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POPULATION AND IDENTIFICATION OF MYCORRHIZAL FUNGI IN ST.
AUGUSTINEGRASS IN FLORIDA AND THEIR EFFECT ON SOILBORNE
WHITNEY COLLEEN ELMORE
A DISSERTATION PRESENTED TO THE GRADUATE SCHOOL
OF THE UNIVERSITY OF FLORIDA IN PARTIAL FULFILLMENT
OF THE REQUIREMENTS FOR THE DEGREE OF
DOCTOR OF PHILOSOPHY
UNIVERSITY OF FLORIDA
This dissertation is dedicated to my family in the memory of my father,
I would like to thank my parents, Malcome and Donna Elmore, for their loving
support and my sister, Emilee. I would also like to acknowledge a very special person,
LaVette Burnette, for all of the patience and caring attention she has shown me for many
years. I would also like to thank Dr. James Kimbrough and his wife, Jane, for their
support, both emotionally and spiritually. I would also like to thank Drs. Jim Graham
and Kevin Kenworthy for agreeing to serve on my graduate committee and for their
willingness to offer advice on my studies. I also owe Dr. Vertigo Moody a big "thank
you" for motivating me to finish my Ph.D. as well as for his technical support in writing.
Additionally, I would like to say a big "thank you" to Dr. Gerald Benny both for
serving on my committee and for his attention in the lab. Dr. Benny is always ready to
help with research, or simply listen to my ramblings about research and politics which I
appreciate greatly. I would like to extend a personal "thank you" to the Department of
Plant Pathology staff, Gail Harris, Lauretta Rahmes, and Donna Perry. These ladies
always have a smile ready and a helping hand for students. I would also like to thank
Eldon Philman and Herman Brown for their assistance in experimental studies at the
greenhouse complex. They seem to always have a good solution or answer to any
problem or question. Finally, I would like to extend my sincerest appreciation to the
Department of Plant Pathology, namely Dr. Gail Wisler, at the University of Florida and
to the Institute of Food and Agricultural Sciences for financial and technical support in
this endeavor. I would not have been able to fulfill my dreams without the help and
support from all of these people.
TABLE OF CONTENTS
A C K N O W L E D G M E N T S ..................................................................................................iii
LIST OF TABLES ..... ......... ........ ....... ... .. .... ........vii
LIST OF FIGURES............... .. ...................... ........viii
A B STRA CT .................. ............ ......................... ................ ........ xii
1 GENERAL IN TRODU CTION ...................................... ...................... ............... 1
M ycorrhizal Types and Phylogeny ......................................................................... 2
Arbuscular M ycorrhiza Physiology ................................ ...............4
A rbuscular M orphology ................................................................. ..............5
M ycorrhizal C olonization ................................................................ ............... 6
M ycorrhizal Rhizosphere Interactions.................. ............................................ .8
Effects of Abiotic Factors on Mycorrhiza ...................................12
Effects of Seasonality on M ycorrhiza.............................. ...... ......... 14
M ycorrhizas in Grasses ................... ..... .................................... ...... 16
2 POPULATION AND IDENTIFICATION OF ARBUSCULAR
MYCORRHIZAL FUNGI IN ST. AUGUSTINEGRASS .......................................24
M materials and M ethods.................................... ............... 25
Results .......................... .. .. ...... ......... .29
Discussion ................... ................. ....... ....... ........34
3 THE EFFECT OF ARBUSCULAR MYCORRHIZAL FUNGI ON
GAEUMANNOMYCES GRAMINIS VAR. GRAMINIS AND RHIZOCTONIA
SOLANI COLONIZATION OF ST. AUGUSTINEGRASS SOD IN NORTH
CEN TRAL FLORID A SOILS ....................................................... 57
M materials and M ethods.................................... ............... 63
Results and Discussion ............................................... ..... .. 65
4 EFFECT OF GLOMUS INTRARADICES ON THE EXTENT OF DISEASE
CAUSED BY GAEUMANNOMYCES GRAMINIS VAR. GRAMINIS AND
RHIZOCTONIA SOLANI IN ST. AUGUSTINEGRASS ...............................74
Material and Methods ...... .................... ......... ........77
Direct Experiments ............. ... ............... ...............77
Indirect Experiments.......... ...... ............... 82
Results .............................. ... ...... ...........84
Direct Experiments ............. ... ............... ...............84
Discussion ................................. ................86
Results .................................... ..... ...........87
Indirect Experiment ...... ......... ..............................87
D iscu ssio n .......... ..... .... ......... ....................... 8 8
5 SUMMARY AND CONCLUSIONS ........................... ...............100
A SELECTIVE MEDIA RECIPES FOR ISOLATION OF G. GRAMINIS VAR.
GRAMINIS AND R. SOLANI FROM PLANT TISSUE .............. ................104
B NUTRIENT SOLUTION (20-0-20) USED IN DIRECT AND INDIRECT
TRIALS DESCRIBED IN CHAPTER 4................................................................105
C RHIZOCTONIA SOLANI AND G. GRAMINIS VAR. GRAMINIS INOCULUM
PRODUCTION PROTOCOLS ....... ................................... 106
D ADDITIONAL DATA ANALYSIS RESULTS REFERENCED IN CHAPTER 4
DIRECT EXPERIM ENTS .............................................. ............... 107
E ADDITIONAL DATA ANALYSIS RESULTS REFERENCED IN CHAPTER 4
INDIRECT EXPERIM ENTS ............................................................. ...............110
F ANALYSIS OF VARIANCE TABLES FOR CHAPTERS 2, 3, AND 4.............115
L IST O F R E F E R E N C E S ........................................................................................... 133
B IO G RA PH ICA L SK ETCH .......................................................................... .......... 150
LIST OF TABLES
2-1. Species of AMF positively identified at each sod farm location from pot cultures
of sorghum-sudangrass within a combination of field and sterile, low P soil..........45
2-2. Evaluation of analysis of variance data for spore density data from each sod farm
location by date..................... ............... ........ 48
2-3. Pearson correlation coefficients (r) for AMF spore density and soil moisture and
2-4. Evaluation of analysis of variance data for percent root length colonized from
each sod farm location................... ... ...................................... ...... 54
2-5. Chemical characteristics of soils sampled for AMF at three north central Florida
sod farm locations during January, April, August, and November 2005..............56
LIST OF FIGURES
2-1 A-C. 'Floratam' St. Augustinegrass sod farms located at (A) Fort McCoy
(Marion County), (B) Lake Butler (Union County), and (C) Starke (Bradford
County) in north central Florida. ............................................................ ............ 40
2-2. Sorghum-sudangrass pot cultures containing 50% (w/w) field soil combined
w ith 50% sterile, low P soil. ............................................................ 41
2-3. Spore extract from field soil following the wet sieving procedure...........................42
2-4 2-7. Stained arbuscular mycorrhizal structures observed within 'Floratam' St.
2-8 2-11. Stained arbuscular morphology types found within 'Floratam'
St. Augustinegrass................... ................. ................. ..........44
2-14 2-19. Arbuscular mycorrhizal fungal spores identified at the Lake
Butler sod farm location......... .. ............................ .. .......46
2-20 2-28. Arbuscular mycorrhizal fungal spores identified at the Fort
M cCoy sod farm location............... .. ........................... .........47
2-29. Spore density with increasing soil moisture levels over a 12-month period at the
Starke sod farm location. ................................................................................... .......51
2-30. Spore density with increasing soil moisture levels over a 12-month period at the
Fort M cCoy sod farm location. .................................... ..................... 51
2-31. Spore density with increasing soil moisture levels over a 12-month period at the
Lake Butler sod farm location. ........................................ ................ 52
2-32. Spore density with increasing soil temperatures over a 12-month period at the
Starke sod farm location. ................................................................................... .......52
2-33. Spore density with increasing soil temperatures over a 12-month period at the
Fort M cCoy sod farm location. .................................... ..................... 53
2-34. Spore density with increasing soil temperatures over a 12-month period at the
Starke sod farm location. ..........................................................................................53
3-1. 'Floratam' St. Augustinegrass sod mat infected with Gaeumannomyces graminis
var. graminis. Insert in bottom right-hand comer depicts underside of a mat
with rotting roots. ....................................................68
3-2 3-3. Comparison of healthy 'Floratam' St. Augustinegrass sod mat and sod
affected by brow n patch. ................................................ ............... 69
3-4. Deeply-lobed hyphopodia isolated from Gaeumannomyces graminis var.
graminis in 'Floratam' St. Augustinegrass sod samples. Scale bar = 40 [im..........70
3-5. Medium isolation plate depicting a Gaeumannomyces graminis var. graminis
colony isolated from 'Floratam' St. Augustinegrass sod samples. Arrow points
to colony. ........................................................70
3-6. Rhizoctoniasolani hyphae isolated from 'Floratam' St. Augustinegrass sod
exhibiting diagnostic 900 branching at constriction points and characteristic
septa. Scale bar = 40 rim. Arrow points to branching pattern..............................71
3-7. Medium isolation plate depicting light brown Rhizoctonia solani colony isolated
from 'Floratam' St. Augustinegrass sod samples...........................71
3-8. Mean percent of Rhizoctonia solani colonization of 'Floratam' St.
Augustinegrass in north central Florida. ................................. ............. 72
3-9. Mean percent of Gaeumannomyces graminis var. graminis colonization of
'Floratam' St. Augustinegrass in north central Florida ................................ 73
4-1. Rhizoctoniasolani isolate (PDC 7884) colony used to prepare inoculum in direct
and indirect experim ents................................... ......... 90
4-2. Gaeumannomyces graminis var. graminis isolate (JK2) used to prepare inoculum
in direct and indirect experim ents. ............................................. 90
4-3. Conetainers filled with low P soil and 'Floratam' St. Augustinegrass sprigs
inoculated in trial 1 of the direct experiment............... ....................................91
4-4. Glomus intraradices isolate (FL 208 A) used in direct and indirect assays to
inoculate 'Floratam' St. Augustinegrass sprigs. ..................................... 91
4-5. Photo showing nylon sleeves and plastic clips used in direct and indirect
experiments to clear and stain root segments from treatment replicates................92
4-6. Photo of mycorrhizal St. Augustinegrass root with arbuscules and intraradical
hypha of Glomus intraradices stained with 0.05% trypan blue from the direct
experiment G. intraradices inoculated control sprigs. ........... ... ..........92
4-7. 'Floratam' St. Augustinegrass sprigs after inoculation with Rhizoctonia solani
depicting disease severity rating scale (1-6)............................ ............. 93
4-8. 'Floratam' St. Augustinegrass sprigs after inoculation with Gaeumannomyces
graminis var. graminis depicting disease severity rating scale (1-6).................. 94
4-9 4-10. Photo depicting re-isolation plates of the two pathogenic isolates used to
challenge Glomus intraradices in both the direct and indirect experimental trials..95
4-11. Photo of the indirect experimental trial 3 containers arranged in a randomized
complete block design with four replicates per treatment..................... ..........96
4-12. Photo showing a close-up view of the experimental units of the indirect
experimental trial 1 depicting the split-root assay................................ ..........96
4-13. Photo showing the split-root assay of the indirect experimental trial 2 after
inoculation with ryegrass seeds inoculated with Gaeumannomyces graminis var.
graminis (JK2). ....................................................97
4-14. The direct effect of G. graminis var. graminis on St. Augustinegrass take-all
root rot disease severity without G. intraradices. ...................................... 98
4-15. The direct effect of G. graminis var. graminis on St. Augustinegrass take-all
root rot disease severity with G. intraradices. ................ .................. ..........98
4-16. The indirect effect of R. solani without G. intraradices on St.
Augustinegrass brown patch disease severity in an adjacent split sprig system....99
D-1. The direct effect of G. intraradices colonization on take-all root rot disease
severity in 'Floratam' St. Augustinegrass. ....................... ............. 107
D-2. The relationship between R. solani colonization and brown patch disease
severity in 'Floratam St. Augustinegrass....................................................... 108
D-3. The relationship between R. solani colonization and G. intraradices on brown
patch disease severity in 'Floratam' St. Augustinegrass...................................... 109
E-1. Photograph depicting a container used in the indirect experiment with drilled
hole and cut to allow for sprig to be inserted without tissue damage.....................110
E-2. The indirect effect of G. graminis var. graminis on take-all root rot disease
severity in 'Floratam' St. Augustinegrass without G. intraradices.................... 111
E-3. The effect of Glomus intraradices colonization on brown patch and take-all root
rot disease severity in 'Floratam' St.Augustinegrass on plants in the split sprig
E-4. The indirect effect of R. solani on disease severity in 'Floratam' St.
Augustinegrass with G. intraradices on an adj acent split sprig system............ 113
E-5. The indirect effect of G. graminis var. graminis on disease severity in
'Floratam' St. Augustinegrass with G. intraradices. .................... 114
F-1. Analysis of variance tables for spore density and percent colonization data in
Chapter 2, and Pearson's product moment correlation coefficients for attempted
correlations between variables and soil chemical characteristics and soil
m oisture and soil tem perature. ....................................................... 120
F-2. Analysis of variance tables for Rhizoctonia solani percent colonization data in
Chapter 3. ............................... ........... 122
F-2. Analysis of variance tables for Gaeumannomyces graminis var. graminis percent
colonization data in Chapter 3 ......................................... ... ..... 126
F-4. Analysis of variance tables for the direct assay in the split-sprig challenge
including Gaeumannomyces graminis var. graminis and Rhizoctonia solani data
in Chapter 4. ......................................................129
F-4. Analysis of variance tables for the indirect assay in the split-sprig challenge
including Gaeumannomyces graminis var. graminis and Rhizoctonia solani data
in Chapter 4. ...................................................... 132
Abstract of Dissertation Presented to the Graduate School
of the University of Florida in Partial Fulfillment of the
Requirements for the Degree of Doctor of Philosophy
POPULATION AND IDENTIFICATION OF MYCORRHIZAL FUNGI IN ST.
AUGUSTINEGRASS IN FLORIDA AND THEIR EFFECT ON SOILBORNE
Whitney Colleen Elmore
Chair: James W. Kimbrough
Major Department: Plant Pathology
Arbuscular mycorrhizal fungi (AMF) are obligate symbionts of more than 90% of
all land plants. Mycorrhizae are documented in many crops as positive associations with
roots of plants that help reduce disease severity soilbome pathogens and increase nutrient
and water uptake while lowering plant stress and ultimately management costs.
However, there is no information concerning the effects of AMF colonization in St.
In Florida, St. Augustinegrass sod production contributes hundreds of millions of
dollars to the economy annually while supplying a product to homeowners and
commercial entities with great aesthetic value. The use of AMF in St. Augustinegrass
sod production has many potential benefits to the sod industry and the environment
including lowered management costs, pesticide use and pollution. In these studies, a
survey of St. Augustinegrass sod farms in north central Florida revealed a moderate level
of AMF colonization as well as a diverse population of AMF species. Direct and indirect
pathogen challenges with the ubiquitous AMF, Glomus intraradices, in St.
Augustinegrass plants suggested a limited role for AMF in lowering disease severity in
two of the more devastating diseases of St. Augustinegrass in Florida, brown patch and
take-all root rot.
While no positive correlation was observed between AMF colonized St.
Augustinegrass plants and the soilborne pathogens Rhizoctonia solani or
Gaeumannomyces graminis var. graminis, effective assays for mycorrhizal St.
Augustinegrass evaluations were developed and foundation information concerning the
association between St. Augustinegrass and AMF provided valuable data, which may
help in the development of future AMF evaluations in St. Augustinegrass field trials and
with other AMF species. These results were the first to suggest an association between
AMF and St. Augustinegrass, and to evaluate their potential effects on disease severity.
"Mykorrhizen" was a term first applied by the German forest pathologist, A.B.
Frank, who described structures in plant roots as "fungus-roots" (1885). Harley (1989)
described them as a mutualistic symbiosis in which a fungus and host exist as one.
Despite minuscule differences in description, mycorrhizas are recognized by scientists as
economically important in most agricultural crops. In fact, the mutually beneficial
relationships are actually three-way associations in which the soil, plant root, and fungus
interact to produce symbiotic effects.
