ecology V31(8), 1933-

1952.

Zucker WV (1983) Tannins: Does structure determine function? An ecological

perspective. The American Naturalist 121(3), 335-365.

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