Download `Evaluation of beneficial bacteria for improved growth and resistance

‘Evaluation of beneficial bacteria for improved
growth and resistance against Fusarium oxysporum
f. sp. zingiberi in ginger (Zingiber officinale Roscoe)’
Emily Rames
Submitted for the fulfilment of the requirements of the
degree of Master of Science by Research.
Faculty of Science, Health and Education
University of the Sunshine Coast
2008
Abstract
Severe losses in productivity in regional ginger cultivation have been caused by
declining soil health and Fusarium oxysporum f. sp. zingiberi (Foz). Ongoing studies
worldwide have demonstrated that plant growth promoting bacteria may increase
yield, improve resistance to disease and reduce the requirement for fertilisers in a
variety of crops of agronomic importance. Initially compost teas and commercial
microbial inoculants were investigated as a source of microorganisms that may be
beneficial to the growth of ginger. Quality control problems and associated safety
issues precluded these inoculants from further study. Accordingly, reference strains
of bacteria and strains isolated from the ginger rhizosphere were tested in
glasshouse conditions for their plant growth promoting ability under reduced levels of
fertiliser application.
Bacillus F2 (field isolate), Azospirillum brasilense Sp7 and
Azospirillum brasilense Sp7 combined with Bacillus coagulans NCTC 10334
significantly increased rhizome fresh weight of micropropagated ginger plants by
40.9%, 45.9% and 50% respectively.
As the buffer used to apply the bacteria
reduced plant growth, results indicated plant stress caused by salt in the buffer was
overcome by the bacterial treatments.
The aforementioned bacteria did not
significantly improve the growth of ginger-plants grown from seed pieces and as
such optimisation of application methods may be required. In a subsequent trial a
dried alginate bead formulation of
Azospirillum brasilense Sp7 significantly
increased rhizome weight of ginger tissue cultured plants compared to application of
a suspension of this bacterium.
The alginate bead material also significantly
improved the growth of micropropagated ginger plants. The importance of including
appropriate controls, to identify effects of buffers and carrier materials on plant
growth was repeatedly demonstrated. Antagonistic activity of bacterial isolates
against Foz was also evaluated. Bacillus subtilis DAR26659 caused lysis of hyphae
and also increased rhizome weight of micropropagated ginger plants by 60%,
although evaluation of in planta biocontrol activity was limited by inconsistent
infection of ginger plants by Foz. Improved productivity of micropropagated ginger
plants might result in increased uptake of this source of disease-free planting
material in the ginger industry.
i
Acknowledgements
I would like to sincerely thank Dr Ipek Kurtboke, my principal supervisor, for giving
me the opportunity to undertake this project.
Thankyou for your patience and
advice.
I would like to gratefully acknowledge the contribution of Sharon Hamill, Department
of Primary Industries and Fisheries, Maroochy Research Station, for her generosity
of time and materials (including ginger tissue culture plants and seed pieces) and
invaluable advice on ginger agronomy and setting up glasshouse trials.
The provision of funding for the project by Natural Resource Management South
East Queensland and project management by Susie Chapman is gratefully
acknowledged. Thankyou to Bob Cameron, Rockcote Industries and other local
industries involved in the project, for their financial and material contributions, useful
discussions and preparation of compost teas. Many thanks to Shane Templeton,
Templeton’s Ginger, for sharing your expertise on ginger cultivation, for provision of
samples and yield estimation.
The goodwill and contribution of other steering
committee members is kindly acknowledged: Brain Stockwell (DPI&F Nambour),
Mick Millington (Milltech Industries), Scott Graham (Natural Resource Management),
Esma Armstrong (Landcare) and Jack Connolly (Maroochy Springs Winery).
Many thanks to Wayne Robinson, University of the Sunshine Coast, for statistics
advice and for checking data analysis. Thanks also to Dr. Mike Smith, Maroochy DPI
& F for useful discussions and literature on the history of ginger cultivation.
Thank you to the technical staff at the University of the Sunshine Coast, in particular
Daniel Powell for technical advice and risk assessments.
Jenny Cobon, DPI&F
Indooroopilly, is kindly acknowledged for performing nematode analyses. Thankyou
to Dr Peter Williamson, DPI & F Toowoomba, for providing wheat seed and advice
on growth conditions.
Thanks also to Maroochy DPI staff for assistance in
harvesting ginger plants. Thanks to my lovely daughter Zoe, for her patience and
being so great.
ii
Table of Contents
Abstract
i
Acknowledgements
ii
Table of Contents
iii
List of Figures
ix
List of Tables
xiii
Chapter 1. Introduction
1
Chapter 2. Literature Review
5
2.1.
Cultivation and global demand for ginger.
2.2.
Impacts of the ginger pathogen Fusarium oxysporum
f. sp. zingiberi in regional production.
2.3.
7
Significance of Meloidogyne root-knot nematodes in
regional ginger cultivation.
2.4.
5
9
Impact of conventional farming practices on soil
biodiversity and the environment.
10
2.5.
Plant-bacteria interactions.
15
2.6.
Beneficial effects of bacteria on plant growth.
17
2.6.1. Nutrient availability and uptake.
18
2.6.2. Plant stimulating hormones and metabolites.
19
2.6.3. Induced resistance against plant pathogens.
21
2.6.4. Antibiotic and lytic enzymes.
27
Use of plant growth promoting bacteria in agriculture.
28
2.7.1. Fluorescent Pseudomonas species.
29
2.7.2. Bacillus species.
32
2.7.3. Phosphate solubilizing bacteria (PSB).
33
2.7.4. Diazotrophic bacteria.
34
2.7.5. Mycorrhizal helper bacteria.
36
Combinations of plant growth promoting bacteria.
37
2.7.
2.8.
iii
2.9.
Production and application of bacterial inoculants.
38
2.9.1. Culture of plant growth promoting bacteria.
38
2.9.2. Application of plant growth promoting bacteria
2.10.
and safety aspects.
38
Conclusion
41
Chapter 3. Assessment of compost tea and commercial microbial
inoculants as a source of beneficial microorganisms for improved
43
growth of ginger.
3.1. Introduction
43
3.2. Materials and Methods
46
3.2.1a. Isolation of bacteria from compost teas and
commercial microbial inoculants.
46
3.2.1b. Phylogenetic analysis and biochemical testing
of selected bacterial isolates.
50
3.2.2 Risk assessment for exposure to pathogenic
organisms in contaminated cultures.
3.3.
51
Results
3.3.1.a. Isolation of bacteria in compost teas and
commercially available mixed microbial inoculants.
52
3.3.1.b. Phylogenetic analysis and biochemical testing
of selected bacterial isolates.
61
3.3.2. Risk assessment for exposure to pathogenic
organisms in contaminated cultures.
65
3.4.
Discussion
65
3.5.
Conclusion
71
Appendix 3.1. Primers used in 16S rDNA PCR
and sequencing.
73
Appendix 3.2. Illustration of CLUSTAL W 2.0 multiple sequence
alignment used to produce a consensus sequence.
Appendix 3.3. 16S rDNA consensus sequences (5’-3’) of
isolates from untreated microbial inoculants.
iv
75
76
Appendix 3.4. Public health significance of human
pathogenic bacteria.
78
Chapter 4. Selection of bacterial isolates for further testing
81
in ginger.
4.1.
Introduction
81
4.2. Materials and Methods
82
4.2.1. Isolation of bacteria from the ginger
rhizosphere and rhizoplane and field observations.
82
4.2.2. Assessment of phosphate solubilizing activity
and growth of bacteria on nitrogen free media.
84
4.2.3. Identification of selected rhizosphere and
4.3.
rhizoplane bacteria.
85
4.2.4. Reference strains of bacteria.
86
Results
86
4.3.1. Field observations and isolation of bacteria
from the ginger rhizosphere and rhizoplane.
86
4.3.2. Assessment of phosphate solubilizing activity of
bacterial isolates and growth on nitrogen free media.
89
4.3.3. Identification of selected rhizosphere and
rhizoplane bacteria.
89
4.4.
Discussion
92
4.5.
Conclusion
95
Appendix 4.1. Media used for the isolation of
rhizosphere and rhizoplane bacteria.
96
Appendix 4.2. 16S rDNA amplification and sequencing
and arbitrarily primed PCR.
98
Appendix 4.3. 16S rDNA sequences (5’- 3’) of bacteria
isolated from ginger roots.
v
99
Chapter 5. Evaluation of plant growth promoting ability of
104
selected bacteria.
5.1. Introduction
104
5.2. Materials and Methods
107
5.2.a. Preparation of Bacterial Inoculants.
107
5.2.b. Plants and Growth Conditions.
108
5.2.c. Experimental Design.
108
5.2.1. Wheat as a model system for testing efficacy
of bacterial inoculants in promoting plant growth.
109
5.2.2. Effect of application method on bacterial induced
growth response in ginger tissue culture plants
(ginger tissue culture trial I).
111
5.2.3. Evaluation of plant growth promoting ability of
additional bacterial strains in ginger tissue culture
plants (ginger tissue culture trial II).
114
5.2.4. Evaluation of plant growth promoting activity
of selected bacteria in ginger grown from seed pieces.
116
5.2.5. Effect of alginate beads for the delivery of
A. brasilense Sp7 on the growth response of ginger
tissue culture plants (alginate bead trial).
5.3. Results
118
120
5.3.1. Wheat as a model system for testing efficacy
of bacterial inoculants in promoting plant growth.
120
5.3.2. Effect of application method on bacterial induced
growth response in ginger tissue culture plants
(ginger tissue culture trial I).
123
5.3.3. Evaluation of plant growth promoting ability of
additional bacterial strains in ginger tissue culture
plants (ginger tissue culture trial II).
vi
127
5.3.4. Evaluation of plant growth promoting activity
of selected bacteria in ginger grown from seed pieces.
133
5.3.5. Effect of alginate beads for the delivery of
A. brasilense Sp7 on the growth response of ginger
tissue culture plants (alginate bead trial).
138
5.4. Discussion
143
5.5. Conclusion
151
Appendix 5.1 Solutions and media.
153
Appendix 5.2. Supplementary data for wheat trial.
155
Appendix 5.3. Supplementary data for ginger
tissue culture trial 1.
156
Appendix 5.4. Supplementary data for ginger tissue
culture trial II.
157
Appendix 5.5. Supplementary data for ginger seed
piece trial.
158
Appendix 5.6. Supplementary data for alginate bead trial.
159
Chapter 6. In vitro and in vivo analysis of interactions between
bacterial isolates and Fusarium oxysporum f. sp. zingiberi.
160
6.1. Introduction
160
6.2. Materials and Methods
162
6.2.a. Fusarium oxysporum cultures.
162
6.2.1 Dual culture assays on agar plates.
162
6.2.2. Dual culture assays on microscope slides.
163
6.2.3. Effect of bacterial treatments on incidence of
Foz infection in ginger tissue culture plants.
163
6.2.4. Experimental design.
165
6.3. Results
167
6.3.1. Dual culture assays on agar plates.
167
6.3.2. Dual culture assays on microscope slides.
172
vii
6.3.3. Effect of bacterial treatments on incidence of Foz
infection in ginger tissue culture plants
182
6.4. Discussion
188
6.5. Conclusion
193
Appendix 6.1. Media
195
Appendix 6.2. Supplementary data for Fusarium trial.
195
Chapter 7. General discussion and conclusion
196
7.1. Discussion
196
7.2. Conclusion
211
References
214
Web based references
268
Note regarding thesis layout: This thesis is divided into chapters, for ease of
interpretation of data by the reader. An introduction, material and methods,
results, discussion and conclusion are provided in each research section. In the
final chapter an overall discussion and conclusion is presented.
viii
List of Figures
Page
Figure 1.
Bacteria isolated from isolated from aerated compost tea
(ACT 1) on a) MacConkey No.3 (faecal coliforms) and
b) Mannitol salt agar (Staphylococcus spp).
53
Figure 2.
Isolation of bacteria from untreated materials used to prepare
microbial cultures.
56
Figure 3.
Laboratory culture of Additive 1 and liquefied compost.
57
Figure 4.
Demonstration of esculin hydrolysis ability of putative
enterococci on bile esculin agar at 44 oC.
58
Figure 5.
Bacteria isolated from microbial cultures produced with
6ppm dissolved oxygen.
59
Figure 6.
Isolation of bacteria on UriSelect4 from microbial inoculants
after overnight culture with aeration and various additives.
60
Figure 7.
a. Genomic DNA extracted from bacteria isolated from
microbial inoculants, 1kb Step Ladder (Promega).
b. 16S rDNA PCR products, Lambda-Hind III DNA
Marker (Promega).
62
Figure 8.
Typical sequence chromatogram obtained following sequencing
of 16S rDNA PCR product, visualised using the FinchTV program.
62
Figure 9.
Biochemical tests performed using Enterotube II.
64
Figure 10.
Genomic fingerprints produced by arbitrarily
primed PCR of gDNA from rhizoplane bacteria.
90
Figure 11.
Acclimatised tissue cultured ginger plants that had
been maintained for several weeks in a growth cabinet.
112
ix
Page
Figure 12.
Ginger seed pieces used for testing of bacterial treatments.
117
Figure 13.
Growth response of wheat plants to introduction of bacterial
treatments as either a seed treatment or seed treatment
as well as soil drenches.
121
Figure 14.
Ginger tissue culture trial I.
124
Figure 15.
Regression analysis assessing stem width as a predictor
of rhizome weight.
126
Figure 16.
Ginger tissue culture II 4 weeks after planting.
128
Figure 17.
Ginger tissue culture II at harvest.
129
Figure 18.
Effect of bacterial treatments on the fresh weight of ginger
issue culture plants (Ginger Tissue Culture Trial II).
131
Figure 19.
Effect of bacterial treatments on the dry weight and
growth parameters of ginger tissue culture plants
(Ginger Tissue Culture Trial II).
132
Figure 20.
Ginger seed piece trial 9 weeks after planting.
133
Figure 21.
Ginger seed piece trial at harvest (17 weeks after planting).
134
Figure 22.
Effect of bacterial treatments on fresh weight of ginger
grown from seed pieces.
136
Figure 23.
Effect of bacterial treatments on the dry weight of ginger
grown from seed pieces.
137
Figure 24.
Alginate beads and their effects on the growth of ginger
tissue culture plants.
139
x
Page
Figure 25.
Effect of an alginate bead formulation of A. brasilense Sp7
on the fresh weight of ginger.
141
Figure 26.
Effect of an alginate bead formulation of A. brasilense Sp7
on the dry weight of ginger.
142
Figure 27.
Effect of bacteria on the in vitro growth of Foz on potato
dextrose agar (PDA) and Waksman agar (WA) plates.
169
Figure 28.
Microscope slide agar film culture of Foz alone or with
bacterial isolates.
172
Figure 29.
Microscope slide agar film culture of Foz under magnification.
173
Figure 30.
Microscope slide agar film culture of Foz and
B. subtilis 26659 under magnification.
174
Figure 31
Microscope slide agar film culture of Foz and
B. subtilis A13 under magnification.
175
Figure 32
Microscope slide agar film culture of Foz and
P. fluorescens under magnification.
176
Figure 33.
Microscope slide agar film culture of Foz and
A. brasilense Sp7 under magnification.
177
Figure 34.
Microscope slide agar film culture of Foz and
B. subtilis 6633 under magnification.
178
Figure 35.
Microscope slide agar film culture of Foz and
B. megaterium 2582 under magnification.
179
Figure 36.
Microscope slide agar film culture of Foz and
Pseudomonas Dz5 under magnification.
180
xi
Page
Figure 37.
Microscope slide agar film culture of Foz and
Dz11 under magnification.
181
Figure 38.
Glasshouse trial assessing effect of bacterial treatments on
growth and infection of ginger tissue culture plants by Foz.
183
Figure 39.
Effect of bacterial treatments on the fresh weight of tissue
cultured ginger plants that were inoculated with Foz.
185
Figure 40.
Effect of bacterial treatments on the dry weight of ginger
tissue cultured plants that had been inoculated with Foz.
186
Figure 41.
Effect of bacterial treatments on growth parameters and
development of symptoms of Foz infection (number of
yellow shoots and rhizome discoloration).
187
.
xii
List of Tables
Page
Table 1.
Induced systemic resistance by various Bacillus spp.
under field or greenhouse conditions.
25
Table 2.
Examples of fluorescent Pseudomonas spp. with biocontrol
activity against different Fusarium oxysporum diseases.
30
Table 3.
Media used for the isolation of bacteria from compost teas
and commercial microbial inoculants.
47
Table 4.
Label description of commercially available microbial
inoculants and substrates used in this study.
48
Table 5.
Conditions used to ferment microbial cultures.
49
Table 6.
Microbiological analysis of fermented materials.
54
Table 7.
Microbiological analysis of untreated materials.
55
Table 8.
Taxonomic assignment of 16S rDNA sequences of bacterial Isolates
from microbial inoculants using Ribosomal Database – II (RDP-II).
63
Table 9.
Biochemical testing using Enterotube II for verification
of 16S rDNA sequencing results.
64
Table 10.
Primers used for 16S rDNA PCR sequencing of inoculant isolates.
73
Table 11.
Types of infections caused by selected human pathogenic bacteria.
80
Table 12.
Media and culture conditions for the isolation of root
associated bacteria.
84
Table 13.
Reference strains of bacteria.
87
xiii
Page
Table 14.
Nematode trophic groups recovered per 100g of fumigated
and non-fumigated soil.
88
Table 15.
Analysis of culturable populations of bacteria associated
with ginger root samples.
91
Table 16.
Phylogenetic analysis of ginger root associated bacteria.
92
Table 17.
Primers used for 16S rDNA analysis and arbitrarily
primed PCR of field isolates.
98
Table 18.
RDPII-Classifier Analysis of 16S rDNA sequences of
field isolates.
103
Table 19.
Treatments and application methods used in the wheat trial.
110
Table 20.1.
Treatments applied in ginger tissue culture trial 1.
113
Table 20.2.
Treatments used in ginger tissue culture trial II .
115
Table 21.
Treatments applied in ginger seed piece trial.
117
Table 22.
Treatments in applied in the alginate bead trial.
119
Table 23.
Analysis of effect of application method for different bacterial
treatments on growth response in wheat by two-way ANOVA.
122
Table 24.
Effect of different bacterial application methods on mean
(± standard deviation) growth parameters in ginger tissue
culture trial I.
125
Table 25.
Effect of a range of bacterial treatments (root dip followed by
soil drenches) on mean (± standard deviation) growth parameters
of ginger tissue culture plants, ginger tissue culture trial II.
130
xiv
Page
Table 26.
Effect of bacterial treatments on the mean growth parameters
of ginger plants grown from seed pieces (± standard deviation).
135
Table 27.
Viable numbers of cells of A. brasilense Sp7 in alginate beads.
140
Table 28.
Effect of alginate carrier material on the mean growth
response (± standard deviation) of ginger tissue culture plants
to introduction of A. brasilense Sp7.
140
Table 29.
Percentage difference in growth parameters of bacterial
treatments compared to the buffer control for wheat.
155
Table 30.
Percentage difference in growth parameters for bacterial
treatments compared to the buffer control in ginger tissue culture trial I.
156
Table 31.
Percentage difference in plant growth parameters for bacterial treatments
compared to the buffer control for ginger tissue culture trial II.
157
Table 32.
Comparison of plant growth in two ginger tissue culture trials
in growth cabinets.
157
Table 33.
Percentage difference in growth parameters of bacterial
treatments compared to buffer control in ginger seed piece trial.
158
Table 34.
Percentage difference in growth parameters of bacterial treatments
compared to the water control in ginger seed piece trial.
158
Table 35.
Percentage difference in growth parameters of bacterial treatments
and controls in the alginate bead trial.
159
Table 36.
Bacterial treatments used in the in planta Foz bioassay.
166
Table 37.
Rating of plants for symptoms of Foz infection.
167
xv
Page
Table 38.
Nature of interaction between bacterial isolates
and Foz on agar plates.
168
Table 39.
Effect of bacterial treatments on mean growth parameters and
incidence of Foz infection (± standard deviation) in tissue
cultured ginger plants (Fusarium trial).
184
Table 40.
Percentage difference between treatment means, demonstrating
the effect of bacterial treatments on the growth of ginger tissue
cultured plants inoculated with Foz.
195
xvi
Chapter 1. Introduction
The value of the Australian ginger industry, centred on the Sunshine Coast region, is
estimated to be greater than $US 40 million (Smith 2004). The majority of Australian
grown ginger is produced in the Caboolture, Sunshine Coast and Gympie areas,
where subtropical conditions are ideal for cultivation (Sanewski 2002).
Local
processor, Buderim Ginger, is renowned for the production of high quality
confectionary ginger products and supplies approximately 40% of this worldwide
market (Smith 2004; Buderim Ginger 2006).
Severe losses in regional production have been caused by the pathogen Fusarium
oxysporum f. sp. zingiberi, which causes a rhizome rot and vascular wilt of ginger
(Stirling 2004). The disease may be transmitted via infected planting material and
once introduced into the soil, may persist for up to ten years (Pegg et al. 1974).
While the use of disease-free planting material has been critical for improved
establishment of ginger crops, once soil has been contaminated, there are no control
measures to prevent infection of plants by soil borne propagules of Foz. Other
practices common to conventional cultivation (monoculture, tillage, soil fumigation
and the application of inorganic fertilisers) are increasingly associated with
decreased productivity and increased disease pressures and emphasises the need
for the implementation of sustainable practices in agriculture (Wolf and Snyder 2003;
Garbeva et al. 2004). Viable alternatives to chemical inputs in agriculture are also
sought as human health and environmental impacts are realised (Wood 1995).
Recent research has targeted the development of integrated pest management
(IPM) in sustainable agricultural systems. As stated by Jacobsen and colleagues
(2004) IPM can be defined as a sustainable approach to managing pests by
combining biological, cultural, physical and chemical tools in a way that minimises
economic, health and environmental risks. Such an approach may include the use
of a diversity of cover crops, organic amendment, conservation tillage, resistant
cultivars and microbial inoculants that benefit plant growth (Jacobsen et al. 2004;
Barker and Koenning 1998; Compant et al. 2005; Whipps 2000; Vessey 2003).
Locally grown ginger cultivars are not resistant to Foz, nor have breeding programs
been established (Smith, personal communication 2007). As means for the control
1
of soil borne propagules of Foz do not exist, the development of microbial inoculants
that afford protection to this disease would be of significant value.
Microbial
inoculants
containing
symbiotic
Rhizobium,
Bradyrhizobium
or
Sinorhizobium spp. have been used successfully for augmented nitrogen nutrition in
leguminous crops for many years (Bullard et al. 2005; Deaker et al. 2004). Such
bacterial species form a symbiosis with legumes and induce the formation of
nodules that fix atmospheric nitrogen (Sprent and Sprent 1990; Zahran et al. 1999).
The use of microbial inoculants that contain plant growth promoting bacteria, that are
symbiotic or free living and function independently of root nodules, has become of
increased importance in providing alternatives in sustainable crop production
systems (Kennedy et al. 2004; Compant et al. 2005; Johansson et al. 2003). Many
thousands of reports have described the mechanism of action and/or use of plant
growth promoting bacteria in a variety of grain and vegetable plants, in greenhouse
and field conditions, for increased yields, improved resistance to disease and a
reduced requirement for inorganic fertilisers (Bashan et al. 2004; Dobbelaere et al.
2003; Okon and Labandera-Gonzales 1994; Vessey 2003; Zehnder et al. 2001;
Jacobsen et al. 2004; Bashan and Holguin 1998).
Plant dependant variables require that the efficacy of a microbial inoculant be
evaluated for different plant and cultivar types (Martin and Bull 2002; Berg et al.
2002; Broadbent et al. 1977). Very few reports have described the use of microbial
inoculants for improving the growth of ginger.
Meena and Mathur (2003)
demonstrated growth promotion and improved resistance to F. solani resulted from
the application of a fluorescent Pseudomonas species (alone or in combination with
the fungal biocontrol agent Trichoderma) to ginger seed pieces and/or soil.
Reference has been made to unpublished studies that used Azospirillum spp. for
improved growth and reduced fertiliser application in ginger cultivation (Nybe and
Raj 2005). The use of B. subtilis for reducing infection of Foz in ginger was reported
by Sharma and Jain (1979), although again experimental details were not described.
While the potential for the use of microbial inoculants to improve growth and
resistance to disease in ginger has been indicated, detailed accounts (such as
strain/species level identification, application methods, number of viable cells,
cultivar tested) of these trials are lacking. In addition, reports describing the activity
2
of plant beneficial bacteria in ginger under Australian regional environmental and
seasonal conditions were not found in searches of literature.
In general, inconsistent performance of microbial inoculants in field conditions is
considered to have limited their widespread implementation in agriculture.
An
increased understanding of which beneficial microorganisms exhibit synergistic
relationships when co-inoculated in plants and the optimisation of delivery methods
are suggested as key areas for improving the efficacy of microbial inoculants in
different soil types and under different environmental and seasonal conditions
(Fravel 2005; Whipps 2000; Guetsky et al. 2004; Kennedy et al. 2004). Several
inoculants, available in the United States, have been registered by the US-EPA and
have been used extensively where no other means for disease control exists (as in
the case of Fusarium oxysporum) or in organic production systems where chemical
controls are limited (US-EPA 2005; Kloepper et al. 2004a; Stockwell and Stack
2007).
A number of commercially available microbial inoculants are also available
in Australia. The use of plant beneficial microorganisms in compost teas, produced
by the fermentation of microflora extracted from composted waste, is also an
increasingly popular practice in agriculture.
Accordingly, the aims of the current research were to assess:
1) The suitability of using commercially available compost tea and liquid based
microbial inoculants to improve the growth of ginger.
Safety issues and quality control problems precluded the use of such inoculants
from further research. Therefore this study focused on:
2) The assessment of the ability of Class I bacteria to promote growth and
resistance against Fusarium oxysporum in ginger. Bacteria evaluated were:
•
Isolated from the rhizosphere of ginger grown in fumigated and non-fumigated
field soil and
3
•
Reference strains with known biocontrol and/or plant growth promoting activity,
obtained from culture collections.
In addition:
•
The effect of different methods of application and concentration of viable cells
of bacteria on plant growth was investigated.
•
Bacteria were assessed for their ability to promote the growth of ginger
cultivated with minimal levels of synthetic fertilisers under greenhouse
conditions.
Bacterial based inoculants were chosen for study, due to their relative ease of
preparation and widely described benefits in many different crops (Lucy et al. 2004).
Bacteria belonging to the genera Azospirillum, Pseudomonas and Bacillus were
selected for testing, as they include the most extensively characterised and studied
plant growth promoting bacteria used to enhance resistance to disease and/ or
increase growth in a variety of plants of agronomic importance (Bashan et al. 2004;
Kloepper et al. 2004; Jacobsen et al. 2004; Mercado-Blanco and Bakker 2007).
4
Chapter 2. Literature Review
2.1. Cultivation and global demand for ginger.
The rhizome or underground stem, of the herbaceous monocotyledon, ginger
(Zingiber officinale Roscoe) is used as a spice, confectionary product, and
component of herbal remedies (Khatun et al. 2003; Smith 2004). Gingerols, pungent
constituents of fresh ginger, were reported to relieve pregnancy, post-operative and
chemotherapy associated nausea in clinical trials (Borrelli et al. 2005; Anderson and
Johnson 2005; Chaiyakunaprik et al. 2006).
Such compounds have also been
investigated for their analgesic, anti-inflammatory, anti-tumorigenic, anti-viral and
anti-coagulative properties (Kim et al. 2005; Nurtjahja et al. 2003; Surh 2002; Surh
1999). Oleoresins, that retain the pungent principle and essential oils are extracted
from rhizomes for use in perfumes, flavourings and essences.