In 1879, de Bary defined symbiosis as "the living together of differently named
organisms," which included both parasitic and beneficial relationships. Later, Raymer
(1927), commenting on the nature of symbionts, acknowledged such partnerships, but did
not provide functional information concerning the fungi involved. However, after many
years of advanced research throughout the 1960's and 70's, the meaning of the
relationship was refined to refer to naturally beneficial relationships exclusively. Most
likely, organisms co-existing became symbiotic as a result of selection pressures exerted
over the course of time (Remy et al., 1994). In fact, it is possible that the movement of
plants from water to land could not have occurred without mycorrhizal associations
(Nicolson, 1975; Pirozynski and Malloch, 1975). It is now recognized that mycorrhizas
are the norm and not the exception within the Kingdom Planta. With ancient lineages
stretching across evolutionary history, Bryophytes, Angiosperms, Pteridophytes, and
some Gymnosperms all possess these associations
(Fitter, 1991), while members of the Brassicaceae seem to evade infection by any type of
mycorrhizal fungi (Gerdemann, 1968), even in close proximity to mycorrhizal plants.
Involved in mycorrhizal symbiosis are members of the fungal taxa Ascomycotina,
Basidiomycotina, Zygomycotina, Deuteromycotina, and Glomeromycota (Schtissler et al.,
2001; Srivastava et al., 1996). Infrequently found living as saprobes, most of these fungi
are widespread across various soil types with strong biotrophic host dependence (Smith
and Read, 1997).
Mycorrhizal Types and Phylogeny
Types of mycorrhizae are divided based on their fungal associations, extent of
root penetration, presence or lack of an external mantle and/or sheath, as well as the intra-
and intercellular structures produced inside of the host root (Srivastava et al., 1996).
Presently, seven types of mycorrhizae are recognized by taxonomists (Bagyaraj, 1991).
The types of mycorrhizae include: Ectomycorrhizae, Ectendomycorrhizae, Arbutoid,
Monotropoid, Ericoid, Orchidoid, and Endomycorrhizae or the vesicular-arbuscular
mycorrhizae (Bagyaraj, 1991). Endomycorrhizae, also known as vesicular-arbuscular
mycorrhizae or VAM, were taxonomically placed within the Order Glomales of the
Phylum Zygomycota based on morphological features of asexual spores resembling
sexual reproductive structures of the Zygomycota. Six genera are recognized within the
Glomales: Glomus, Sclerocystis, Gigaspora, Scutellospora, Acaulospora, and
Entrophospora (Morton and Benny, 1990). In 2001, Schussler et al., using information
provided by small subunit rRNA gene sequences, proposed a new Phylum, to separate
arbuscular mycorrhizal fungi from other fungal groups in a monophyletic clade.
Schussler et al. (2001) suggested that they be removed from the Zygomycota and placed
into a newly erected Phylum Glomeromycota. Small subunit rRNA gene sequencing also
placed Geosiphon pyriformis, an endocytobiotic fungus, which is a distant relative of the
arbuscular mycorrhizal fungi, within this new Phylum (Schussler et al., 2001). Within
the same article, Schussler et al. (2001) also suggested that the Glomus genus be emended
to include the termination -eraceae, with the family named Glomeraceae and the higher
taxon names reflecting this change with Glomerales. Furthermore, Schussler et al.
(2001) suggested three new orders, mostly diverged from the Ascomycetes and
Basidiomycetes, be recognized as well. These are the Archaeosporales, Diversisporales,
and the Paraglomerales. Based on a combination of molecular, ecological, and
morphological characteristics, these fungi can now be separated from other fungal
groups. The use of molecular techniques such as small subunit rRNA sequencing has led
to the recent introduction of other species within the genus Glomus. Walker et al. (2004)
and Rani et al. (2004) also used this technology to add Glomus hyderabadensis from
India, and a new genus Gerdemannia, to the growing list of arbuscular mycorrhizal fungi
collected and speciated around the world. Based on their distinct molecular differences
from the Zygomycota and placement into a new phylum, Goto and Maia (2005) recently
suggested that spores of the arbuscular mycorrhizal fungi be referred to as
glomerospores. Indeed, these spores are not chlamydospores, conidia, or azygospores, so
differentiation based on molecularly distinct features is pertinent.
Forming vesicles and arbuscules within cortical root cells, fungi of the
Glomeromycota produce aseptate hyphae without the presence of a sheath or mantle.
Gigaspora and Scutellospora produce arbuscules only within roots and vesicles only
within the soil, and, therefore, the vesicular-arbuscular mycorrhizal term has been
emended to simply read as arbuscular mycorrhizae. The name was amended simply
because arbuscules are the most basic and one of the few commonalities between the
members of the group (Morton and Benny, 1990). Taylor et al. (1995) proposed that
Glomites be included as a new fossil genus of Glomales, and two years later, Wu and Lin
(1997) added another genus, Jimtrappea. However, these two genera are not widely
accepted. Currently, there are about 150 recognized species described within the
Glomales, of which only a few have been carefully studied and recognized as endo-
mycorrhizal (Morton and Bentivenga, 1994; Morton and Benny, 1990; Morton et al.,
1992; Pirozynski and Dalpe, 1989; and Stuessy, 1992). Glomeromycota are not known to
produce sexual reproductive spores and, therefore, are characterized and classified by
their resting structures. These structures vary in wall characteristics, size, shape, and
color (Morton et al., 1992; Morton and Bentivenga, 1994; and Morton and Benny, 1990).
Arbuscular Mycorrhiza Physiology
The most widespread of the mycorrhizae, both geographically and among species,
the arbuscular mycorrhizae occur frequently in the top 15-30 cm of cultivated soil
(Bagyaraj, 1991). Arbuscular mycorrhizae-forming fungi colonize and form associations
with most agriculturally and horticulturally important plant species, from fruit and forest
trees to shrubs and grasses. Unlike other mycorrhizae, these associations do not typically
lead to noticeable external morphological changes in plant roots, and they cannot be
observed easily without staining procedures (Phillips and Hayman, 1970). In most cases,
plants which have formed associations with other types of mycorrhizal fungi, such as
basidiomycetes and ascomycetes, do not form relationships with arbuscular mycorrhizae.
From the standpoint of the fungus, host specificity exists while the opposite view would
be held about the host due to the wide host range of most of the arbuscular mycorrhizal
fungi (Gerdemann, 1955). Their limited capacity to be grown from spores, vesicles, or
hyphae from root residue has led to special methodologies in order to maintain strains
and for taxonomic evaluation. Typically, single spore types are cultivated in "pot
cultures" on plant roots so that characteristics of spores, their mode of colonization, and
effects on plant growth can be studied (Smith and Read, 1997).
Named by Gallaud (1905) for the structures formed inside cortical root cells,
arbuscules are similar to branched haustoria, which form early on in the association
between plant root and the repeatingly branched fungal hyphae. Baylis (1975) and St.
John (1980) suggested that the form of the root system is a defining factor in the extent to
how plants react, nutritionally, and in growth to mycorrhizal colonization. Evolving
across phylogenetic lines many times, it appears that dicotyledons have a large incidence
of associations with fungal species which form mycorrhizal associations, with very few
being non-mycorrhizal in nature (Trappe, 1987). In comparison, the lines of
monocotyledons studied by Cronquist (1981) are heavily mycorrhizal, with arbuscular
mycorrhizas predominating except in the Orchidaceae, which have mycorrhizas formed
by Basidiomycetes. In plants forming primarily magnolioid type roots, with wide
diameters up to 1.5 mm, slow growth habits, and little root-hair development,
mycorrhizas are usually well accepted and form greatly receptive relationships. On the
other hand, roots that are primarily fine and rapidly growing with long root-hairs lack the
same responsiveness (Baylis, 1975; St. John, 1980). Mycorrhizal relationships were first
described by the type of colonization patterns, referred to as either Arum- or Paris-type
(Gallaud, 1904). In fact, there appears to be a continuum between the two forms, with
intermediate types along the way.
The Arum-type, which was considered the most common association, develops
primarily within cultivated crops and consists of intercellular hyphae and arbuscules. In
contrast, the Paris-type of symbiosis involving intercellular hyphae, arbusculate coils,
and hyphal coils, typically develops within forest trees and herbs (Dickson, 2004). In
surveys of mycorrhizal plants and trees from both natural and cultivated environments, it
appears that most plant families are dominated by only one symbiotic type (Smith and
Smith, 1997). There are, however, a few plant families that appear to possess
intermediate forms of the colonization types, including the Poaceae (Smith and Smith,
1997). In an extensive survey of various plant families and mycorrhizal fungi, eight
distinct classes of colonization types were found along a continuum ranging from the
Paris- to Arum-type (Dickson, 2004). Most researchers agree that one fungus can form
either type of arbuscular colonization with most of the specificity in structure dependent
upon the host plant (Barrett, 1958; Gerdemann, 1965). Brundett and Kendrick (1988)
commented on the presence of intercellular spaces within the host root cortex as being the
main factor influencing arbuscular type. Conversely, in tomato, Cavagnaro et al. (2001)
suggested that the colonization type was dependent on both the host and fungus involved.
In mycorrhizal colonization, the host plasmalemma is invaginated with the
encroaching arbuscules. These are physiologically active sites for nutrient translocation,
for 4-6 days, within the roots (Bracker and Littlefield, 1973; Brundett et al., 1984).
Arbuscules are important sites for P exchange for plants under deficient conditions
(Simth and Read, 1997). The vesicles, which are small and usually dark, globular or
spherical structures, form later in the association and arise from swelling of terminal and
intercalary hyphal cells. Vesicles act as storage sites for lipids (Srivastava et al., 1997).
Transversing long distances of soil beyond nutrient depletion zones and reaching areas
untouched by growth limited root hairs, the external hyphae absorb nutrients such as P
and make it available to plants, rendering these plants more equipped to survive nutrient
competitions (Nicolson, 1967). Once the fungal hyphae and plant roots become closely
associated in space, a functionally and structurally complex symbiotic relationship is
formed between the compatible organisms.
Formed only on unsuberized root tissue, certain areas of the root are more readily
colonized even though mycorrhizae can develop on any portion of young root tissue
(Brundett and Kendrick, 1990). Based on mathematical and geometrical models, root
tissue directly behind the meristematic area is considerably more susceptible to
penetration and colonization when compared to other root segments (Garriock et al.,
1989; Bonfante-Fasolo et al., 1990). This area of discrete colonization was described
earlier as the mycorrhizal infection zone by Marks and Foster (1973), who considered the
area to be "non-static," thus growing with the root. Furthermore, Brundett and Kendrick
(1990) found that the fungus penetrates and colonizes root cells with little or no suberin
deposition, which has been shown to occur just prior to or after fungal penetration.
Usually, epidermal and outermost cortical cell colonization is minimal with the
intercellular hyphae formed in the inner cortex and the majority of the colonization is
deep within the cortex where arbuscules are formed (Srivastava et al., 1997).
With the aid of cellulolytic and pectinolytic enzymes produced by the fungus,
direct penetration of the outermost cell wall is the preferred mode of hyphal entry (Jarvis
et al., 1988). Physiochemical aspects of the epidermal cell wall seem to be the primary
reasons for preferential site penetration (Jarvis et al., 1988). After cell to cell contact
between fungus and host, the external mycelia swell to form defined appresoria (20-40
lm in length). Within these appresoria, infection hyphae are formed and penetrate host
cell walls (Garriock et al., 1989). Once penetration has occurred via mechanical and
enzymatic interactions, the host's plasmalemma appears to extend around the fungus
(Bracker and Littlefield, 1973). Arbuscule formation takes between 4-5 days after which
extramatrical hyphae occurs promoting new penetration sites (Brundett et al., 1984).
Arbuscules are major contributors to the transfer of nutrients, in particular sugars,
between the plant to fungus and inorganic materials, mainly P, from the fungus to the
plant (Smith and Gianinazzi-Pearson, 1988).
Mycorrhizal Rhizosphere Interactions
A necessary component of plant life, the macro element P, occurs as part of DNA
and RNA nuclei and as part of plant membranes as phospholipids (Griffiths and
Caldwell, 1992; Smith and Read, 1997). Present in high amounts within active
meristematic regions as part of nuclear proteins and as part of ADP, ATP, NADP, and
NAD, P is partly responsible for oxidation-reduction reactions such as respiration,
nitrogen and fat metabolism, and photosynthesis, which are necessary for life (Beever
and Burns, 1980; Munns and Mosse, 1980). Symptoms of deficiency often include
purple or red leaf pigmentation, dead and/or necrotic leaves, petioles, and fruits,
premature leaf drop, stunting, and poor vascular tissue development (Srivastava et al.,
1997). An important aspect of arbuscular mycorrhizal associations is the increase in P
uptake by the plant.
The importance of arbuscular mycorrhizal fungi for P absorption was first
suggested by Baylis (1959) and then Gerdemann (1964). Later, Baylis (1967), Daft and
Nicolson (1966), Holevas (1966), and Murdoch et al. (1967) provided advanced
information showing the close association between mycorrhizas and P nutrition of the
host. Interestingly, Mosse (1973) once remarked that more than one quarter of
mycorrhizal text is devoted to P research. In fact, Sanders and Tinker (1973) stated that
"the value of these mycorrhizas for the phosphate nutrition of plants in deficient
environments may rival that of Rhizobium in nitrogen." Obviously, such a strong
statement must be supported by an abundance of research. As mycorrhizal research
progressed during the last three decades, P research remained an important topic. For
instance, in 1986, Gianinazzi-Pearson and Gianinazzi studied the kinetic associations
between P concentration in soil solutions and its effect on root and shoot tissues, while
Young et al. (1986) evaluated the effect of arbuscular mycorrhizal fungi inoculation on
soybean yield and P utilization in tropical soils. Later, Koide (1991) determined that it is
the variation among plant species in phenological, morphological, and physiological traits
that influence P demand and supply which are directly connected to potential response of
mycorrhizal associations. Once absorbed, P is allocated for plant functions or stored for
later use (Cox and Sanders, 1974). Since P deficiency is caused by both P availability
and plant demand, mycorrhizal associations can have various effects based on the plant
species (Koide, 1991).
In low P soils, mycorrhizal plants have an advantage over non-mycorrhizal plants
with root to shoot ratios lowered and shoot fresh weight to dry weight ratios higher in
mycorrhizal plants (Tinker, 1978). The plant's growth rate is influenced by interactions
in mycorrhizal colonization such as nutritional, and non-nutritional, physiological effects,
such as pH, temperature, microbial turnover, phosphatase activity, soil and plant
moisture, and/or iron (Fe) or aluminum (Al) chelate concentration (Nye and Tinker,
1977; Rusell, 1973). In P deficient soils, studies have shown that plant species with few
root hairs are strongly mycorrhizal, providing evidence that root anatomy has a strong
correlation to mycorrhizal colonization (Crush, 1974; Baylis, 1975).
Smith and Read (1997) wrote "the focus (of current research) is on P uptake, as
well as on the uptake of other nutrients for which there is now unequivocal evidence of
mycorrhizal involvement." Furthermore, they noted that "there is excellent evidence to
demonstrate that external hyphae of VA mycorrhizal fungi absorb non-mobile nutrients
(P, Zn, Cu) from soil and translocate them rapidly to the plants, thus overcoming
problems of depletion in the rhizosphere which arise as a consequence of uptake by
roots." Throughout the 1960's, reviews of the occurrence of arbuscular mycorrhizal
colonized plants and anatomy were the norm in mycorrhizal research (Smith and Read,
1997). There had been little mention of mineral nutrition until Mosse (1957) released
details of an experiment with apple seedlings which provided evidence for increased
amounts of potassium (K), iron (Fe), and (copper) Cu in mycorrhizal plant tissue versus
noninoculated control plants. Other researchers such as Gerdemann (1964) established
that P tissue concentrations were also higher in mycorrhizal plants, although the
mechanisms were not yet clearly understood. Mosse (1973) reported a shift in
mycorrhizal research from pot experiments to study the anatomy of arbuscular
mycorrhizal fungi to that of plant growth and P uptake. Now, the mechanisms underlying
the mycorrhizal effect on P uptake are coming to light including extraradical hyphae
growing into soil not already colonized by roots; hyphae that are more effective than
roots, due to size and spatial distribution, in competing with free-living microorganisms
or mineralized or solubilized P; the kinetics of P uptake into hyphae may differ from
roots; and that mycorrhizal roots can use sources of P in soil that are not plant available
(Smith and Read, 1997).