Ginger has been cultivated since ancient times although it is not known in a wild
state (Purseglove 1972). This plant is thought to have originated in Southeast Asia
and this area still produces the majority of ginger demanded by worldwide markets
(Purseglove 1972; Smith 2004). The Australian industry has established as a well
renowned producer of high quality confectionary ginger and demands for fresh
market ginger in local markets have also increased in recent times (Smith et al.
2004; Stirling 2004; Connell and Jordan 1971; Leverington 1975). Ginger is typically
grown as an annual crop where day length and temperature determine periods of
growth and senescence (Groszmann 1954).
In the Sunshine Coast region the senescent period of ginger, accompanied by death
of above ground plant material, occurs in July-August (Groszmann 1954). Following
this, the rhizome may be harvested (August to September) and cut into “seed
pieces”, each containing at least one bud (growing point), for vegetative propagation
in the following crop (Sanewski et al. 1996; Ravindran et al. 2005; Stirling 2004).
The seed pieces are typically planted from late August till October, within six weeks
of harvesting (Whiley 1974; Groszmann 1954).
5
The branched ginger rhizome, which functions as a storage organ, grows
horizontally beneath the soil surface (Ravindran et al. 2005; Purseglove 1972). The
first order branch grows out from the apical bud of the seed piece and produces a
leafy, aerial pseudostem.
Formation of further buds may be followed the
successive development of rhizome branches (second order, third order branches
etc.) (Ravindran et al. 2005; Lee et al. 1981). A number of these rhizome branches
may also produce one or more aerial pseudostems (Lee et al. 1981).
The ginger
plant produces flowers from March to April, although seed is rarely set (Whiley
1974).
In the conventional cultivation of ginger, in order to augment soil organic matter and
nitrogen, stimulate microbial populations antagonistic to ginger pathogens and
enhance soil structure, poultry manure is often incorporated with cover crops several
months prior to planting (Whiley 1974; Stirling 1989). The soil is also worked to a
fine tilth to form a seedbed just prior to planting and herbicides are typically applied
for weed control (Broadley 2005). A high level of irrigation is required during ginger
cultivation, in particular to prevent sunburn that occurs in new leaves during early
growth when temperatures exceed 32oC and to delay fibre development (Broadley
2005; Stirling 2004). In an analysis of the effect of various levels of nitrogenous
fertilisers in regional ginger cultivation, Lee and colleagues (1981) demonstrated that
maximal yields could be obtained with the application of 200 to 300 kg /ha of
ammonium nitrate throughout the growing season. However, inorganic nitrogen,
super-phosphate and potassium fertilisers are applied at rates of up to 750kg, 1
tonne and 210kg per hectare respectively, over the season (Broadley 2005; Whiley
1974).
The two predominant varieties of ginger grown in the regional industry are the
Queensland and Canton cultivars (Sanewski 1994; Stirling 2004).
Immature
rhizomes of the Queensland cultivar have a lemony aroma and are harvested for
confectionary markets from February till March (Connell and Jordan 1970; Sanewski
1994). The time of this early harvest is dependant on fibre development, beginning
when the rhizome is 45% fibre free and ending when the fibre free content reaches
35% (Whiley 1980). Rhizomes grown beyond this are used fresh as a culinary
product or dried for use as a spice or extraction of oleoresins and oils. The Canton
6
variety of ginger, which has larger knobs, is grown for the late harvest of rhizomes in
April or over fifteen months after planting (Stirling 2004; Sanewski 1994). The longer
growth period of late harvest ginger is associated with increased susceptibility to the
two major pathogens that affect regional production, Fusarium oxysporum forma
specialis zingiberi and Meloidogyne root-knot nematodes (Stirling 2001; Stirling
2004).
2.2. Impacts of the ginger pathogen Fusarium oxysporum f. sp. zingiberi in
regional production.
Infection of ginger by the fungal pathogen, Fusarium oxysporum forma specialis
zingiberi (Foz) may manifest as a severe rhizome rot, stunting of plants, yellowing of
leaves and a vascular wilt that results in plant death (Pegg et al. 1974). The informal
designation of Fusarium oxysporum into form species is based the on specificity of
the host plant infection, that is Foz is known to only infect ginger (Burgess et al.
1981). The mycelium of Fusarium oxysporum produces micro- and macro-conidia
that are usually associated with short-term survival. Long-term survival of Fusarium
oxysporum is facilitated by the production of chlamydospores formed in plant tissue
or by soil borne mycelium (Burgess et al. 1981).
While infection by Fusarium
oxysporum diseases often occurs through roots, entry of Foz is reported to be via
rhizome cracks or wounds in ginger
(Pegg et al. 1974; Burgess et al. 1981).
Infected rhizomes display an internal brown discolouration.
As the disease
progresses, rhizomes become shrivelled until eventually only a shell and fibrous
tissue remain (Pegg et al. 1974). Fungal growth may result in occlusion of the
vascular system, which is associated with yellowing and death of shoots.
While symptoms of Foz infection (or Fusarium yellows) are typically more prominent
in late harvest ginger, if Foz affected planting material is used, under conducive
conditions a rapid progression of rotting may prevent germination or a yellow shoot
may be produced that undergoes premature death (Pegg et al. 1974). In the late
1990s Foz was implicated in high incidences of rhizome rotting or plants that
displayed leaf yellowing and died, causing poor establishment in around one third of
regional farms (Stirling 2004). The emergence of this disease earlier in the season
7
was associated with planting of seed pieces infected with Foz; increased inoculum
levels in soil (as a result of rhizomes being left in the ground for longer periods due
to increased demand for fresh ginger); increased wounding of rhizomes as a result
of mechanisation of farming practices; environmental factors (high levels of rainfall
and soil moisture particularly when Foz occurred with Erwinia chrysanthemi); and
successive cropping of ginger/poor rotational practices (Stirling 2004).
Vegetative
compatibility analyses (that demonstrate sexual/DNA compatibility) indicated that
Foz isolates in the Sunshine Coast region belonged to the same group (Stirling
2004). This is consistent with the hypothesis that Foz was spread throughout the
industry from a source of infected planting material (Stirling 2004).
The use of disease free ginger planting material has been critical in reducing losses
caused by Foz. This has included discarding of seed pieces that display brown
internal discolouration and chemical disinfection of cutting implements during
preparation of planting material (Stirling 2004).
Disease free plantlets have been
produced via tissue culture based propagation of ginger (Smith and Hamill 1996).
Such plantlets have been used to establish “mother blocks” that supply growers with
planting material of low disease incidence. These sites have been established on
land that has not previously grown Foz affected ginger. Similarly, farm equipment
and machinery must not have been contaminated with Foz.
Currently there is
insufficient clean planting material to supply the entire industry (Stirling 2004).
A further measure to reduce the incidence of Foz infection of plants has included
treatment of seed pieces with the fungicide, benomyl, prior to planting (Pegg et al.
1974; Stirling 2004).
This may provide protection against infestation of seed
surfaces in the initial stages of growth, but fungicide treatment does not provide
protection to the newly formed rhizome that grows out from the seed piece (Stirling
2001).
Once introduced into soil, Foz may survive as a saprophyte, produce
resistant chlamydospores and persist for up to ten years (Pegg et al. 1974; Khatun
et al. 2003). Measures to control the infection of ginger by soil borne propagules of
Foz are not known.
8
Fusarium often occurs as a “disease-complex” with parasitic root-knot nematodes,
where lesions caused by nematodes are a possible route of entry for Fusarium
(Back et al. 2002).
2.3. Significance of Meloidogyne root-knot nematodes in regional ginger
cultivation.
Plant parasitic root-knot nematodes (RKN) Meloidogyne javanica and Meloidogyne
incognita may be transmitted on ginger planting material but are described as soil
borne pathogens that may be present in virgin soil (O’Brien and Stirling 1991; Stirling
1994). Larvae of RKN invade plant roots (and eventually rhizomes) and induce the
formation of giant cells and galls. Galling of roots at early harvest may reduce yields
and by late harvest galls on the rhizome may result in further losses (O’Brien and
Stirling 1991).
Conventional control measures for root-knot nematodes in regional ginger
production include crop rotation and the incorporation of cover crops with poultry
manure (150m3 /Ha) prior to planting and the use of nematode free planting material
(Stirling 1989; Stirling and Nikulin 1998). Where ginger is cropped in successive
years due to economic demands, preplant nematacides are frequently applied as an
assurance against devastating losses that may be caused by root-knot nematodes
(Stirling and Nikulin 1998; Stirling 1994).
The volatile-fumigant nematacide,
metham sodium is registered for use in the ginger industry. Methyl isothiocyanate
(MITC) is liberated on contact of metham sodium with the soil (Gerstl et al. 1977).
MITC is distributed via the gaseous phase and water films in soil and is biocidal to
many nematodes, as well as bacteria, fungi and weeds (O’Brien and Stirling 1991;
Collins et al. 2006; Gerstl et al. 1977).
Increased growth of plants may occur in response to soil fumigation that is not
always be explained by pathogen reduction alone (O’Brien and Stirling 1998; Martin
and Bull 2002). Bacterial populations with enhanced biodegradation capacity may
increase following treatment of soil with metham sodium and increased resilience of
soil microflora may result in decreased effectiveness of such nematacides over time
(Kapouzas et al. 2005; Ibekwe et al. 2001; Ibekwe et al. 2004; Matthiessen et al.
2004). The use of these nematacides is also being phased out due to long-term
9
adverse effects on soil biodiversity and their mammalian toxicity (Macalady et al.
1998; Klose et al. 2006).
2.4.
Impact of conventional farming practices on soil biodiversity and the
environment.
High input farming, involving soil fumigation, inorganic fertiliser application,
monoculture (the growth of one plant) and tillage, as used in the conventional
cultivation of ginger, may impact negatively on soil biodiversity (Garbeva et al. 2004;
Johansson and Finlay 2004; Wardle 1995; Bunemann et al. 2006; Wolf and Synder
2003). As stated by Griffiths et al. (1997) classical concepts of diversity involve
species richness, evenness and composition (i.e. the number of different species
present, the distribution of each species and the type and relative contribution of the
particular species present). Bacterial richness and evenness can be described by
Hill’s diversity numbers N1 and N2 (Ludwig and Reynolds 1988). The Shannon
Weaver Diversity Index (Shannon and Weaver 1949) and the maturity index of
nematode communities have also been used to estimate the biodiversity of soil
(Bongers 1990).
Soil micro-organisms, namely bacteria, actinomycetes and fungi, are integral in
processes that affect plant growth including: 1) formation of soil aggregates, by
production of extracellular polysaccharides and mucilages, that affects soil porosity
and thus water drainage/infiltration and exchange of gases (release of CO2 and
influx of O2); 2) toxin degradation/bioremediation; 3) biological nitrogen fixation; 4)
resistance to plant pathogens; 5) production of plant stimulating hormones and; 6)
nutrient cycling/mobilisation in soil (Subba Rao 1999; Garbeva et al. 2004; Wood
1995; Wolf and Synder 2003; Kennedy 1999; Gentry et al. 2004; Dobbelaere et al.
2003; Nehl and Knox 2006; Saleh-Lakha and Glick 2007; Prosser 2007). Following
decomposition of plant, animal and microbial residues by soil micro-organisms,
nutrients are released from organic matter (mineralisation) in plant available forms or
more often immobilised in microbial biomass; humic substances (more persistent
reserves of organic matter) are also formed (Wolf and Synder 2003; Powers and
McSorley 2000).
These microorganisms serve as a substrate for higher order
10
trophic groups present in soil, such as nematodes and protozoa (that mineralise
significant quantities of N) that are in turn fed upon by other predatory nematodes,
mesofauna and macrofauna. The term “soil food web” has been used to describe
the flow of nutrients and energy between these trophic levels and is of importance in
soil ecosystem functioning and productivity (Schoener 1989; Ingham et al. 1985; De
Angelis 1992; Wardle 1995; Powers and McSorley 2000).
The sensitivity of soil organisms to perturbance, for example due to agronomical
practices, may result in reduced complexity and functional competence of soil food
webs and negatively impact on soil health. As stated by Doran and Parkin (1996)
soil health can be defined as the capacity of a soil to function within an ecosystem,
to sustain biological productivity, maintain environmental quality and promote plant
and animal health. Nematode community populations have been correlated to soil
physical, chemical and biological properties and therefore have been used as an
indicator of soil health, which is closely tied to the concept of biodiversity (Pattison et
al. 2004; Stirling et al. 2005; Neher 2001; Ritz and Trudgill 1999). The utility of
nematode trophic groups as a biological indicator of soil health results from the
reliance of population levels on other microorganisms present and the variable
growth rate of different groups. For example, the prevalence of higher order trophic
groups (predatory nematodes, fungivores and bacterivores) is dependant on levels
of organisms on which they feed (nematodes, fungi and bacteria respectively).
Bacterivores are small, have a short generation time (few days) and typically
colonise disturbed habitats (Stirling 2005). The larger omnivorous and predatory
nematodes have a longer life cycle and may take months to years to re-establish
following soil disturbance (Stirling 2005).
Plant parasitic nematodes may
accumulate when nematode diversity is reduced, for example due to agronomical
measures (particularly tillage and N fertiliser application) and may indicate poor
ecosystem health (Berkelmans et al. 2003; Sarathchandra et al. 2001).
Calculation of bacterial, fungal and nematode biomass (mass of living cells) has also
been used to assess impacts of agronomical measures on soil biodiversity, although
this approach may not reflect structural or functional alterations in microbial
communities, for example due to fertiliser application and tillage (Wardle 1995;
Bunemann et al. 2006; Mazzola 2004). Therefore assessment of biomass is often
11
used in conjunction with methods, such as measurement of fluorescein diacetate
hydrolysis (to assess microbial activity) and culture based methods (Bunemann et
al. 2006; Stirling et al. 2005; Chen et al. 1988). Cultivation-dependant methods,
which rely on the ability of bacteria and fungi to grow on laboratory media, have only
enabled characterisation of between 0.1 and 5% of soil microorganisms (Amann et
al. 1995; Ovreas and Torsvik 1998; Rondon et al. 1999; Rondon et al. 2000).
The
presence of high numbers of residual spores of bacteria and fungi is a further
limitation in the use of culture-based analyses for analyses of community
populations (Mazzola 2004).
Techniques that have been used for the taxonomic analysis of bacteria recovered by
culture based analyses include: 1) biochemical analysis, for example by
commercially available API™ test strips and Biolog™ plates; 2) Phospholipid fatty
acid analysis (PLFA) or fatty acid methyl ester analysis (FAME) that rely on the use
of signature molecules for identification and 3) Sequencing of the small subunit
ribosomal RNA gene (16S rDNA) (Garbeva et al. 2004; Wunsche et al. 1997;
Mazzola 2004; Spratt 2004; Kirk et al. 2004).
The rRNA gene displays a low level
of evolutionary divergence and has been sequenced in many microorganisms (Lane
et al. 1985; Weisburg et al. 1991). Compilations of these sequences at the
Ribosomal Database Project and Genbank have enabled comparative phylogenetic
analysis of bacterial isolates (Cole et al. 2005; Altschul et al. 1997; Neefs et al.
1991). The polymerase chain reaction (PCR) has been used to amplify 16S rRNA
genes from community DNA samples, which is then used to construct clone libraries
or separated on high-resolution gels, to enable the identification of unculturable
bacteria and generate community level genetic fingerprints (Teidje et al. 1999;
Nakatsu 2007; Heuer and Smalla 1997). Such methods that enable separation of
rDNA
fragments
from
a
mixed
sample
include
denaturing
gradient
gel
electrophoresis (DGGE), single stranded conformational polymorphism (SSCP) and
terminal restriction fragment length polymorphism (T-RFLP) (Fischer and Lerman
1983; Muyzer et al. 1993; Liu et al. 1997; Dunbar et al. 2000; Schweiger and Tebbe
1998).
DDGE has been employed to demonstrate alterations and reduced
complexity of microbial communities in response to tillage, cultivation, fumigants and
fertiliser application (van Elsas et al. 2002; Garbeva et al. 2006; Marschner et al.
2003; Ibekwe et al. 2001; Dungan et al. 2003; Peixoto et al. 2006).
12
Biases inherent in these molecular methods may result from: 1) preferential
amplification of templates present in high copy number (dominant populations), thus
group specific primers may be needed for the detection of bacteria present at lower
frequencies; 2) inefficient lysis of resistant bacteria during DNA extraction; 3)
production of one band/sequence by a number of closely related species or 4) the
production of more than one band by a single strain of bacteria due to rDNA
heterogeneity (Garbeva et al. 2004; Mazzola 2004). Thus the characterisation of
soil microbial communities, that may contain 109 bacteria and 104 bacterial species
per gram (Torsvik et al. 1996; Amann et al. 1995; Torsvik et al. 1990) is a difficult
task, where molecular methods such as DGGE represent primarily dominant
populations and a very large number of clones may be needed to be screened in
libraries in detailed analysis. In contrast to labour and cost intensive construction
and screening of cDNA libraries, high throughput micro-array technology, using
taxon specific 16S rDNA probes, is expected to greatly facilitate future research that
aims to characterise complex microbial populations (Sanguin et al. 2006; Sessitsch
et al. 2006; Gentry et al. 2006).
A range of methods has been devised for assessment of functional activities of soil
microbial communities.
These methods include: 1) Use of primers in the
aforementioned molecular methods that are targeted toward functional genes, such
as those that encode nitrogenase enzymes (involved in nitrogen fixation) or antibiotic
synthetic loci; 2) Construction of metagenomic libraries (bacterial artificial
chromosome or fosmid libraries) using DNA extracted from an environmental sample
and sequence analysis to yield information such as enzymatic activities present and;
3) Use of BIOLOG Ecoplates® for profiling of community metabolic functions (as an
indicator of metabolic diversity) based on carbon source utilisation by culturable
bacteria (Rondon et al. 2000; Torsvik and Ovreas 2002; Leveau 2007; Jenkins et al.
2004; Mazzola 2004; Palojarvi et al. 1997). The latter method has been used to
demonstrate that reduced microbial diversity may result from cultivation, tillage and
N fertiliser application (Yan et al. 2000; Diosma et al. 2006; McCraig et al. 2001;
Lupwayi et al. 2001; Chen et al. 2007; Larkin 2003).
13
Thus a range of methods has been used to demonstrate that soil biodiversity may be
compromised by agronomical practices. It is also known that soil microorganisms
play an important role in processes such as nutrient cycling, formation of soil
aggregates and disease suppression.
Hence reduced soil biodiversity resulting
from intensive agricultural practices may: 1) negatively impact on soil physical and
chemical properties and reduce the availability of macro-elements (carbon, nitrogen,
phosphorus) and micronutrients (eg. iron, magnesium, molybdenum, copper, zinc)
required for plant growth; 2) favour the accumulation of pests and deleterious
bacteria (which have a negative impact on plant growth) and; 3) culminate in a
decline in soil quality, reduced yield of crops and decreased nutritional value of the
food (Johansson and Finlay 2004; Garbeva et al. 2004; Garbeva et al. 2006; van
Elas et al. 2002; Martin and Bull 2002; Suslow and Schroth 1982; Nehl et al. 1997;
Ogut and Er 2006; Kennedy 1999; van Bruggen et al. 2006; Krull et al. 2003).
As well as being detrimental to soil microorganisms commonly used inorganic
nitrogen and phosphorus fertilisers may be leached from the soil into ground and
surface waters, encouraging weed and algal infestations (Bunemann et al. 2006;
Yan et al. 2000; Diosma et al. 2006; Wood 1995; Hart et al. 2004; Powers and
McSorley 2000). It is estimated that the world- wide usage of nitrogenous fertilisers
is in the order of 60 million tonnes per annum (Woods 1995). As discussed by
Kennedy and colleagues (2004), following denitrification and volatilisation of
nitrogenous fertilisers, greenhouse gases N2O, NH3 and NO are produced. The
global warming potential of N2O is approximately 300 times that of CO2 and
therefore measures that facilitate a reduced reliance of crop production systems on
applied nitrogen are of significance (Venterea et al. 2005; IPCC 1996).
In order to address human health and environmental concerns associated with
intensive farming and to achieve sustainable practices in agriculture, integrated pest
management is being targeted, which may involve conservation tillage, organic
amendment, crop rotation, a diversity of cover crops and introduction of plant
beneficial bacteria. Such bacteria may reduce required inputs of synthetic fertilisers,
promote plant growth and/or assist in the management of phytopathogens
(Jacobsen et al. 2004; Okon and Labandera-Gonzalez 1994; Vessey 2003;
Dobbelaere et al. 2001; Fravel 2005).
14
2.5. Plant-bacteria interactions.
The interaction between plants and microorganisms begins at germination.
As
discussed in a recent review by Nelson (2004), carbon-containing exudates released
during seed germination may stimulate proliferation of microbes carried on and in
the seed and in soil surrounding the seed (the spermosphere).
Germination of
Fusarium and Pythium was demonstrated to occur maximally in response to the
release of seed exudates; fatty acid metabolising bacteria present in disease
suppressive composts inhibited this interaction and subsequent infection of
cottonseed by Pythium (McKellar and Nelson 2003, Hoitink and Boehm 1999, Short
and Lacey 1974).
Similarly microbial proliferation in soil may be stimulated by root exudates, which
may include carbohydrates, carboxylic acids and amino acids, as well as enzymes,
sterols, fatty acids and growth factors (Uren 2001; Pinton et al. 2001). Up to 40% of
photosynthetic carbon may be lost by plant root exudation, although basal levels of
exudation have been estimated at 3 to 5 % (Lynch and Whipps 1991; Pinton et al.
2001). The process of rhizodeposition includes substances released by plant-root
exudation and plant residues (Brimecombe et al. 2001; Jones et al. 2004). As such
the soil surrounding the plant roots is different in chemical, biochemical and
microbiological composition from the bulk soil; this volume of soil is called the
rhizosphere (Uren 2001; Pinton et al. 2001; Smalla et al. 2001).
Bacteria that are
able to compete and establish in the rhizosphere have been termed rhizobacteria.
Bacteria that have been frequently isolated from the rhizosphere of different plants
include
species
of
Pseudomonas,
Flavobacteria,
Alcigenes,
Arthobacter,
Comamonas, Agrobacterium and Rhizobium (Pinton et al. 2001; Peixoto et al. 1994;
Whipps 2001; Mazzola 1999; Costa et al. 2006).
Plant root exudation may be induced by rhizobacteria in a species-specific way
(Bolton et al. 1993; Chanway et al. 1988; Merharg and Zillham 1995; Phillips et al.
2004). Such observations have lead to the suggestion that a co-evolution of plants
and rhizobacteria has been concomitant with the development of selective root
15
exudation that favours the development of beneficial rhizosphere populations (Bolton
et al. 1993; Phillips et al. 2004).
Patterns of plant exudation may vary between plant species and cultivars, which
may explain the plant specific nature of rhizosphere communities (Ryan et al. 2001;
Van der Krift et al. 2001). Grayston and colleagues (1998) analysed metabolic
profiles of microbial communities associated with wheat, ryegrass, bentgrass and
clover in two different soil types.
Results demonstrated differences in root
associated microbial communities between different crops and uncultivated soil.
Berg and colleagues (2002) also demonstrated that Verticillium antagonists in the
rhizosphere of potato, strawberry and oilseed rape were plant specific. Due to the
plant specific nature of rhizodeposition, conditions might be more favorable for a
selected population of micro-organisms and this might be one of the reasons why
growing one plant (monoculture) can allow: i) the accumulation of deleterious
rhizobacteria and pests; ii) a reduction in micro-organisms that suppress disease
and iii) a reason that a diversity of cover crops is important in integrated approaches
to sustainable farming (Garbeva et al. 2006; Garbeva et al. 2004; Mazzola 1999;
Mazzola 2004; Lupwayi et al. 2004; Larkin 2003).
In addition, changes in plant exudation over the plant growth cycle is linked to
concurrent differences in bacteria detected in the rhizosphere (Atkinson and Watson
2000; Smit et al. 2001).
In analyses of culturable population isolated from the rice
rhizosphere, Mew and co-workers (1994)
observed that an increase in
Pseudomonas population densities corresponded to decreased population levels of
Bacillus as the crop progressed. Conversely in the wheat plant, the incidence of
P. putida in the rhizosphere was shown to decline with the age of the plant, while the
reverse was true for Bacillus (Wong 1994). Thus variations in bacterial populations
over the growth cycle may be plant dependant.
Microbial populations associated with a given plant type may also be influenced by
soil type, which is determined by factors such as parent material, relative proportions
of sand, silt and clay, levels of organic matter and pH (Harpstead et al. 2001; Ulrich
and Becker 2006).
In some cases, soil type may be the major determinant of
16
microbial communities in the spermosphere and rhizosphere (Buyer et al. 1999;
Latour et al. 1996). Garbeva and co-workers (2004) discuss that in other instances
plant type and cultural practices are the dominant “driving forces” that determine
microbial communities present in the rhizosphere.
Considering the vast number of species of bacteria found in soil, only a minority is
reported to be able to compete in the rhizosphere, and an even more select
population is able to attach to the root surface (rhizoplane) (Van Tran et al. 1994). A
further subset of these bacteria is able to penetrate plant roots and survive in plant
internal tissues; such species that do not have a negative effect on plant growth are
termed endophytes (Sturz and Nowak 2000; Hallman and Quadt-Hallman 1997).
Endophytic bacteria may be transmitted on vegetative planting material or seed;
alternatively these bacteria may originate from the spermosphere or rhizosphere and
enter the plant via natural wounds that occur during plant growth (for example sites
of lateral root emergence and radicle germination), stomata, epidermal junctions,
root hairs or by an active process, where plant cell walls are degraded by cellulytic
and pectinolytic enzymes (Sprent and de Faria 1988; Huang 1986; Mahaffee et al.
1997b; Patriquin and Dobereiner 1978; Reinhold and Hurek 1988; Hallman et al.
1997; Ryan et al. 2008).
Colonisation by endophytic bacteria is: i) typically in
intercellular spaces and the vasculature, although specific bacteria may be found in
intracellular locations and ii) may be restricted (for example to the root cortex) or
systemic (Hurek et al. 1994; James et al. 1994; Mahaffee et al. 1997; Patriquin and
Dobereiner 1978; Quadt-Hallmann et al. 1997).
Endophytes may differ from rhizosphere and non-rhizosphere strains of the same
species. As with rhizosphere bacteria, endophyte populations may vary with soil
type and may be specific to the host plant (Zinniel et al. 2002; Siciliano et al. 2001;
Conn and Franco 2004).
17
2.6. Beneficial effects of bacteria on plant growth.
Plant associated bacteria may have beneficial, inhibitory or neutral effects on plant
growth (Dobbelaere et al. 2003). Kloepper et al. (1980) used the term “plant growth
promoting rhizobacteria” to describe bacteria having a stimulatory effect on plant
growth.
The synonymous term, plant growth promoting bacteria (PGPB) that
includes rhizosphere and endophytic bacteria is selected for further use in this
manuscript (Bashan and Holgiun 1998; Compant et al. 2005). Such bacteria may
increase the availability of plant nutrients and/or stimulate root proliferation, resulting
in increased water and nutrient uptake and yield of crops. Increased plant vigour,
induced systemic resistance and/or antagonism of plant pathogens by PGPB may
reduce disease incidence and further improve yields in crop production. (Whipps
2001; Vessey 2003; Bashan 1998; Bashan et al. 2004; Compant et al. 2005; Lucy et
al. 2004; Sturz and Nowak 2000).
2.6.1. Nutrient availability and uptake.
Controlling the availability of iron, nutrient mobilisation, biological nitrogen fixation
(discussed later) and increasing nutrient uptake include mechanisms by which
bacteria may affect plant growth. Certain bacteria, including strains of fluorescent
pseudomonads and Azospirillum species, produce sideophores that sequester iron
in the rhizosphere, making it unavailable to other microbes but often also usable by
plants (O’Sullivan and O’Gara 1992; Bloemberg and Lugtenberg 2001; MercadoBlanco and Bakker 2007). Production of organic acids or phosphatases by certain
bacteria may mobilise nutrients such as phosphorus (Rodriguez et al. 2006; see
below). PGPB may also competitively exclude pathogens by utilising nutrients
exuded by plants and occupying a similar niche on plant roots.
For example
Bolwerk et al. (2003) used fluorescently labelled Pseudomonas spp. and Fusarium
oxysporum f. sp. lycopersici (Fol) to demonstrate that both microorganisms occupied
the same niche on tomato root cells (intercellular junctions), a site of plant exudation.
Occupation of these sites by the antibiotic producing Pseudomonas chlororaphis
strain reduced colonisation by Fol.
Similarly Van Dijk and Nelson (2000)
18
demonstrated that metabolism of fatty acids in the cottonseed spermosphere by
Enterobacter cloacae prevented germination of sporangia of Pythium ultimatum.
In addition PGPB may stimulate increased uptake of macronutrients and
micronutrients.
For instance Esitken and associates (2006) demonstrated that (in
addition to increased yields) N, P, K Fe, Zn and Mn were elevated in leaves of sweet
cherry (Prunus avium) following the application of specific Bacillus and
Pseudomonas strains.
In corn (Zea mays) and Sorghum bicolour, Azospirillum
brasilense Sp7 and Azospirillum brasilense Cd increased K+ (19% to 32%) and
H2PO4 (50% to 66 %) in root segments (Lin et al. 1983).
Ogut and Er (2006)
demonstrated A. brasilense Sp7 increased concentrations of Mn, Zn and Cu in bean
grown with supplementary P (25kg/ha).
The ability of PGPB to increase
micronutrient or macronutrient content of plants has also been reported in barley
(Hordeum vulgare), raspberry, bean, wheat, cucumber and many other plants
(Canbolat et al. 2006; Orhan et al. 2006; Bashan et al. 2004). Increased nutrient
uptake may result from enhanced proton efflux from roots, regulation of specific ion
transport channels or increased root growth induced by phytohormones (Bashan et
al. 1989; Amooaghaie et al. 2002; Vessey 2003; Hamdia et al. 2004; Bertrand et al.
2000).
2.6.2. Plant stimulating hormones and metabolites.
Plant growth promoting bacteria may produce hormones, such as auxins (indole 3acetic acid) and gibberellins, which stimulate plant growth (Woodward and Bartel
2005; Steenhoudt and Vanderleyden 2000; Yanni et al. 2001).
Bottini and
colleagues (2004) reviewed the promotion of many facets of plant growth by
gibberellins, including germination, root growth and stem proliferation. Production of
gibberellins by strains of Bacillus macroides, Bacillus subtilis, Azospirillum brasilense
and Azospirillum lipoferum has been documented (Joo et al. 2004; Bottini et al.
1989; Janzen et al. 1992).
Specific strains of Azospirillum brasilense and Pseudomonas fluorescens have
demonstrated the capacity to excrete indole acetic acid, an important auxin in most
19
plants (Dobbelaere et al. 1999; Barbieri et al. 1986; Benizri et al. 1998; Woodward
and Bartel 2005; Spaepen et al. 2007). Auxins have been shown to enhance the
activity of plasma membrane ATPase, which leads to acidification of the extracellular
space, allowing loosening of the cell wall thus enabling cell expansion and
subsequently division (Hager 2003). In addition, many other aspects of plant growth
and development are affected by auxins, which may induce changes in gene
expression, protein phosphorylation, production of certain plant hormones (for
example ethylene and gibberellic acid) and repression of cytokinin biosynthesis
(Woodward and Bartle 2005; Spaepen et al. 2007).
IAA plays an important role in
inducing root (and/or root hair) development and proliferation, resulting in an
increased surface area from which the plant can take up nutrients (Skoog and Miller
1957; Dobbelaere et al. 1999). This may be reflected as an improved nutrient and
water status of the plant, increased yields, increased resistance to disease and a
reduced requirement for synthetic fertilisers (Bashan et al. 2004; Okon and
Labandera-Gonzalez 1994; Vessey 2003; Dobbelaere et al. 1999).
Ryu and colleagues (2003) demonstrated the bacterial volatile metabolites, acetonin
and
2,3-butanediol,
produced
by
Bacillus
subtilis
GB03
and
Bacillus
amyloliquiefaciens IN937a promoted the growth of Arabidopsis thaliana seedlings.
Petri dishes with a centre partition were used to spatially separate seedlings and the
bacterial strains. The positive effect on plant growth was mimicked by exposure of
the seedlings to extracted bacterial volatiles and synthetic 2,3-butanediol.
Use of
various Arabidopsis mutants indicated that volatile metabolites of B. subtilis GB03
induced growth promotion via cytokinin-dependant signalling pathways. Cytokinins
are known to stimulate plant cell division (Lynch 1985). Growth promotion induced
by B. amyloliquefaciens IN937a was independent of known pathways (ethylene,
gibberellic acid, cytokinins and brassinosteroids) although studies with auxin
deficient mutants were not conclusive. These volatiles were later demonstrated to
confer resistance to disease in Arabidopsis (Ryu et al. 2004).
Many processes in plants are regulated by the volatile hormone ethylene, including
seed emergence, fruit ripening, senescence and defence responses against plant
pathogens (Abeles et al. 1992; Saleh-Lakha and Glick 2007; van Loon et al. 2006).
While generation of ethylene is usually associated with alleviation of stress, high
20
levels may inhibit root growth and are associated with decreased functional capacity
of plants in abiotic or biotic stress (Saleem et al. 2007).
The enzyme 1-
aminocyclopropane-1-carboxylate (ACC) deaminase degrades ACC a precursor of
ethylene, thereby regulating levels of this hormone (Glick et al. 1998; Argueso et al.
2007). PGPB that produce ACC deaminase may lower plant ethylene and exert a
positive effect on plant growth and resistance to stress (Saleem et al. 2007). In
contrast, Ribaudo and co-workers (2006) associated increased levels of indole-3acetic acid (IAA) and ethylene with the enhanced growth response of tomato
following the introduction of A. brasilense FT 326.
Application of ethephon, a
compound that releases ethylene, produced increased root and root hair growth
similarly to the bacterium. In addition, when binding of ethylene to its receptor was
inhibited, the growth promoting effects of the bacterial strain were ameliorated.
Therefore results suggested that the A. brasilense FT 326 induced growth response
was mediated via an ethylene dependant-signalling pathway, which may have been
activated by IAA, a positive regulator of ethylene synthesis (Abeles et al. 1992;
Kende 1993).
Hence complex mechanisms may be involved in the plant response
to ethylene, as low levels are required for many plant growth processes (explaining
positive effects of bacteria such as FT 326), while high levels are inhibitory to plant
growth (explaining positive effects of ACC deaminase excreting bacteria). Similar
complexities have been encountered in elucidation of the role of ethylene as a
modulator of plant defence responses against phytopathogens (van Loon et al.
2006).
2.6.3. Induced resistance against plant pathogens.
Innate plant defence mechanisms may be activated following infection by pathogenic
organisms (Pieterse and van Loon 2004).
These mechanisms may be triggered
following the recognition of pathogen virulence factors (encoded by avirulence
genes) by plant resistance (R) factors (encoded by R-genes), which is known as a
compatible interaction (Montesinos et al. 2002; Nimchuk et al. 2003).
Defence
responses may include localised electrolyte leakage, an oxidative burst and an
accumulation of salicylic acid.
Systemically, salicylic acid-, ethylene- and/or
jasmonic acid-dependant signalling pathways mediate the hypersensitive response
and transcription of defence-related proteins. Inducible pathogenesis-related (PR)
21
proteins and defence factors include superoxide dismutase, proteinase inhibitors,
monoxygenases, lignin, proteinase inhibitors, phenolic compounds, callose,
antimicrobial compounds (such as phytoalexins and phenolics), chitinases and
glucanases (van Loon et al. 2006b; Montesinos et al. 2002).
Suppression of plant
defence responses or failure of plant encoded R factors to recognise pathogen
virulence factors (an “incompatible response) may result in disease progression
(Nomura et al. 2005; Nimchuk et al. 2003).
Micro-array analyses have
demonstrated that the expression of hundreds of genes may be affected in
compatible and incompatible responses (Tao et al. 2003, van Wees et al. 2003). de
Vos and colleagues (2006) analysed gene expression profiles in Arabidopsis
following attack by a number of different pathogen types. Results indicated that
pathogen-specific responses may be induced in Arabidopsis, for example in
transcription of genes induced by jasmonic acid and cross-communication between
signalling pathways mediated by jasmonic (JA) acid, salicylic acid (SA) and
ethylene. Further, the signalling pathway activated may be dependant on plant type,
for instance resistance to Botrytis cinerea was shown to occur through pathways
mediated by salicylic acid in tomato, but not tobacco (Achuo et al. 2004). While
defence responses may be activated by the application of salicylic acid or ethylene,
disease progression may be stimulated when ethylene is applied after manifestation
of disease symptoms and ethylene is a virulence factor of a number of plant
pathogens (Elad 1993; Chague et al. 2006; Weingart et al. 2001; Marco and Levy
1979).
Thus the nature of the response to ethylene may depend on time of
exposure and pathways activated in a particular plant by specific pathogens (van
Loon et al. 2006b). Hence mechanisms involved in the activation of plant defence
responses following pathogen challenge are highly complex and not completely
elucidated.
Following the induction of plant defence responses by a compatible interaction with
a necrotizing pathogen, PR-proteins remain elevated and resistance to further
infection by a wide variety of pathogens is enhanced, a phenomenon known as
systemic acquired resistance (SAR) (Ryals et al. 1996; Sticher et al. 1997; Durrant
and Dong 2004; de Vos et al. 2006). In order for SAR to be expressed,
responsiveness to salicylic acid and NPR1 (Non-expressor of pathogenesis-related
genes1) is required (Pieterse and van Loon 2004; Ryals et al. 1996; Dempsey et al.
22
1999). NPR1 is involved in the regulation of transcription of SAR induced genes and
co-ordinating signalling pathways dependant on SA and JA. When SAR is activated,
upon subsequent pathogen challenge or abiotic stress, enhanced induction of the
hypersensitivity response occurs and transcription of PR-proteins is potentiated (Kuc
1995; Conrath et al. 2006; Durrant and Dong 2004). As stated by Conrath and coworkers (2006) the physiological condition in which plants are able to better or more
rapidly mount defence responses, or both, to biotic and abiotic stress is called the
“primed state”. Priming may result from post-translational modification (for example
phosphorylation), elevated levels or increased activity of proteins and transcription
factors that are integral in mechanisms of plant defence and stress alleviation
(Conrath et al. 2002; Conrath et al. 2006; van Loon et al. 2006b).
Priming of plant defence responses and augmented resistance to disease is also
known to occur in induced systemic resistance (ISR) that is elicited by certain nonpathogenic rhizobacteria and endophytes (van Loon et al. 2006b; Conrath et al.
2006). As with SAR, the primed state of ISR results in faster and higher levels of
expression of plant defence genes upon pathogen challenge (Verhagen et al. 2004;
de Vos et al. 2006; van Loon et al. 2006b; Conrath et al. 2006; Maleck et al. 2000).
ISR, demonstrated by the spatial separation of eliciting rhizobacteria and challenging
pathogens, may induce changes in the plant, such as strengthening of cell walls,
which reduces the ability of pathogens to invade tissues (Siddiqui and Shaukat
2002; Zehnder et al. 2001; Ryu et al. 2004). Specific strains of Bacillus spp. and
fluorescent Pseudomonas spp. are known to induce ISR in a variety of different
plants.
The signalling pathways activated in ISR may depend on the eliciting
rhizobacteria.
In the case of fluorescent Pseudomonas species, ISR may be elicited in response to
salicylic acid, 2,4 – diacetylphloroglucinol (DAPG) or sideophores produced by the
bacteria or due the presence of bacterial lipopolysaccharide and flagellins (van Loon
et al. 1998; Bakker et al. 2007; Siqqiqui et al. 2001).
The signalling pathways
involved in ISR by Pseudomonas fluorescens WCS417r have been extensively
investigated in the model plant species, Arabidopsis thaliana. In this plant SAR is
activated via salicylic acid-dependant pathways and results in the systemic
accumulation of PR-proteins, while ISR elicited by WCS417r was associated with
23
elevated levels of PR-proteins only in roots and not in leaves; was not mediated via
salicylic-acid dependant pathways; required functional ethylene and jasmonic acid
responses and; may be associated with increased sensitivity to jasmonic acid and
ethylene and increased ethylene production following challenge inoculation (Pieterse
et al. 1996; Verhagen et al. 2004; van Loon et al. 2006b; Pieterse et al. 2000;
Pieterse et al. 2002). A range of pathogen types (Fusarium oxysporum, Alternaria
brassicicola and Peronospora parasitica) was resisted in Arabidopsis when ISR was
elicited by P. fluorescens WCS417r (Pieterse et al. 1998; Pieterse et al. 1996; Hase
et al. 2003). ISR was also elicited by WCS417r in a variety of plant types (tomato,
bean, carnation and radish), although the pathogens that are resisted may be plantspecific (Hase et al. 2003).
While transcriptome analysis demonstrated that ISR was not associated with an
accumulation of PR-proteins in leaves of Arabidopsis, following challenge inoculation
with the leaf pathogen Pseudomonas syringae augmented expression of 81 genes
occurred in the leaves and approximately one third of those were specific to
WCS417r inoculated plants (Verhagen et al. 2004). Conversely Cartieaux and
colleagues (2003) demonstrated increased transcription of defence related proteins
was much greater in leaves than in roots when ISR was elicited by in Pseudomonas
thivervalensis MLG45 in an Arabidopsis mutant that was not responsive to ethylene
and jasmonic acid. Further evidence that different pathways may be involved in ISR
depending on the eliciting Pseudomonas spp. was provided by the demonstration
that P. aeruginosa and P. fluorescens P3 may induce resistance via salicylic aciddependant pathways (de Meyer and Hofte1997; Maurhofer et al. 1998).
Different signalling pathways have also been implicated in ISR by Bacillus spp.
depending of the eliciting strain. For example Ryu and associates (2004) analysed
signalling pathways involved in ISR in Arabidopsis by Bacillus spp. against Erwinia
carotovora. Volatiles produced by B. subtilis GB03 elicited ISR via an ethylenedependant pathway (in contrast to the cytokinin-dependant growth promotion). In
contrast, induced resistance activated by volatiles of B. amyloliquifaciens IN937 was
independent of ethylene. ISR by both GB03 and IN937 was not dependent on SA,
JA or NPR1. Therefore ISR elicited by the bacterial volatiles was suggested to have
24
operated through a novel signalling pathway or alternatively is explained by
redundancy between different signalling pathways.
A number of different Bacillus spp. have been shown to elicit ISR against a range of
foliar pathogens, viral insect, diseases, fungal and bacterial pathogens in
greenhouse and field conditions as indicated in Table 1 (Kloepper et al. 2004).
25
Table 1. Induced systemic resistance by various Bacillus spp. under field or greenhouse conditions.
Eliciting Bacillus
spp.
Test Plant
Pathogen resisted
B. pumilus SE34
Bean
B. pumilus SE34
Pea
F. oxysporum f. sp. pisi Cell walls strengthened due to via callose apposition and elevated
phenolics.
F. oxysporum f. sp. pisi Restricted infiltration of pathogen, strengthening of cell walls via
callose apposition and elevated phenolics.
Leon-Kloosterziel et al.
2005
Benhamou et al. 1996
B. pumilus SE34
Tobacco
Peronospora tabacina Growth promotion; reduced sporulation of the pathogen.
Zhang et al. 2004
B. pumilus SE34
Tomato
Phytopthora infestans Increased plant height and weight; Ethylene and jasmonic-acid
dependant; Salicylic acid independent.
Yan et al. 2002
B. pumilus SE34
Tomato
F. oxysporum f. sp.
lycopersici
Deposition of polymorphic, osmophilic and amorphous material
that reduced pathogen colonisation.
Benhamou et al. 1998
B. pumilus 230-6
B. pumilus SE34
Sugar beet
Tobacco
Botrytis cinerea
Increased chitinase and glucanase, elevated peroxidase activity.
Ethylene and jasmonic-acid dependant; Salicylic acid independent.
Bargabus et al. 2004
Zhang et al. 2002
B. pumilus SE34
NPR1-, jasmonic acid- and ethylene-dependant.
Ryu et al. 2003
B. pumilus T4
Arabidopsis Pseudomonas
syringae
Arabidopsis P. syringae
Not dependant on jasmonic acid or NPR1.
Ryu et al. 2003
B. pumilus SE34
Tobacco
P. syringae
Salicylic acid-dependant, activation of PR-1.
Park and Kloepper 2000
B. pumilus T4
Tobacco
Peronospora tabacina Growth promotion; reduced sporulation of the pathogen.
B. pumilus INR7
Cucumber
Erwinia trachiephila
Growth promotion.
Zehnder et al. 2001
B. pumilus INR7
Cucumber
P. syringae
Growth promotion, increased yield.
Zehnder et al. 2001
Independent of ethylene SA, JA and NPR1 pathways.
Ryu et al. 2004
Ethylene dependent; independent of SA, JA and NPR1 pathways.
Ryu et al. 2004
B. amyloliquifaciens Arabidopsis P. syringae
IN937
B. subtilis GB03
Arabidopsis P. syringae
Pathways, mechanisms, results
26
Reference
Zhang et al. 2004
2.6.4. Antibiotic and lytic enzymes.
Lytic enzymes, such as chitinases and glucanases, secreted by certain bacteria
including specific Bacillus and Pseudomonas species, degrade chitin and glucans
present in fungal cell walls (Gooday 1990; de Boer et al. 1998; Bargabus et al.
2004).
Chitinolytic activity of bacteria can result in hyphal lysis and reduced
infection of plants by fungal plant pathogens, although production of antibiotics
and/or gulcanases or proteases may also be required for biocontrol activity of
certain bacteria (Mitchell and Alexander 1961; Whipps 2001).
The production of antibiotic compounds by soil microorganisms is described as a
natural defence mechanism to aid in their survival in soil (Mazzola et al. 1992).
Many Bacillus and Pseudomonas spp. have been shown to secrete a wide variety
of antifungal metabolites. Those produced by certain Bacillus spp. include cyclic
lipopeptides (CLPs) belonging to the iturin, surfactin and fengycin families. These
antifungal metabolites have a lipid moiety and may insert into cell membranes
forming ion-conducting pores; this results in increased permeability to K+ and other
ions and membrane destabilisation which may cause cell death (Maget-Dana et al.
1992; Maget-Dana and Peypoux 1994; Heerklotz and Seelig 2001; Deleu et al.
2005; Grau et al. 2000; Sheppard et al. 1991; Montesino 2007; Mizumoto et al.
2006; Vanittanakom et al. 1986).
Antifungal cyclic lipopeptides produced by fluorescent Pseudomonas spp. are also
structurally diverse and belong to the viscosin, amphisin and syringomycin families
(Raaijmakers et al. 2006; de Bruijn et al. 2007).
In addition to surfactant
properties, these CLPs may be involved in biofilm formation and motility, which
may contribute to biocontrol traits of the bacteria (Raaijmakers et al. 2006; Nielsen
et al. 2002; Nielsen et al. 2005; Andersen et al. 2003).
The contribution of these antifungal peptides to the biocontrol activity of relevant
bacterial strains has been demonstrated by i) loss of biocontrol ability in mutants
deficient in the production of these lipopeptides; ii) attainment of biocontrol activity
by introduction of biosynthetic genes into strains that do not naturally suppress
27
disease; iii) enhanced biocontrol activity by increased expression of antifungal
lipopeptides and iv) induction of disease suppression by application of cell free
supernatants containing these lipopeptides (Koumoutsi et al. 2004; Bais et al.
2004; Romero et al. 2007; Leclere et al. 2005; Asaka and Shoda 1996; Bolwerk et
al. 2003).
Further, research has demonstrated that following introduction of
producing bacterial strains, these fungitoxic lipopeptides may be detected at
concentrations that are inhibitory to fungal pathogens in soil or plants (Romero et
al. 2007; Toure et al. 2004; Ongena et al. 2005; Cazorla et al. 2007). Recently the
application of purified surfactin and (to lesser extent) fengycin were also shown to
elicit induced systemic in bean against Botrytis cinerea similarly to the inoculated
B. subtilis strain (Ongena et al. 2007).
Similarly, the antifungal metabolite 2,4 – diacetylphloroglucinol (DAPG) produced
by certain fluorescent Pseudomonas spp. is required for activation of ISR by these
bacteria (Bakker et al. 2007).
Other antifungal metabolites produced by
fluorescent pseudomonads may include one or a combination of phenazine,
phenazine-1-carboxylic acid (PCA), pyoluteorin and pyrrolnitrin (Thomashow and
Weller 1990; Raaijmakers and Weller 1998; O’Sullivan and O’Gara 1992; Mazzola
et al. 1992; Loper and Gross 2007; Chin-A-Woeng et al. 2003).
Phenazine and
pyrrolnitrin compounds may acts as electron shuttles and disrupt hyphal growth or
oxidative phosphorylation (Bolwerk et al. 2003; Chin-A-Woeng et al. 2003; Tripathi
and Gottlieb 1969).
2.7. Use of plant growth promoting bacteria in agriculture.
A significant potential exists for the management of soil-borne plant diseases,
decreasing reliance on applied fertilisers and increasing crop productivity by the
application of plant growth promoting bacteria (Duffy and Defago 1999; Lucy et al.
2004; Castro-Sowinski et al. 2007; Dobbelaere et al. 2001; Cocking 2005; Sturz
and Nowak 2000).
The term “biofertiliser” has been used to describe formulations of plant growth
promoting bacteria that may increase the availability of nutrients in forms usable by
plants and/or produce substances that stimulate root proliferation, resulting in
28
enhanced nutrient uptake, plant vigour and yield of crops, often at reduced rates of
inorganic fertiliser application; increased plant vigour may also suppress disease
(Vessey 2003; Hafeez et al. 2006; Kennedy et al. 2004). Bacteria used to target
plant pathogens by antagonism (antibiosis or parasitism) or that induce systemic
resistance have been referred to as biocontrol PGPB or biopesticides (Bashan
1998; US-EPA 2005).
An objective of inoculation with PGPB, as described by Van Tran and co-workers
(1994) is to displace deleterious rhizobacteria with species that are beneficial to
plant growth.
Bacterial inoculants that augment populations of fluorescent
Pseudomonas spp., Bacillus spp, Azospirillum spp, phosphate solubilizing bacteria
and/or mycorrhizal helper bacteria have been applied to agronomically important
crops in order to establish such a “beneficial rhizosphere” (Atkinson and Watson
2000). The establishment of beneficial rhizosphere populations may assist in the
expression of the full genetic potential of plants (Cook 2000).
2.7.1. Fluorescent Pseudomonas species.
Fluorescent Pseudomonas (RNA group 1) species produce sideophores, present
as yellow-green pigments that fluoresce under UV light (Elliot 1958) and include
Pseudomonas fluorescens, Pseudomonas putida and Pseudomonas aeruginosa
(Mercado-Blanco and Bakker 2007). Sequestration of iron by these sideophores
may result in suppression of plant disease by certain strains of fluorescent
pseudomonads (O’Sullivan and O’Gara 1992). As discussed earlier, sideophores,
flagellins, 2,4-DAPG and LPS may induce systemic resistance in plants.
Competitive exclusion and antibiosis may also be involved in the biocontrol activity
of fluorescent Pseudomonas spp. (Lugtenberg and Dekkers 1999; Bolwerk et al.
2003; Haas and Defago 2005).
Meena and Mathur (2003) tested the ability of a fluorescent Pseudomonas spp. to
reduce rhizome rot of ginger caused by Fusarium solani.
The bacteria were
applied to seed pieces (although the concentration of cells used was not reported)
and the plants were grown in autoclaved garden soil.
29
The application of
Trichoderma spp. to the seed and the soil was more effective in reducing the
incidence of rhizome rotting and promoting plant growth than application of the
Pseudomonas spp. to the seed alone, although the application of the bacteria to
the soil was not tested. Further, the actual species used in this study was not
described. Numerous other reports have documented the ability of fluorescent
Pseudomonas species to reduce infection of different plants by F. oxysporum,
typical examples are listed in Table 2.
Table 2.
Examples of fluorescent Pseudomonas spp. with biocontrol activity
against different Fusarium oxysporum diseases.
Fluorescent
Pseudomonas species
Form species of
Fusarium oxysporum
Plant
Reference
P. chlororaphis PCL 1391 radicus-lycopersici
Tomato
P. fluorescens
raphani
Radish
Chin-A-Woeng et al.
1998
de Boer et al. 1999
P. fluorescens 63-28
pisi
Pea
Benhamou et al. 1996
P. putida (two strains)
melonis
Musk-melon Bora et al. 2004
P. fluorescens WCS365
radicus-lycopersici
Tomato
Dekkers et al. 2000
P. putida WCS538
dianthi
Carnation
Lemanceau et al. 1992
Increased populations of fluorescent Pseudomonads have often observed in the
rhizosphere of diseased plants (Mazzola and Cook 1991). The colonisation of
hyphae of Fusarium oxysporum f. sp. lycopersici (Fol) by certain fluorescent
Pseudomonas species, that also produce antifungal metabolites and lytic
enzymes, may contribute to the biocontrol traits of these bacteria (Bolwerk et al.
2003).
It was also shown that fusaric acid produced by Fol served as a
chemoattractant stimulating motility of P. fluorescens WCS635 toward and
colonisation of the hyphae of the fungus (de Weert et al. 2004). Colonisation of
hyphae is also required for biocontrol activity of P. putida 06909 against
Phytopthora parasitica (Yang et al. 1994). Ahn and co-workers (2006) further
demonstrated that genes that were up regulated in P. putida 06909 when the
30
bacterium colonised the hyphae of Phytopthora were primarily involved in
carbohydrate metabolism and membrane transport. This suggests that following
lysis of fungal hyphae released nutrients may be used by the bacterium as a
substrate; the term bacterial mycophagy has been suggested to describe this
phenomenon (Ahn et al. 2006; Kamilova et al. 2007).
An accumulation of antibiotic producing fluorescent pseudomonads that occurs in
continuous cropping of wheat and pea plants has been associated with the
development of soil suppressiveness to Take-all (Gaeumannomyces graminis var.
tritici) and Fusarium wilt respectively (Raaijmakers and Weller 1997; de Souza et
al. 2003; Landa et al. 2002).
The role of antibiotic producing fluorescent
pseudomonads in the latter example of “natural” biocontrol has been demonstrated
by the development of suppression to Fusarium following introduction of these
bacteria.
A number of different DAPG genotypes have been identified among fluorescent
pseudomonad populations. Picard et al. (2000) observed that DAPG genotypes
may vary with growth stage of the plant.
Raaijmakers and Weller (2001)
demonstrated that bacterial strains with different DAPG genotypes varied in their
rhizosphere colonisation efficiency and ability to inhibit Fusarium.
Landa and
colleagues (2002) reported that in the pea plant, certain genotypes were able to
maintain threshold levels in the rhizosphere over multiple growing seasons, while
other genotypes typically declined in the RS with crop cycling.
Raaijmakers and
Weller (2001) maintain that further investigation of the genotypic basis for root
colonising ability might have potential for reducing the observed variability in
performance of inoculants of fluorescent pseudomonads in the field. As discussed
by Kloepper and colleagues (2004), difficulties in formulating fluorescent
pseudomonads, due to their sensitivity to desiccation, may have also limited
commercialisation of these bacteria.