Hyphal pathways between plants may offer links for soil-derived nutrient transfer,
as is the case with plant-derived carbon (C), which can have important roles in the inter
plant and species competition in the environment (Smith and Read, 1997). Enzymes are
not the only substances produced by arbuscular mycorrhizal fungi. An Iron-containing
glycoproteinaceous substance called glomalin, produced by these fungi, is deposited in
soils (Rilling et al., 2003). Glomalin is considered to be linked to soil Carbon storage due
to its effect on soil aggregation (Rilling et al., 2003). Consistently correlated with soil
aggregate water stability, glomalin is involved in C and N content as well as being useful
as a potential land-use change indicator (Rilling et al., 2003). After many years of
taxonomic research with proteins and soil stability, micronutrient uptake research has
increased following studies by Mosse (1957), Daft et al. (1975), and Gildon and Tinker
(1983) where uptake of Cu and zinc (Zn) were observed in apples and maize when
inoculated with arbuscular mycorrhizal fungi. The uptake of other micronutrients is not
well documented, however, Marschner and Dell (1994) observed that the uptake of
manganese (Mn) is usually reduced by mycorrhizal associations. Occasionally, instances
of increased K concentrations in plant tissues have been reported, which is to be expected
given the immobility of the K ion within the soil matrix (Srivastava et al., 1997).
Conversely, with increased P uptake as well as other nutrients in mycorrhizal plants
comes the risk of accumulating toxic elemental levels. With improved P nutrition and
plant growth, the uptake of heavy metals per plant is greatly increased as demonstrated
by El-Kherbawy et al. (1989) on alfalfa inoculated with arbuscular mycorrhizae in
various soil pH levels with and without rhizobia.
Effects of Abiotic Factors on Mycorrhiza
Many climatic and physiochemical or abiotic features of the soil influence
arbuscular mycorrhizal establishment, growth and benefit. For instance, light, which is
not directly required by mycorrhizas in some cases, is essential for the host to thrive and
translocate photosynthates to the root, which in turn provides a home for mycorrhizal
fungi. In other cases, arbuscular mycorrhizal fungi are stimulated by light to increase
root colonization and spore production as well as plant response to mycorrhizal
colonization (Furlan and Fortin, 1973; Hayman, 1974).
The rate of photosynthesis and translocation of its products are heavily influenced
by air temperature (Furlan and Fortin, 1973; Hayman, 1974). By increasing air
temperature to 260 C an increase in plant growth is typical (Hayman, 1974). Soil
temperatures also influence mycorrhizal development at all stages: spore germination,
hyphal penetration, and proliferation within cortical root cells (Schenck and Schroder,
1974; Smith and Be, 1979). Optimal temperatures vary for spore germination between
species and other stages in development. The ability of the arbuscular mycorrhizal spores
to survive following host death or harvest is also dependent on soil temperature, though
also affected by soil texture (Bowen, 1980).
Soil pH is an additional determinant factor in mycorrhizal growth and
development. The efficiency of the mycorrhizae is directly determined by its ability to
adapt to soil pH. Soil pH affects both spore germination and hyphal development (Angle
and Heckman, 1986; Green et al., 1976). The interaction of soil pH and mycorrhizal
development is difficult with soil type, plant and fungal species and P forms involved.
Typically, mycorrhizas are able to colonize and grow well in soils of pH 5.6 to 7.0, but
not in soils of pH 3.3 to 4.4, as reported by Hayman and Mosse (1971).
Generally, mycorrhizas are not found within aquatic conditions, due to a
reduction in colonization, however, some aquatic plants are commonly mycorrhizal, such
as Lobelia dortmanna L. and Eichhornia crassipes [Martius] Solms (Read et al., 1976).
Conversely, most plants found within drought are typically mycorrhizal, which aids in
their survival in harsh conditions (Sondergaard et al., 1977). Arbuscular mycorrhizal
colonization of roots affects many mechanisms in plant water determination. Root
hydraulic conductivity, leaf gas exchange and expansion, phytohormone regulation, and
leaf conductants are all affected by interactions with arbuscular mycorrhizas (Gogala,
1991; Hardie and Leyton, 1981; Koide, 1985; Nelson, 1987; Auge et al., 1986). Fungal
mycelium is involved in the transport of water especially at low soil potentials, which has
made arbuscular mycorrhizae colonization and development a hot research topic in arid
and tropical landscapes (Faber et al., 1991).
Mycorrhizal roots and organic matter content play important roles in arbuscular
mycorrhizal survival and development as well. Organic root debris may act as a reserve
for soil inocula (Warner and Mosse, 1980), while in arid areas contact between
susceptible plant roots and colonized root residue is considered by Rivas et al. (1990) to
be the most important means for mycorrhizal dissemination when little water is available
for spore transport. Soil structure, pH, water, and nutrient availability are all affected by
organic matter content, thus influencing mycorrhizal associations (Khan, 1974; Daniels
and Trappe, 1980; Johnston, 1949). For instance, Johnston (1949) suggested that organic
materials such as manures can enhance tropical soil mycorrhizas in cotton stands. And,
Sheikh et al. (1975) reported that spore population and organic matter content were
positively correlated in soils with 1-2% organic matter, but low in soils with 0.5%
organic matter or less. Organic matter and root residue are important ecologically as part
of the three-way soil, plant and fungal mycorrhizal relationship.
Effects of Seasonality on Mycorrhiza
Seasonality is another abiotic contributor to arbuscular mycorrhizal colonization.
Seasonality has been shown to affect spore production as a function of host and climate
(Hetrick, 1984), while seasonal patterns can be correlated with P availability and soil
water potential in combination with host growth stages, other biotic and abiotic factors,
and management practices such as fertilization (Cade-Menun et al., 1991; Yocums,
1985). Hayman (1975) demonstrated that fertilizers such as P and Nitrogen (N) could
potentially reduce spore number and fungal colonization with N having a more
detrimental effect than P. Despite the possibility for soil chemical treatment injury,
arbuscular mycorrhizae can be found in fertile soils, which Hayman et al. (1976)
contributed to other factors such as host species, soil type, and management practices
influencing fungal survival and development.
As previously mentioned, management practices such as pesticide applications, in
particular, fungicides, may inhibit the effect of arbuscular mycorrhizal fungal sporulation
and colonization (Nemec and O'Bannon, 1979; El-Giahmi et al., 1976). Rhodes and
Larsen (1979) examined arbuscular mycorrhizae of turfgrasses in field and greenhouse
conditions. The researchers discovered that when fungicides were applied to bentgrass,
infection averaged 9 to 17%, however, in non-treated field plots, the roots were infected
at a rate of 40-60 percent. The same observation was reported in the greenhouse
evaluations, with one fungicide, PCNB, totally eliminating mycorrhizae (Rhodes and
Larsen, 1979). Conversely, DBCP, a nematicide, has actually been reported by Bird et al.
(1974) to enhance arbuscular mycorrhizal development.
It is imperative to mention that mycorrhizal interactions lie along a continuum
from mutualistic to parasitic based on the cost to benefit ratio colonization. Obviously,
mycorrhizal associations can be mutualistic, but they can also be parasitic, commensal,
amensal, and even neutral in nature (Johnson et al., 1997). Where, along this continuum
the association will fall, depends on a complex hierarchy mediated by biotic and abiotic
factors within the rhizosphere and ecosystem being affected. No doubt, this range of
mycorrhizal associations is greatly affected by time and space. The complexity of
mycorrhizal investigations is ultimately confounded by the fact that the plant and fungal
perspective on costs to benefits differs greatly from situation to situation (Johnson et al.,
With this in mind, Ryan and Graham (2002) presented the point-of-view that
arbuscular mycorrhizal fungi do not play such a vital role in production agricultural
systems, in relation to nutrition and growth, simply because the high cost of energy from
the plant to support the fungal invader outweighs the benefits of that association. This
outcome is not beneficial in terms of crop production and may, in fact, be detrimental.
Nonetheless, those production systems not considered to be within a natural or traditional
cultivated production system, such as sod, still need much attention where mycorrhizal
symbiosis is concerned before a definitive yes or no can be applied to functional use of
mycorrhizal fungi. Conversely, in 1997, Srivastava et al. concluded that "there is little
doubt that vesicular arbuscular mycorrhizae fungi will emerge as a potential tool for
improving crop plants in the years to come." These opinions, in conjunction with the
increased concern for environmental quality and sustainable technologies warrants an
examination of more specific research reports in agricultural crops. In this review, the
concentration is on turfgrass research.
Mycorrhizas in Grasses
There has been a considerable amount of research on mycorrhizal fungi
associated with grasses (Hetrick et al., 1988, 1991; Trappe, 1981; Bethlenfalvay et al.,
1984). Though much of the work conducted on grasses was begun in the 1970's,
Nicolson (1955) examined mycotrophic nature in grasses and later (Nicolson, 1956) with
mycorrhizae in both grasses and cereals. These first studies in grasses and cereals were
mainly concentrated on the ecological aspects of mycorrhizal infection. In fact, it was
not until Nicolson (1956) showed diagrammatically that external hypha penetrate the root
hairs or epidermal cells and spread throughout the cortex of grasses. Additionally,
Nicolson noted that arbuscules form later in the inner cortical layers, which was valuable
information in the study of grasses and their mycorrhizal partners.
In experiments on fescue (Festuca ovina L.), cocksfoot (Dactylis glomerata L.),
sand fescue (Festuca rubra var. arenaria L.), and marram grass (Ammophila arenaria L.:
Link), Nicolson (1956) found that mycorrhizal infection was prevalent throughout a wide
range of different habitats and soil types, although the incidence of infection varied
greatly between habitats and communities. With a lull in ecological studies throughout
the 1960's, environmental issues surpassed many of the more basic research topics. In
1979, Rhodes and Larsen examined the effects of fungicides on mycorrhizal development
in cool-season turfgrasses. Again, Rhodes and Larsen (1981) conducted a similar study,
where the effects of fungicides on bentgrasses and the mycorrhizal fungus, Glomus
fasciculatus, were explored. Arbuscular mycorrhizas of 'Penncross' creeping bentgrass
(Agrostispalustris Huds.) were studied in greenhouse experiments to evaluate popular
fungicides, such as, chloroneb and maneb, which did not affect mycorrhizal development.
However, foliar applications of PCNB, chlorothanil, bayleton, anilazine, benomyl, and
chloroneb at various weeks after inoculation with Glomusfasciculatus resulted in
significantly reduced mycorrhizal colonization, thus limiting their beneficial effects.
Later, studies of mycorrhizas in turfgrasses seemed to swing back toward
ecological studies with the introduction of seasonal and edaphic variation of arbuscular
mycorrhizal infection (Rabatin, 1979). In a population survey, Rabatin (1979) sampled
for Glomus tenuis infection in Panicum virgatum L., Poa compressa L., Poapratensis L.,
Poapalustris L., Phleum pratense L., and Festuca etalior L., all cool-season meadow
grasses. Rabatin (1979) determined that the greatest percentage of root infection by this
fungus occurred in grass roots from dry, P deficient fields. Moreover, the percent of
infection was lowest in the cool, wet months of the spring. Thus, Rabatin (1979)
concluded that mycorrhizal infection tends to be greater in drier, P deficient soils versus
wet or flooded conditions.
Bagyaraj et al. (1980) concluded that a study of the spread of mycorrhizas from
the site of infection along the root to deeper soil layers was necessary to provide
important information for plant inoculations. This was done in grasses since the roots
grow out of the inoculated sites quickly. Researchers collected root samples from various
depths and found that roots at 3 4 and 8 9 cm were mycorrhizal at 45 days after
inoculation. However, when roots were collected from deeper layers, the roots were only
mycorrhizal after 75 days. The research lead Bagyaraj et al. (1980) to conclude that
mycorrhizal infection of warm-season grasses such as Sudangrass (Sorghum bicolor L.:
Moench), was spread to deeper layers by mycelial growth through the root, which was
helpful information when researching inoculation methodologies important in such
experiments as population surveys where pot cultures are a necessary to speciate the
fungi collected. In an attempt to determine the distribution and occurrence of
mycorrhizal fungi in Florida's agricultural crops, Schenck and Smith (1981) examined
bahiagrass (Paspalum notatum Fluigge) and digitgrass (Digitaria decumbens Stent)
among 30 Cucurbitaceae, Leguminosae, Solanaceae, and Vitaceae crops. In a population
survey, the authors found that mycorrhizal fungi in Glomus occurred most frequently in
Florida, with species of Gigaspora found regularly in central and south Florida and
Entrophospora collected only once (Schenck and Smith, 1981). Furthermore,
Acaulospora was found in the highest frequency in the grasses evaluated. In this
instance, there was no correlation among species or genera occurrence and the available
soil P or soil pH.
In another study, endomycorrhizas and bacterial populations were examined in
three cool-season grasses. Agrostis tenuis Sibth., Deschampsiaflexuosa L.: Trin., and
Festuca ovina L., were collected and examined by Lawley et al., (1982) for mycorrhizal
associations. In this case, the researchers noticed that mycorrhizal abundance was lowest
when Agrostis species were partnered with other plants and highest when partnered with
Finally, Sylvia and Burks (1988) began working with grasses other than those
only found in cool-season climates. Beach erosion in coastal areas became a major
economic concern in the late 1980's; beach grasses such as sea oats (Uniola paniculata
L.) were often utilized to restore southeastern beaches to slow loss of sand. It was
unclear whether or not these grasses relied on arbuscular mycorrhizal associations for
survival in the harsh climate. Sylvia and Burks (1988) found that isolates of Glomus
deserticola and G. etunicatum significantly increased the dry mass, height, and P content
of the sea oats, while other isolates had little or no effect.
In the search for a better host for inoculum production, compared to the traditional
bahiagrass, Sreenivasa and Bagyaraj (1988) evaluated seven grasses for their ability to
quickly produce large masses of mycorrhizal spores for inoculations. Grasses such as
guinea grass (Panicum maximum Jacq.) and rhodes grass (Chloris gayana Kunth) were
studied and all were found to be mycorrhizal. However, the highest root colonization
was observed in the rhodes grass, as well as the highest production of spores and
infective propagules. Studies on other warm-season grasses such as St. Augustinegrass
(Stenotaphrum secundatum [Walt.] Kunze), Centipedegrass (Eremochloa ophiuroides
[Munro] Hack.), or even bermudagrass (Cynodon dactylis L.: Pers.) have not been
In studies of the difference in responses of C3 and C4 grasses to P fertility and
mycorrhizal symbiosis, Hetrick et al. (1990) showed that warm-season grasses such as
big bluestem (Andropogon geradii Vitm.) and indian grass (Sorghastrum nutans L.:
Nash), responded positively to mycorrhizae or P fertilization, or mycorrhization in cool-
season grasses, such as perennial ryegrass (Lolium perenne L.). In warm-season grasses,
there was a positive relationship between root colonization and dry weight, with an
inverse relationship between mycorrhizal root colonization and P fertilization. The
evaluation provided evidence that the C3 and C4 grasses display profoundly different
nutrient acquisition strategies (Hetrick et al., 1990b).
The effect of mycorrhizal symbiosis on regrowth of rhizomes of big bluestem was
assessed as a function of clipping tolerance (Hetrick et al., 1990a). Mycorrhizal clipped
plants were larger than nonmycorrhizal clipped plants, but the effect diminished with
successive clippings as did mycorrhizal root colonization. This information on clipping
tolerance indicates that mycorrhizal turfgrasses respond similarly when clipped or mowed
under constant turf management.