Despite this, there are currently three
biopesticide formulations of fluorescent Pseudomonas spp. registered by the USEPA (Stockwell and Stack 2007).
31
2.7.2. Bacillus species.
Bacillus species may produce highly resistant endospores and have therefore
been formulated with relative ease in commercial biopesticide preparations
(Schisler et al. 2004). The ability of certain strains of B. subtilis, B. pumilus, B.
cereus, B. amyloliquefaciens, B. licheniformis, B. simplex, B. firmus and
B. sphaericus to inhibit plant pathogens and promote plant growth has been
reported (Kloepper et al. 2004; Zehnder et al. 2001; Manjula and Podile 2005;
Jacobsen et al. 2004; Gutierrez-Manero et al. 2001; Barneix et al. 2005).
B. subtilis strain A13 was isolated from the lysed mycelium of Sclertonia rolfsii by
Broadbent and colleagues in Australia (1971), and upon inoculation improved
growth of wheat, barley, oats, carrots and several nursery plants (Merriman et al.
1974; Broadbent et al. 1977). This strain is antagonistic to a wide range of fungal
wheat pathogens in vitro and produces moderate amounts of gibberellin
(Broadbent et al. 1977; Broadbent et al. 1971).
B. subtilis A13 was used to
inoculate peanut seeds in the United States, where earlier emergence, increased
root proliferation, enhanced nutritional status and improved plant vigour/robustness
were associated with a reduced incidence of Rhizoctonia solani induced cankers
and increased yields (Turner and Backman 1991).
In this case, there was a
decrease in effectiveness of the treatment over successive crop cycles, where
colonisation was detected in non-inoculated controls and intercropped winter
wheat (Turner and Backman 1991).
B. subtilis A13 was one of the first commercialised biopesticides in the United
States, where it was applied as a seed treatment (along with fungicides) for
suppression of soil-borne fungal pathogens (Zehnder et al. 2001; Backman et al.
1994).
B. subtilis A13 was host passaged in cotton and the derivative, B. subtilis
GB03 is marketed under the trade name Kodiak™. In the USA this formulation of
B. subtilis endospores has been used extensively in cotton crops for the
suppression of Fusarium oxysporum f. sp. vasinfectum (Jacobsen et al. 2004;
Backman et al. 1994). Kodiak™ is also approved by the US-EPA for use in seed
and pod vegetable crops as a biological fungicide seed treatment (US-EPA 2005).
32
This B. subtilis formulation is also used for its growth promoting effects in cotton,
vegetables, small grain, peanut, soybean, and corn (Brannen and Kenny 1997).
Sharma and Jain (1977) reported growth promotion and reduced incidence of Foz
in ginger following the application of B. subtilis strain-1.
The bacterium was
applied to the soil and rhizomes in greenhouse conditions.
Details such as
methods used to identify the bacteria, the number of cells used and the time
course of the experiment were not described.
Thus data on the efficacy of
B. subtilis in promoting growth and resistance to disease in ginger is lacking and
not extensive.
As discussed earlier, eliciting ISR and antibiotic production are mechanisms by
which Bacillus spp. may reduce disease and increase yield in crop production.
Certain strains have been shown to also produce volatiles, lytic enzymes and/or
gibberellins that may be involved in growth promotion or disease resistance
(Schallmey et al. 2004; Jetiyanon and Kloepper 2002; Kloepper et al. 2004; Ryu et
al. 2004).
2.7.3. Phosphate solubilizing bacteria (PSB).
When applied in soluble forms, phosphate is readily fixed and precipitated in soil,
becoming unavailable to plants and accumulating in many agricultural soils (Toro
et al. 1997). Phosphate-solubilizing bacteria (PSB) through their ability to solubilize
applied phosphate and other indigenous organic sources may release phosphate
ions, which might in turn be assimilated by plants or other beneficial microbes (eg.
Mycorrhizal fungi). Strains of B. subtilis, B. megaterium, B. polymxa, B. sphaericus,
B. brevis, B. thuringiensis, Enterobacter spp. and Agrobacterium radiobacter have
demonstrated the in vitro ability to solubilize phosphates, which may be related to
the production of phosphatases or organic acids (de Fretis et al. 1997; Belimov et
al. 1995; Toro et al. 1997; Rodriguez et al. 2006). Groups of PSB were shown to
be stimulated in the rhizosphere in two of four soils amended with compost
(Marcos et al. 1995). Some PSB are reported to be synergistic in co-culture with
nitrogen-fixing bacteria (Belimov et al. 1995; Rojas et al. 2001).
33
2.7.4. Diazotrophic bacteria.
It has been estimated that in worldwide terms, biological nitrogen fixation adds
approximately 175 million tons of nitrogen to the soil each year (Orhan et al. 2006;
Dobereiner 1997). The biological fixation of atmospheric nitrogen (and conversion
into plant available ammonia) is carried out by nitrogenase enzymes of
diazotrophic bacteria (Bashan et al. 2004; Dobereiner 1995; Sprent and Sprent
1990). These nitrogenase enzymes may be inactivated by high levels of NH3,
which may be one reason why effects of inoculation with nitrogen-fixing bacteria
are often more pronounced under reduced levels of nitrogen fertiliser application
(Dobbelaere et al. 2001; Okon and Labandera-Gonzalez 1994; Dobereiner and
Pedrosa 1987; Kennedy and Islam 2001). Examples of free-living diazotrophic
bacteria include Bacillus spp., Azotobacter spp., Herbaspirillum spp., Klebsiella
spp., Azocarcus spp., Acetobacter spp. and Azospirillum spp. (Dobereiner 1995;
Dobereiner 1997; Kennedy et al. 2001; Dobbelaere et al. 2003; Sprent and Sprent
1990; Steenhoudt and Vanderleyden 2000; Kovtunovych et al. 1999; Dong et al.
2003). These bacteria are able to fix nitrogen independently of nodules, in contrast
to Rhizobium species (Kennedy et al. 1997; Sprent and Sprent 1990).
Growth promotion following the introduction of Azospirillum brasilense and
Azospirillum lipoferum in cereal and other plants has been extensively documented
(Okon and Labandera-Gonzalez 1994; Bashan et al. 2004; Dobbelaere et al.
2001).
The increase in plant growth following inoculation of wheat with
microaerophilic Azospirillum species was often linked to the production of plant
growth promoting substances such indole acetic acid rather than nitrogen fixation.
Mutant strains of Azospirillum brasilense deficient in auxin production lost the
ability to induce growth promotion in wheat (Barbieri and Galli 1993). Dobbelaere
and colleagues (1999) demonstrated that an increase in root hair formation in
wheat following inoculation with auxin producing Azospirillum brasilense Sp7,
could be mimicked by the application of indole acetic acid.
In addition,
Azospirillum mutants defective in production of nitrogenase enzymes maintained
the ability to promote wheat growth (Dobbelaere et al. 2003).
Two reasons
associated with the failure to demonstrate nitrogen fixation in mechanisms of plant
growth promotion by Azospirillum species include: i) most species attach to root
34
surfaces, where oxygen may inactivate nitrogenase enzymes and ii) ammonia
derived from nitrogen fixation is not typically excreted by Azospirillum spp. (Rao et
al. 1998; Bashan et al. 2004; Kennedy et al. 1997).
Ammonia excreting Gluconacetobacter diazotrophicus that occupies an endophytic
niche in sugarcane and grasses (where oxygen is more limiting) contributed
significantly to the nitrogen content of sterile sugar cane plants; in addition the
nitrogenase enzymes of this bacterium may be more tolerant to low levels of
oxygen (Cojho et al. 1993; Sevilla et al. 2001; Boddey et al. 2003; Cocking 2005;
Cavalcante and Dobereiner 1988). G. diazotrophicus has also been found as an
endophyte in carrot, beetroot, coffee, radish, pineapple, rice and banana
(Madhaiyan et al. 2004; Herandez et al. 2000; Muthukumarasamy et al. 2002;
Saravanan et al. 2007). Cocking et al. (2006) demonstrated that following the
introduction of low levels of G. diazotrophicus, this bacterium was detected
intracellularlly in Arabidopsis, maize, rice, wheat, tomato and oilseed rape under in
vitro conditions.
In this intracellular location, the bacterium would be ideally
positioned to contribute directly to the nitrogen content of plants. Certain strains of
Gluconacetobacter may also be antagonistic toward Fusarium and other fungal
pathogens (Mehnaz and Lazarovits 2006).
Herbaspirillum spp. are also primarily endophytic diazotrophs, that were first
isolated in sugarcane, maize and sorghum and were later found in other cereals,
banana and pineapple (Baldini et al. 1986; Weber et al. 1999; Cruz et al. 2001;
Baldini et al. 1992; Oliveras et al. 1996).
In 30 day-old rice seedlings, a
Herbaspirillum sp. was shown to contribute 31-54% of plant N by biological fixation
even under high levels of N fertiliser application (Baldini et al. 2000).
Whether directly supplying the plant with biologically fixed nitrogen or indirectly
enabling more efficient use of applied fertilisers by stimulating root growth,
inoculation with diazotrophic bacteria may greatly reduce required inputs of
nitrogenous fertilisers. Up to fifty percent less nitrogen fertiliser input was required
following inoculation of monocots, grains, grasses and a variety of vegetable crops
with Azospirillum species, Herbaspirillum spp. or Gluconacetobacter spp. (Bashan
et al. 2004; Cocking 2005; Kennedy et al. 2004; Okon and Labandera-Gonzalez
35
1994; Steenhoudt and Vanderleyden 2000; Dobbelaere et al. 2001). Reduced
application of inorganic fertilisers may benefit other soil microorganisms, such as
mycorrhizal fungi, which may also interact synergistically with Azospirillum species
(Johansson et al. 2004).
2.7.5. Mycorrhizal helper bacteria.
Mycorrhizal fungi are an important component of the rhizosphere in many plants,
forming an obligate symbiosis and playing a key role in the supply of phosphorus in
plant available forms (Martin 2001; Rilig 2004; Barea et al. 2002). The intraradical
hyphae of arbuscular mycorrhizal fungi (AMF) are able to penetrate root surfaces,
forming “arbuscules” in the cytoplasm of root cortical cells, while the hyphae
external to the root surface (extraradical hyphae) increase the area of soil from
which plants are able to obtain nutrients (Johannson et al. 2004; Finlay 2004).
AMF also play an important role in improving soil structure and decrease the ability
of fungal pathogens to reach the rhizoplane (Johansson et al. 2004; Thomas et al.
1993; Schreiner et al. 1997; Filion et al. 1999; Andrade et al. 1998; Azcon-Aguilar
and Barea 1996). Plant exudation, bacterial populations and nutrient availability in
the rhizosphere can be altered by AMF colonisation (Timonen and Marschner
2006).
Mycorrhiza helper bacteria (MHB) may stimulate colonisation, sporulation and
growth of mycorrhizal fungi, by mechanisms that include production of plant cell
wall degrading enzymes and/or by increasing nutrients available to AMF (Garbaye
1994; Frey-Klett et al. 2007; Bianciotto and Bonfante 2002).
Such beneficial
associations have been reported for nitrogen-fixing bacteria (including Azospirillum
spp.), phosphate solubilizing bacteria and AMF (Budi et al. 1999; Toro 1997; Barea
et al. 2002; Voplin and Kapulnik 1994).
Exopolysaccharides of Azospirillum
brasilense and Rhizobium leguminosarum species were shown to be involved in
the attachment of the bacteria to the hyphae of mycorrhizal fungi (Biancototto et al.
2001).
In cucumber plants, bacteria isolated most often in association with AMF
included Pseudomonas, Arthrobacter, Burkholderia and Paenibacillus (MansfieldGiese 2002).
Schreiner and colleagues (1997) also found Pseudomonas and
36
Arthrobacter spp. associated with AMF in soybean grown in phosphorus deficient
soil.
A Paenibacillus sp. was isolated from the mycorrhizal-rhizosphere of Sorghum
bicolour. The isolate displayed the ability to stimulate mycorrhization and inhibit
fungal pathogens
(Budi et al. 1999).
Bacillus coagulans is reported to be a
mycorrhizal helper bacteria for mulberry and papaya (Mamatha 2002). A
synergistic effect of inoculation with PGPB and AMF in suppression of plant
pathogens has been demonstrated (Siddiqui and Mahmood 1995; Schelkle and
Peterson 1996).
Frey-Klett and colleagues (2007) recently reviewed further
examples of MHB, their in vitro inhibition of phytopathogens and potential for use
as inoculants to promote mycorrhization in crop production systems.
2.8. Combinations of plant growth promoting bacteria.
There is increased interest in using combinations of strains or species that act via
complementary modes of action to improve reliability of PGPB (Jetiyanon and
Kloepper 2002; De Boer et al. 1999; Whipps 2001). Jetiyanon and Kloepper (2002)
aimed to find suitable combinations for eliciting ISR in 4 different host plants with
different diseases in greenhouse conditions.
Four mixtures and one individual
strain were successful (three combinations involving two different B. pumilus
strains and one combination of B. amyloliquifaciens and B. pumilus).
These
combinations were later tested in field trials over two seasons and the combination
of B. amyloliquifaciens and B. pumilus was reported to be successful against
Sclerotium rolfsii infection of tomato, cucumber mosaic virus and Colletotrichum
gloeosporioides disease of long cayenne pepper (Jetiyanon et al. 2003).
A further successful combination of strains was found when B. subtilis GB03 was
combined with B. amyloliquefaciens IN937a.
The two bacteria elicit ISR and
growth promotion via different mechanisms (Ryu et al. 2004; Ryu et al. 2003).
GB03 and IN937a have been formulated, using a chitosan based carrier, to
produce a commercial product (Bioyield™) that is used to promote growth and
resistance to disease in tomato, cucumber, pepper, tobacco in the USA (Raupach
37
and Kloepper 1998; Raupach and Kloepper 2000; Murphy et al. 2003; Anith et al.
2004; Kloepper et al. 2004b; Kokalis-Burelle et al. 2005).
Combination of nitrogen fixing bacteria with phosphate solubilizing bacteria was
reported to have a synergistic effect on the growth of sugar beet, barley and a
variety of vegetable crops (Sahin et al. 2004; Belimov et al. 1995; El-Komy 2004;
Bashan et al. 2004).
2.9. Production and application of bacterial inoculants.
2.9.1. Culture of plant growth promoting bacteria.
Conditions used in the culture of PGPB can have important effects on the
production of compounds relevant to plant growth and disease suppression. For
example indole acetic acid production by Pseudomonas strains and Azospirillum
spp. was found to be increased by addition of L-tryptophan and glucose to the
growth media (Benizri et al. 1998; Thuler et al. 2003; Prinsen et al. 1993). The
production of antibiotic compounds produced by P. fluorescens CHAO, isolated
from a disease suppressive soil in Switzerland, was found to be variable
depending on fermentation time and additives to culture media; DAPG was
stimulated by zinc and ammonium molybdate and glucose; Pyoluteorin was
stimulated by zinc, cobalt and glycerol; Pyrrolnitrin production was increased by
cobalt, fructose, mannitol and a mixture of zinc and ammonium molybdate (Duffy
and Defago 1999). Hamid and colleagues (2003) demonstrated an increase in
nematacidal activity of P. fluorescens CHAO by adding zinc and ammonium
molybdate to the culture medium or soil.
2.9.2. Application of plant growth promoting bacteria and safety aspects.
The application of microorganisms as a seed treatment is extensively reported and
has practical advantages in terms of production and application inputs (Martin and
Bull 2002; Cakemaker et al. 2001; Dobbelaere et al. 2002; Raaijmakers and Weller
38
1998).
Other methods of introducing plant growth promoting bacteria have
included application of inoculants to soil, dipping plant roots into a bacterial
suspension and introduction to the growth medium of tissue culture plants in vitro
(Zehnder et al. 2001; Roncato-Maccari et al. 2003; Jetiyanon et al. 2003; Rapauch
and Kloepper 1998; Njoloma et al. 2006; Kadouri et al. 2003; Munoz-Rojas and
Caballero-Mellado 2003; Akkopru and Demir 2005; Nowak 1998). Critical research
for the optimisation of inoculation methods may improve the reliability of
biofertilisers (Kennedy et al. 2004).
The reapplication of bacteria as a soil drench throughout the growing season or the
use of a carrier based system, may assist in maintaining threshold populations of
bacteria to improve the consistency of the plant growth response to inoculation
(Kloepper et al. 2004; Martin and Bull 2002; Zhang et al. 2004).
Difficulties
associated with the broad acre application of liquid inoculants, as stated by Martin
and Bull (2002) include sanitation of fertigation lines and the large volume of
inoculum required. In addition, the soil is described as a harsh environment and
the form in which the inoculum is applied can greatly determine its effectiveness
(Fravel 2005; Gentry et al. 2004; Zohar-Perez et al. 2005).
In greenhouse experiments using Serratia marcescens, a dramatic difference in
the ability to suppress bean disease was observed depending on whether a liquid
preparation (10% disease reduction) or alginate beads (40-60% disease reduction)
were used to apply the bacterium (Zohar-Perez et al. 2005).
Formulation of
Azospirillum spp. as dried alginate beads may increase survival of the bacterium
when it is applied to the soil (Fages 1992; Bashan 1986a). This may result from
the ability of such carrier materials to protect microorganisms from mechanical
damage, UV light and predation from other organisms (Zohar-Perez et al. 2005).
Alginate is a non-toxic, biodegradable and naturally occurring polymer extracted
from kelp.
Formulation of bacterial cells into alginate beads allows for the
continual release of the bacteria as the carrier degrades, which can reduce
required application rates of applied bacteria (Bashan 1998).
Bashan and
Gonzalez (1999) demonstrated that cells of A. brasilense encapsulated in dried
alginate beads remained viable for over fourteen years. Thus, when compared to
the use of a liquid suspension of bacteria that must be applied within hours of
39
preparation to maintain viability, the use of alginate beads provides a more
practical method of production and application.
Further, the formulation of
microorganisms into a stable form may provide opportunity to test inoculants for
purity and dry products may be less susceptible to contamination (Fravel 2005).
The use of carrier materials may also allow for containment of the microorganisms
to prevent their dissipation to the atmosphere and ground water (Gentry et al.
2004; Zohar-Perez et al. 2005). The addition of chitin, chitosan or humic acids has
further improved the efficiency of some bacterial formulations (Bashan et al. 2002;
Zehnder et al. 2001; Young et al. 2006).
The US-EPA has established production and application requirements for nonpathogenic biopesticides such that risks to human health and the environment are
expected to be mitigated (US-EPA 2005).
This has included requirements for
ensuring the absence of indicator pathogens in inoculants by culture based
methods; RAPD analyses for strain verification; the use of personal protective
equipment during production and application of micro-organisms; disposal and
application guidelines; and toxicology testing to demonstrate that non-target effects
do not occur in fish, bird and insect species. Thus the use of inoculants of PGPB
containing non-pathogenic microorganisms found naturally in soils and on plant
roots is not associated with adverse environmental or human health impacts.
Although, as discussed by Miller and Aplet (1993), a microorganism should be
considered natural within the context of the ecological niche in which it is found in
nature. Additional safety aspects potentially associated with the use of biocontrol
bacteria include displacement of, or toxigenicity or pathogenicity toward non-target
microorganisms (Cook et al. 1996).
Conn and Franco (2004) demonstrated the
utility of molecular methods (TRFLP) for documenting the effect of a microbial
inoculant on endophytic actinomycetes populations in wheat. Such methods may
be implemented to ensure that soil biodiversity is not compromised following the
application of inoculants. The establishment of many introduced microorganisms in
soil has proven to be difficult, where populations typically decline with time and
distance from the point of introduction (Whipps 1999; Jacoud et al. 1998; Smith et
al. 1984). As such often transitory (and minor) effects of inoculants on the
indigenous soil microflora have been documented (Cook et al. 1996; Girlanda et al.
2001; Thirup et al. 2001; Castro-Sowinski et al. 2007; Herschkovitz et al. 2005).
40
This is in contrast to the longer-term alterations in soil microbiota that may result
from cultivation and intensive agricultural practices such as tillage and pesticide
application (Cook et al. 1996; Garbeva et al. 2004; Wardle 1995; Buckley and
Schmidt 2001; Berkelmans et al. 2002).
2.10. Conclusion
Deleterious impacts of intensive agricultural practices on soil microorganisms may
result in degraded soils, increased disease pressures, reduced yields and a
decline in the nutritional content of food. In contrast, the agricultural application of
non-pathogenic plant growth promoting bacteria, found naturally in the rhizosphere
and within plant roots or tissues, is not associated with adverse human health or
environmental impacts.
A plethora of literature has documented the use and modes of action of PGPB,
particularly Bacillus spp., Azospirillum spp. and Pseudomonas spp., which improve
growth and resistance to disease and/or reduce fertiliser requirements in many
agronomically important crops. However, there is a scarcity of literature describing
the application of plant-beneficial bacteria to ginger, and examples in subtropical
regional conditions were not found.
Despite an abundance of research that has established the paradigm for the
implementation of PGPB in agriculture, variable performance of inoculants in field
conditions is considered to have limited their commercial viability in many
instances.
Optimisation of application methods and the use of combinations of
bacterial strains are suggested as key areas for improved consistency of bacterial
inoculants in field conditions (Kennedy et al. 2004; Kloepper et al. 2004).
An
improved understanding of plant dependency on symbiotic, rhizospheric and
endophytic bacteria and environmental influences may also allow for more
effective PGPB inoculation strategies to be developed (Martin and Bull 2002;
Johansson and Finlay 2004; Ross et al. 2002; Akkopru and Demir 2005; CastroSowinski et al. 2007; Duffy and Defago 1999).
Research that addresses the
aforementioned topics may enable inoculants of PGPB to be effectively
implemented to assist in the management of plant health in integrated approaches
41
to sustainable agriculture and could be of significant value where no means of
disease control exists, as for example in Fusarium yellows of ginger (Jetiyanon et
al. 2003; Fravel 2005; Bashan et al. 2004; Atkinson and Watson 2000; Sturz and
Nowak 2000; Cook 2000).
42
Chapter 3. Assessment of compost tea and commercial microbial inoculants
as a source of beneficial microorganisms for improved growth of ginger.
3.1. Introduction
Use of compost tea as a source of beneficial organisms for improved soil health,
plant nutrition and suppression of plant pathogens in agriculture (and the home
garden) has increased in recent times (CTTFR 2004; Scheurell and Mahaffee 2002;
Ingram and Millner 2007). Compost tea is produced by mixing compost with water
and encouraging the growth of the extracted microflora by the addition of substrates
such as sugars (eg. molasses) and proteins (Ingham 2004). Additives such as
guano, kelp, humic acids and rock dusts may be incorporated for their potential
benefits on plant growth and to promote fermentation.
The addition of growth
substrates and additives may alter the efficacy of compost teas in suppressing plant
pathogens.
For example, Scheurell and Mahaffee (2004) demonstrated that
compost tea produced with humic acids and kelp, but not with molasses, suppressed
Pythium ultimum in cucumber. Air is actively circulated through such preparations,
typically in open containers, over 24-36 hours to produce aerated compost tea.
Non-aerated compost tea may be left to stand for at least three days (Ingham 2004).
Hundreds of litres per hectare are applied to crops on a weekly to fortnightly basis,
via foliar sprays or soil drenches (Ingram 2004; Sturz et al. 2006).
Reports in peer-reviewed literature have described the use of compost tea for the
suppression of plant pathogens, although inconsistent results have often been
reported (Scheuerell and Mahaffee 2002; Litterick and Harrier 2004; Haggag and
Saber 2007; Scheuerell and Mahaffee 2004). Grower testimonials constitute the
majority of evidence that supports the use of compost teas (CTTFR 2004). Compost
teas are purported to contain billions of beneficial bacteria and fungi and are typically
assessed by counting the number of active cells per ml or by calculating biomass.
Analyses that document the actual type of bacteria of fungi present in compost teas
are lacking.
Compost, a source of microorganisms in these teas, is typically
produced by thermophilic composting of plant and/or animal waste. Thermophilic
43
composting requires the maintenance of aerobic conditions and that a temperature
of 57 oC be reached for at least 3 days throughout the pile (Ingham 2004). While
composting is a “process to significantly reduce pathogens” Christensen and
colleagues (2002) reported that Enterococcus faecalis and Escherichia coli were
detected in finished compost. Klebsiella pneumoniae, Clostridium botulinum,
Pseudomonas aeruginosa, Enterobacter spp. and Legionella spp. have also been
isolated from composts (Bohnel and Lube 2000; Droffner et al. 1995; Hassen et al.
2001; Lasaridi et al. 2006; Hughes and Steele 1994; Boutler et al. 2002). Data on
the survival of pathogenic organisms in cooler zones of the compost pile is also
lacking (Christensen et al. 2002).
Thus enteric and other human pathogenic
bacteria may be present in composted materials, which are used to prepare compost
teas.
As a means of quality control, levels of indicator organisms are typically monitored in
order to determine whether conditions that enable transmission, survival or growth of
pathogens have occurred (Hocking 2002). While E. coli and Salmonella species
have frequently been monitored as indicators of enteric contamination, enterococci
are increasingly preferred as indicator organisms due to their longer persistence in
the environment that is typical of more resistant pathogens (US-EPA 2002;
Kinzelman et al. 2003). The presence of indicator pathogens can predict a larger
population of pathogens (Sidhu et al. 1999).
The maintenance of aerobic conditions (aerated tea) and heterotrophic diversity is
purported to prevent growth of human pathogenic microorganisms in compost tea
(Ingham 2004).
The growth of pathogenic and phytotoxin producing species may
be expected following the formation of anaerobic conditions, for example due to the
addition of too much growth substrate or insufficient aeration of the brew and is
detected by a foul smelling tea (Ingham 2004; Ingham 1998). A study by Duffy and
colleagues (2004) demonstrated that E. coli 0157 and Salmonella enterica grew in
compost tea when molasses concentrations greater than 0.2% were used. The
Agricultural Research Service (USA) reported similar findings, where the growth of
44
pathogenic indicator organisms in the presence of soluble carbon additives was not
prevented by heterotrophic diversity in aerated teas (Ingram et al. 2005).
The USA National Organic Standards Board (NOSB) recommended quality
assurance testing to demonstrate that a specific production system can generate
compost tea that meets microbial quality guidelines of 126 colony forming units
(CFU) E. coli /100ml or 33 CFU enterococci /100ml. A 90/120 day (winter/summer)
withholding period for compost teas made with additives was introduced. The foliar
application of manure extract or tea and of compost leachate was prohibited and
otherwise restricted to a 90/120 day withholding period (CTTFR 2004).
Canadian
government authorities recommended similar withholding periods and that compost
tea is not applied to edible portions of plants (Ministry of Agriculture and Lands,
MOAL 2005).
The NOSB recognised an urgent need to evaluate the potential for compost tea to
contaminate crops with food and water borne pathogens of concern, such as
E. coli 0157, Salmonella spp., Cryptosporidium parvum, Giardia Lamblia, Ascaris,
Klebsiella species, Staphylococcus spp., Proteus spp., Enterobacter spp.,
Clostridium perfringens and Burkholderia spp. (CTTFR 2004).
An increasing
incidence of food borne disease worldwide and in Australia is increasingly
associated with domestic and imported fruits and vegetables, where untreated or
improperly treated manure and unsanitary irrigation water are implicated as sources
of contaminants (Beuchat and Ryu 1997; Hocking 2002; Aruscavage et al. 2006;
Solomon et al. 2002; Solomon et al. 2006; Buck et al. 2003; US-FDA 1998).
The objective of this part of the study was to assess the microbial nature of these
popularised compost teas to be potentially used in further research for application to
ginger crops for improved productivity. Locally produced samples were first analysed
to determine whether human pathogens could be detected and whether such brews
met microbial guidelines proposed by the NOSB and water quality standards set by
the Queensland Environmental Protection Agency for the use of recycled water in
45
agriculture (QLD-EPA 2005).
A further aim included analysis of the types of
beneficial bacteria present in compost tea. Following the detection of enteric bacteria
in aerated compost tea, further analyses were undertaken to assess untreated
source materials (liquefied compost and additives) used to prepare the compost tea
and these materials cultured with aseptic technique. Subsequently commercially
available mixed microbial inoculants (purported to contain non-pathogenic
organisms) fermented similarly to compost tea and a range of different growth
substrates were evaluated.
3.2. Materials and Methods
3.2.1a. Isolation of bacteria from compost teas and commercial microbial
inoculants.
Selective or chromogenic media used to isolate bacteria from compost tea and
commercial microbial inoculants are described in Table 3. Ten fold serial dilutions of
the following materials were performed in sterile phosphate buffered saline (PBS:
137 mM NaCl, 2.7 mM KCl, 10 mM Na2HPO4, 2mM KH2PO4, pH 7.4) for plating on
selective media: (i) untreated compost tea starting materials (commercially available
liquefied compost, growth substrates and additives) and commercially available
mixed microbial inoculants (two batches of two different inoculants); (ii) The
aforementioned materials cultured overnight under laboratory conditions in sterile
media (heart serum infusion, HSI) using aseptic technique; (iii) Aerated compost
teas or microbial brews (utilising commercial mixed microbial inocula), produced in
an open air container with turbulent aeration for 24 hours. Three aerated microbial
brews produced on independent occasions were analysed; the dissolved oxygen
content of the second brew was confirmed to be >6ppm, measured hourly for 24
hours. Further description of the components and methods used in the preparation
of different microbial cultures is listed in Tables 4 and 5.
In laboratory-based
analyses, plates and cultures were incubated at 37oC for 24-48 hours.
46
Table 3. Media used for the isolation of bacteria from compost teas and commercial
microbial inoculants.
Media
Composition of Media
Target Bacteria
MacConkey No.3#
Oxoid premix (Australia).
Faecal coliforms; E. coli and
Enterobacter appear pink to red.
Colourless colonies are produced
by Shigella flexneri.
E. coli chromogenic
media
Oxoid premix (Australia).
E. coli turns bluish/purple.
E. coli 0157 selective
media
Biomerieux premix (France).
E. coli 0157.
Bile esculin agar#
Per litre: 20g oxall, agar 15g,
pancreatic digest of gelatin
5g, beef extract 3g, esculin
1g, ferric citrate 0.5g, horse
serum 50ml, pH 6.8
Presumptive identification of
enterococci based on esculin
hydrolysis, that turns the media
black, at 44oC.
Mannitol Salt agar#
Per litre: 1g beef extract, 10g
peptone, 75g NaCl, 10g dMannitol, 0.025g phenol red,
15g agar; pH 7.4
Fermentation of mannitol by
pathogenic Staphylococcus spp.,
such as S. aureus turns the media
from red to yellow.
UriSelect 4
Biorad prepared agar plates
Differentiation of common urinary
pathogens E. coli, Enterococcus
faecalis, Klebsiella pneumoniae,
Proteus and Staphylococcus
based on colony colour*
SS Agar
Biorad prepared agar plates
Salmonella enteritidis and Shigella
spp.**
*On UriSelect4 E. coli and E. coli 0157 produce pink colonies indicative of betagalactosidase activity; Enterococcus faecalis colonies turn blue due to beta-glucosidase
activity; Klebsiella/Enterobacter/Serratia appear as violet colonies, although strains with
weak beta-galactosidase activity may be turquoise; Bacterial colonies that turn orange
possess tryptophan deaminase activity and those that do not exhibit a colour change on
addition of Kovacs reagent are potential Proteus mirabilis spp. (Biorad™ UriSelect 4 pack
insert).
** On this medium Salmonella enteriditis produces colonies with a black centre; Shigella
sonnei and Enterobacter aerogenes appear pink to red; Shigella flexneri develop a light pink
colour (Biorad ™ SS-Agar pack insert).
#
Atlas 1993.
47
Table 4. Label description of commercially available microbial inoculants and
substrates used in this study.
Commercial Product
Product Components
Microbial inoculant 1
Bacillus subtilis, Bacillus megaterium, Azotobacter
vinelandii, Pseudomonas fluorescens, Pseudomonas
putida, Rhizobium japonicum, Pseudomonas stutzeri,
Rhizobium leguminosarum, Streptomyces albidoflavus,
Streptomyces cellulosae, Chaetomium globosum,
Saccharomyces cerevisiae, Trichoderma lignorum,
Trichoderma harzianum, Gliocladium virens
Microbial inoculant 2
Trichoderma lignorum, Trichoderma harzianum,
Gliocladium virens, Bacillus subtilis.
Liquefied compost
Micronised, liquefied and stabilised thermophilic
compost.
Substrate 1
Soybean meal.
Substrate 2
Various amino acids, fatty acids, carbohydrates,
minerals and plant extracts.
Substrate 3
Autoclaved growth substrate.
Additive 1 (also used as a
substrate)
Aloe vera, fulvic acid, humic acid, kelp, fish emulsion,
fish oil, liquid vermicast, and various minerals.
Additive 2
Liquefied fish solids, kelp, fulvic acid and triacontanol.
48
Table 5. Conditions used to ferment microbial cultures.
Type of culture
Culture components
Culture conditions
Laboratory culture
of liquefied compost
(compost*)
Aseptic technique was
used to prepare the culture.
Flasks were incubated at
37oC overnight with
shaking.
Aerated compost
tea 2 (ACT2)
The liquefied compost was
diluted 1:5 with sterile PBS.
25ml of sterile heart serum
infusion (HIS) was inoculated
with 200uL of the diluted
material.
The material was diluted 1:5 with
sterile PBS. 25ml of sterile heart
serum infusion (HIS) was
inoculated with 200uL of the
diluted material.
200L water*, 2L liquefied
compost, 2L additive 1, 1kg
humus, 1kg leaf mulch, 2L
vermicast.
10L water, 100ml liquefied
compost, 100ml additive 1.
Aerated microbial
brew 1 (AMB1)
10L water, 12.5 g microbial
inoculant 1, 75g substrate 1.
Aerated microbial
brew 2 (AMB2)
10L water, 12.5 g microbial
inoculant 1, 100ml substrate 2,
50g sugar.
Aerated microbial
brew 3 (AMB3)
15L water, 12.5g microbial
inoculant1, 150ml substrate 2.
Aerated for 24 hours in a
20L container, dissolved
oxygen measured hourly
(>6ppm).
Aerated for 24 hours in a
20L container, dissolved
oxygen measured hourly
(>6ppm).
Aerated for 24 hours in a
20L container, dissolved
oxygen measured hourly
(>6ppm).
Aerated for 24 hours in a
20L container.
Aerated microbial
brew 4 (AMB4)
15L water, 12.5g microbial
inoculant1, 150ml additive 2.
Aerated for 24 hours in a
20L container.
Aerated microbial
brew 5 (AMB5)
15L water, 12.5g microbial
inoculant1, 150ml substrate 3.
Aerated for 24 hours in a
20L container.
Aerated microbial
brew 6 (AMB6)
15L water, 15g microbial
inoculant 2, 150ml substrate 3.
Aerated for 24 hours in a
20L container.
Laboratory culture
of additive 1
Aerated compost
tea 1 (ACT1)
Aseptic technique was
used to prepare the culture.
Flasks were incubated at
37oC overnight with
shaking.
Aerated for 24 hours.
*Tap water used in aerated brews was allowed to dechlorinate for at least one hour
prior to the addition of brew components.
49
3.2.1.b. Phylogenetic analysis and biochemical testing of selected bacterial
isolates.
Putative isolates of Enterococcus spp. (blue colonies recovered on UriSelect4 and
red colonies from m-Enterococcus selective medium) were transferred to bile esculin
plates and incubated at 44oC for 1 hour, 2 hours or overnight (Atlas 1993).
For phylogenetic analysis of selected bacteria partial sequencing of the 16S rRNA
gene was undertaken (Weisberg et al. 1991). Ten representative bacteria, isolated
from the untreated commercial microbial inoculants, were purified by streaking on
Uriselect4 or nutrient agar at least 3 times. Total genomic DNA was extracted from
a single colony using the ChargeSwitch gDNA Mini Bacteria Kit (Invitrogen,
California) according to manufacture’s instructions.
The 16S rRNA gene was
amplified by the polymerase chain reaction (PCR) using primers 518F and 1513R
(Appendix 3.1). The 50µL PCR reaction included 1X PCR Supermix (20mM TrisHCl, 5mM KCL, 1.5mM, 200µM of each dNTP, 1 U Taq DNA polymerase; Invitrogen,
California), 0.5µM of each primer, 2.5 µL genomic DNA and an additional 0.5mM
MgCl2 to produce a final concentration of 2mM MgCl2. The PCR reaction was
performed with an Eppendorf Mastercycler Thermal Cycler. Cycling conditions
included an initial denaturation step at 94 oC for 4 min; 25 cycles of 94 oC for 30 sec,
55 oC for 30 sec, 72 oC for 90 sec; and a final extension at
72 oC for 10 min. PCR
products were visualised on a 1.5% agarose gel stained with ethidium bromide and
purified using a QIAquick PCR Purification Kit (QIAGEN, Clifton Hill, Australia).
Primers 518F and 926R were used to sequence both strands of DNA using ABI
PRISM Big Dye Terminator Sequencing Chemistry (Version 3.1) (Australian
Genome Research Facility, Brisbane, Australia). Sequences were visualised and
edited using the program Finch TV (GeoSpiza).
A consensus sequence was
determined
by
following
alignment
of
sequences
the
ClustalW
program
(http://www.ebi.ac.uk/clustalw/). The Basic Local Alignment Search Tool (BLASTn)
at
the
National
Centre
for
Biotechnology
Information
(http://www.ncbi.nml.nih.gov/BLAST) was used to compare ~500bp of DNA
50
sequence to those in GenBank.
Sequences were additionally analysed using
Ribosomal Database Project (RDP-II) programs SeqMatch, to determine nearest
neighbours and Classifier, for assignment of taxonomic hierarchy (Michigan State
University, East Lansing, Michigan; Maidak et al. 2000).
Twenty representative bacteria isolated on UriSelect4 from aerated compost tea and
microbial brews (where dissolved oxygen content was measured hourly) were
purified as described above.
Templates for 16S rDNA PCR were prepared as
described by Raaijmakers and Weller (1997). Briefly two bacterial colonies were
resuspended in 100 µL of lysis solution (0.05M NaOH, 0.25% SDS) and heated to
100oC for 15 minutes. The suspension was centrifuged at high speed for 1 minute.
The supernatant was diluted 1:50 in sterile milliQ water for use in the 16S rDNA
PCR as described earlier. The PCR product was purified and sequenced using
primer 926R using ABI PRISM Big Dye Terminator Sequencing Chemistry (Version
3.1) with an ABI3730XL by Macrogen Inc (Seoul, Korea). Sequence analysis was
performed as described earlier.
Biochemical testing was undertaken for confirmation of 16S rDNA sequencing
results. Fifteen biochemical reactions were performed using Enterotube II (Becton
Dickson) according to manufacture’s instructions.
Independent microbiological
analyses of the two mixed microbial inoculants (two batches) were also performed
by two different commercial laboratories in Queensland.
3.2.2.
Risk
assessment
for
exposure
to
pathogenic
organisms
in
contaminated cultures.
In accordance with guidelines of the Queensland “Workplace Health and Safety Risk
Management Advisory Standard 2000” (QLD-WHSRMAS 2000), the risk level
associated with exposure to pathogenic organisms that could be present in
contaminated microbial fermentations had to be ascertained.
Risk assessments
were performed using HAZNET software of the National Safety Council of Australia.
51
In undertaking the risk assessment process, the likelihood that an event could occur
and the seriousness of consequences was taken into account in determining the risk
level associated with a relevant activity/process. According to the Workplace Health
and
Safety
Risk
Management
Advisory
Standard
2000,
permissible
activities/processes are those where a hazardous situation is not likely to be
encountered and are associated with a low level of risk. A moderate level of risk
requires correction, but is not an emergency.
A high or very high-risk
activity/process must be ceased immediately and is described as an urgent situation.
A hierarchal method for reducing the risk level associated with an activity/process is
required and includes elimination, substitution, isolation, minimisation, administration
and use of personal protective equipment, in order of effectiveness.
Risk was assessed for exposure to pathogenic organisms (Klebsiella pneumoniae or
Enterobacteriaceae) whilst sampling the microbial fermentations due to formation of
aerosols produced by the turbulent aeration of the brews and contact with the brews.
Risk assessments for the spray application of Class 2 (capable of causing human
disease) and Class1 (do not usually cause human disease) biological organisms
were also undertaken.
The effect of implementation of personal protective
equipment and containment of fermentations in bioreactors on the risk level of
relevant activities was determined.
3.3. Results
3.3.1.a.
Isolation bacteria from compost teas and commercially available
mixed microbial inoculants.
Colonies typical of faecal coliforms and Staphylococcus aureus were isolated from
aerated compost tea on MacConkey No.3 and mannitol salt agar respectively
(Table 6, Figure 1).
When the untreated liquefied compost material (used for
preparation of the tea) was analysed colonies typical of faecal coliforms, S. aureus
52
Klebsiella/Enterobacter/Serratia and Enterococcus faecalis were produced (Table 7,
Figure 2). Bacteria isolated from a tea additive (additive 1) were typical of S. aureus
and E. faecalis (Table 7, Figure 2). Similar bacteria were also isolated from liquefied
compost and additive 1 which were cultured in laboratory media using aseptic
technique (Table 6, Figure 3). Out of the forty blue colonies isolated on UriSelect 4
from laboratory cultures of liquefied compost that were transferred onto bile esculin
agar, only 24 produced a black halo after overnight incubation (Figure 4). Isolates
that produced a black halo were putative enterococci. From Additive 1 cultured
under laboratory conditions, 48 colonies were picked and 45 produced a black halo
(94%).
Next, in factory-produced cultures, where dissolved oxygen was confirmed hourly to
be greater than 6ppm (purported to inhibit the growth of pathogenic organisms),
colonies typical of indicator pathogens (K/E/S group and enterococci) were still
isolated from cultures prepared with either compost or microbial inoculants and
various substrates (ACT2, AMB1, AMB2; Table 6, Figure 5).
A number of
commercially available growth substrates were then identified in which enteric
contaminants were not detected (Substrate 2, Substrate 3, and Additive 2).
Pathogenic indicators were still isolated from factory-produced cultures using these
substrates/additives along with commercial microbial inoculants (AMB3-6; Table 6,
Figure 6). Further analyses suggested that the contaminants were present in the
untreated microbial inoculants (Table 7).
Figure 1. Bacteria isolated from isolated from aerated compost tea (ACT 1) on a)
MacConkey No.3 (faecal coliforms) and b) Mannitol salt agar (Staphylococcus spp).
a
b
53
b
S-S Agar (pink to red
colonies: S.sonnei
/Enterobacter)
S-S Agar (light pink
colonies: S.flexneri)
UriSelect4 white
colonies
UriSelect4 orange
colonies
UriSelect4 purple
colonies
(likely K/E/S group)
UriSelect4 blue
colonies
E. coli chromogenic
media
Mannitol salt agar
(Staphylococcus)
MacConkey #3 (Faecal
coliforms)
Nutrient agar
(culturable bacteria)
Microbial Culture
Table 6. Microbiological analysis of fermented materials.
compost* 7.3 x 107 1.7 x 107 9.1 x 106 3.8 x 102 3.7 x 107 7.0 x 105 >1 X 106 1.0 X 106
ACT1
2.7 x 109 1.5 x 106 3.8 x 103
ACT2
2.6 x 106 5 x104
AMB1
>1 x 106 >1 x 106 >1 x 106 >1 x 106
AMB2
5 x 103
5 x 102
AMB3
1 x 104
1.5 x 103
1 x 104
AMB4
1 x 104
65
1.5 x 104
AMB5
5 x 104
5 x 104
1 x 104
1 x 102
2 x 105
2.5 x 104 8 x 103
4 x 102
>1 x 106
AMB6
Water**
3 x 105
N/D
N/D
N/D
N/D
*Liquefied compost material cultured under laboratory conditions
** Water that was aerated for 24 hours (without additives)
ACT aerated compost tea; AMB aerated microbial brew; N/D not detected; all values
represent CFU/ml.
ACT1: Liquefied compost, Additive 1, humus, leaf mulch, vermicast.
ACT2: Liquefied compost, Additive 1; AMB1: Microbial inoculant 1, substrate 1.
AMB2: Microbial inoculant 1, substrate 2, sugar.
AMB3: Microbial inoculant1, substrate 2. AMB4: Microbial inoculant1, Additive 2.
AMB5: Microbial inoculant1, substrate 3.
AMB6: Microbial inoculant 2, substrate 3.
Additive1: Aloe vera, fulvic acid, humic acid, kelp, fish emulsion, fish oil, liquid
vermicast, and various minerals; Additive 2: Fish solids, kelp, fulvic acid and
triacontanol; Substrate 1: soybean meal; Substrate 2: Various amino acids, fatty
acids, carbohydrates, minerals and plant extracts; Substrate 3: autoclaved
substrate. Microbial inoculant 1: twenty different bacteria and four fungi; Microbial
inoculant 2: three Trichoderma strains and B. subtilis.
54
>1 x 105
UriSelect4 white colonies
(potential Staphylococcus
spp.)
1.4 x 105
3.7 x 107 7 x 105
>1X106
>1 X 106
5 x 106
1 X 106
>1 X 106
N/D
N/D
UriSelect4 blue colonies
E. coli chromogenic media
Mannitol salt agar
(Staphylococcus spp.)
MacConkey #3 (faecal
coliforms)
4.4 x 108 N/D
UriSelect4 orange colonies
(potential Proteus spp.)
Additive 1
1.4 x 105 >1 x 105 N/D
UriSelect4 purple colonies
(potential K/E/S* group)
Liquefied
compost
Nutrient agar (culturable
bacteria)
Material
Table 7. Microbiological analysis of untreated materials.
Additive 2
N/D
N/D
Substrate 1
1.5 x 103 2.5 x 102 5 x 102
1 x 102
Substrate 2
N/D
N/D
N/D
N/D
Substrate 3
N/D
N/D
N/D
N/D
Microbial
1.5 x 106
1.8 x 106
Inoculum 1
Microbial
1.3 x 106 2.2 x 107 2.3x 107 2.8 x 107
Inoculum 2
N/D not detected; all values represent CFU/ml, except microbial inocula 1 and 2,
where count represents CFU/g.
* K/E/S: Klebsiella/Enterobacter/Serratia group.
Additive1: Aloe vera, fulvic acid, humic acid, kelp, fish emulsion, fish oil, liquid
vermicast, and various minerals (also used as a growth substrate); Additive 2
Liquefied fish solids, kelp, fulvic acid and triacontanol; Substrate 1: soybean meal;
Substrate 2: Various amino acids, fatty acids, carbohydrates, minerals and plant
extracts; Substrate 3: autoclaved substrate.
Microbial Inoculant 1: Bacillus subtilis, Bacillus megaterium, Azotobacter vinelandii,
Pseudomonas fluorescens, Pseudomonas putida, Rhizobium japonicum,
Pseudomonas stutzeri, Rhizobium leguminosarum, Streptomyces albidoflavus,
Streptomyces cellulosae, Chaetomium globosum, Saccharomyces cerevisiae,
Trichoderma lignorum, Trichoderma harzianum, Gliocladium virens.
Microbial Inoculant 2: Trichoderma lignorum, Trichoderma harzianum, Gliocladium
virens, Bacillus subtilis.
55
Figure 2. Isolation of bacteria from untreated materials used to prepare
microbial cultures. a) Additive 1 plated on UriSelect4; b) Additive 1 plated on
mannitol salt agar; c) Liquefied compost plated on MacConkey No. 3; d) Liquefied
compost plated on mannitol salt agar.
a
b
c
d
56
Figure 3. Laboratory culture of Additive 1 and liquefied compost. a) Additive 1
plated on Uri Select4; b) Liquefied Compost plated on Uri Select4; c) Liquefied
Compost (10-4 dilution) plated on MacConkey No.3; d) Liquefied Compost (10-5
dilution) plated on MacConkey No.3.
a
b
Liquefied
Compost
Additive1
10-3, 10-4,10-
Liquefied
Compost
10-3, 10-4, 10-5
Liquefied
Compost
c
57
d
Figure 4. Demonstration of esculin hydrolysis ability of putative enterococci on bile
esculin agar at 44 oC.
58
Figure 5. Bacteria isolated from microbial cultures produced with 6ppm-dissolved
oxygen. a) ACT2 (Liquefied compost + Additive 1) plated on UriSelect4; b) ACT2
(Liquefied compost + Additive 1) plated on SS-Agar; c) AMB1 (Microbial Inoculant 1
+ Substrate 1) plated on UriSelect4; d) AMB1 (Microbial Inoculant 1 + Substrate 1)
plated on SS-Agar; e) AMB2 (Microbial Inoculant 1 + Substrate 2 + sugar) plated on
UriSelect4; f) AMB2 (Microbial Inoculant 1 + Substrate 2 + sugar) plated on SSAgar.
a
b
c
d
e
f
59
Figure 6. Isolation of bacteria on UriSelect4 from microbial inoculants after
overnight culture with aeration and various additives. a) Control treatments including
water aerated overnight and dilution buffer; b) AMB3 (Microbial Inoculant +
Substrate 2); c) AMB5 (Microbial Inoculant 1 + Substrate 3); d) AMB6 (Microbial
Inoculant 2 + Substrate 3).
a
b
c
d
60
3.3.2a. Phylogenetic analysis and biochemical testing of selected bacterial
isolates.
Extraction of genomic DNA from bacterial isolates from the microbial inoculants
produced a single clean band on an agarose gel (Figure 7a). Specificity of 16S
rDNA PCR was demonstrated by the production of a single band on an agarose gel
and the absence of a band in the negative control (Figure 7b). The chromatogram
produced by sequencing of PCR products consisted of clear and distinct peaks with
few ambiguous bases (Figure 8).
Completely homologous sequences were
produced following alignment of DNA sequences of sense and (reversecomplemented) anti-sense strands (Appendix 3.2-3.3). Phylogenetic analysis based
on 16S rDNA sequences indicated that 9 out of 10 bacteria isolated on UriSelect 4
media from untreated microbial inocula belonged to the Enterobacteriaceae family
(100% confidence according to the RDP-Classifier program): the majority of isolates
were placed in the Klebsiella pneumoniae/Enterobacter cloacae/Enterobacter
dissolvens group; two isolates were most closely related to Pantoea spp.; one
isolate displayed a high homology with Bacillus pumilus (Table 8).
Biochemical
testing of isolates using Enterotube II generally verified 16S rDNA sequencing
identification, confirming the presence of Klebsiella pneumoniae and Enterobacter
cloacae in untreated mixed microbial inoculants (Table 9, Figure 9).
Independent testing by a commercial laboratory confirmed Enterobacteriaceae
levels of greater than 2000 CFU/gram were present in untreated microbial
inoculants. API testing undertaken by a second commercial laboratory indicated
Pseudomonas
aeruginosa,
Photobacterium
damsela,
Bacillus
cereus
and
Stenotrophomonas maltophilia were also present in the untreated commercial
microbial inoculants.
16S rDNA sequencing of bacterial isolates recovered on Uri Select 4, from an
aerated microbial fermentation (AMB1) indicated that species most closely related to
61
Pseudomonas aeruginosa, Bacillus cereus, Brevundimonas spp., Pseudomonas
stutzeri, Bacillus pumilus and Bacillus subtilis were present in dominant populations.
a
b
Figure 7. a) Genomic DNA extracted from bacteria isolated from microbial
inoculants, 1kb Step Ladder (Promega). b) 16S rDNA PCR products, Lambda-Hind
III DNA Marker (Promega).
Figure 8. Typical sequence chromatogram obtained following sequencing of 16S
rDNA PCR products, visualised using the FinchTV program.
62
Table 8. Taxonomic assignment of bacterial isolates from microbial inoculants using
Ribosomal Database – II (RDP-II).**
Isolate Family*
Code
Genus*
Most Closely Related Species# S_ab scorey
Bacillus (92%)
Bacillus pumilus
4
Bacillaceae (92%)
1
7
Enterobacteriaceae (100%) Klebsiella (49%)
Enterobacter hormaechei,
Enterobacter cloacae, Pantoea
agglomerans
8
Enterobacteriaceae (100%) Klebsiella (84%)
Klebsiella pneumoniae,
Enterobacter cloacae,
Enterobacter sakazaki
0.982-1.000
10
Enterobacteriaceae (100%) Klebsiella (86%)
Klebsiella pneumoniae,
Enterobacter cloacae
0.987-1.000
12
Enterobacteriaceae (100%) Klebsiella (90%)
Klebsiella pneumoniae,
Enterobacter cloacae
0.984-1.000
13
Enterobacteriaceae (100%) Klebsiella (58%)
Klebsiella pneumoniae,
Enterobacter cloacae,
Enterobacter sakazaki
0.984-1.000
14
Enterobacteriaceae (100%) Klebsiella (69%)
Klebsiella pneumoniae,
Enterobacter cloacae
0.974-1.000
16
Enterobacteriaceae (100%) Klebsiella (63%)
Klebsiella pneumoniae,
Enterobacter cloacae, Kluyerva
intermedia
1.000
17
Enterobacteriaceae (100%) Pantoea (84%)
Pantoea ananatis, Pantoea
agglomerans, Pantoea stewartii
0.976
18
Enterobacteriaceae (100%) Erwinia (51%)
Erwinia soli, Pantoea ananatis,
Pantoea stewartii
0.997-1.000
1.000
*
RDP-II Classifier, Naïve Bayesian rRNA Classifier Version 2.0, taxonomic
assignment (confidence threshold).
#
RDP-II Seqmatch most closely related species.
Y
RDP-II Seqmatch S_ab score is calculated based on the number of unique
oligos in the query sequence compared to sequences in the RDP-II database; the
closer the score is to 1.000, the higher the degree of sequence similarity (i.e. the
same unique oligos).
** RDP-II and BLASTn results were in agreement.
63
Table 9. Biochemical testing using Enterotube II for verification of 16S rDNA
sequencing results.
Isolate
Code
Colony Source
colour on
UriSelect4
Sequence Identification Enterotube Identification
7
Blue
Microbial inoculant 1
Klebsiella/Enterobacter
Klebsiella pneumoniae
subspecies ozaenae
12
Purple
Microbial inoculant 2
Enterobacter cloacae
Enterobacter cloacae
16
Purple
Microbial inoculant 2
Enterobacter cloacae
Enterobacter cloacae
18
Purple
Microbial inoculant 2
Pantoea species
Klebsiella pneumoniae
subspecies ozaenae
Figure 9. Biochemical tests performed using Enterotube II.
64
3.3.2.
Risk
assessment
for
exposure
to
pathogenic
organisms
in
contaminated cultures.
Risk
assessment
Enterobacteriaceae
indicated
that
(opportunistic
exposure
pathogens)
to
or
bioaerosols
contact
while
containing
sampling
contaminated microbial brews was associated with a high level of risk. Sampling
activity associated with exposure to bioaerosols containing Klebsiella pneumoniae
was determined to be of very high risk.
High and very high levels of risk are
unacceptable and required cessation of the activity in accordance with the
Workplace Health and Safety Act 1995 (WHSA 1995). The implementation of risk
control measures including HEPA filtered respirators, skin protection and eyewear,
reduced the health risk associated with sampling of microbial fermentations
containing human pathogenic organisms to moderate upon reassessment.
moderate risk level still required correction.