Hetrick et al. (1991) compared the root architecture of five warm and five cool-
season grasses in an attempt to evaluate whether mycorrhizal symbiosis confers a greater
tolerance to drought, soilborne disease, vigor, and yield through direct or indirect
improved nutritional status of the host plant. The cool-season grasses had significantly
more primary and secondary roots than the warm-season grasses and the diameter of
those roots was smaller than that of the warm-season grasses. The mycorrhizas did not
affect the number or diameter of cool-season grass roots, however, the warm-season
grasses did respond to mycorrhizal inoculation. Additionally, the root length was
significantly increased in the warm-season grasses with mycorrhizal infection when
compared to the cool-season grasses. Through the aid of topological analysis of root
architecture, mycorrhizal symbiosis was shown to inhibit root branching in warm-season
grasses, but had no effect on cool-season grass rooting (Hetrick et al., 1991). The
researchers concluded that mycorrhizal-dependent warm-season grasses have unique root
architecture, allowing energy to be conserved for root development, while the less
dependent cool-season grasses do not exhibit the same benefits of mycorrhizal infection.
In studies designed to determine the dependence of warm-season grasses on
arbuscular mycorrhizae and relationships between mycorrhizae and P availability and
plant density, Brejda et al. (1993) and Hetrick et al. (1994) evaluated sand bluestem
(Andropogon geradii var. paucipilus Nash), switchgrass (Panicum virgatum L.), and
Canada wild rye (Elymus canadensis L.).
The popular cool-season grasses, creeping bentgrass (Agrostis stolonifera L.) and
Kentucky bluegrass (Poa pratensis L.) were evaluated in relation to the impact of
arbuscular mycorrhizae and P status on plant growth (Charest et al., 1997). The authors
revealed that as mycorrhizal infection increased in the grasses, root colonization
increased to more than 40% with lowered P fertilization. This information could be
particularity helpful in warm-season grasses where P may have a major impact in soils,
such as those found throughout Florida. The researchers of this study concluded that
arbuscular mycorrhizal symbiosis could be considered as a potential fertilizer reduction
agent (Charest et al., 1997).
More recently, mycorrhizal symbiosis and fertilizer relationships have dominated
arbuscular mycorrhizal research; however, the majority of this work has concerned cool
and warm-season prairie grasses. The emphasis of molecular technologies has resulted in
less applied types of research being performed with grasses and mycorrhizas. Using
terminal restriction fragment length polymorphism (T-RFLP), Vandenkoornhuyse et al.
(2003) assessed the diversity of arbuscular mycorrhizal fungi in various cool-season
grasses, which co-occurred in the same research plots. Based on a clone library, the level
of diversity was consistent with past studies; showing that mycorrhizae fungal host-plant
preference exists, even between grass species.
Obviously, there is limited information on warm-season turfgrasses when
compared to the warm-season prairie and cool-season meadow grasses. In the Southeast,
warm-season turfgrasses are highly valued for their drought resistance, aesthetic
importance and generally low maintenance on some home lawns, golf courses, soccer,
and football fields. Species such as bermuda, St. Augustinegrass, seashore paspalum
(Paspalum vaginatum Swartz), zoysia (Zoysia sp.) bahia, and centipede are used in
landscapes throughout Florida. St. Augustinegrass is dominant residential species in
Florida (Trenholm, 2004). Haydu et al., (2002) estimated that 36% of the total lawn
acreage in Florida, or 1.5 million acres, was comprised of St. Augustinegrass in 1996.
Valued for its shade tolerance, ability to adapt to various soils, and color, St.
Augustinegrass cultivars such as 'Floratam', 'bitterblue', 'Raleigh', and 'Floratine'
became popular with home owners. Chinch bug resistant 'Floratam' quickly became the
number one cultivar upon its release in the 1970's. St. Augustinegrass is a desirable
species home lawn, however problems with disease susceptibility can be devastating.
Two examples are brown patch (Rhizoctonia solani Ktuhn) and take-all root rot
(Gaeumannomyces graminis (Sacc.) Arx & D. Olivier var. graminis).
To date, research evaluating the potential benefit of mycorrhizae in St.
Augustinegrass has been neglected such as reduced fertilizer use and production cost.
The method of production of St. Augustinegrass may result in limited benefits of
mycorrhizal research. St. Augustinegrass is produced vegetatively as sod throughout the
southeast. Once or twice a year, the sod is harvested leaving "ribbons" or strips of grass
behind. These ribbons are responsible for re-growth, through stolons, of the sod field.
Harvesting cycles would make lengthy mycorrhizal studies difficult. An extensive
survey of this plant system in relation to the arbuscular mycorrhizal fungi is warranted.
The overall objective of this research is to investigate the impact of mycorrhizal
fungi on warm-season turfgrasses in Florida. A survey of the population and
identification of arbuscular mycorrhizal fungi associated with St. Augustinegrass roots in
Florida sod is provided in Chapter II. In Chapter III, a survey of root pathogens is
explored in relation to arbuscular mycorrhizal colonization in sod production fields.
Chapter IV includes studies designed to determine whether or not arbuscular mycorrhizal
fungi affect root disease caused by pathogenic isolates of R. solani and G. graminis var.
graminis, and if potential affects are direct fungal interactions or indirect systemically
acquired mechanisms of resistance. In Chapter V, a general summary and conclusions
concerning arbuscular mycorrhizal fungi in St. Augustinegrass in Florida are provided.
POPULATION AND IDENTIFICATION OF ARBUSCULAR MYCORRHIZAL
FUNGI IN ST. AUGUSTINEGRASS
There is no information regarding arbuscular mycorrhizal fungi (AMF) in the
popular warm-season St. Augustinegrass (Stenotaphrum secundatum). In Florida, St.
Augustinegrass sod is a valuable commodity in home lawns and commercial landscapes.
'Floratam' the most common and widely adaptable cultivar is extensively used across the
state. It is also the primary cultivar grown in Florida for sod. In north central Florida,
sod production is increasing and growers are eager to increase production and lower
pesticide and fertilizer inputs. No information exists about mycorrhizas in this species.
The information is potentially useful in sod management to reduce disease severity,
chemical usage, and other production costs. In most cases, AMF populations are
decreased by agricultural practices are associated with conventional farming. St.
Augustinegrass sod production is unique in that it is not a traditional or natural plant
system. Currently, no information is available to growers to make informed decision
about inoculation with these fungi. The feasibility of inoculation studies for nutrient
acquisition, pesticide, and disease management can be performed using mycorrhizal fungi
more efficiently in the future once St. Augustinegrass is determined to be mycorrhizal.
Of current interest to mycorrhizal researchers is the ecology of mycorrhizal
populations and their benefit to both organic and more conventional cropping systems.
Information from less natural and conventional systems like St. Augustinegrass
sod is timely and could shed light on a little known ty cropping method. Mycorrhizal
systems and those interactions within it are complex and require extensive evaluation,
especially in crops not yet known to possess such associations. This evaluation may
supply valuable answers about mycorrhizal ecology. The objective of this study is to
determine if AMF colonize St. Augustinegrass, to what extent, and to identify the
Materials and Methods
Sampling.|| 'Floratam' St. Augustinegrass plant roots and associated soil were
collected monthly from three sod farms in three counties (Marion, Bradford, and Union)
in north central Florida from December 2004 through December 2005 with the exception
of July. Each of the sod farms had been cropped with 'Floratam' St. Augustinegrass for
12 years or more (Fig. 2-1 A-C).
Ten subsamples of soil were taken from three (3 m2) plots per sod farm with a
1.27 cm diameter soil probe to a depth of approximately 15 cm as suggested by Brundrett
et al. (1995). Root samples from each plot were extracted with a small hand trowel.
Subsamples of roots and soil from each plot were pooled, resulting in three separate
composite plot samples per location. Root samples were placed into plastic ziplock bags
separate from soil samples and stored at room temperature for approximately 1 d prior to
spore extraction and root manipulation for mycorrhizal evaluation. Approximately 200 g
of field soil from each plot were combined with 200 g of a low P, low organic matter soil
mined from the UF/IFAS Plant Science Research and Education Unit in Citra, Florida.
This soil was then potted into 10 cm clay pots sown with sorghum-sudangrass hybrid
seed (Sorghum bicolor [L.] Moench x Sorghum sudanense) cv. Summergrazer III. Low P
soil was used in pot cultures to enhance sporulation of potentially cryptic species in order
to facilitate their recovery and identification (Fig. 2-2).
The cultures were incubated for 60 d at 20-25 C with 12 h artificial light
(day/night). The seed was surface-sterilized using a 10% sodium hypochlorite and
deionized water solution for 30 sec and rinsed for 1 min with sterile deionized water prior
to planting. The pot cultures received a Peter's 20-0-20 (Spectrum Group, St. Louis,
MO) nutrient solution, devoid of P, every two weeks. Approximately 90 d later, single
spores from the field soil pot cultures were selected from spore extracts (Fig. 2-3). This
process was accomplished by wet sieving, decanting (Gerdemann and Nicolson, 1963),
and 40% sucrose (v/v) centrifugation (Jenkins, 1964). These spores were used to
inoculate sterile, low P soil (Citra, Florida) and sorghum-sudangrass hybrid seed for
spore production and subsequent identification of the sporulating AMF as suggested by
Gerdemann and Trappe (1974). The soil was sterilized twice for 90 min at 121 C at 15
psi for two consecutive days. Samples of field soil were also submitted to the IFAS
Extension Soil Testing Laboratory in Gainesville, Florida on a tri-monthly basis for soil
nutrient composition and pH testing. Soil pH, from all three fields, ranged from 5.6 to
7.0 during the 12 month sampling period. Phosphorous levels ranged from 5 to 119 ppm.
Root preparation. 11 Young, healthy-appearing fibrous roots were rinsed in tap water
and separated with a scalpel from the plant crown and/or seminal roots. Selected roots
were cut into 1-2 cm long segments and cleared of cell and wall components in 10%
KOH (w/v) under pressure in an autoclave for approximately 20 min (Brundrett et al.,
1996). The root segments were cooled, then rinsed in tap water, and placed into hot
0.05% trypan blue with glycerol overnight to stain mycorrhizal structures (Bevege, 1968;
Phillips and Hayman, 1970; Kormanik and McGraw, 1982). Excess stain was rinsed
from the root segments with tap water and then mounted in water on glass slides to view
vesicles and arbuscules. Slight pressure applied to the cover slip, with occasional heating
over an alcohol burner, aided in flattening the root segments adequately for microscopic
evaluation of mycorrhizal structures in root cells.
One hundred root segments were evaluated per sample for intensity of
colonization and to identify any variations in arbuscular morphology which might exist.
Mycorrhizal structures on glass slides were viewed with a Nikon Optiphot compound
microscope at 200, 400, and 1000x magnifications, and photographs were taken with a
Nikon CoolPix 990 digital camera. In order to judge the amount of mycorrhizal root
colonization, the grid line intersect method was used to estimate the total root length
colonized by AMF (Newman, 1966; Tennant, 1975; Giovannetti and Mosse, 1980).
Spore extractions. I Mycorrhizal spores were extracted by wet sieving and decanting
by mixing 100 g of air-dried sample soil with 300 ml of tap water, blending at low speed
in a commercial Waring blender for 1 min, and then allowed to settle for 1 min. The
supernatant was then passed through a series of Tyler 250, 125, and 38 mrn mesh sieves
(Daniels and Skipper, 1982). The remaining fraction was rinsed with tap water to remove
sediment and any organic materials left behind. The fraction was decanted into 50 ml
centrifuge tubes containing a 40% sucrose/deionized water solution (w/v) (Jenkins,
1964). The tubes were centrifuged for 3 min at 2,000 rpm in a Dynac III centrifuge. The
supernatant, containing the spores, was decanted off the top of the tube into a 38 .im
mesh sieve and rinsed to remove the sucrose. The extracted spores were collected in a
9 cm Petri dish with tap water rinse and viewed with a Zeiss dissecting scope.
Mycorrhizal spore densities were enumerated by using an ocular field method described
in the International Culture Collection of (Vesicular) Arbuscular Mycorrhizal Fungi for
high spore densities (Morton, 2005).
Intact and parasite free spores were selected using a Gilson 20 .il pipetman.
These spores were used to inoculate 10 cm diameter clay pots containing the low P,
sterile soil (as described above) and planted with surface-sterilized sorghum-sudangrass
hybrid seed. The monocultures were kept at 20-25 C for approximately 60 d. At that
point, any spores that had been produced as a result of the inoculations were extracted as
previously mentioned, and used to inoculate another crop of sorghum-sudangrass in
sterilized, low P soil. The second generation of monocultures were then maintained for
60-90 d and processed for spore extraction and mycorrhizal identification.
Arbuscular mycorrhizal fungi tleutificutioo ||I Identification of the mycorrhizal fungi
associated with St. Augustinegrass was accomplished by selecting healthy, single spores
with a 20 [il Pipetman and mounting in either sterile, deionized water or (1:1 v/v) PVLG
(polyvinyl alcohol-lactic acid) + Melzer's reagent (Khalil et al., 1992). The spores were
then viewed at 200, 400, and 1000x using a Nikon compound microscope and identified.
Using arbuscular mycorrhizal descriptions by Schenck and Perez (1988), a tentative
determination to genus was made based on the average measurement of 20 similar spores
per pot. The species was determined based on taxonomic descriptions from the INVAM
Species Guide (Schenck and Perez, 1988). Identifying characteristics of the
monocultured spores, such as spore wall number and width, hyphal appendages, the
presence or absence of germ shields, approximate overall spore diameter and color in
reagents, were used as described by Schenck and Perez (1988).
Statistical analysis. |I Spore density and percent colonization data were analyzed using
the General Linear Model procedure (SAS Institute, Version 9.0, 2004) (Appendix F-l).
The survey was performed using a random model in a randomized complete block design
with multiple samplings at multiple locations. The percent root colonization data were
transformed with the arcsine square root transformation prior to an analysis of variance
due to distribution of propagules within soil being highly variable resulting in a non-
normal frequency of distribution points (St. John and Hunt, 1983; Friese and Koske,
1991). Spore density data were transformed to their natural log prior to analysis of
variance to prevent violation of the assumption of normal distribution. Significant
interactions were separated using Tukey's Studentized Range Distribution test.
Correlations between percent colonization or spore density data, with soil nutrient
composition, and percent colonization to spore density were done in SAS using Pearson
product-moment correlation coefficients. Regression analyses also were performed with
the regression procedure in SAS.
Root Evaluation. || Roots, collected from sod fields evaluated in this survey revealed
the first evidence of an interaction between AMF and St. Augustinegrass. In stained
roots mounted on glass slides, AMF structures such as internal vesicles, intra and
extraradical hypha, and an assortment of arbuscular types were observed. Bulbous
appressoria (Fig. 2-4) were noted at inoculation points along the length of the root, giving
rise to carbohydrate storage vesicles of various shapes within cortical root cells (Figs. 2-
5, 2-6). Copious amounts of intra and extraradical hypha were observed within and along
the outer surface of root tissue (Fig. 2-7). Most notably, a variety of arbuscular types
were observed within the cortical root cells. Arbuscules, or haustoria-like structures,
have been categorized into two morphological types (Gallaud, 1904); Arum- and Paris-
types. These intercellular mycorrhizal structures are the presumed active fungal sites of
nutrient translocation between host and fungus (Bracker and Littlefield, 1973; Brundett et
In this study, field grown plant roots were found to contain both the Arum- and
Paris- type of arbuscules along with a variety of intermediate Arum- morphologies.
Intermediate forms of the Arum- type found in cortical root cells of St. Augustinegrass
sod plants ranged from a typical "feathery" form (Fig. 2-8) extending from intracellular
hypha to a "dense-compact" form between cells of conjoined intercellular hyphae (Fig. 2-
9). A "grainy" form (Fig. 2-10) was also found in cortical root cells on several occasions.
This could be a collapsing arbuscule instead of an intermediate arbuscular form. The
Paris-type arbuscule found in St. Augustinegrass plant roots shows a typical arbusculate
coil (Fig. 2-11) in the root cell, while intermediate forms were not observed. An unusual
structure was found along intercellular hyphae that resembled a hyphal mat with a
mantle-like appearance often found in conjunction with certain types of ectomycorrhizas
(Fig. 2-12). This may be a new arbuscular form found in the Poaceae. This structure
was only observed once in St. Augustinegrass plants harvested in April 2005 at the Fort
Spore density evaluation. | Further evidence supporting an interaction between AMF
and St. Augustinegrass was observed outside the root within the rhizosphere. AMF
spores clinging to epidermal tissue on roots were frequently observed in field samples
and in pot cultures using field soil from each farm location and sorghum-sudangrass as
the trap plant. The three sod farms sampled in this survey have been cropped solely in
'Floratam' St. Augustinegrass sod for more than 12 years. Weeds are heavily controlled
with herbicides at each location. The AMF spores recovered from field soil are entirely
dependent upon the St. Augustinegrass plants because they are obligate heterotrophs.
The limited availability of other plant species at each location, and the availability of
numerous spore types for pot culturing and subsequent AMF identification, provides
adequate evidence of AMF colonizing St. Augustinegrass plants in North Central Florida
Additional mycorrhizal structures such as auxiliary cells were frequently
observed in slide mounts of spores from both pot cultures and field soil (Fig. 2-13).
Selected single spores that appeared non-parasitized and viable, were chosen under light
microscopy for culturing in sterile, low P soil in order to obtain consistent spore
structures compatible with identification procedures. Spores, retrieved from pot cultures
were used as sieved soil sub-cultures to produce another generation of spores capable of
being readily identified from their morphological structures according to Schenck and
Perez (1988). Table 2-1 lists the species of AMF positively identified from sub-cultures
of soil from each location over a year-long period.
Species of Glomus were the most commonly encountered AMF in north central
Florida soils at each location. At the Lake Butler location, Glomus species included: G.
etunicatum Becker & Gerdemann (Fig. 2-14), G. intraradices Schenck & Smith (Fig. 2-
15, 2-16), G. reticulatum Bhattacharjee & Mukerji (Fig. 2-17, 2-18), and G. uggregutimln
Schenck & Smith (Fig. 2-19). Glomus species isolated at the Fort McCoy location
included: G. ambisporum Smith & Schenck (Fig. 2-20), G. formosanum Wu & Chen
(Fig. 2-21), G. macrocarpum Tulasne & Tulasne (Fig. 2-22), G. gerdemannii Rose,
Daniels & Trappe (Fig. 2-23), G. intraradices, and G. etunicatum.
Acaulospora spinosa Walker & Trappe (Fig. 2-24) and an unidentified species of
Scutellospora were isolated at Lake Butler. Additional AMF genera were found at Fort
McCoy including: Entrophospora infrequens [Hall] Ames & Schneider (Fig. 2-25), A.
denticulata Sieverding & Toro (Fig. 2-26), A. lacunosa Morton (Fig. 2-27), and
Scutellospora minute [Ferr. & Herr.] Walker & Sanders (Fig. 2-28). The Starke location
was unusual in species diversity with only 3 species isolated: Glomus etunicatum, G.
intraradices, and Scutellospora minute. One unique spore type was found at the Fort
McCoy location, but could not be grown in a pot culture successfully. The unidentified
spore type was observed on two occasions during the late spring of 2005 in very small
numbers and appeared to be either a species of Acaulospora or Entrophospora based on
morphology. Without a sufficient number of cultivated spores for microscopic
evaluation, positive identification of the species was not possible.
Sieving field soil from each location not only yielded spores for pot culturing, but
also enabled a numerical count of spore density, which is a good indicator of the
infectivity of the AMF in the soil and their level of activity in the rhizosphere. The total
spore density at the three locations ranged from 78 to 2,132 spores per 100 g of dry soil
(non-transformed data). Spore density but did not vary among or within sod farm
locations (P < 0.0001), indicating that variations in soil factors did not significantly affect
AMF spore production between locations from December 2004 through December 2005
(Table 2-2). Spore production did vary significantly (P < 0.0001) between monthly
sampling, which suggested a possible seasonal influence on spore production. Greater
spore density totals occurred in soils collected during the warmer summer and fall
months, as compared to, lowered spore production occurring in the cooler months of
winter and spring. Total spore density in December 2004 was significantly lower when
compared to December 2005. This might be explained by increased rainfall, prior to the
sampling period, in north central Florida during the 2004 hurricane season.
With spore densities varying between dates, analysis of variance for these points
showed a significant date by location interaction (P < 0.05) indicating that seasonal
effects and unknown variations in site-related effects might measurably influence the
total spore density. In this survey, rainfall and soil moisture where positively correlated
to spore density (Table 2-3).
Based on the regression equations, a quadratic response was generated in total
spore density to soil moisture at each location. Spore density at the Starke location
increased at soil moisture levels between 0 and 2 cm, but declined until soil moisture
levels reached 6 cm where another increase was observed (Fig. 2-29). Above 9 cm a
decrease in spore density occurred (r=0.73). The same general response to soil moisture
was noted at the Fort McCoy location except where soil moisture declined to
approximately 8 cm (r=0.61) (Fig. 2-30). At the Lake Butler location, spore density
increased slightly until soil moisture levels reached 7 to 8 cm when a slight decline in
spore density was observed (r=0.68) (Fig. 2-31). This lends credibility to the theory that
excessive rainfall during the hurricane season of 2004 lowered spore production in
December of that year.
A quadratic response was also produced in total spore density to temperature at
each location. Spore density at the Starke location (r= 0.60) (Fig. 2-32) decreased from
15 C until the temperature reached 20 C. Between 20 C and 28-29 C a gradual increase
in spore density was observed until the temperature reached 30 C. At that point there was
another gradual decrease in spore density, which seemed to level off near 35 C. At the
Fort McCoy location (r=0.84) a gradual increase in spore density was observed until the
temperature was approximately 28-29 C, then a decline was noted (Fig. 2-33). At the
Lake Butler location (r=-0.59) a slight increase in spore density occurred across all
temperature ranges (Fig. 2-34). Based on these data, it appears that soil temperatures
above 28-30 C have a detrimental effect on the AMF. In addition, this temperature range
might also damage host root tissue.
Percent colonization evaluation. |I Percent root length colonized by AMF yielded no
significant difference among or within location differences, but there was a significant
date interaction (P < 0.0001). Colonization was generally highest in the cooler months of
winter and spring, with lower colonization occurring in the warmer summer and fall
months except in December 2005, when colonization was the lowest. The amount of root
length colonized ranged from 13 to 39% across the sampling dates (non-transformed
data). No correlation was found between temperature and soil moisture in relation to
percent root length colonized (Table 2-4).
Dickson (2004) suggested an Arum-Paris continuum of mycorrhizal symbioses in
a survey of 12 colonized plant families, with arbuscule formation dependent on the
fungus as well as the host plant. Most mycorrhizal angiosperms were once thought to
only produce the Arum-type of arbuscule, which consists of both intercellular hyphae and
arbuscules, while most angiosperms and bryophytes were thought to only produce the
Paris-type with intercellular hyphae and arbuscular coils (Dickson, 2004). The majority
of scientific research has been conducted on flowering plants versus trees and bryophytes
causing these fallacies to be argued as fact until Smith and Smith (1997) produced a
comprehensive list of plant families that included their arbuscule types. The list showed
that the Paris-type is in fact most common among all plant families and that,
"intermediate" or transitional arbuscular morphotypes were observed in some plant
species. One genus (Ranunculus) forms both types within the same plant (Smith and
Experiments on maize (Zea mays) and the tuliptree (Liriodendron tulipfera),
among many others, revealed that AMF can form either type of arbusculate structure
based on the host plant (Barrett, 1958; Gerdemann, 1965). In a field experiment using
tomatoes (Lycopersicon esculentum) and other annual crops, investigators found that
arbuscule morphology is actually dependent on intercellular spaces in cortical root cells
(Brundrett and Kendrick, 1988; Cavagnaro et al., 2001). Intermediate forms of the Arum
and Paris-type arbuscules are common in certain plant families such as those described in
three cultivars of flax (Linum usitatissimum), which Dickson et al. (2003) referred to as
arbuscules "in pairs in adjacent longitudinally arranged cortical cells arising from a
single, radial intercellular hyphae."
On rare occasions, both arbuscule types (Arum and Paris) occur in the same plant
species, which Smith and Smith (1997) noted in the family Poaceae. The Paris- and
Arum- types were found in millet, ryegrass, and wheat. In addition, a series of
intermediate forms between the two main types of arbuscules were also observed. The
same can be said for St. Augustinegrass plants in relation to AMF colonization. In field
studies, environmental effects may interact to influence fungal and plant response to the
mycorrhizal interaction. Sylvia et al. (1993) suggested that even in the presence of high
amounts of soil P, water stress and pesticide applications can have extensive effects on
mycorrhizal response. Rabatin (1979) noted that soil moisture may have the greatest
effect on the degree of infection of Glomus species in field situations. Furthermore, the
stages of plant development (Saif and Khan, 1975) as well as temperature (Giovannetti,
1985; Schenck and Kinloch, 1980; Smith & Smith, 1997; Sylvia, 1986) all play a major
role in mycorrhizal activity.
In this survey, AM fungi preferred warmer months for spore production and
cooler months for colonization of St. Augustinegrass plants. In the north central region
of Florida, St. Augustinegrass does not usually go completely dormant in cooler
temperatures, and there is usually some plant activity during the winter months especially
in the roots where AMF colonization occurs. This increase in colonization during cooler
temperatures may be an effort to preserve valuable carbon and energy reserves for future
spore production. Subsequent proliferation in the warmer months, while the plant host is
most active, would provide more carbohydrates from a symbiotic interaction (Johnson et
al., 1997). It is also possible that AMF are actually acting as a parasite in the winter
months when colonization is highest while the plant is less active.
During less than optimal winter growing conditions, the St. Augustinegrass plant
is less able to defend itself against infection and colonization due to lowered metabolic
activity. Johnson et al. (1997) suggested a mycorrhizal continuum ranging from
mutualistic to parasitic in some managed habitats where humans unknowingly altered the
association through management regimes. Another possibility is environmentally
induced parasitism due to morphological, phenological, and physiological differences in
the symbionts which may influence the mycorrhizal association (Johnson et al., 1997).
Conversely, in natural habitats, mycorrhizal associations have evolved over many
years to encourage fitness in the plant and the fungus making the interaction continually
mutual (Johnson et al., 1997). St. Augustinegrass sod systems are not traditional
cropping systems needing continual management inputs from man, nor are they a natural,
non-impacted habitat. St. Augustinegrass sod could be referred to as a non-conventional
cropping system due to minimal inputs after harvesting where ribbons of grass are left
behind for re-growth. Cloned host plants are in constant supply in sod fields providing
the AMF with a dependable host, but when the plant is semi-dormant throughout the
winter months the fungi may actually pose a threat to the health of the plant because net
costs in carbon might then exceed net benefits in some situations. For example, during
instances of lowered metabolic activity in the winter, plants lower photosynthetic ability
and subsequent output and will not benefit from the added benefits of a mutual
interaction. Acquisition of nutrients and water is less important during these times, but
St. Augustinegrass may be harmed by the loss of stored carbon to AMF. Throughout the
year, there are potential times when the interaction between plant and AMF is such that
the symbiosis might actually be neutral in nature (Johnson et al., 1997).
An attempt was made in this survey to correlate spore density to the percent root
length colonized, but no correlation was found. Some researchers have reported a
correlation between the two variables (Giovannetti, 1985; Miller et al., 1979) while
others have observed no such relationship (Giovannetti and Nicolson, 1983; Hayman and
Stovold, 1979). This is most likely due to the vast variations observed in soils, plant
species and their developmental stage, and fungal specificity. Many mycorrhizal studies
suggest a significant interaction with soil P where spore production or colonization is
lowered by increasing levels of P. Correlations between soil chemical characteristics
such as P content to spore density and percent root colonization have been reported in
grasses (Brejda et al., 1993). Others suggest that mycorrhizal ecology plays less of a
role. P content in south Florida soils had no effect on AMF in tropical forage legume
pastures (Medina-Gonzalez et al., 1988), nor did potassium or pH in studies of cultivated
soils (Abbott and Robson, 1977; Hayman, 1978).
In this survey, soil samples from each location were evaluated during the months
of January, April, August, and November 2005 in an attempt to correlate soil Mg, Ca, K,
P, soil pH, and organic matter percentage to spore density and/or percent root length
colonized, but a correlation was not observed (Table 5). One theory to explain the lack of
correlation between AMF and P content, in this case, might be explained by asexual
organisms, without the cost of sexual reproduction and consequently no genetic
variability, and having scores of mutations that accumulate over a long period of time
(Helgason and Fitter, 2005). The Glomeromycota possess ancient asexual lineages
(Gandolfi et al., 2003). This apparent genetic isolation would presumably cause
mutations to allow for some adaptations such as P tolerance. In AMF the coenocytic
mycelium is multinucleate providing a set of mutations within the DNA of all nuclei
(Helgason and Fitter, 2005). Reductions in fitness due to a lack of genetic variability due
to asexual reproduction may never be noticed in AMF because mutated, non-functional
genes from one nuclear lineage might be subjugated by functional alleles on another
nucleus (Helgason and Fitter, 2005).
Arbuscular mycorrhizal fungi in these sod fields are secluded, thus reducing
genetic variability, so it is possible that the ancient fungi are capable of evolving and
adapting through mutations to tolerate large amounts of added nutrients like P. P is
widely used in large amounts in St. Augustinegrass to promote root growth and health for
winter survival and spring green-up. Through years of isolation in sod fields and large
applications of P on a frequent basis, these fungi might have evolved a mechanism
through spontaneous mutation to tolerate elevated P levels. This is speculation, but the
lack of spore density and percent colonization variable correlation to P levels could be
due to genetic mutation in the fungi within these fields leading to a significant adaptation
and evolutionary event.
Overall root colonization and spore density were low to moderate, which suggests
that the AMF populating St. Augustinegrass sod production soils are moderately active.
This situation might lend itself to field inoculation where AMF could potentially provide
a level of root disease protection, which might lower pesticide use and cost. It could also
lead to increased and more efficient P acquisition and use when combined with more
conducive management strategies. On the other hand, inoculation with AMF might be
ineffective in situations where genetic isolation combined with perennial cropping and
moderate to heavy fertilizer inputs are unavoidable for proper management.
Figure 2-1 A-C. 'Floratam' St. Augustinegrass sod farms located at (A) Fort McCoy
(Marion County), (B) Lake Butler (Union County), and (C) Starke (Bradford
County) in north central Florida.
Fig. 2-2. Sorghum-sudangrass pot cultures containing 50% (w/w) field soil combined
with 50% sterile, low P soil.
Fig. 2-3. Spore extract from field soil following the wet sieving procedure.
Figs. 2-4 2-7. Stained arbuscular mycorrhizal structures observed within 'Floratam' St.
Fig. 2-4. Bulbous appressoria found originating from extraradical hypha.
Bar = 40 im.
Fig. 2-5. Circular type of AMF vesicle stained with trypan blue. Bar = 40 rim.
Fig. 2-6. Oblong type of AMF vesicle stained with trypan blue. Bar = 40 im.
Fig. 2-7. Extraradical hyphae observed with light microscopy infecting
and colonizing roots. Bar = 20 rim.
Figs. 2-8 2-11. Stained arbuscular morphology types found within 'Floratam'
Fig. 2-8. Feathery form of the Arum-type arbuscule morphology, stained
with trypan blue, within cortical root cells. Bar = 40 inm.
Fig. 2-9. Dense and compacted Arum-type arbuscule morphology stained
with trypan blue. Bar = 40 inm.
Fig. 2-10. Grainy or collapsing Arum-type arbuscule morphology stained
with trypan blue. Bar = 40 inm.
Fig. 2-11. Paris-type coiled arbuscule, stained with trypan blue, within
cortical root cells. Bar = 40 inm.
Fig. 2-12. Net-like AMF structure observed in roots across adjacent
cortical root cells. Bar = 20 inm.