A
The use of a bioreactor (enclosed
vessel) to contain bioaerosols reduced the risk level of this activity to low. While
these measures reduced risks associated with production of inoculants, risk
assessment for the spray application of Class 2 biological organisms (that are
capable of causing human disease) indicated a high level of risk is encountered
even if skin, respiratory and eye protection is used.
Conversely the spray
application of Class 1 organisms (not likely to cause human disease), where
personal protective equipment is employed (P2 mask, goggles and skin protection)
was associated with a low level of risk (assessed for small scale application only).
3.4. Discussion
Microbiological analysis indicated that enteric bacteria, that may be present in
materials used to prepare compost tea and in commercial microbial inoculants, may
also be detected at high levels in aerated cultures produced under non-sterile
conditions. Further analyses by 16S rDNA sequencing and/or biochemical testing
demonstrated contaminants present in commercial inoculants included Class II
65
organisms Klebsiella pneumoniae subspecies ozaenae, Enterobacter cloacae,
Stenotrophomonas maltophilia, Pseudomonas aeruginosa and Photobacterium
damsela.
While many of these opportunistic pathogens have plant growth
promoting and/or biocontrol activities (Berg et al. 2005), such organisms may cause
human disease, particularly in susceptible populations such as immunocompromised individuals, the elderly, neonates, patients undergoing advanced
medical procedures or those with a predisposing illness (Black 1999). It has been
estimated that such susceptible groups may account for up to 25% of the human
population (Matthews 2006).
The clinical significance of pathogens isolated is
described in Appendix 3.4.
The results of the current study are in agreement with findings of Sturz and coworkers (2006) who also detected Klebsiella (and Escherichia) species in a
commercial compost tea preparation.
Following the application of the compost tea
to potato plants by means of a foliar spray (690L/Ha) populations of phylloplane
bacteria were assessed. Both Klebsiella and Escherichia species were detected in
leaf washings, but were absent in control treatments. Salmonella species were also
detected in the preparation but were not detected on leaves.
A commercial
preparation of powdered kelp was also assessed. When this product was mixed in a
tank with well water, the dominant bacterial population detected was Pseudomonas
aeruginosa (Appendix 3.4) and this bacterium was also detected in analyses of
phylloplane bacteria after application of the kelp treatment (Sturz et al. 2006).
Other recent research assessed the propagation of food-borne pathogens
E. coli 0157, Salmonella enterica and Enterococcus spp. in aerated and non-aerated
compost tea (Ingram and Milner 2007). Findings supported the results of this study,
in that both faecal coliforms and food borne pathogens grew in compost tea
produced with additives, such as kelp, humic acids and rock dusts, with or without
the addition of molasses.
The growth of pathogenic indicator organisms was
demonstrated in additives alone that are typically added to microbial brews for soil
health benefits and to promote fermentation (Ingram and Milner 2007). Similarly in
66
the present study, high levels of bacteria were detected in additives and substrates
used for the production of compost tea/microbial cultures. The use of such additives
in the culture of microbial inoculants, particularly under non-sterile conditions, might
result in the final product not having the desired microbial composition.
Furthermore, the culture of numerous strains together may result in competitive
inhibition amongst the different bacteria, so that not all strains would be present in
the end product of fermentation.
As suggested by Kennedy and colleagues,
different strains of bacteria should be cultured separately and then combined at the
final stage of production.
Ingram and Milner (2007) also reported that when enteric contaminants were not
detected in the untreated compost material, the growth of pathogenic indicator
organisms did not occur in compost teas that were prepared without additives (such
as kelp and molasses).
In these compost teas produced without additives,
comparable populations of aerobic heterotrophs were supported by nutrients present
in the compost. Further research is required to assess the efficacy of compost teas
produced without additives to enhance plant growth (Ingram and Milner 2007), as
Scheurell and Mahaffe (2004) showed that the disease suppressive capacity of
compost tea could depend on the additives used in preparation. Perhaps a system
where materials such as kelp, humic acids and rock dusts are applied separately to
the compost tea and without fermentation may assist in reducing levels of human
pathogenic contaminants in compost teas. The fate of other compost resistant
pathogens, such as Legionella spp., P. aeruginosa and Clostridium perfringens in
uncontrolled microbial culturing remains to be thoroughly assessed. It is noteworthy
that in the current study, the commonly used indicator E. coli was detected at very
low levels in the presence of other pathogenic contaminants. Thus enterococci or
Klebsiella spp. might be more suitable indicators for the monitoring of pathogens in
compost tea/microbial cultures.
The requirement to test individual compost tea
production systems, including water quality is vitally important, as unsanitary
irrigation water or improperly treated manures are increasingly implicated in the
67
transmission of food-borne pathogens on fresh produce (CTTFR 2004; Brandl
2006).
Risk analysis undertaken in this study indicated that exposure to bioaerosols
containing human pathogenic bacteria, particularly those that are transmissible via
the respiratory route (i.e. Klebsiella pneumonia), generated in the production and
application of contaminated microbial brews or contact with these fermentations, is
associated with an unacceptable level of risk.
Bioaerosols are dusts or droplets of
water in air that may contain fungi, mycotoxins, bacteria, enzymes and endotoxins
(Millner et al. 1994).
Long-term exposure to bioaerosols containing bacterial
lipopolysaccharides was associated with an inflammation response of the upper
airways, allergic alveolitis, diseases of the skin, hypersensitivity pneumonitis and
gastro-intestinal infections in studies of compost and wastewater treatment workers
(Milner et al. 1994; Hansen et al. 2003; Bunger et al. 2000; Ivens et al. 1999).
Therefore respiratory protection should be used to prevent inhalation of bioaerosols
that contain both pathogenic and non-pathogenic organisms during the production
and application of microbial cultures.
The implementation of personal protective equipment (HEPA filtered respirator, eye
goggles, gloves and skin protection), reduced risks associated with sampling of
cultures containing Enterobacteriaceae or K. pneumoniae to moderate, which still
required correction.
While containment of aerosols in an enclosed bioreactor
reduced the health risk of the sampling activity to low, this equipment was not
available at the time the experiments were performed. Therefore further sampling of
these microbial fermentations was not permitted under University of the Sunshine
Coast Occupational Health and Safety Policies that are in accordance with the
Workplace Health and Safety Act 1995.
As such, commercial inoculants and
compost fermentations were precluded from further study that included analysis of
beneficial bacteria populations.
Furthermore, it was determined that the spray
application of microbial cultures containing Enterobacteriaceae or K. pneumoniae
was associated with a high level of risk, even when personal protective equipment
68
was used.
The risk assessment outcomes support concerns expressed by The
National Organic Program of the United States Department of Agriculture, who
stated that the microbial composition of compost teas are difficult to ascertain and
control and we are concerned that applying compost teas could impose a risk to
human health …. additional input from the NOSB and the agricultural research
community (is required) before deciding whether these materials should be
prohibited in organic production or whether restrictions on their use are appropriate
(NOP 2007).
The size and distance travelled by bioaerosols produced via the spray application of
compost teas has not been reported.
Paez-Rubio and co-workers (2005)
demonstrated that bioaerosols containing up to 109 enteric bacteria /m3 of air were
produced when effluent wastewater was applied to land by flood irrigation.
Bioaerosols containing up to 104 bacteria per cubic metre of air were detected 46 m
downwind from the point of spray application of wastewater that contained 2.4 x 105
CFU/ml (Bausam et al. 1981; Teltsch and Katzenelson 1977). In addition, the spray
application of wastewater effluent was associated with increased incidence of
communicable disease in an adjacent community (Katzenelson et al. 1976). Thus
the spray application of compost tea containing up to 109 CFU/ml may produce
bioaerosols of an even greater magnitude and the distance travelled by these
bioaerosols remains to be assessed.
The potential for production and application of contaminated brews (in the order of
100 -1000L) to increase pathogen loads in the environment is a serious threat which
needs to be managed in order to prevent increased incidence of human disease.
Many of the pathogenic organisms present in the microbial cultures evaluated are
also found in animal manures and increased microbial pollution of surface waters
has been associated with leaching and run-off following the land application of
livestock wastes (Tyrrel and Quinton 2003, Pell 1997; Jones 2003; USDA/FDA 1998;
Entry and Farmer 2001). Levels of indicator pathogens in recreational waters have
been correlated to incidences of swimming-related gastro-intestinal illnesses (Cabelli
69
et al. 1979; Kinzelman et al. 2003; Eyles et al. 2003). In order to reduce pollution of
waterways and reduce the potential for the contamination of fresh produce by foodborne pathogens in crop production systems, composting of animal manures prior to
land application, manure storage facilities, testing of sanitary quality of irrigation
water and withholding periods between manure application and harvesting have
been recommended as a “good agricultural practices” by the United States Food and
Drug Administration (US-FDA 1998; Meals and Braun 2006; Matthews 2006). Thus
leaching and run-off following the application of compost teas/microbial cultures
containing pathogenic organisms might add to problems of increased environmental
microbial pollution, which is a human health concern. Consideration should be also
be given to the safe disposal of contaminated cultures to minimise run off into
waterways. In fact the safe disposal of hundreds to thousands of litres of cultures
that contain pathogenic and/or phytotoxin producing bacteria may be problematic.
While similar organisms may be present in manures and compost teas, there are
important differences in the way these two materials are applied, where bioaerosols
produced via the spray application of compost tea might provide additional routes for
the dissemination and transmission of pathogenic organisms.
The detection in this study, of the fish pathogen and human flesh-eating bacteria,
Photobacterium damsela (Yamane et al. 2004; Osorio et al. 1999) and Klebsiella
pneumoniae that is associated with severe morbidity (Bingen et al. 1993; Podschun
and Ullmann 1998), in a commercial microbial inoculant highlights the potential
danger that is present due to lack of regulation and quality control in the Australian
biofertiliser industry; and supports the argument of Kennedy and colleagues (2004)
that an infrastructure closely linked to the biofertiliser application industry allowing
research to improve inoculant quality and quality control of current production as well
as of stored commercial products is considered essential (as stated).
The
establishment of standards requiring minimal numbers of Rhizobium and limited
levels of contaminants has been associated with the success of commercial
inoculants in the legume industry (Bullard et al. 2005; Deaker et al. 2004). Improved
performance and safety of microbial inoculants targeted to broader agricultural
70
industries might be achieved in Australia by adopting standards such as those
stipulated in US EPA regulations for biopesticides; including confirmation of the
absence of pathogenic contaminants on selective media, strain verification and the
use of personal protective equipment during production and application of inoculants
(US-EPA 2005). These measures are expected to mitigate risks to human health in
the use of biopesticides formulated with non-pathogenic microorganisms.
In
agreement with this assertion, are risk assessments undertaken in the current study,
where the application of non-pathogenic organisms while using PPE was associated
with a low level of risk. Therefore further research aimed to investigate the potential
of pure cultures of non-pathogenic bacteria to promote growth and resistance to
disease in ginger, under greenhouse conditions.
3.5. Conclusion
In conclusion, results of this study demonstrated that human pathogenic bacteria
may be present in compost tea and commercial microbial inoculants. Exposure to
pathogens in microbial cultures prepared from these materials, via bioaerosols or
direct contact, was associated with a high level of risk, which was not mitigated by
the use of PPE and thus precluded these inoculants from further study. This
demonstrates the necessity for quality control standards in the commercial
biofertiliser industries and in compost tea production systems in Australia to avoid
contamination of fresh produce with pathogenic organisms and; to minimise the
dispersal and transmission of pathogens via bioaerosols and leaching into
watercourses. Users of compost teas in Australia were observed to perceive that
microbial cultures prepared from compost are completely safe. Risks associated
with active sniffing of microbial solutions containing human pathogenic organisms
and the need to use skin, respiratory and eye protection (even though all risks are
not mitigated) in the production and application of compost needs to be more widely
promoted.
71
Further research on the use of microorganisms to promote growth and resistance to
disease in ginger therefore aimed to use only pure cultures of non-pathogenic
bacteria produced with aseptic technique as this was associated with a low level of
risk.
72
Appendix 3.1. Primers used in 16S rDNA PCR and sequencing.
Table 10. Primers used for 16S rDNA PCR sequencing of inoculant isolates.
Primer Primer Sequence (5’- 3’)
Code*
Reference
Primer Synthesis
27F
GAGAGTTTGATCCTGGCTCAG
Girfoni et al. 1995;
Weisburg et al. 1991.
Sigma-Genosys
(Australia).
518F
CCAGCAGCCGCGGTAATACG
926R
CCGTCAATTCCTTTGAGTTT
Lu et al. 2000; Lane et Invitrogen (South
al. 1985.
Australia) and
Sigma-Genosys
(Australia).
Schwieger and Tebbe Invitrogen (South
1998; Lane et al. 1985. Australia) and
Sigma-Genosys
(Australia).
Girfoni et al. 1995;
Sigma-Genosys
Weisburg et al. 1991. (Australia).
1492R ACGGCTACCTTGTTACGACTT
1513R ATCGGCTACCTTGTTACGACTTC Lu et al. 2000.
Invitrogen (South
Australia).
* For 16S rDNA primers, numbers indicate relative position of primers along the16S
rRNA gene, according to numbering in E. coli.
73
Appendix 3.2. Illustration of CLUSTAL W 2.0 multiple sequence alignment used to produce a consensus
sequence *.
4_U1
4_U3
GCGGTTTCTTAAGTCTGATGTGAAAGCCCCCGGCTCAACCGGGGAGGGTCATTGGAAACT 60
------------------------------------------------------------------------------------GGGGAGGGTCATTGGAAACT 20
4_U1
4_U3
GGGAAACTTGAGTGCAGAAGAGGAGAGTGGAATTCCACGTGTAGCGGTGAAATGCGTAGA 120
GGGAAACTTGAGTGCAGAAGAGGAGAGTGGAATTCCACGTGTAGCGGTGAAATGCGTAGA 80
4_U1
4_U3
GATGTGGAGGAACACCAGTGGCGAAGGCGACTCTCTGGTCT---------------------------------------- 161
GATGTGGAGGAACACCAGTGGCGAAGGCGACTCTCTGGTCTGTAACTGACGCTGAGGAGC 140
4_U1
4_U3
-----------------------------------------------------------------------------------------------------------------------------GAAAGCGTGGGGAGCGAACAGGATTAGATACCCTGGTAGTCCACGCCGTAAACGATGAGT 200
4_U1
4U3
-----------------------------------------------------------------------------------GCTAAGTGTTAGGGGGTTTCCGCCCCTTAGTGCTGCAGCTA 241
*100% sequence homology demonstrates accuracy and reproducibility of the sequencing procedure.
75
Appendix 3.3. 16S rDNA consensus sequences (5’-3’) of isolates from untreated
microbial inoculants.
>4_consensus
GCGGTTTCTTAAGTCTGATGTGAAAGCCCCCGGCTCAACCGGGGAGGGTCAT
TGGAAACTGGGAAACTTGAGTGCAGAAGAGGAGAGTGGAATTCCACGTGTAG
CGGTGAAATGCGTAGAGATGTGGAGGAACACCAGTGGCGAAGGCGACTCTCT
GGTCTGTAACTGACGCTGAGGAGCGAAAGCGTGGGGAGCGAACAGGATTAGA
TACCCTGGTAGTCCACGCCGTAAACGATGAGTGCTAAGTGTTAGGGGGTTTCC
GCCCCTTAGTGCTGCAGCTA
>7U3_consensus
TCTGTCAAGTCGGATGTGAAATCCCCGGGCTCAACCTGGGAACTGCATTCGAA
ACTGGCAGGCTAGAGTCTTGTAGAGGGGGGTAGAATTCCAGGTGTAGCGGTG
AAATGCGTAGAGATCTGGAGGAATACCGGTGGCGAAGGCGGCCCCCTGGACA
AAGACTGACGCTCAGGTGCGAAAGCGTGGGGAGCAAACAGGATTAGATACCC
TGGTAGTCCACGCCGTAAACGATGTCGACTTGGAGGTTGTGCCCTTGAGGCG
TGGCTTCCGGAGC
>8_consensus
TAAAGCGCACGCAGGCGGTCTGTCAAGTCGGATGTGAAATCCCCGGGCTCAA
CCTGGGAACTGCATTCGAAACTGGCAGGCTGGAGTCTTGTAGAGGGGGGTAG
AATTCCAGGTGTAGCGGTGAAATGCGTAGAGATCTGGAGGAATACCGGTGGC
GAAGGCGGCCCCCTGGACAAAGACTGACGCTCAGGTGCGAAAGCGTGGGGA
GCAAACAGGATTAGATACCCTGGTAGTCCACGCCGTAAACGATGTCGATTTGG
AGGTTGTGCCCTTGAGGCGTGGCT
>10_consensus
CGGAGGGTGCAAGCGTTAATCGGAATTACTGGGCGTAAAGCGCACGCAGGCG
GTCTGTCAAGTCGGATGTGAAATCCCCGGGCTCAACCTGGGAACTGCATTCGA
AACTGGCAGGCTGGAGTCTTGTAGAGGGGGGTAGAATTCCAGGTGTAGCGGT
GAAATGCGTAGAGATCTGGAGGAATACCGGTGGCGAAGGCGGCCCCCTGGA
CAAAGACTGACGCTCAGGTGCGAAAGCGTGGGGAGCAAACAGGATTAGATAC
CCTGGTAGTCCACGCCGTAAACGATGTCGATTTGGAGGTTGTGCC
>12_consensus
GGTCTGTCAAGTCGGATGTGAAATCCCCGGGCTCAACCTGGGAACTGCATTC
GAAACTGGCAGGCTGGAGTCTTGTAGAGGGGGGTAGAATTCCAGGTGTAGCG
GTGAAATGCGTAGAGATCTGGAGGAATACCGGTGGCGAAGGCGGCCCCCTG
GACAAAGACTGACGCTCAGGTGCGAAAGCGTGGGGAGCAAACAGGATTAGAT
ACCCTGGTAGTCCACGCCGTAAACGATGTCGATTTGGAGGTTGTGCCCTTGAG
GC
>13_consensus
AAGCGCACGCAGGCGGTCTGTCAAGTCGGATGTGAAATCCCCGGGCTCAACC
TGGGAACTGCATTCGAAACTGGCAGGCTGGAGTCTTGTAGAGGGGGGTAGAA
TTCCAGGTGTAGCGGTGAAATGCGTAGAGATCTGGAGGAATACCGGTGGCGA
AGGCGGCCCCCTGGACAAAGACTGACGCTCAGGTGCGAAAGCGTGGGGAGC
AAACAGGATTAGATACCCTGGTAGTCCACGCCGTAAACGATGTCGAT
>14_consensus
CTGGCAGGCTGGAGTCTTGTAGAGGGGGGTAGAATTCCAGGTGTAGCGGTGA
AATGCGTAGAGATCTGGAGGAATACCGGTGGCGAAGGCGGCCCCCTGGACAA
76
AGACTGACGCTCAGGTGCGAAAGCGTGGGGAGCAAACAGGATTAGATACCCT
GGTAGTCCACGCCGTAAACGATGTCGATTTGGAGGTTGTG
>16_consensus
CTGGGCGTAAAGCGCACGCAGGCGGTCTGTCAAGTCGGATGTGAAATCCCCG
GGCTCAACCTGGGAACTGCATTCGAAACTGGCAGGCTAGAGTCTTGTAGAGG
GGGGTAGAATTCCAGGTGTAGCGGTGAAATGCGTAGAGATCTGGAGGAATAC
CGGTGGCGAAGGCGGCCCCCTGGACAAAGACTGACGCTCAGGTGCGAAAGC
GTGGGGAGCAAACAGGATTAGATACCCTGGTAGTCCACGCCGTAAACGATGT
C
>17_consensus
GGTGCAAGCGTTAATCGGAATTACTGGGCGTAAAGCGCACGCAGGCGGTCTG
TTAAGTCAGATGTGAAATCCCCGGGCTTAACCTGGGAACTGCATTTGAAACTG
GCAGGCTTGAGTCTCGTAGAGGGGGGTAGAATTCCAGGTGTAGCGGTGAAAT
GCGTAGAGATCTGGAGGAATACCGGTGGCGAAGGCGGCCCCCTGGACGAAG
ACTGACGCTCAGGTGCGAAAGCGTGGGGAGCAAACAGGATTAGATACCCTGG
TAGTCCACGCCGTAAACGATGTCGACTTGGAGGC
>18_U3
GGAATTACTGGGCGTAAAGCGCACGCAGGCGGTCTGTTAAGTCAGATGTGAA
ATCCCCGGGCTTAACCTGGGAACTGCATTTGAAACTGGCAGGCTTGAGTCTCG
TAGAGGGGGGTAGAATTCCAGGTGTAGCGGTGAAATGCGTAGAGATCTGGAG
GAATACCGGTGGCGAAGGCGGCCCCCTGGACGAAGACTGACGCTCAGGTGC
GAAAGCGTGGGGAGCAAACAGGATTAGATACCCTGGTAGTCCACGCCGTAAA
CGATGTCGACTTGGAGGCTGTTCCCTTGAGGAGTGG
77
Appendix 3.4. Public Health Significance of Human Pathogenic Bacteria.
Opportunistic pathogens, often a part of the gastrointestinal microflora, are
implicated as causes of disease in: i) immuno-compromised or severely
malnourished individuals (for example AIDS, chemotherapy/radiation); ii) cases of
pre-exiting diseases (such as diabetes mellitus and cystic fibrosis); iii) advanced
age, pregnancy and neonates; iv) incidences where the organism is introduced into
sites where it is not usually resident and; instances where the normal microflora has
been disturbed (Black 1999).
Soil, plant and water habitats may serve as reservoirs of these organisms
(Podschun et al. 2001; Berg et al. 2005).
The way in which virulence factors
determine pathogenicity and differ in environmental and clinical isolates is generally
poorly understood for many opportunistic pathogens (Berg et al. 2005). Although
environmental strains of Klebsiella spp. and Pseudomonas aeruginosa have been
shown to carry virulence factors and to be indistinguishable from clinical isolates
(Podschun et al. 2001; Wolfgang et al. 2003; Morgan et al. 1999). Clinical isolates
with plant growth promoting properties and the ability to fix nitrogen have also been
reported (Dong et al. 2003; Lovtunovych et al. 1999).
The frequency of nosocomial (hospital acquired) infections caused by opportunistic
pathogens has increased in recent times, which may be in part due to increased
numbers of people undergoing advanced medical procedures.
Nosocomial
infections are increasingly difficult to treat due to emerging antibiotic resistance and
are often associated with severe morbidity and mortality (Bingen et al. 1994; AGAR
2003; Bell et al. 1998). Species that have become increasingly difficult to treat due
to emerging antibiotic resistance include methicillin resistant Staphylococcus
aureus, vancomycin resistant Enterococcus, Klebsiella spp. and Enterobacter
(AGAR 2003).
Antibiotic resistance factors are often plasmid encoded.
The
horizontal transfer of resistance plasmids between different pathogenic genera has
been demonstrated in sewage effluent, biofilms, rhizospheres and soil
(Arana
2001; Chee-Sanford et al. 2001; Hausner and Wuertz 1999; Molbank et al. 2007).
78
Infections caused by relevant human pathogenic bacteria are described in Table
11. Community-onset bloodstream infections often associated with high mortality
rates include those caused by S. aureus, E. coli, coagulase-negative staphylococci,
Enterococcus spp. Klebsiella spp. and P. aeruginosa (Diekema et al. 2003).
79
Table 11. Types of infections caused by selected human pathogenic bacteria*.
Organism
AR#
Klebsiella
pneumoniae
+
Pneumonia,
bloodstream,
meningitis, liver
abscess.
UTI, bacteremia,
pneumonia
Extremely virulent
pathogen, infections
cause severe morbidity
and mortality in the aged,
newborn and immunocompromised.
Enterobacter spp.
+
………..
Bloodstream,
pulmonary, wound and
urinary infections.
Emerging pathogen.
Enterococcus
faecalis
+
………..
Endocarditis,
bloodstream, UTI and
wound infections.
Becoming major
nosocomial pathogen.
Community
Infections
Nosocomial
Infections
Further description
Hemorrhagic colitis ………..
and haemolytic
uremic syndrome.
Food borne pathogen;
vero toxigenic: small
numbers, often below
conventional detection
limits can cause disease.
Acute ulcerative
keratitis (contact
lens users), blood
stream infections
(burn victims).
Nosocomial infections
(eg. bacteremia in
neonates and
immunocompromised),
lung infections (cystic
fibrosis).
Cause of morbidity and
mortality in cystic fibrosis
and hospitalised patients.
Stenotrophomonas +
maltophilia
………..
Bloodstream, eye,
respiratory and central
nervous system
infections
High mutational rate,
emergent pathogen with
broad spectrum antibiotic
resistance
Pantoea
agglomerans
Infections:
………..
bloodstream,
abscesses,
joint/bones, urinary
tract
Escherichia coli
0157
Pseudomonas
aeruginosa
+
Paediatric infections;
community acquired
infections following
trauma (by vegetation or
clinical procedures)
*(Podschun and Ullman 1998; Sanders and Sanders 1997; Chow et al. 1991; Kaye
et al. 2001; Bell et al. 1998; Cetinkaya et al. 2000; Eaton and Gasson 2001;
Altekruse et al. 1997; Armstrong et al. 1996; Bingen et al. 1995; Pai et al. 2004;
Lyczak et al. 2000; Carmeli et al. 1999; Berg et al. 1999; Garrison et al. 1996;
Denton and Kerr 1998; Denton et al. 1998; Cruz et al. 2007).
# AR: antibiotic resistance. UTI: urinary tract infection.
80
Chapter 4. Selection of bacterial isolates for further testing in ginger .
4.1. Introduction
There is an abundance of literature that has described the use of different strains of
Pseudomonas, Bacillus and Azospirillum to promote growth and resistance to
disease in crops of agronomic importance (Kloepper et al. 2004; Jacobsen et al.
2004; Mercado-Blanco and Bakker 2007; Bakker et al. 2007; Whipps 2001;
Dobbeleare et al. 2001; Steenhoudt and Vanderleyden 2000). Consideration of the
strain concept is paramount in the use of these bacteria to enhance plant growth.
Minor genetic variation between strains of the same species can determine
phenotypic traits including specificity of plant-bacteria interactions, as well as plant
growth promoting and biocontrol activities (Kloepper 1996). Colonisation of plant
roots or tissues is essential for plant growth promoting bacteria to exert beneficial
effects on plant growth via phytostimulatory hormones, antibiosis or induced
systemic resistance (O’Sullivan and O’Gara 1992). Thus the isolation of PGPB from
the rhizosphere or plant tissues may assist in: i) the separation of plant-adapted
strains from diverse indigenous microbial populations present in soil; ii) identify
which bacteria are naturally associated with different plant types and: iii) provide
strains for testing that are adapted to regional conditions. This may aid in the
selection of bacteria that more reliably induce plant growth promotion and disease
resistance in field conditions (Kennedy et al. 2004; Gunarto et al. 1999; Fisher et al.
2006; Fravel 2005).
The use of combinations of beneficial strains that act via complementary
mechanisms or that occupy a different niche on the root surface may also increase
the efficacy of bacterial inoculation in different soil types and under different
environmental and seasonal conditions (Fravel 2005; Whipps 2001; Guetsky et al.
2004).
For example combination of nitrogen fixing bacteria with phosphate
solubilizing bacteria had a synergistic effect on the growth of sugar beet, barley and
a variety of vegetable crops (Sahin et al. 2004; Belimov et al. 1995; El-Komy 2004;
Bashan et al. 2004). It has also been proposed that microorganisms implicated in
the improved plant growth response to soil fumigation might be used for plant
inoculation (Martin and Bull 2002).