Fig. 2-13. Auxiliary cells of an AMF observed in spore extracts from field
soil. Bar = 40 nm.
Table 2-1. Species of AMF positively identified at each sod farm location from pot
cultures of sorghum-sudangrass within a combination of field and sterile, low
Figs. 2-14 2-19. Arbuscular mycorrhizal fungal spores identified at the Lake
Butler sod farm location.
Fig. 2-14. A spore of Glomus etunicatum stained in Melzer's reagent. Bar = 20 [im.
Fig. 2-15. A spore of G. intraradices in deionized water. Bar = 20 [im.
Fig. 2-16. Spore wall morphology of G. intraradices spore stained in
Melzer's reagent (arrows point to cell wall layers). Bar = 40 [im.
Fig. 2-17. A spore of G. reticulatum in deionized water. Bar = 20 ism.
Fig. 2-18. Spore wall morphology of G. reticulatum in deionized water
(arrows point to cell wall layers). Bar = 40 ism.
Fig. 2-19. A broken spore of G. aggregatilni in Melzer's reagent. Bar = 20 ism.
20 1_ 22
26 -_ 27 28
Figs. 2-20 2-28. Arbuscular mycorrhizal fungal spores identified at the Fort
McCoy sod farm location.
Fig. 2-20. A spore of Glomus ambisporum stained in Melzer's reagent. Bar = 20 rim.
Fig. 2-21. A spore of G. formosanum stained in Melzer's reagent. Bar = 20 rim..
Fig. 2-22. A spore of G. macrocarpum stained in Melzer's reagent. Bar = 20 rim.
Fig. 2-23. A spore of G. gerdemannii stained in Melzer's reagent. Bar = 20 rim.
Fig. 2-24. A spore of Acaulospora spinosa stained in Melzer's reagent. Bar = 20 rim.
Fig. 2-25. A spore of Entrophospora infrequens stained in Melzer's reagent.
Bar = 20 rm.
Fig. 2-26. A spore of A. denticulata stained in Melzer's reagent. Bar = 20 rim.
Fig. 2-27. A spore of A. lacunosa stained in Melzer's reagent. Bar = 20 rim.
Fig. 2-28. A spore of Scutellospora minute stained in Melzer's reagent. Bar = 20 rm.
Table 2-2. Evaluation of analysis of variance data for spore density data from each sod
farm location by date.
Dec. '04 Fort McCoy
March '05 Fort McCoy
April'05 Fort McCoy
May'05 Fort McCoy
June'05 Fort McCoy
Aug'05 Fort McCoy
Sept'05 Fort McCoy
Total Spore Density
mean 5.00 d
mean 5.38 cd
mean 5.71 bcd
mean 5.72 bcd
mean 6.55 a
mean 6.73 a
mean 6.66 a
mean 6.26 ab
mean 6.05 abc
Nov'05 Fort McCoy
mean 6.14 abc
mean 6.51 a
mean 5.99 abc
tEach value is the average of three sample plots/location (10 sub-samples/plot).
Means followed by the same letter are not significantly different according to
Tukey's (HSD) Studentized Range Test (P = 0.0001).
Table 2-3. Pearson correlation coefficients
(r) for AMF spore density and soil moisture
*** Significant at P = 0.0001, respectively.
f Percolon = percent root length colonized; Sporeden = spore density;
Rainfall = amount of rainfall in month preceding sampling date;
Soiltemp = soil temperature for sampling date.
Soil Moisture (cm)
Fig. 2-29. Spore density with increasing soil moisture levels over a 12-month period at
the Starke sod farm location.
Soil Moisture (cm)
Fig. 2-30. Spore density with increasing soil moisture levels over a 12-month period at
the Fort McCoy sod farm location.
y = -0.0264X4 + 0.4465x3 2.3561x2 + 4.4331x + 3.4988
y = -0.0067X4 + 0.1113x3 0.6217x2 + 1.5001x + 4.6121
R2 = 0.6136
Soil Moisture (cm)
Fig. 2-31. Spore density with increasing soil moisture levels over a 12-month period at
the Lake Butler sod farm location.
y = -3E-06x6 + 0.0004X5 0.028x4 + 0.9465x3 17.252x2 + 160.12x 583.17
R = 0.6062
15 20 25 30 3!
Soil Temperature (C)
Fig. 2-32. Spore density with increasing soil temperatures over a 12-month period at the
Starke sod farm location.
y = 0.0118x4 0.195x3 + 0.9987x2 1.4413x + 5.8091
R2 = 0.6888
y = 0.0002X4 0.0206X3 + 0.8144x2 13.691x + 88.104
R2 = 0.8455
Soil Temperature (C)
Fig. 2-33. Spore density with increasing soil temperatures over a 12-month period at the
Fort McCoy sod farm location.
Fig. 2-34. Spore density with increasing soil temperatures over a 12-month period at the
Starke sod farm location.
STy = 1 E-05x4 + 0.0009x3 0.1124x2 + 3.1385x 20.969
R2 = 0.5939
15 20 25 30 35
Soil Temperature (C)
Table 2-4. Evaluation of analysis of variance data for percent root length colonized from
each sod farm location.
Jan'05 Fort McCoy
Feb '05 Fort McCoy
March'05 Fort McCoy
April '05 Fort McCoy
May'05 Fort McCoy
June'05 Fort McCoy
Aug'05 Fort McCoy
Sept '05 Fort McCoy
Oct '05 Fort McCoy
Nov '05 Fort McCoy
mean 26.13 ab1
mean 29.01 a
mean 28.85 a
mean 24.84 abc
mean 28.58 a
mean 26.73 ab
mean 25.92 ab
mean 22.63 bcd
mean 21.68 bcd
mean 19.50 cd
Lake Butler 20.94
mean 19.91 cd
Dec '05 Fort McCoy 18.87
Lake Butler 19.90
mean 18.68 d
tEach value is the average of three sample plots/location (10 sub-
Means followed by the same letter are not significantly different according
to Tukey's (HSD) Studentized Range Test (P = 0.0001).
Table 2-5. Chemical characteristics of soils sampled for AMF at three north central
Florida sod farm locations during January, April, August, and November
Soil Nutrient Levels
tSoil pH, nutrient level, and organic matter content based on the mean of three composite
OM = Organic matter content.
*FM = Fort McCoy location.
**LB = Lake Butler location.
THE EFFECT OF ARBUSCULAR MYCORRHIZAL FUNGI ON
GAEUMANNOMYCES GRAMINIS VAR. GRAMINIS AND RHIZOCTONIA
SOLANI COLONIZATION OF ST. AUGUSTINEGRASS SOD IN NORTH CENTRAL
Take-all root rot and brown patch are two of the more common and devastating
diseases of St. Augustinegrass sod throughout Florida. Take-all root rot, caused by
Gaeumannomyces graminis (Sacc.) Arx & D. Olivier var. graminis, is a disease of both
grasses and cereals (Nilsson, 1969; Huber and McCay-Buis, 1993). Take-all root rot was
first described in Sweden in the early 1800's infecting grasses (Mathre, 1992). It is one
of several G. graminis varieties which infect many important crops worldwide (Rovira
and Whitehead, 1983). This particular variety of the fungus infects all cultivars of St.
Augustinegrass (Elliott, 1995; Datnoff et al., 1997). In the late 1980's, large, chlorotic
patches of St. Augustinegrass were observed on sod farms in South Florida and were
confirmed as the first disease symptoms of G. graminis var. graminis infection observed
in this species (Elliott, 1993). The disease was found in St. Augustinegrass throughout
Alabama, Florida, and Texas (Fig. 3-1) and it is notably more severe in the summer and
fall months, especially during periods of increased precipitation (Elliott, 1993). Early
studies suggested that the fungus preferred alkaline or high pH soil, mild winters, thatch-
accumulation and frequent light irrigation, however the conditions that predisposed the
stand to disease or prompted disease escape are not known (Guyette, 1994).
Management recommendations included elimination of low areas where water
accumulates, watering only when needed, and the use of pH decreasing
fertilizers in the fall, as well as thatch prevention and aeration (Guyette, 1994).
Fungicides were recommended as preventative but not curative treatments, which limited
management options to growers (Guyette, 1994). The effect of systemic fungicides on G.
graminis var. graminis infection and colonization of turfgrasses was evaluated; but
results indicated that preventative and/or curative rates of fungicides did not limit take-all
root rot disease or increase turfgrass quality (Elliott, 1995). Biological controls were
explored in an attempt to decrease take-all root rot in wheat and turfgrasses. The effects
of bacterial isolates, actinomycetes, and fluorescent pseudomonads on the roots of wheat
were evaluated as antagonists against G. graminis var. tritici (Sivasithamparam and
Parker, 1978). These organisms make up a large portion of the microbial community of
soils and researchers expected their production of antibiotics or toxic metabolites would
inhibit take-all in wheat in suppressive soils. While combinations of these
microorganisms reduced disease, none were successful alone (Sivasithamparam and
Parker, 1978). To date, no effective curative or preventative controls for take-all root rot
are recognized for use in St. Augustinegrass.
In order to determine the impact of arbuscular mycorrhizal fungi (AMF) on take-
all root rot in St. Augustinegrass sod, it is necessary to accurately diagnose G. graminis
var. graminis and determine its population within the field. The diagnosis of take-all root
rot involves several characteristics and diagnostic tools for isolation and identification.
The pathogen is somewhat elusive and may be easily confused with other fungi if the
scientist is not familiar with the morphology of the fungus and patterns of infection. The
ascomycete, G. graminis var. graminis, is classified in the order Diaporthales because it
produces ascospores in black, flask-shaped, ostiolate perithecia, which are fully enclosed
and lined with hyaline periphyses (Landschoot, 1997; Walker, 1973). The perithecia are
typically 200-400 [m x 150-300 C[m in length, with the neck portion 100-400 C[m in
length and 70-100 C[m wide (Landschoot, 1997). The asci, clavate in shape, are
unitunicate, are formed in a hymenium, and range in length from 80-140 C[m and 10-15
[lm in width. The apex of the ascus, which has a refractive apical ring, is generally
yellowish en masse. Each ascospore is typically 70-110 C[m in length, 2-4 C[m in width
and they usually contain 3-8 septa, but there may be 11 or 12 septa produced. The
anamorphic state, which is rarely observed, is a Philaphora species that produces conidia
5-14 C[m in length x 2-4 C[m in width. The use of conidia as taxonomic criterion is not
recommended due to variation between isolates and their non-descript morphology. In
culture, mycelia range from short to aerial, white to gray, green to brown, or black
Dark runner hyphae are typically observed on and around the crown portion of the
plant, with extension onto the stem and stolons. The roots usually have relatively fewer
dark surface runner hyphae, compared to the foliar portion of the plant, which may
remain green. Instead of dark runner hyphae, the roots are often covered with dark
brown to black lesions and subsurface hyaline hyphae. The cortical browning of roots is
thought to be a host defense mechanism, while the discoloration of shoots is a necrotic
symptom of disease (Penrose, 1992). The name "take-all root rot," implies that the roots
are the first plant parts to be severely affected whether facilitated by feeding damage
from nematodes or mole crickets, mechanical damage from sod production, cultural
techniques, or through natural openings.
After the initial invasion, the seminal roots are colonized internally by more
hyaline and infectious, secondary hyphae usually right behind the root tip (Henson et al.,
1999; Gilligan, 1983), which is were AMF usually colonize root tissue. Pathogenic
colonization causes an occlusion of vascular tissues resulting in the characteristic gradual
decline in plant health and potential death. Dark runner hyphae may continue up the
plant in search of more juvenile and susceptible tissue while producing deeply lobed and
The hyphopodia are considered by most as superficial hyphal structures (Henson
et al., 1999) since they originate from the hyphae, however they behave much in the same
way as appressoria, which develop from the germ tube of germinating fungi providing
infection pressure and anchoring the fungus to plant tissue (Agrios, 2004). Hyphopodia
cluster and develop into an infection cushion which provides the added structural stability
while helping to maintain the turgor pressure required for colonization (Henson et al.,
1999). The force of exertion of G. graminis var. graminis is associated with reduced cell
wall permeability, turgor, and wall rigidity (Bastmeyer et al., 2002). The deeply lobed
hyphopodia are unique to G. graminis var. graminis and may exist to allow the fungus to
overcome plant resistance mechanisms. Plants of St. Augustinegrass may benefit from
AMF colonization in the presence of Gaeumannomyces graminis var. graminis. But, it is
possible for AMF to have a negative impact on plants in some situations, or they may
even be neutral in nature (Johnson et al., 1997).
Brown patch or Rhizoctonia blight, caused by Rhizoctonia solani Ktuhn (Figs. 3-2,
3-3), is most active in St. Augustinegrass from November to May when temperatures
25 C and below (Elliott and Simone, 2001). Brown patch is typically worse in periods of
excessive rainfall or irrigation, or when grass leaves remain wet for more than 48 hours
(Elliott and Simone, 2001). In the field, small chlorotic patches of sod gradually turn
brown as infected leaf blades die, hence the name brown patch (Elliott and Simone,
2001). As patches expand, they may coalesce into large rings of yellow-brown sod with
dark and wilted margins. It is not uncommon for sod to appear green and healthy in the
center of the rings. Grass blades are killed near the crown due to restriction of water and
nutrient transport, which creates a dark rot near the base of the blade. Infected blades can
easily be pulled from the leaf sheath due to the soft rot (Elliott and Simone, 2001). Most
usually the stolons and leaves are affected more than the roots themselves. A barrage of
chemical controls, such as azoxystrobin, fluotanil, and mancozeb offer effective brown
patch control when used as preventatives. Cultural controls include irrigating only when
necessary between 2 and 8 AM and removal of mower clippings from the site. However,
the use of quick release nitrogen during periods of R. solani activity seems most
beneficial (Elliott and Simone, 2001). The use of chemicals in sod production has been
controlled in recent years and these restrictions will continue according to state and
federal regulations. Effective disease prevention strategies including the use of biological
controls, such as AMF, are essential research objectives in an industry where quality is of
utmost importance to buyers and growers.
Brown patch was first described in St. Augustinegrass in the 1980's (Hurd and
Grisham, 1983; Martin and Lucas, 1984) as an aerial type of pathogen common to a
variety of crops including corn, soybean, and rice (Sneh et al., 1991). Other pathogenic
species of Rhizoctonia affecting St. Augustinegrass include R. oryzae Ryker & Gooch
and R. zeae Voorhees which cause a sheath rot or spot, but the two species are rare
(Martin and Lucas, 1984; Haygood and Martin, 1990). The telomorph, Thanatephorus
cucumeris Frank, is assigned to the Basidiomycota (Ainsworth et al., 1973). Mycelia of
R. solani appear buff to dark brown in culture with irregularly shaped light to dark brown
sclerotia (Sneh et al., 1991). Rhizoctonia solani is identified by its characteristic right
angle (900) branching between the primary and secondary hypha (Duggar, 1915) with
branches forming acute (450) angles to main hypha (Butler and Bracker, 1970).
Identification is made easier by the presence of a septum at the branches near hyphal
constrictions at the base of right angles (Duggar, 1915). Additionally, the older, main
runner hypha of R. solani are more than 7 [im in diameter with more than two nuclei per
cell (Sneh et al., 1991).
Arbuscular mycorrhizal fungi have been associated with increased nutrient and
water acquisition in plants for many years. Mycorrhizal symbiosis often results in
increased plant vigor and the use of AMF has been studied in many crops as potential
antagonists to root pathogens (Schenck, 1987; Sylvia and Williams, 1992; Smith and
Read, 1997; Yao et al., 2002). Glomus etunicatum Becker & Gerdemann and G.
intraradices Schenck & Smith are two of the more common AMF species investigated as
potential biological controls and chemical alternatives against R. solani in crops such as
potato (Yao et al., 2002) and species of Fusarium in tomato crops and alfalfa (Caron et
al., 1986; Hwang et al., 1992). In several cases, G. intraradices provided significant
control of soilborne pathogens (Niemira et al., 1996; Khalil et al., 1994; Viyanak and
Bagyaraj, 1990). Newsham et al., (1995) reported that mycorrhizal fungi are capable of
protecting annual grasses from soilborne fungi. In other surveys, researchers found that
G. intraradices significantly reduced take-all root rot caused by G. graminis var.
graminis in cool- season bentgrasses on greens with low soil P levels (Koske et al., 1995).