81
Few reports have described the use of plant growth promoting bacteria for
improved growth and disease resistance in ginger (Meena and Mathur
2003;Sharma and Jain 1979) and details such as species level identification,
application methods or concentration of viable cells used were not described.
Accordingly, initial objectives included obtaining potentially beneficial bacteria by
isolation from roots of ginger plants growing in fumigated and non-fumigated soils.
To assess soil health in the plots used for bacterial isolations predatory, fungalfeeding, bacterial-feeding, omnivorous and plant-parasitic nematodes were counted
(Stirling 2005; Patisson et al.2004).
Another source of potential PGPB were
reference strains, with demonstrated root colonising ability, plant-growth promoting
or biocontol potential, available from various culture collections.
The bacteria
targeted included strains of Azospirillum, Bacillus and Pseudomonas, the most
commonly used and studied PGPB, that are able to improve disease resistance
and/or promote growth in a wide variety of different plant types (Kloepper et al.
2004; Jacobsen et al. 2004; Mercado-Blanco and Bakker 2007; Bakker et al. 2007;
Whipps 2001; Dobbeleare et al. 2001; Steenhoudt and Vanderleyden 2000). The
phosphate solubilizing ability of bacterial strains was assessed to identify strains
that might provide benefits when co-inoculated with other PGPB in ginger.
4.2. Materials and Methods
4.2.1. Isolation of bacteria from the ginger rhizosphere and rhizoplane and
field observations.
4.2.1.a. Field observations and sampling.
Soil and ginger root samples were collected from a 1.5 Ha site at Eumundi,
Queensland (GPS 26 29’39S, 152 57’11E) used for commercial cultivation of early
harvest ginger cv. Queensland. Poultry manure and cover crops (Saia Oats) were
incorporated approximately eight weeks prior to planting. The red clay loam was
also tilled to form a seed-bed for planting. Half of the plot was fumigated with
metham sodium and the remainder was not fumigated; otherwise both plots
received equivalent treatment, including irrigation and applied nitrogen, phosphorus
82
and potassium fertilisers. Ginger was planted in October and harvested in March
(2006). Yield was estimated as tonnage of rhizomes produced per acre.
Ten plants were collected from both fumigated and untreated plots in January and
March 2006. Plants were dug up with a trowel so that roots remained intact and
shaken to remove loosely adhering soil (Loper et al. 1985; McSpadden Gardener et
al. 2001). Roots were collected into UV-irradiated plastic zip-lock bags. A soil
sample beneath the plant (10-25cm deep) was collected with a trowel for analysis
of nematode populations (Stirling 1994).
Samples were combined to form a
composite sample. Nematodes were extracted in a Whitehead tray for counting of
plant parasitic and community nematode populations (Whitehead and Hemming
1965).
4.2.1.b. Isolation of rhizosphere and rhizoplane bacteria.
Rhizosphere suspensions were prepared as described by Raaijmakers and Weller
(2001) with minor modifications and using aseptic technique. Roots were cut with
into 5cm segments in a Petri dish using a scalpel blade. Samples were transferred
to a 100ml Schott bottle and weighed. Sterile phosphate buffered saline (PBS) was
added to the sample (10ml/gram of root). Samples were incubated with agitation
on a Griffin shaker (Stuart Scientific, 250 rpm) for 20 minutes.
The resulting
rhizosphere suspension was serially diluted in sterile PBS (ten fold dilutions) and
200 µL of each dilution was plated in triplicate on the following media (Table 12).
King's B medium (King et al. 1954) was used to isolate fluorescent Pseudomonas
species. For the enrichment of heat resistant endospore forming bacteria such as
Bacillus species a sub-sample of rhizosphere suspension was heated to 80oC for
20 minutes and plated on nutrient agar (Krause et al. 2003).
The method used to prepare rhizoplane suspensions was adapted from Ross et al.
2000 and McSpadden-Gardener and Weller (2001) as follows. Roots were washed
in eight changes of sterile PBS to remove visible soil particles. The washed roots
were then macerated in sterile PBS, incubated on a Griffin shaker for 30 minutes
and sonicated for 60 seconds (Mazzola and Cook 1991).
83
Serial dilutions and
plating on media were performed as described earlier. This rhizoplane suspension
was also plated on Congo red media for the isolation of diazotrophic bacteria
attached to the root surface (Rodriguez-Caceres 1982; Muthukumar et al. 2001).
Table 12. Media and Culture conditions for the isolation of root associated bacteria.
Culture Media* Bacteria Isolated
Incubation Temp,
Time
King’s B agar
28oC, 48 hours
Fluorescence under UV
light.
35oC, 72 hours
Growth on NFb Media.
Fluorescent
Pseudomonas
Congo Red Agar Diazotrophic bacteria
Confirmatory
Analyses
Endospore forming
28oC, 48 hours
Phosphate solubilizing
bacteria (heat treated
activity.
sample).
*References and composition of media is listed in Appendix 4.1. Cycloheximide
and nystatin (50ppm) were added to all media to inhibit growth of fungi.
Nutrient Agar
4.2.2. Assessment of phosphate solubilizing activity and growth of bacteria
on nitrogen free media.
For confirmation of the ability to grow on nitrogen free media isolates recovered on
Congo red medium were transferred to New Fabian b (NFb) agar (Appendix 4.1;
Dobereiner 1995) and incubated at 35oC for five days.
Isolates recovered on
Congo red medium that were also able to grow on NFb media were selected for
phylogenetic analysis.
Endospore forming bacteria from rhizoplane and rhizosphere suspensions were
transferred to Pikovskaya media (Appendix 4.1; Johri et al. 1999) to assess in vitro
phosphate solubilization ability.
The phosphate solubilizing ability of reference
strains (section 4.2.2) was also assessed on PVK media.
incubated at 30oC for two weeks.
84
PVK plates were
4.2.3. Identification of selected rhizosphere and rhizoplane bacteria.
Pure cultures of bacteria were obtained by streaking on the isolation media or
nutrient agar a least three times or until pure. A single colony of selected bacteria
was used to inoculate nutrient broth amended with yeast extract (Appendix 4.1) and
grown overnight with shaking (~100rpm, 28oC).
Bacterial genomic DNA was
extracted from cells of the overnight cultures using the ChargeSwitch gDNA Mini
Bacteria Kit (Invitrogen, California) according to manufacture’s instructions.
Arbitrarily primed PCR was used to generate a genomic fingerprint of potential
diazotrophic rhizoplane bacteria so that different species might be identified
(Gunarto et al. 1999).
The PCR reaction mixture consisted of 1X PCR Supermix
(20mM Tris-HCl, 5mM KCL, 1.5mM, 200µM of each dNTP, 1 U Taq DNA
polymerase; Invitrogen, California), 5 pmol of primer OPT-08 (Appendix 4.2), an
additional 1mM MgCl2 (final concentration 2.5mM) and 100ng DNA. Thirty cycles of
94 oC for 1 min, 36 oC for 1 min and 72 oC for 2 min were performed in an Eppendorf
Mastercyler thermal cycler. PCR products were visualised on a 1.5% agarose gel
stained with ethidium bromide. The reaction was repeated three times to determine
consistency of banding patterns produced.
For phylogenetic analysis of selected bacteria partial sequencing of the 16S rRNA
gene was undertaken (Weisberg et al. 1991).
Primers 27F and 1492R (Appendix
4.2) were used to amplify the 16S rRNA gene by PCR. The 50µL PCR reaction
included 1X PCR Supermix (20mM Tris-HCl, 5mM KCL, 1.5mM, 200µM of each
dNTP, 1 U Taq DNA polymerase; Invitrogen, California), 0.5µM of each primer, 1 µL
of diluted genomic DNA (~10 ng) and an additional 0.5mM MgCl2 to produce a final
concentration of 2mM MgCl2. Cycling conditions included an initial denaturation
step at 94 oC for 4 min; 25 cycles of 94 oC for 30 sec, 55 oC for 30 sec, 72 oC for 90
sec; and a final extension at 72 oC for 10 min. PCR products were visualised on a
1.5% agarose gel stained with ethidium bromide.
85
The PCR products were purified and then sequenced with an ABI3730XL using ABI
PRISM Big Dye Terminator Sequencing Chemistry (Version 3.1) by Macrogen Inc
(Korea). The primers 518F and 926R were used to sequence both strands of the
DNA (Appendix 4.2).
The sequence chromatograms were visualised using the
program Finch TV (GeoSpiza). A consensus sequence was determined following
alignment of sequences by the ClustalW program (http://www.ebi.ac.uk/clustalw/).
The Basic Local Alignment Search Tool (BLASTn) at the National Centre for
Biotechnology Information (http://www.ncbi.nml.nih.gov/BLAST) was used to
compare ~500bp of DNA sequence to those in GenBank.
Sequences were
additionally analysed using Ribosomal Database Project (RDP-II) programs
SeqMatch, to determine nearest neighbours and Classifier, for assignment of
taxonomic hierarchy (http://rdp.cme.msu.edu/ Michigin State University, East
Lansing, Michigan; Maidak et al. 2000).
4.2.4. Reference strains of bacteria.
Reference strains of bacteria used in this study are described in Table 13. Bacteria
were maintained on nutrient agar (Oxoid) or NBY agar (Appendix 4.1) and were
stored in 20% glycerol, 80% nutrient broth at –80oC.
4.3. Results
4.3.1.
Field observations and isolation of bacteria from the ginger
rhizosphere and rhizoplane.
4.3.1.a. Field observations and sampling.
In soils used for the isolation of rhizosphere bacteria, root-knot nematodes were not
detected in either fumigated or non-fumigated plots in early January 2006. A low
incidence of Fusarium wilt was observed in both plots.
Differences in nematode
trophic groups between fumigated and non-fumigated soil are described in Table
14.
86
Table 13. Reference Strains of Bacteria.
Bacterial Strain
Source
Described Activity
References
Bacillus subtilis DAR26694
(strain A13)
Orange Agricultural Institute (NSW
DPI).
Isolated from a vegetable garden loam;
Growth promotion, biocontrol activity
against Fusarium oxysporum and a range
of fungal plant pathogens.
Broadbent et al. 1977; Broadbent et al.
1971; Merriman et al. 1974; Turner and
Backman 1991; Zehnder et al. 2001;
Brannen and Kenny 1997.
Bacillus subtilis DAR26659
Orange Agricultural Institute (NSW
DPI).
Isolated from diseased wheat seed; In
vitro antagonism of Alternaria alternata
from wheat.
Noble, personal communication 2007.
Azospirillum brasilense Sp7
(ATCC* 29145)
Australian Collection of
Microorganisms (University of Qld
Brisbane, QLD).
Isolated form the rhizosphere of Digitaria Tarrand et al. 1978; Okon and
decumbens; Growth promotion, improved Labrandera-Gondalez 1994; Dobbeleare
water status and improved nutrient uptake. et al. 1999; Lin et al. 1983.
Azosprillum lipoferum Br-17
(ATCC* 29709)
Australian Collection of
Microorganisms (University of
Queensland, Brisbane, QLD).
Isolated from the roots of maize that had Tarrand et al. 1978; de Oliveira Pinheiro
been surface sterilised; Was combined
et al. 2002; Okon and Labranderawith A. brasilense Cd to form the
Gondalez 1994.
commercial product Zea-Nit™ which
promoted growth and reduced required N
input by 35-40% in maize cultivation.
Bacillus subtilis ATCC* 6633
Australian Collection of
Microorganisms (University of
Queensland, Brisbane, QLD).
L form bacteria that produces antifungal
peptides mycosubtilin, surfactin and
rhizocticin A. Growth promotion in
Chinese cabbage, but not in pepper;
inhibition of Botrytis cinerea.
Leclere et al. 2005; Kugler et al. 1990;
Leenders et al. 1999; Walker et al. 2002;
Allan 1991.
Pseudomonas putida KT2442
Teaching Collection, Faculty of
Science, Health and Education,
University of the Sunshine Coast
(FoSHE, USC Sippy Downs, QLD).
Teaching Collection (FoSHE, USC
Sippy Downs, QLD).
Rifampicin resistant derivative of P. putida
KT2440, which is an efficient coloniser of
roots and degrader of pollutants; biosafety
strain.
Wild type strain.
Jiminez et al. 2002; Nakazawa 2002;
Molina et al. 2005; Timmis 2002; Nelson
et al. 2002; Espinosa-Urgel et al. 2000;
Espinosa-Urgel et al. 2002.
Bacillus coagulans NCTC**
10334
Teaching Collection (FoSHE, USC
Sippy Downs, QLD).
Type strain.
Sarles and Hammer 1931.
Bacillus megaterium NCTC**
10342
Teaching Collection (FoSHE, USC
Sippy Downs, QLD).
Type strain.
Lawrence and Ford 1916.
Pseudomonas fluorescens
*ATCC American Type Culture Collection, Manassas USA ; **NCTC National Collection of Type Cultures, London UK
87
Table 14. Nematode trophic groups recovered per 100g of fumigated and nonfumigated soil.
Fumigated
Non-fumigated
Soil
Soil
Trophic Group
Genera
Number Total
Number Total
Plant Parasitic nematodes
Not detected
0
0
0
0
Bacterial feeding nematodes
Rhabditis
2
21
0
5
Teratocephalus
1
0
Pridmatolaimus
8
5
Alaimus
10
0
Tylenchus 1
5
Aphelenchus
0
5
Aphelenchoides
0
4
Tipyla
0
Fungal feeding nematodes
Predatory nematodes
5
6
Monchus ventral form 6
Omnivorous nematodes
Dorylaimus
4
Eudorylaimus
0
Total nematodes
6
1
15
12
11
4
16
18
2
36
50
The yield of rhizomes produced in fumigated and non-fumigated plots was 18.9
tonne/acre and 14.3 tonne/acre respectively. Statistical analysis of the increased
yield of 24.3% in the fumigated plot was confounded by lack of replication, due to
practical limitations.
4.3.1.b. Isolation of rhizosphere and rhizoplane bacteria.
Bacteria producing a yellow/green pigment that fluoresced under UV light, typical
of fluorescent Pseudomonas, were dominant on King’s B agar in ginger
rhizosphere suspensions from non-fumigated soil in January 2006 (Table 15). In
contrast, the rhizosphere suspension that was obtained from ginger roots growing
in fumigated soil at this time and plated on King’s B agar was dominated by white
88
colonies that did not fluoresce under UV light (Table 15); these white colonies were
selected for phylogenetic analysis. In March (2006), bacteria with a yellow-green
fluorescent pigment were not isolated from the ginger rhizosphere on King’s B
agar, neither in non-fumigated nor fumigated soil (Table 15).
4.3.2. Assessment of phosphate solubilizing activity of bacterial isolates and
growth on nitrogen free media.
Bacteria obtained by heating rhizosphere and rhizoplane suspensions to 80 oC did
not produce cleared zones on PVK media, indicating that phosphate-solubilizing
activity was not detected in these strains. B. coagulans NCTC 10334 produced a
cleared zone on PVK media, which indicated the bacteria solubilized phosphate in
vitro.
Bacteria isolated from the ginger rhizoplane on Congo red media, which were also
able to grow on NFb media, were selected for arbitrarily primed PCR and
phylogenetic analysis.
4.3.3. Identification of selected rhizosphere and rhizoplane bacteria.
16S rDNA sequencing indicated that bacterial isolates (white colonies) that were
dominant on King’s B agar in January (2006) in fumigated soil were most closely
related to Bacillus simplex and Bacillus macroides, herein referred to as Bacillus
F1 and Bacillus F2 (Table 15-16).
Banding patterns produced by arbitrarily primed PCR of gDNA from bacteria
isolated on Congo red medium (in March 2006) is illustrated in Figure 10.
Selected isolates were identified by 16S rDNA sequencing. (Table 15-16). Isolates
of Acidovorax spp. produced similar genomic fingerprints. Generally the banding
pattern produced in arbitrarily primed PCR was unique to the different genera
identified, although two closely related Pseudomonas spp. produced different
fingerprints.
89
Figure 10. Genomic fingerprints produced by arbitrarily primed PCR of gDNA from
rhizoplane bacteria *.
*1.
2.
3.
4.
5.
6.
7.
8.
9.
10.
11.
12.
13.
14.
15.
100b ladder (Promega)
Bacillus subtilis (reference strain)
Dz2
Dz4 (Acidovorax spp.)
Dz5 (Pseudomonas spp.)
Dz7
Dz9 (Acidovorax spp.)
Dz10
Dz11
Dz12
Dz13 (Pseudomonas spp.)
Dz14 (Rhizobium spp.)
Dz17
Negative control
Lambda standard (Promega).
90
Table 15. Analysis of culturable populations of bacteria associated with ginger
root samples.
Sample
Type
Soil
type
Isolation
Media
CFU/gram
of root
Colony
Morphology
Tentative
Identification
Rhizosphere
(January)
F
King’s B agar
3.8 x 105
White, nonfluorescent
Bacillus simplex/Bacillus
macroides*
Rhizosphere
(January)
NF
King’s B agar
1.7 x 105
Yellow/green
fluorescent
Fluorescent Pseudomonas**
Rhizosphere
(March)
F
King’s B agar
2.6 x 105
White, nonfluorescent
Undetermined
Rhizosphere
(March)
NF
King’s B agar
1.9 x 105
White, nonfluorescent
Undetermined
Rhizoplane
(March)
F
Congo red
4.3 x 104
Pink, red, pink Acidovorax spp.,
with red centre Pseudomonas spp.*
Rhizoplane
(March)
NF
Congo red
1.2 x 103
Pink, red, pink Pseudomonas spp.,
with red centre Rhizobium spp.,
Aneurinibacillus spp.*
*Tentative identification based on analysis of 16S rDNA sequence.
** Tentative identification based on fluorescence under UV on King’s B Agar.
91
Table 16. Phylogenetic analysis of ginger root associated bacteria*.
Isolate Soil
Code Type#
Genus
Most Closely Related Species
Identification
F1
F
Bacillus
B. simplex/B. macroides
F2
F
Bacillus
B. simplex/B. macroides
Dz1
F
Pseudomonas P. koreensis/P. putida/P. chlororaphis/P. borealis/P. corrugata
Dz3
F
Acidovorax
Acidovorax spp./A. delafieldii
Dz4
F
Acidovorax
Acidovorax sp/A. temperans
Dz5
F
Pseudomonas P. kilonensis/P. putida/P. fluorescens/P. jessenii/P. corrugata
Dz9
F
Acidovorax
Acidovorax spp./A. delafieldii
Dz13 NF#
Pseudomonas P. kilonensis/P. putida/P. fluorescens/P. jessenii/P. corrugata
Dz14 NF
Rhizobium
Dz20 NF
Aneurinibacillus A. aneurinilyticus/A. migulanus/A. danicus/A. terranovensis
R. radiobacter/R. larrymoorei/R. daejeonense/Agrobacterium spp.
* DNA sequences are listed in Appendix 4.3.
# NF Non-fumigated; F Fumigated soil
4.4. Discussion
Analyses of nematode populations in soils used for isolation of ginger rootassociated bacteria illustrated differences in the biodiversity of fumigated and nonfumigated soil. Increased levels of bacterivores were observed in the fumigated
plot; these nematodes have a short generation time and typically colonise
disturbed habitats (Stirling 2005). Omnivorous and predatory nematodes have a
longer life cycle and may take months to years to re-establish following soil
disturbance (Stirling 2005). Therefore it is not unusual that these nematodes were
detected in non-fumigated but not in fumigated soil. The presence of fungivores in
non-fumigated soil may have also indicated higher levels of fungi in this soil. The
increased diversity of nematode trophic groups would typically imply that the nonfumigated soil is a more healthy soil (Pattison et al. 2004). However, yield was
increased by 23% in the fumigated plot compared to the non-treated plot. As plant
92
parasitic nematodes were not detected in either plot and a low incidence of
Fusarium yellows was observed, other biological factors may have contributed to
the improved growth of ginger in response to soil fumigation.
Such biological
factors may have included a reduction in deleterious microorganisms that
negatively impact on plant growth or an increase in species that promote plant
growth (Martin and Bull 2002). In the present study, plating of rhizosphere soil
(collected in January 2006) on King’s B agar indicated Bacillus spp. (Bacillus
simplex/macroides) were increased in fumigated soil, while in non-fumigated soil,
bacteria typical of fluorescent pseudomonads were prevalent. Bacillus spp. with
enhanced biodegradation capabilities are among bacteria shown to be increased in
fumigated soils (Ibekwe et al. 2004; Warton et al. 2001). Certain strains of Bacillus
macroides have been reported to enhance plant growth by production of the
phytostimulator gibberellin (Joo et al. 2004; Joo et al. 2005). Strains of Bacillus
simplex have been reported to improve growth and nitrogen content of wheat and
reduce required application rates of synthetic fertiliser (Barniex et al. 2005).
Therefore these Bacillus strains were selected for inoculation of ginger plants.
Further analyses indicated that in rhizosphere soil collected in March (2006) and
plated on King’s B agar, white non-fluorescent bacteria were dominant in both
fumigated
and
non-fumigated
soil.
As
colonies
typical
of
fluorescent
Pseudomonas were not detected in rhizosphere soil at the later time, results
indicated that (in non-fumigated soil) fluorescent pseudomonads were more
prevalent in the ginger rhizosphere during the earlier stages of growth then
declined as the crop progressed. Similarly, Wong (1994) demonstrated that in the
wheat rhizosphere populations of fluorescent Pseudomonas species declined and
Bacillus species increased with progression through the growth cycle of this plant.
Such differences were associated with changes in root exudation with plant age.
In the current study, Pseudomonas spp. were recovered on nitrogen free media
from the rhizoplane samples of ginger roots collected in March (2006) in both
fumigated and non-fumigated soil. Thus while fluorescent Pseudomonas spp. may
not have been dominant in rhizosphere soil collected at this time, they were
associated with the ginger root surface. Other bacteria detected in rhizoplane
samples that grew on nitrogen free media included Acidovorax spp. and
93
Rhizobium/Agrobacterium.
Acidovorax species have been previously detected in
the rhizosphere and within plant tissues (Cirou et al. 2007; Harichova et al. 2006).
Certain strains of Acidovorax spp. have shown to be active in processes of
nitrogen fixation and bioremediation, although some members of this species are
plant pathogenic (Fegan 2007; Monferran et al. 2005; Nestler et al. 2007; Ohtsubo
et al. 2006). Members of the Rhizobium-Agrobacterium group have also been
detected in rhizosphere, rhizoplane and endophyte populations in a variety of
plants (Hallman et al. 2001; Reddy et al. 1997; Kennedy et al. 1997). Rhizobium
species may promote growth in non-legumes, as a result of production of
phytostimulatory hormones, or enhance resistance to disease by inducing systemic
resistance (Yanni et al. 1997; Hasky-Gunther et al. 1998; Hallmann et al. 2001;
Reitz et al. 2000; Antoun et al. 2004; Mabrouk et al. 2007). The utility of arbitrarily
primed PCR for producing a genetic fingerprint that may enable different bacteria
to be identified was demonstrated, although only a small number of isolates could
be analysed due to time limitations.
Such techniques that produce a genetic
fingerprint of bacteria may be also be useful for monitoring introduced strains. For
example, by employing a variety of primers Brousseau and colleagues (1993) used
arbitrarily primed PCR to distinguish introduced strains of insecticidal B.
thuringensis from indigenous strains.
Phosphate solubilizing ability has been demonstrated by many different Bacillus
species (de Fretis et al. 1997; Belimov et al. 1995; Toro et al. 1997). Phosphatesolubilizing activity was not detected in the heat resistant fraction (that selects for
endospore forming bacteria such as Bacillus) of ginger root-associated bacteria.
This may indicate that microorganisms other than Bacillus spp. solubilize
phosphate in association with the ginger root. As the reference strain B. coagulans
NCTC 10334 demonstrated in vitro phosphate solubilizing activity, this bacterium
was selected for future trials that included dual inoculation with Azospirillum
reference strains, to determine if a synergistic effect of the combination of bacteria
on the growth of ginger might occur.
94
4.5. Conclusion
In conclusion, culture-based analyses of bacteria associated with the ginger root
indicated that colonisation might be influenced by agronomic measures
(fumigation) and growth stage of the plant. In non-fumigated soil, populations of
fluorescent Pseudomonas were detected at high levels in the earlier stages of the
ginger growth cycle and declined as the crop progressed. Field isolates including
Bacillus F1 and Bacillus F2 were increased in the rhizosphere of ginger grown in
fumigated soil. On the basis of the above observations Bacillus F1 and Bacillus
F2, along with reference strains of Bacillus, Pseudomonas Azospirillum were
selected for assessment of growth promoting and/or biocontrol activities they might
have in ginger.
95
Appendix 4.1. Media used for the isolation of rhizosphere and rhizoplane
bacteria.
King’s Agar B (King et al. 1954).
King’s B agar was prepared from a premixed powder (Fluka). Per litre the medium
contained mixed peptone 20g, dipotassium hydrogen phosphate 1.5g, magnesium
sulphate 1.5g, agar 10g, glycerol 10ml, pH 7.2.
Congo Red Medium (Bastarrachea et al. 1988; Rodriguez-Caceres 1982;
Muthukumar et al. 2001).
Per litre the medium contained 5g KH2PO4, 0.2 g MgSO4.7H2O, 0.1g NaCl, 0.5g
yeast extract, 15mg FeCl3.6H2O, 5g DL-malic acid, 4.8g KOH and 20g agar, pH
7.0. Following sterilisation of the media, 37ug/ml of sterile aqueous Congo red was
added.
Pikovskaya (PVK) medium (El-Komy 2004; Johri et al. 1999).
Per litre the medium contained glucose 10g, Ca3(PO4)2 5g, (NH4)2SO4 0.5g, yeast
extract 0.5g, NaCl 0.2g, KCl 0.2g, MgSO4.7H20 0.1g; MnSO4.H20 0.002g,
FeSO4.H20 0.002g, agar 17g; pH7.0.
New Fabian broth (NFb) Media (Dobereiner 1995).
Per litre this media contained D,L-Malic acid 5g (Fluka), K2HPO4 0.5g,
MgSO4.7H2O 0.2g, NaCl 0.1g, CaCl2.2H2O 0.02g, minor element solution 2ml,
bromothymol blue 2ml (0.5% solution in 0.2M KOH; 50mg/10ml), FeCl2 10mg,
vitamin solution 1ml,
agar 15g.
So that iron and salts did not precipitate,
ingredients were added in the sequence listed.
Per 10ml, the minor element
solution contained CuSO4.H2O 4mg, ZnSo4.7H2O 1.2mg, Na2MoO4.2H2O 10mg
and MnSO4.H2O 15mg. Per 10ml the Vitamin Solution contained biotin 1mg and
pyridoxol-HCl 2mg; this solution was filter sterilised and added to the media after
autoclaving.
NBY (Nutrient Broth-Yeast Extract Medium: Vidaver 1967; Kim et al. 1997).
Overnight cultures of bacteria were prepared in sterile NBY, that contained per litre
nutrient broth 8g (Sigma-Aldrich), yeast extract 2g (Fluka), K2HPO4 2g, KH2PO4
96
0.5g, glucose 5g and MgSO4.7H2O 0.25g; 15g of agar was added for preparation
of plates. Glucose (Sigma-Aldrich) was added as a 10% filter sterilised solution
after autoclaving of the media.
Antibiotic preparation
Cycloheximide (Sigma-Aldrich) was dissolved in 100% ethanol (50 mg/ml) and
filter sterilised. Nystatin (Sigma-Aldrich) was dissolved at 50mg/ml in methanol.
Antibiotics were added to media after autoclaving and cooling.
97
Appendix 4.2. 16S rDNA amplification and sequencing and arbitrarily primed
PCR.