Reductions in take-all disease severity in mycorrhizal wheat may be due to
increased P uptake, increased root cell wall lignification, pathogen exclusion, production
of antagonistic compounds, or altered root exudates (Graham and Menge, 1992).
However, baseline information concerning pathogen colonization and potential effects of
AMF on disease in the field is necessary before experiments concerning mechanisms of
resistance and inoculation can be undertaken.
The objective of this survey was to determine the extent of R. solani and G.
graminis var. graminis colonization in production fields of 'Floratam' St. Augustinegrass
sod in north central Florida and to determine whether populations of AMF are having any
effect on disease incidence in the field. Many researchers may feel that the effects of
AMF in turfgrass systems may be outweighed by the benefits of added nutrients,
pesticides, and irrigation. However, in St. Augustinegrass sod systems where inputs are
limited, AMF may serve a greater role in plant resistance to soilborne pathogens or soil
Materials and Methods
Root Pathogen Sampling. 'Floratam' St. Augustinegrass stolons and roots were
collected on a bimonthly basis from the three north central Florida sod farms described in
chapter 2 in January through December 2005. The roots and stolons were surveyed for
take-all root rot and brown patch. From each of the three (3 m2) plots described in
chapter 2, ten subsamples of root and stolon tissue (1-5 cm above the crown) were
randomly dissected from collected plants and cut into 100 pieces of tissue 2-5 cm in
length, in order to quantify the extent of root rot disease and to isolate and identify the
causal organisms. The pieces were washed, surface-sterilized for 1 min in a 10% sodium
hypochlorite and deionized water solution, rinsed twice for 1 min with sterile deionized
water, and blotted dry.
Pathogen Identification. Forty pieces of tissue from each of the 100 segments/plot were
randomly selected for isolation of G. graminis var. graminis and forty for isolation ofR.
solani and aseptically plated into selective agar media (Appendix- A) in 15 x 100 mm
Petri dishes. Selective media (Appendix A) were used to isolate the pathogens from
tissue and to slow growth of other soilborne fungi not associated with diseased tissue.
The Petri dishes were incubated at 24 C under a 12 h diurnal cycle. Fungal growth was
monitored by light microscopy for 5-8 d or until opportunistic fungal growth required
colony transfer to sterile media, in order to isolate the desired root pathogens. Samples of
fungal colonies suspected of being R. solani or G. graminis var. graminis were mounted
in water on glass slides and viewed with a Nikon Optiphot compound microscope to
identify fungal structures microscopically. Gaeumannomyces graminis var. graminis
colonies were readily identified in media by the presence of deeply-lobed hyphopodia
(Figs. 3-4, 3-5) within melanized mycelium (Landschoot, 1997). Rhizoctonia solani
colonies (Figs. 3-6, 3-7) were identified based on the auburn to light brown color and 900
branching of the mycelium (Sneh et al., 1991).
Pathogen Quantification and Statistical Analysis. The number of colonies of G.
graminis var. graminis and R. solani observed emerging from root or stolon pieces were
used to quantify the amount of infection of these root pathogens at each sod farm
location (Figs. 3-5, 3-6). The mean colonization data were expressed as the percentage of
sampled root or stolon pieces colonized by G. graminis var. graminis or R. solani on
selective agar media (Appendix A). The survey was performed using a random model in
a randomized complete block design with multiple samplings at multiple locations. The
percent colonization data were analysed using the Generalized Linear Model (SAS
Institute, Version 9.0, 2004) (Appendix F-2; Appendix F-3). Arbuscular mycorrhizal
fungi sampling data, as described in chapter 2, were used in this survey since root
pathogen sampling occurred simultaneously in the same plot locations as the survey of
AMF in the previous chapter. Significant interactions (P < 0.05) were separated using
Tukey's Studentized Range Distribution test, and correlations between AMF percent
colonization and spore density to percent colonization of each root pathogen were done in
SAS using Pearson product-moment correlation coefficients.
Results and Discussion
No correlation between AMF spore density or percent colonization in relation to
R. solani or G. graminis var. graminis colonization were found. Additionally, no location
effects were detected in the analysis of variance among or within the sampling months (P
< 0.001). However, pathogen colonization did vary significantly between sampling
months (P < 0.001), which suggested a seasonal influence on pathogen activity in north
central Florida soils at each sod farm location. Mean values of root colonization by R.
solani were greatest in December 2004 at 24.40% and lowest in June 2005 at 10.71
percent (Fig. 3-8). The warmer months of June and August had the lowest R. solani
colonization percentages but the values were not significantly different
from values in March, January, or October. The cooler months of December and April
had the highest percentages of R. solani, although the April mean was not significantly
different (P < 0.05) from October, January, or March (Fig. 3-8). This finding is not
surprising since R. solani has optimal growth below 26 C therefore it is typically more
active in cooler weather (Elliott and Simone, 2001). Interestingly, as noted in chapter 2,
AMF spore density (Table 2-2) was generally lowest during the cooler months of
December, January, and April and highest during warmer weather, with percent
colonization highest during the cooler months when R. solani is most active in these soils
Mean values of root colonization by G. graminis var. graminis were highest in the
warmer months of August 2005 at 20.01% and lowest in December 2004 at 5.35 percent
(Fig. 3-9). The months of August, June, and October had the highest percentages of G.
graminis var. graminis colonization, with the lowest mean values occurring in December,
January, March, and April. However, there were no significant differences (P < 0.05)
between mean values in June and October, or October, April, March, and January.
Again, this finding is not surprising because G. graminis var. graminis is most active in
warm, markedly wet conditions where there is excessive thatch accumulation (Elliott,
1993; Guyette, 1994). During the warm, humid days of summer, St. Augustinegrass sod
is often heavily irrigated and mowed, which produces favorable growth conditions for G.
graminis var. graminis because of surplus moisture and accumulating clippings which
add to thatch layers. In this survey, the pathogen is most active during periods when
AMF percent colonization is lowest suggesting a limited role for AMF in take-all root rot
disease suppression in these soils. More controlled studies might shed light on potential
AMF effects on soilborne pathogens which may be confounded during field evaluations
due to rhizosphere variability and environmental effects. If these criteria can be
evaluated under less variable conditions, beneficial AMF effects could be evaluated and
perhaps manipulated for optimal disease suppression and concurrent decreases in
Fig. 3-1. 'Floratam' St. Augustinegrass sod mat infected with Gaeumannomyces
graminis var. graminis. Insert in bottom right-hand corner depicts underside
of a mat with rotting roots.
Figs. 3-2 3-3. Comparison of healthy 'Floratam' St. Augustinegrass sod mat and sod
affected by brown patch.
Fig. 3-2. Healthy 'Floratam' St. Augustinegrass sod mat.
Fig. 3-3. 'Floratam' St. Augustinegrass sod mat infected with R. solani causing brown
Fig. 3-4. Deeply-lobed hyphopodia isolated from Gaeumannomyces graminis var.
graminis in 'Floratam' St. Augustinegrass sod samples. Scale bar = 40 nm.
Fig. 3-5. Medium isolation plate depicting a Gaeumannomyces graminis var. graminis
colony isolated from 'Floratam' St. Augustinegrass sod samples. Arrow
points to colony.
Fig. 3-6. Rhizoctonia solani hyphae isolated from 'Floratam' St. Augustinegrass sod
exhibiting diagnostic 900 branching at constriction points and characteristic
septa. Scale bar = 40 inm. Arrow points to branching pattern.
Fig. 3-7. Medium isolation plate depicting light brown Rhizoctonia solani colony
isolated from 'Floratam' St. Augustinegrass sod samples. Arrows point to
Dec Jan April March June Aug Oct
04 05 05 05 05 05 05
Fig. 3-8. Mean percent of Rhizoctonia solani colonization of 'Floratam' St.
Augustinegrass in north central Florida. Means followed by the same number
are not significantly different according to the Tukey's mean separation test (P
< 0.05). The percent colonization is based on the mean number of colonies
where R. solani was recovered.
.5 ab ab ab
Dec Jan April March June Aug Oct
04 05 05 05 05 05 05
Fig. 3-9. Mean percent of Gaeumannomyces graminis var. graminis colonization of
'Floratam' St. Augustinegrass in north central Florida. Means followed by the
same number are not significantly different according to the Tukey's mean
separation test (P < 0.05). The percent colonization is based on the mean
number of colonies where G. graminis var. graminis was recovered.
EFFECT OF GLOMUS INTRARADICES ON THE EXTENT OF DISEASE CAUSED
BY GAEUMANNOMYCES GRAMINIS VAR. GRAMINIS AND RHIZOCTONIA
SOLANI IN ST. AUGUSTINEGRASS
Arbuscular mycorrhizal fungi (AMF) are widespread symbionts in the majority of
plant species; and are associated with increased plant vigor via improved nutrient uptake,
especially P, and increased water acquisition (Smith and Read, 1997). The beneficial
effects of AMF on crop yield have been thoroughly documented (Harley and Smith,
1983). There is much debate on whether or not AMF alter plant resistance to pathogens
by an indirect mechanism or simply interact directly with the pathogens themselves.
When AMF act as pathogen antagonists, there are likely one or more mechanisms
of resistance. For example, AMF may be deterring pathogen infection by increasing
plant vigor through improved nutrient acquisition, the AMF themselves may be
producing anti- microbial metabolites, or the AMF may be stimulating the plant's own
natural defense response to colonization by increasing phytoalexin production (Schenck,
1970). Previous studies have indicated that AMF symbiosis greatly improves plant
resistance to abiotic pressures such as water stress (Sylvia and Williams, 1992) and
transplant shock (Menge et al., 1978) in various crops. AMF have also been evaluated as
biological controls against biotic stresses such as bacterial pathogens (Weaver and
Wehunt, 1975), parasitic nematodes (Baltruschat et al., 1973; Schenck and Kellam,
1978), viral pathogens (Daft and Okusanya, 1973; Giannakis and Sanders, 1989), and
soilborne fungal pathogens (Jeffries, 1987; Schenck, 1987; Hooker et al., 1994;
Linderman, 1994; Azc6n-Aquilar and Barea, 1996).
The vast majority of evaluations concerning the effects of AMF on disease severity
involve fungal pathogens (Schenck and Kellam, 1978). The first report of an interaction
between mycorrhizal fungi and fungal pathogens involved soybean (Glycine max L.
Merr) and Phytophthora root rot, where the mycorrhizal plants actually had higher rates
of disease versus the nonmycorrhizal plants (Ross, 1972). In other reports, AMF had no
effect on disease at all (Ramirez, 1974; Sherinkina, 1975). Depending on the stage of
host plant development, plant and mycorrhizal fungal species, and the complexities
between biotic and abiotic rhizosphere factors, there is evidence that mycorrhizal
interactions lie along a continuum ranging from mutualistic to parasitic, commensal,
amensal, and potentially even neutral (Johnson et al., 1997). However, there are many
reports of mycorrhizal colonization reducing disease severity in many plant systems such
as pea, tomato, soybean, wheat, and peanut involving such fungal pathogens as Fusarium
solani Mart. (Sacc.), G. graminis (Sacc.) Arx & Olivier var. tritici J. Walker, Sclerotium
rolfsii (Sacc.), Pythium spp., Phytophthora parasitica Dastur, and R. solani Ktuhn
(Graham and Menge, 1992; Dehne, 1982; Krishna and Bagyaraj, 1983; Zambolim and
Schenck, 1983; Hedge and Rai, 1984; Vigo et al., 2000; Yao et al., 2002).
In fact, the effects of mycorrhizal colonization on disease severity is potentially so
important that Newsham et al. (1995) suggested that the benefits of AMF to disease
suppression may be as important as the nutritional benefits derived from the symbiosis in
some instances. For example, in temperate grasslands, the effects of a direct AMF
interaction with root pathogens reduced disease severity and increased plant vigor and
fecundity greatly (Newsham et al., 1995). Soilborne pathogen suppression by AMF
includes both physical and physiological mechanisms (Sharma et al., 1992). Physical
plant defense responses against pathogen penetration are: increased lignification (Dehne
and Schoenbeck, 1978), greater mechanical strength and nutrient flow within vascular
systems (Schoenbeck, 1979), and direct competition with the pathogen for cortical
infection courts and resources (Graham, 2001). Becker (1976) observed that pathogen
penetration of root cells was directly reduced by the presence of AMF and not indirectly
by a systemic plant resistance based on thickening cell walls. In some cases the direct
influence of AMF may be the only reason for observations of disease resistance. It is
important to establish whether or not particular plant systems benefit, suffer, or remain
unaltered by mycorrhizal colonization. If the relationship appears to be beneficial,
Gerdemann (1975) remarked that the effect of mycorrhizal fungi on disease should be
determined whether resistance is due to direct or indirect mechanisms.
The host-pathogen relationship can be greatly influenced by indirect or
physiological effects of AMF through increased P nutrition, enhanced mycorrhizal root
growth which aids in disease escape, or up-regulation of pathogenesis-related proteins
(Gianinazzi-Pearson and Gianinazzi, 1989; Blee and Anderson, 2000; Graham, 2001).
AMF may also be responsible for lowering disease severity in complex reactions
involving host physiology such as the production of rhizosphere leachates from
mycorrhizal plant roots. These leachates have been observed to substantially limit the
production of zoospores and sporangia of Phytophthora cinnamomi Ronds in sweet corn
and chrysanthemum (Meyer and Linderman, 1986).
There appears to be no information concerning the effects of AMF, if any, on
disease severity in St. Augustinegrass. If there is a direct or indirect beneficial effect of
AMF on disease severity of St. Augustinegrass in relation to brown patch or take-all root
rot, several questions will remain concerning the actual mechanism of observed
resistance. However, without basic information and techniques to differentiate between
direct and indirect effects and to determine what extent disease severity may or may not
be lowered, further evaluations would not be warranted.
The economic importance of AMF in soils of north central Florida St.
Augustinegrass sod fields may be considerable where diseases such as brown patch and
take-all root rot reduce harvestable hectares. Arbuscular mycorrhizal fungi can stimulate
plant vigor and possibly interact directly or indirectly with soilborne pathogens to limit
disease. AMF have been observed colonizing St. Augustinegrass (see Chapter 2), and
they might benefit sod production. The potential AMF benefits to sod growers include
reduced loss of sod and revenue to soilborne pathogens, and lowered management costs
through reduced fungicide use. The potential advantages of AMF inoculation or field
manipulation with specialized techniques may also benefit the environment by decreasing
soil and water pollution through reduced of fungicide use. For these reasons, it is prudent
to evaluate the potential benefits of AMF to disease resistance whether by direct or
indirect mechanisms in St. Augustinegrass sod. As part of ongoing research on the effect
of AMF on disease severity in St. Augustinegrass, the objective of this study was to
determine the effect of G. intraradices, on St. Augustinegrass in disease development by
challenging it both directly and indirectly with G. graminis var. graminis or R. solani.
Material and Methods
St. Augustinegrass Sprig Propagation and Stock Plants.- 'Floratam' St.
Augustinegrass sprigs having no apparent signs or symptoms of disease were obtained
from Hendrick's Turf Farm (Lake Butler, Florida). The sprigs were rooted in flat, plastic
nursery trays or 18 cm clay pots in a sterilized Arrodondo fine sand medium
supplemented with a nutrient solution (Appendix B) every three weeks. The sprigs were
grown and maintained in a growth chamber at 25-27 C under cool-white fluorescent
bulbs with irradiance at 25 ilE/m2/s and a 15 h photoperiod/day. Sprigs were watered
every other day throughout the experimental period with water adjusted to pH 6.0-6.5.
After approximately 6 weeks of propagation, selected sprigs, not in direct contact with
soil, were excised from the edge of the flat trays and replanted as sterile stock plantlets.
These sub-cultured plants were maintained as described above until additional sprigs, not
touching the soil and hanging from the edge of the tray, were collected for
R. solani Inoculum Production.- A virulent strain of R. solani (PDC 7884) (Fig. 4-1)
isolated from diseased St. Augustinegrass submitted by a homeowner in Leon County,
Florida was provided by the Plant Disease Clinic (Institute of Food and Agricultural
Sciences, University of Florida, Gainesville, Florida). The isolate was cultured at 4 C
and stored on potato dextrose agar (Difco Laboratories, Inc., Detroit, Michigan) for
approximately 2 weeks. An oat (Avena sativa L.) inoculum was prepared according to
Sneh et al. (1991) and Gaskill (1968) with modifications (Appendix C) and inoculated
with agar plugs from actively growing R. solani (PDC 7884) mycelium or with sterile
agar plugs (control). The inoculum substrate was incubated at 21 C with a 12 h
photoperiod for 4 weeks and shaken 2-3 times/week to prevent packing of the oat seeds.
The inoculated seeds were then air-dried, sealed in plastic zip-lock bags, and stored at
room temperature until use.
G. graminis var. graminis Inoculum Production.- A virulent strain of G. graminis var.
graminis (JK2) was collected and identified from diseased St. Augustinegrass (Fig. 4-2)
from the lawn of Dr. James Kimbrough (Gainesville, Florida) and isolated on selective
media amended with antibiotics (Appendix A). Actively growing G. graminis var.
graminis mycelium from a single Petri dish was chopped and combined with sterilized
ryegrass seed as described by Datnoff and Elliott (1997) with modification (Appendix C).
The inoculated flasks of sterile ryegrass seed substrate and uninoculated control flasks
were incubated in total darkness at 21 C for 4 weeks prior to use. The flasks were shaken
2-3 times/week to prevent packing of the inoculated ryegrass seed.
Mycorrhization of 'Floratam' St. Augustinegrass Sprigs.- Sprigs of 'Floratam' St.
Augustinegrass were selected from the edge of sterile stock plants in flat trays, as
previously described. Sprigs were inspected visually for any signs or symptoms of
potential pathogens or diseases, and if healthy, were selected for experimental use. The
sprigs were then planted into 6.8 cm wide by 18 cm deep containers (Steuwe and Sons,
Inc., Corvallis, Oregon) filled with a sterilized low P soil, as mentioned in Chapter 2 (Fig.
4-3). The sprigs were then placed in a controlled growth room with a 15 h photoperiod/d
at 21-25 C, watered daily with pH adjusted 6.0-6.5 deionized water, and maintained for
approximately 3 weeks to allow root development to occur and transplant shock to
subside. After the 3 week growth period, the sprigs, with approximately 8 cm of root
length, were inoculated with approximately 20 spores of G. intraradices (FL 208A) (Fig.
4-4) obtained from the INVAM Culture Collection (Morgantown, West Virginia) or
noninoculated water controls. The FL 208A isolate was selected because it was first
isolated in a citrus grove in central Florida, near Orlando, in 1978 in 7.0-7.5 pH soil,
which is similar to that of the sod fields in north central Florida. The sprigs were then
acclimatized for approximately 4 weeks in the growth room to allow the AMF time to
colonize the sprig roots, which was determined at 2 and 4 weeks in extra experimental
Pathogen Inoculation.- The AMF colonized sprigs were inoculated with either R. solani
(PDC 7884) or the G. graminis var. graminis (JK2) isolate or uninoculated as controls by
gently pushing the soil aside to expose a portion of the roots near the crown of the sprig.
Approximately 3-5 infected seeds of either the R. solani inoculated oat substrate or G.
graminis var. graminis inoculated ryegrass seed substrate were placed equidistant from
the crown in each container at a 1-2 cm distance from the plant. The soil was carefully
replaced following inoculation. Inoculated sprigs were maintained in the growth room
for approximately 4 weeks with a 15 h photoperiod/d at 21-25 C. Each cone was
supplied with a nutrient solution devoid of P on two occasions at 50 ml/conetainer
(Appendix B). Plants were watered daily with 50 ml water/conetainer adjusted to 6.0-6.5
Mycorrhizal Evaluation.- Roots from the sprigs were rinsed in tap water and separated
with a scalpel from the plant crown. Selected roots were cut into 1-2 cm long segments,
put into porous nylon sleeves, inserted in small, plastic clips (Fig. 4-5), and the cell and
wall components cleared in 10% KOH (w/v) under pressure in an autoclave for
approximately 20 min at 121 C psi (Brundrett et al., 1996). The root segments were
cooled, then rinsed in tap water, and placed into 0.05% trypan blue in 25% glycerol
overnight to stain mycorrhizal structures (Bevenge, 1968; Phillips and Hayman, 1970;
Kormanik and McGraw, 1982). Excess stain was rinsed from the root segments with tap
water and then the roots were mounted in water on glass slides to view vesicles,
intraradical hyphae, and arbuscules (Fig. 4-6).
Root segments from each replicate were pooled from each treatment, and
evaluated for intensity of colonization. Mycorrhizal structures on glass slides were
viewed with a Nikon Optiphot compound microscope at 200, 400, and 1000x
magnifications, and photographs were taken with a Nikon CoolPix 990 digital camera. In
order to judge the amount of mycorrhizal root colonization, the grid line intersect method
was used to approximate the total root length colonized by AMF (Newman, 1966;
Tennant, 1975; Giovannetti and Mosse, 1980).
Direct Experiment Disease Assessment.- Disease severity (root and shoot rot) was rated
at the conclusion of a 3 week growth period on both the AMF inoculated, pathogen
inoculated, and control sprigs. Disease severity was assessed using an arbitrary disease
scale from 1 to 6 with 1 = no symptoms of disease; 2 = 1-25% disease; 3 = 26-50%
disease; 4 = 51-75% disease; 5 = 76-100% disease; and 6 = plant death (Figs. 4-7; 4-8).
The presence of either the R. solani or G. graminis var. graminis pathogens on each
infected sprig was confirmed by re-isolation of each pathogen (Figs. 4-9; 4-10) on
selective media (Appendix A). For each sprig, the percent colonization of the pathogen
and/or AMF was recorded as described in Chapters 2 and 3.
Direct Experiment Design and Statistical Analysis.- The experiment was performed using
a factorial arrangement (1 cultivar of St. Augustinegrass) x (1 AMF + uninfected
pathogen control) x (1 R. solani-infected + 1 AMF) x (1 R. solani- infected AMF) and
(1 G. graminis var. graminis- infected + 1 AMF) x (1 G. graminis var. graminis AMF)
and (uninfected pathogen control + uninoculated AMF control) in a randomized complete
block design with four replicates/treatment (Fig. 4-11). Regression analyses were
performed with the regression procedure in SAS (SAS Institute, 2004) (Appendix F-4).
All data presented are the means of four replicates. As there were no differences between
trials based on the ANOVA, all data presented were combined for the purpose of
presenting the results and discussion more easily.
St. Augustinegrass sprigs were produced and maintained in the same manner as described
above in the Direct Experiment section as were mycorrhization and pathogen inoculum
production, inoculation, and quantification. However, in this experiment, the potential
effects of indirect AMF interactions with soilborne pathogens were evaluated instead of
the potential direct impacts of mycorrhization. Instead of a direct challenge between
AMF and pathogen in one container, indirect effects were investigated using a split-root
Indirect AMF Challenge Split-Root Assay.- Sterile, 4 week old 'Floratam' St.
Augustinegrass sprigs with approximately 8 cm of healthy root tissue were placed into
two adjacent containers with one rooted end of the sprig in one container and the other
rooted end in another container (Fig. 4-12). Holes (1 cm in diameter) were drilled 2.5
cm from the top of each 6.5 cm wide by 18 cm deep container (Steuwe and Sons, Inc.,
Corvallis, Oregon) prior to planting, on one side of the container (Appendix E- 1). A cut
was made from the top of the drilled hole to the top of each container to allow the sprig
to be inserted into the hole without tissue damage. Sprigs were planted into containers
filled with sterile low P soil as previously described and maintained in the growth room
for 3 weeks to limit transplant shock and acclimatize the sprigs. Sprigs were then
inoculated with the G. intraradices isolate (FL 208A) as described in the direct
experiment above, or a control substrate in one container, with either the G. graminis
var. graminis isolate (JK2) or R. solani isolate (PDC 7884) inoculated or an uninoculated
control substrate in the adjacent container occupied by the other rooted end of that same
sprig (Fig. 4-13). The containers were watered daily with 50 ml water/conetainer
adjusted to pH 6.0-6.5 and supplied with a nutrient solution on two occasions (Appendix
B). The sprigs were maintained for 3 weeks in the growth chamber at 21-25 C with a 15
h photoperiod. The sprigs were visually inspected every 2-3 d for the presence of
invading pathogenic mycelia along the stolon portion of the sprig to prevent cross
contamination. The presence of the pathogen used to inoculate one container was not
observed in any of the adjacent experimental units containersr) based on the lack of
recovery of the pathogen from adjacent containers by selective media isolation
(Appendix A). The stolon portion spanning the distance between the two adjacent
containers was approximately 5 cm in length. Percent G. intraradices colonization was
measured using the gridline intersect method described in the previous section.
Indirect Experiment Design and Statistical Analysis.- The experiment was performed
using a factorial arrangement (1 cultivar of St. Augustinegrass) x (1 AMF + uninfected
pathogen control) x (1 R. solani-infected + 1 AMF) x (1 R. solani- infected AMF) and
(1 G. graminis var. graminis- infected + 1 AMF) x (1 G. graminis var. graminis AMF)
and (uninfected pathogen control + uninoculated AMF control) split-root assay in a
randomized complete block design with four replicates. The entire experiment was setup
three times from January May 2006. Regression analyses were performed with the
regression procedure in SAS (SAS Institute, 2004) (Appendix F-5). All data presented
are the means of four replicates/treatment. No differences were found between trials
based on the ANOVA, therefore, data were pooled for analysis.
Mycorrhizal Colonization.- In the direct experiment, mean values of root colonization by
the AMF, Glomus intraradices, were 10% for the R. solani- infect + AMF treatment,
11.3% for the AMF inoculated control treatment (no pathogen), and 11.7% for the G.
graminis var. graminis-infected + AMF treatment, respectively, after mycorrhizal
inoculation. Root colonization of AMF was not significantly affected by the direct
presence of either pathogen nor did the AMF control treatment (no pathogen) have any
direct effect, either positive or negative, on disease severity itself (Appendix D-1). In this
study, the colonization of plants by AMF, G. intraradices, apparently had a neutral effect
on the St. Augustinegrass plants without the direct presence of either pathogen nor did
the AMF affect plant growth.
Disease Development.- The direct effect of G. intraradices on brown patch (caused by R.
solani) disease severity was evaluated by first investigating the relationship of the R.
solani- infected control (no AMF) treatment (Appendix D-2) to disease severity. The
mean percent colonization of the R. solani-infected control treatment was 60%, but the
disease severity (mean = 3.8 on a scale of 1 to 6) was not significantly correlated with the
mean colonization percentage of R. solani using the regression procedure in SAS (SAS
Institute, 2004). Since there was no definitive relationship between plant disease
severity and the percentage of R. solani colonization with this treatment, there was no
need to assume that G. intraradices in the R.solani-infected + AMF treatment would have
a beneficial effect on disease severity. This was supported by the regression analysis
comparing the relationship of disease severity to percent R. solani colonization (mean
colonization = 57%) in the R. solani- infected + AMF treatment (Appendix D-3) where
disease severity (3.3 on a scale of 1 to 6) was not correlated to the mean percentage of
AMF colonization (mean colonization = 18%). In this study, the AMF treatments had no
effect on disease severity in the direct presence of R. solani regardless of the mean
colonization of the pathogen or AMF.
The direct effect of G. intraradices on disease severity was also evaluated in this
study for take-all root rot caused by G. graminis var. graminis. Based on regression, the
relationship between disease severity and the G. graminis var. graminis- infected control
(no AMF), it appears that the pathogen (mean colonization = 42.8%) had a significant
relationship (r2 = 0.65) with disease severity (2.4 on a scale of 1 to 6). This model shows
that as disease severity increases so does G. graminis var. graminis percent colonization
in a direct pathogenicity challenge (Fig. 4-14). This finding suggests that the AMF could
potentially have a direct effect on disease severity and that the relationship could be
evaluated since the percent colonization of G. graminis var. graminis had a measurable
effect on disease severity. The regression analysis of disease severity (mean = 3.3 on a
scale of 1 to 6) to the G. graminis var. gramninis- infected + AMF inoculated treatment
revealed a highly correlated relationship between the treatment and disease severity (r2 =
0.81). As disease severity increased according to this treatment, so did the percent
colonization of G. graminis var. graminis even in the direct presence of AMF (mean =
8.6%) (Fig. 4-15). There was no apparent reduction or increase in disease severity.
Therefore, the AMF have no direct beneficial effect on take-all root rot disease severity.
Additionally, the AMF treatment alone could not be correlated to a reduction in percent
G. graminis var. graminis colonization (data not shown) nor did the treatment have a
direct effect on lowering take-all root rot disease severity since the disease severity trend
did not differ from that of the G. graminis var. graminis- infected AMF treatment.
Since disease severity was not affected by G. intraradices in the G. graminis var.
graminis- infected + AMF treatment or correlated to the percent of G. graminis var.
graminis colonization in the control uninoculated with AMF, it appears that the AMF
colonization had no direct negative or positive impact on the pathogen or disease
severity. In this study, the interaction between AMF and the plant in the direct presence
of the pathogens, G. graminis var. graminis and R. solani would thus be considered
neutral in nature.
More importantly, this study demonstrates that mycorrhization with the AMF, G.
intraradices, did not reduce development of R. solani or G. graminis var. graminis in
direct contact nor did the AMF treatment reduce or increase disease severity of brown
patch or take-all root rot in 'Floratam' St. Augustinegrass, as has been observed in other
mycorrhizal studies (Ross, 1972; St. Arnaud et al., 1994; Mark and Cassells, 1996).
Arbuscular mycorrhizal fungi have been associated with increased disease severity in
some instances with R. solani, so analysis based on this assumption was as necessary as
assuming the AMF treatment would lower disease severity (Ramirez, 1974; Sherinkina,
1975; Johnson et al., 1997; Yao et al., 2002). No beneficial effects of AMF inoculation
on take-all root rot or brown patch disease severity in St. Augustinegrass were observed.
This is perhaps due to the relatively low levels of mycorrhizal root colonization. Possibly
AMF inoculation would be more beneficial to plants with a higher level of mycorrhizal
In summary, the results show that the purported beneficial effects of direct AVIMF
interactions with plant roots such as increased cell wall lignification or the production of
antagonistic mycorrhizal root exudates did not play a role in this study (Becker, 1976;
Dehne and Schoenbeck, 1978; Graham, 2001). Thus, inoculation with G. intraradices
will not improve disease severity or reduce disease development. The effects of such an
interaction within field trials could potentially yield contradictory results, and the
microbial and environmental variability within the rhizosphere would make such
experiments difficult at best.
In order to thoroughly evaluate the potential effects of AMF on disease severity
and/or soilborne pathogen development, another series of studies involving a more
indirect method was performed simultaneously with the direct experiment described
above. This assay was designed to isolate potential systemic resistance responses from
mycorrhization which have been documented (Gianinazzi-Pearson and Gianinazzi, 1989;
Blee and Anderson, 2000; Graham, 2001).
In this assay, the R. solani control (no AMF) treatment revealed a significant
correlation between pathogen colonization and disease severity (Fig. 4-16). In this
instance, as percent colonization of R. solani (mean = 54.9%) increased so did disease
severity (mean = 3.5 on a scale of 1 to 6; r2 = 0.75). Since there was a significant
relationship between the pathogen and disease severity, the regression procedure in SAS
was also used to analyze the indirect effects of the R. solani + G. intraradices treatment
on disease severity. The combination of this pathogen and AMF in an indirect assay,
where one container was inoculated with R. solani and the other container containing