Table 17. Primers used for 16S rDNA analysis and arbitrarily primed PCR of field
isolates.
Primer
Code*
Primer Sequence (5’- 3’)
Reference
Primer Synthesis
27F
GAGAGTTTGATCCTGGCTCAG Girfoni et al. 1995;
Weisburg et al. 1991.
518F
CCAGCAGCCGCGGTAATACG
Lu et al. 2000; Lane et Invitrogen (South
al. 1985.
Australia) and SigmaGenosys (Australia).
926R
CCGTCAATTCCTTTGAGTTT
Schwieger and Tebbe Invitrogen (South
1998; Lane et al. 1985. Australia) and SigmaGenosys (Australia).
1492R
ACGGCTACCTTGTTACGACTT
Girfoni et al. 1995;
Weisburg et al. 1991.
Sigma-Genosys
(Australia).
OPT-08
GACCAATGCC
Gunarto et al. 1999.
Sigma-Genosys
(Australia).
Sigma-Genosys
(Australia).
* For 16S rDNA primers, numbers indicate relative position of primers along
the16S rRNA gene, according to numbering in E. coli.
98
Appendix 4.3. 16S rDNA sequences (5’- 3’) of bacteria isolated from ginger
roots.
>F1_U1
AACTGGGGAACTTGAGTGCAGAAGAGGAAAGATGGAATTCCAAGTGTAGCGG
TGAAATGCGTAGAGATTTGGAGGAACACCAGTGGCGAAGGCGACTTTCTGGT
CTGTAACTGACACTGAGGCGCGAAAGCGTGGGGAGCAAACAGGATTAGATAC
CCTGGTAGTCCACGCCGTAAACGATGAGTGCTAAGTGTTAGAGGGTTTCCGC
CCTTTAGTGCTGCAGCTAACGCATTAAGCACTCCGCCTGGGGAGTACGGCCG
CAAGGCTGAAACTCAAAGGAATTGACGGGGGCCCGCACAAGCGGTGGAGCA
TGTGGTTTAATTCGAAGCAACGCGAAGAACCTTACCAGGTCTTGACATCCTCT
GACAACCCTAGAGATAGGGCTTTCCCCTTCGGGGGACAGAGTGACAGGTGG
TGCATGGTTGTC
>F1_U3
ACGGGAGGCAGCAGTAGGGAATCTTCCGCAATGGACGAAAGTCTGACGGAG
CAACGCCGCGTGAACGAAGAAGGCCTTCGGGTCGTAAAGTTCTGTTGTTAGG
GAAGAACAAGTACCAGAGTAACTGCTGGTACCTTGACGGTACCTAACCAGAA
AGCCACGGCTAACTACGTGCCAGCAGCCGCGGTAATACGTAGGTGGCAAGC
GTTGTCCGGAATTATTGGGCGTAAAGCGCGCGCAGGTGGTTCCTTAAGTCTG
ATGTGAAAGCCCACGGCTCAACCGTGGAGGGTCATTGGAAACTGGGGAACTT
GAGTGCAGAAGAGGAAAGTGGAATTCCAAGTGTAGCGGTGAAATGCGTAGAG
ATTTGGAGGAACACCAGTGGCGAAGGCGACTTTCTGGTCTGTAACTGACACT
GAGGCGCGAAAGCGTGGGGAGCAAACAGGATTAGATATCCCTGGTAGTCCA
CGCCGTAAACGATGAGTGCTAAGTGTTAGA
>F2_U1
TGGAATTCCAAGTGTAGCGGTGAAATGCGTAGAGATTTGGAGGAACACCAGT
GGCGAAGGCGACTTTCTGGTCTGTAACTGACACTGAGGCGCGAAAGCGTGG
GGAGCAAACAGGATTAGATACCCTGGTAGTCCACGCCGTAAACGATGAGTGC
TAAGTGTTAGAGGGTTTCCGCCCTTTAGTGCTGCAGCTAACGCATTAAGCACT
CCGCCTGGGGAGTACGGCCGCAAGGCTGAAACTCAAAGGAATTGACGGGGG
CCCGCACAAGCGGTGGAGCATGTGGTTTAATTCGAAGCAACGCGAAGAACCT
TACCAGGTCTTGACATCCTCTGACAACCCTAGAGATAGGGCTTTCCCCTTCGG
GGGACAGAGTGA
>F2_U3
TGTTGTTAGGGAAGAACAAGTACCAGAGTAACTGCTGGTACCTTGACGGTAC
CTAACCAGAAAGCCACGGCTAACTACGTGCCAGCAGCCGCGGTAATACGTAG
GTGGCAAGCGTTGTCCGGAATTATTGGGCGTAAAGCGCGCGCAGGTGGTTC
CTTAAGTCTGATGTGAAAGCCCACGGCTCAACCGTGGAGGGTCATTGGAAAC
TGGGGAACTTGAGTGCAGAAGAGGAAAGTGGAATTCCAAGTGTAGCGGTGAA
ATGCGTAGAGATTTGGAGGAACACCAGTGGCGAAGGCGACTTTCTGGTCTGT
AACTGACACTGAGGCGCGAAAGCGTGGGGAGC
>Dz1_U1
AGGGTGGTGGAATTTCCTGTGTAGCGGTGAAATGCGTAGATATAGGAAGGAA
CACCAGTGGCGAAGGCGACCACCTGGACTGATACTGACACTGAGGTGCGAA
AGCGTGGGGAGCAAACAGGATTAGATACCCTGGTAGTCCACGCCGTAAACGA
TGTCAACTAGCCGTTGGGAGCCTTGAGCTCTTAGTGGCGCAGCTAACGCATT
AAGTTGACCGCCTGGGGAGTACGGCCGCAAGGTTAAAACTCAAATGAATTGA
CGGGGGCCCGCACAAGCGGTGGAGCATGTGGTTTAATTCGAAGCAACGCGA
99
AGAACCTTACCAGGCCTTGACATCCAATGAACTTTCCAGAGATGGATTGGTGC
CTTCGGGAACATTGAGACAGGTGCTGCATGGCTGTCGTCAGCTCGTGTCGTG
AGATGTTGGGTTAAGTCCCGTAACGAGCGCAACCCTTGTCCTTAGTTACCA
>Dz1_U3
ACGGGAGGCAGCAGTGGGGAATATTGGACAATGGGCGAAAGCCTGATCCAG
CCATGCCGCGTGTGTGAAGAAGGTCTTCGGATTGTAAAGCACTTTAAGTTGG
GAGGAAGGGTTGTAGATTAATACTCTGCAATTTTGACGTTACCGACAGAATAA
GCACCGGCTAACTCTGTGCCAGCAGCCGCGGTAATACAGAGGGTGCAAGCG
TTAATCGGAATTACTGGGCGTAAAGCGCGCGTAGGTGGTTTGTTAAGTTGGAT
GTGAAATCCCCGGGCTCAACCTGGGAACTGCATCCAAAACTGGCAAGCTAGA
GTATGGTAGAGGGTGGTGGAATTTCCTGTGTAGCGGTGAAATGCGTAGATAT
AGGAAGGAACACCAGTGGCGAAGGCGACCACCTGGACTGATACTGACACTG
AGGTGCGAAAGCGTGGGGAGCAAACAGGATTAGATACCCTGGTAGTCCACG
CCGTAAACGATGTC
>Dz3_U1
ACTGCATTTGTGACTGCATAGCTAGAGTACGGCAGAGGGGGATGGAATTCCG
CGTGTAGCAGTGAAATGCGTAGATATGCGGAGGAACACCGATGGCGAAGGC
AATCCCCTGGGCCTGTACTGACGCTCATGCACGAAAGCGTGGGGAGCAAACA
GGATTAGATACCCTGGTAGTCCACGCCCTAAACGATGTCAACTGGTTGTTGG
GTCTTCACTGACTCAGTAACGAAGCTAACGCGTGAAGTTGACCGCCTGGGGA
GTACGGCCGCAAGGTTGAAACTCAAAGGAATTGACGGGGACCCGCACAAGC
GGTGGATGATGTGGTTTAATTCGATGCAACGCGAAAAACCTTACCCACCTTTG
ACATGTACGGAATCCTTTAGAGATAGAGGAGTGCTC
>Dz3_U3
TGGGGTCCATGACGGTACCGTAAGAATAAGCACCGGCTAACTACGTGCCAGC
AGCCGCGGTAATACGTAGGGTGCGAGCGTTAATCGGAATTACTGGGCGTAAA
GCGTGCGCAGGCGGTTATGTAAGACAGATGTGAAATCCCCGGGCTCAACCTG
GGAACTGCATTTGTGACTGCATAGCTAGAGTACGGCAGAGGGGGATGGAATT
CCGCGTGTAGCAGTGAAATGCGTAGATATGCGGAGGAACACCGATGGCGAA
GGCAATCCCCTGGGCCTGTACTGACGCTCATGCACGAAAGCGTGGGGAGCA
AACAGGATTAGATACCCTGGTAGTCCACGCCCTAAACGATGTCAACTGGTTGT
TGGGTCTTCACTG
>Dz4_U1
GATGTGAAATCCCCGGGCTCAACCTGGGAACTGCATTTGTGACTGCATAGCT
AGAGTACGGCAGAGGGGGATGGAATTCCGCGTGTAGCAGTGAAATGCGTAG
ATATGCGGAGGAACACCGATGGCGAAGGCAATCCCCTGGGCCTGTACTGAC
GCTCATGCACGAAAGCGTGGGGAGCAAACAGGATTAGATACCCTGGTAGTCC
ACGCCCTAAACGATGTCAACTGGTTGTTGGGTCTTCACTGACTCAGTAACGAA
GCTAACGCGTGAAGTTGACCGCCTGGGGAGTACGGCCGCAAGGTTGAAACT
CAAAGGAATTGACGGGGACCCGCACAAGCGGTGGATGATGTGGTTTAATTCG
ATGCAACGCGAAAAACCTTACCCACCTTTGACATGTACGGAATCCTTTAGAGA
TAGAGGAGTGCTCGAAAGAGAGCCGTAACACAGGTGCTGCATGGCTGTCGTC
AGCTCGTGTCGTG
Dz4_U3
GGGGTCCATGACGGTACCGTAAGAATAAGCACCGGCTAACTACGTGCCAGCA
GCCGCGGTAATACGTAGGGTGCGAGCGTTAATCGGAATTACTGGGCGTAAAG
CGTGCGCAGGCGGTTATGTAAGACAGATGTGAAATCCCCGGGCTCAACCTGG
GAACTGCATTTGTGACTGCATAGCTAGAGTACGGCAGAGGGGGATGGAATTC
CGCGTGTAGCAGTGAAATGCGTAGATATGCGGAGGAACACCGATGGCGAAG
GCAATCCCCTGGGCCTGTACTGACGCTCATGCACGAAAGCGTGGGGAGCAA
100
ACAGGATTAGATACCCCTGGTAGTCCACGCCCTAAACGATGTCAACTGGTTGT
T
>Dz5
TTAACCTAATACGTTAGTGTTTTGACGTTACCGACAGAATAAGCACCGGCTAA
CTNTGTGCCAGCAGCCGCGGTAATACAGAGGGTGCAAGCGTTAATCGGAATT
ACTGGGCGTAAAGCGCGCGTAGGTGGTTTGTTAAGTTGGATGTGAAAGCCCC
GGGCTCAACCTGGGAACTGCATTCAAAACTGACAAGCTAGAGTATGGTAGAG
GGTGGTNGAATTTCCTGTGTAGCGGTGAAATGCGTAGATATAGGAAGGAACA
CCAGTGGCGAAGGCGACCACCTGGACTGATACTGACACTGAGGTGCGAAAG
CGTGGGGAGCAAACAGGATTAGATACCCTGGTAGTCCACGCCG
>Dz9_U1
ATGTAAGACAGATGTGAAATCCCCGGGCTCAACCTGGGAACTGCATTTGTGA
CTGCATAGCTAGAGTACGGCAGAGGGGGATGGAATTCCGCGTGTAGCAGTG
AAATGCGTAGATATGCGGAGGAACACCGATGGCGAAGGCAATCCCCTGGGC
CTGTACTGACGCTCATGCACGAAAGCGTGGGGAGCAAACAGGATTAGATACC
CTGGTAGTCCACGCCCTAAACGATGTCAACTGGTTGTTGGGTCTTCACTGACT
CAGTAACGAAGCTAACGCGTGAAGTTGACCGCCTGGGGAGTACGGCCGCAA
GGTTGAAACTCAAAGGAATTGACGGGGACCCGCACAAGCGGTGGATGATGT
GGTTTAATTCGATGCAACGCGAAAAACCTTACCCACCTTTGACATGTACGGAA
TCCTTTAGAGATAGAGGAGTGCTCNAAAGAGAGCCGTAACACAGGTGCTGCA
TGGCTGTCGTCAGCTCGTGTCGTGAGATGT
>Dz9_U3
GTAAGAATAAGCACCGGCTAACTACGTGCCAGCAGCCGCGGTAATACGTAGG
GTGCGAGCGTTAATCGGAATTACTGGGCGTAAAGCGTGCGCAGGCGGTTATG
TAAGACAGATGTGAAATCCCCGGGCTCAACCTGGGAACTGCATTTGTGACTG
CATAGCTAGAGTACGGCAGAGGGGGATGGAATTCCGCGTGTAGCAGTGAAAT
GCGTAGATATGCGGAGGAACACCGATGGCGAAGGCAATCCCCTGGGCCTGT
ACTGACGCTCATGCACGAAAGCGTGGGGAGCAAACAGGATTAGATACCCTGG
TAGTCCACGCCCTAAACGATGTCAACTGGTTGTTGGGTCTTCAC
>Dz13_U1
TGTTAANTTGGATGTGAAAGCCCCGGGCTCAACCTGGGAACTGCATTCAAAA
CTGACAAGCTAGAGTATGGTAGAGGGTGGTGGAATTTCCTGTGTAGCGGTGA
AATGCGTAGATATAGGAAGGAACACCAGTGGCGAAGGCGACCACCTGGACT
GATACTGACACTGAGGTGCGAAAGCGTGGGGAGCAAACAGGATTAGATACCC
TGGTAGTCCACGCCGTAAACGATGTCAACTAGCCGTTGGGAGCCTTGAGCTC
TTAGTGGCGCAGCTAACGCATTAAGTTGACCGCCTGGGGAGTACGGCCGCAA
GGTTAAAACTCAAATGAATTGACGGGGGCCCGCACAAGCGGTGGAGCATGTG
GTTTAATTCGAAGCAACNCGAAGAACCTTACCAGGCCTTGACATCCAATGAAC
TTTCCAGAGATGGATTGGTGCCTTCGGGAACATTGAGACAGGTGCTGCATGG
CTGTCGTCAGCTCGTGTCGTGAGATGTTGGGT
>Dz13_U3
GCCAGCAGCCGCGGTAATACAGAGGGTGCAAGCGTTAATCGGAATTACTGG
GCGTAAAGCGCGCGTAGGTGGTTTGTTAAGTTGGATGTGAAAGCCCCGGGCT
CAACCTGGGAACTGCATTCAAAACTGACAAGCTAGAGTATGGTAGAGGGTGG
TGGAATTTCCTGTGTAGCGGTGAAATGCGTAGATATAGGAAGGAACACCAGT
GGCGAAGGCGACCACCTGGACTGATACTGACACTGAGGTGCGAAAGCGTGG
GGAGCAAACAGGATTAGATACCCTGGTAGT
>Dz14_U1
101
CGGATATTTAAGTCAGGGGTGAAATCCCAGAGCTCAACTCTGGAACTGCCTTT
GATACTGGGTATCTTGAGTATGGAAGAGGTAAGTGGAATTGCGAGTGTAGAG
GTGAAATTCGTAGATATTCGCAGGAACACCAGTGGCGAAGGCGGCTTACTGG
TCCATTACTGACGCTGAGGTGCGAAAGCGTGGGGAGCAAACAGGATTAGATA
CCCTGGTAGTCCACGCCGTAAACGATGAATGTTAGCCGTCGGGCAGTATACT
GTTCGGTGGCGCAGCTAACGCATTAAACATTCCGCCTGGGGAGTACGGTCGC
AAGATTAAAACTCAAAGGAATTGACGGGGGCCCGCACAAGCGGTGGAGCATG
TGGTTTAATTCGAAGCAACGCGCAGAACCTTACCAGCTCTTGACATTCGGGGT
ATGGTCATTGGAGACGATGACCTTCAGTTCGGCTGGCCCTAGAACAGGTGCT
GCATGGCTG
>Dz14_U3
CCATGCCGCGTGAGTGATGAAGGCCTTAGGGTTGTAAAGCTCTTTCACCGAT
GAAGATAATGACGGTAGTCGGAGAAGAAGCCCCGGCTAACTTCGTGCCAGCA
GCCGCGGTAATACGAAGGGGGCTAGCGTTGTTCGGAATTACTGGGCGTAAA
GCGCACGTAGGCGGATATTTAAGTCAGGGGTGAAATCCCAGAGCTCAACTCT
GGAACTGCCTTTGATACTGGGTATCTTGAGTATGGAAGAGGTAAGTGGAATTG
CGAGTGTAGAGGTGAAATTCGTAGATATTCGCAGGAACACCAGTGGCGAAGG
CGGCTTACTGGTCCATTACTGACGCTGAGGTGCGAAAGCGTGGGGAGCAAA
CAGGATTAGATACCCTGGTAGTCCACGCCGTAAACGATGAATGTTAGCCGTC
GGGCAGTATACTGTTCGGTG
>Dz20_U1
AGATGTGGAGGAACACCCGTGGCGAAGGCGGCTCTCTGGCCTGTAACTGAC
GCTGAGGCGCNAAAGCGTGGGGAGCGAACAGGATTAGATACCCTGGTAGTC
CACGCCGTAAACGTTGAGTGCTAGGTGTTGGGGACTCCAATCCTCANTGCCG
CAGCTAACGCAATAAGCACTCCGCCTGGGGAGTACGGCCGCAAGGCTGAAA
CTCAAAGGAATTGACGGGGACCCNCACAAGCGGTGGAGCATGTGGTTTAATT
CGAAGCAACNCGAA
>Dz20_U3
TACCAGGGGTATCCTAATCCCTGNTTCGCTCCCCACGCTTTCGCGCCTCAGC
GTCAGTTACAGGCCAGAGAGCCGCCTTCGCCACGGGTGTTCCTCCACATCTC
TACGCNTTTCACCGCTACACGTGGAATTCCGNTCTCCTCTCCTGCACTCAAGC
TTCCCAGTTTCAAGTGGCCCTCCACGGTTGAGCCGTGGGCTTTCACACCTGA
CTTAAGAAGCCGCCTGCGCGCGCTTTACGCCCAATAATTCCGGACAACGCTT
GCCCCCTACNTATTACCGCGGCTGCTGGCACGTAGTTAGCCGGGGCTTTCTC
GTTAGGTACCGTCAGACCGGGAGGTCATCCCGG
102
Table 18. RDPII-Classifier Analysis of 16S rDNA sequences of field isolates.
F1_U3 Root[100%] Bacteria[100%] Firmicutes[100%] Bacilli[100%]
Bacillales[100%] Bacillaceae[100%] Bacillus[100%]
F2 Root[100%] Bacteria[100%] Firmicutes[100%] Bacilli[100%]
Bacillales[100%] Bacillaceae[100%] Bacillus[100%]
Dz1 Root[100%] Bacteria[100%] Proteobacteria[100%]
Gammaproteobacteria[100%] Pseudomonadales[100%]
Pseudomonadaceae[100%] Pseudomonas[100%]
Dz3 Root[100%] Bacteria[100%] Proteobacteria[100%]
Betaproteobacteria[100%] Burkholderiales[100%] Comamonadaceae[100%]
Acidovorax[100%]
Dz4 Root[100%] Bacteria[100%] Proteobacteria[100%]
Betaproteobacteria[100%] Burkholderiales[100%] Comamonadaceae[100%]
Acidovorax[100%]
Dz5 Root[100%] Bacteria[100%] Proteobacteria[100%]
Gammaproteobacteria[100%] Pseudomonadales[100%]
Pseudomonadaceae[100%] Pseudomonas[100%]
Dz9 Root[100%] Bacteria[100%] Proteobacteria[100%]
Betaproteobacteria[100%] Burkholderiales[100%] Comamonadaceae[100%]
Acidovorax[100%]
Dz13_U1 Root[100%] Bacteria[100%] Proteobacteria[100%]
Gammaproteobacteria[100%] Pseudomonadales[100%]
Pseudomonadaceae[100%] Pseudomonas[100%]
Dz14_U1 Root[100%] Bacteria[100%] Proteobacteria[100%]
Alphaproteobacteria[100%] Rhizobiales[100%] Rhizobiaceae[100%]
Rhizobium[100%]
Dz20_U1 Root[100%] Bacteria[100%] Firmicutes[100%] Bacilli[100%]
Bacillales[100%] Paenibacillaceae[100%] Aneurinibacillus[100%]
103
Chapter 5. Evaluation of plant growth promoting ability of selected bacteria.
5.1. Introduction
The application of plant growth promoting bacteria (PBPB) has resulted in increased
plant growth and reduced requirements for synthetic fertilisers in crop production
(Bashan et al. 2004; Dobbelaere et al. 2003; Esitken et al. 2006; Jacobsen et al. 2004;
Mercado-Blanco and Bakker 2007; Cocking 2006). The activity of many PGPB may be
more pronounced or only occur under reduced levels of fertiliser application (Dobbelaere
et al. 2001; Okon and Labandera-Gonzalez 1994).
While inconsistencies in the
performance of PGPB in field conditions is considered to have limited their widespread
use in commercial applications, combinations of bacterial strains that have different
modes of action and optimisation of application methods may improve the plant growth
response following the introduction of PGPB (Kennedy et al. 2004; Kloepper et al. 2004;
de Boer et al. 1999).
Methods used to introduce PGPB to plants have included application to seed material,
soil and/or plant roots (Akkopru and Demir 2005; Munoz-Rojas and Caballero-Mellado
2003; Zehnder et al. 2001; Roncato-Maccari et al. 2003; Jetiyanon et al. 2003; Rapauch
and Kloepper 1998; Njoloma et al. 2006; Kadouri et al. 2003). The resultant plant growth
response may be influenced by concentration of viable cells, frequency of application and
formulation of introduced bacteria (Bashan 1998; Fages 1992; Schisler et al. 2004;
Moenne-Loccoz et al. 1999; Bressan and Borges 2004). The establishment of minimal
threshold levels of introduced PGPB may be required to induce a consistent plant growth
response, while too high a concentration of introduced bacteria may have a negative
effect on plant growth (Dobbelaere et al. 1999; Lucy et al. 2004; Okon and LabanderaGonzalez 1994; Landa et al. 2003; Raaijmakers and Weller 1998).
Due to practical advantages, PGPB have been frequently applied as a seed treatment
(Bashan 1986b). Sticker agents such as methylcellulose and gum arabic have been
used to promote adhesion of bacteria to seed surfaces (de Souza et al. 2003; Siqqiqui et
al. 2001; Raaijmakers and Weller 2001; Deaker et al. 2004). Bacteria have also been
104
formulated into preparations, such as powders and alginate micro-beads for application
as a seed treatment (Bora et al. 2004; Bashan et al. 2002). The re-application of PGPB
to the soil during the growth plant cycle may aid in the maintenance of threshold
populations of bacteria required to induce a consistent and continuing plant growth
response (Kloepper et al. 2004; Zhang et al. 2004; Martin and Bull 2002).
Bacteria have been applied to soil as a liquid suspension or by use of a carrier material,
such as alginate or vermiculite (Nautiyal et al. 2002; Bashan 1986a; Landa et al. 2003;
Bapat and Shah 2000; Jetiyanon et al. 2003; Graham-Weiss et al. 1987).
When
compared to the use of a liquid suspension of bacteria that must be applied within hours
of preparation to maintain viability, the use of a carrier such as alginate beads may
provide a more practical and effective method of production and application, as microbial
cells may be stable within the carrier for many years and these cells are continuously
released as the alginate beads degrade (Bashan 1986a; Bashan and Gonzalez 1998;
van Elsas et al. 1992; Chang and Park 2000; Zohar-Perez et al. 2005; Young et al.
2006).
Given the establishment of optimal concentrations of introduced bacteria (106-107 CFU
per plant or seed), a negative growth response to introduction of Azospirillum spp. has
not been reported and this bacterium may promote plant growth in a non-specific way
(Bashan et al. 2004; Okon and Labandera-Gonzalez 1994; Dobbelaere et al. 1999;
Dobbelaere et al. 2001; Saleena et al. 2002; Lin et al. 1983; Larraburu et al. 2007;
Somers and Vanderleyden 2004). In contrast, plant or cultivar dependant responses
may occur following the introduction of certain strains of other bacteria, such as
Pseudomonas spp. and Bacillus spp., where a positive effect on plant growth may be
seen in one type of plant and a negative or lack of response may be produced in a
different plant type (Broadbent et al. 1977). A wider range of concentrations of cells of
Bacillus spp. and Pseudomonas spp. (105-1011 CFU/ml) has been used for plant/soil
inoculation (Siddiqui et al. 2001; Akkopru and Demir 2005; Jetiyanon and Kloepper 2002;
Jetiyanon et al. 2003; Broadbent et al. 1977; Joo et al. 2004; Domenech et al. 2006;
Ahmed 2003).
105
Considering the above information, in this study various bacterial strains and application
methods were assessed as few reports have described the use of beneficial bacteria to
improve the growth of ginger.
The study of ginger grown from seed pieces, the
conventional planting material used in commercial production, is restricted by seasonal
conditions that determine planting time. In the Sunshine Coast region, short day length
and cooler temperatures cause a senescent period of ginger in July-August (Groszmann
1954). Following senescence, harvested ginger rhizomes may be cut into seed pieces
for replanting from August to October annually (Whiley 1974). Therefore prior to testing
of bacterial treatments in ginger grown from seed pieces, wheat and then ginger tissue
culture plants were used to assess the efficacy of combinations of bacteria and different
application methods. Wheat was chosen as an indicator plant as the use of PGPB in this
model system has been extensively documented and effects of treatments can be
determined within weeks (Lin et al. 1983; Bashan et al. 1987). Micropropagated ginger
plants are used to establish sites that provide industry with a source of Foz free planting
material, in order to reduce losses caused by seed-borne propagules of this pathogen.
First generation tissue culture plants have a much smaller rhizome than plants grown
from seed pieces, and produce relatively higher amounts of roots and shoots (vegetative
growth) (Smith and Hamill 1996).
Rhizomes of first generation tissue culture plants
selected for replanting (use as seed pieces), must be of sufficient size in order to
produce a second-generation plant that has a large rhizome (comparable to
conventionally propagated plants), otherwise vegetative growth and inferior rhizome size
persists in second generation plants (Hamill, personal communication 2006).
The
identification of measures which improve rhizome growth in micropropagated ginger
plants may improve productivity and result in increased uptake of this source of clean
planting material in the ginger industry.
The most effective bacterial treatments in ginger tissue culture plants and further
combinations of these bacteria were then tested for their ability to promote the growth of
ginger grown from seed pieces. An additional trial assessing the efficacy of alginate
beads as a carrier material for the introduction of the type strain A. brasilense Sp7 in
tissue cultured ginger plantlets was also undertaken.
106
5.2. Materials and Methods
5.2.a. Preparation of bacterial inoculants.
Bacterial cultures (described in Chapter 4) were streaked onto nutrient agar from a
glycerol stock (-80oC) at least every four weeks. Cultures were incubated at 28oC to
30oC, with the exception of B. coagulans NCTC 10334 that was incubated at 37oC.
A single colony from a nutrient agar plate was used to inoculate 3ml of NBY* broth (NBY
broth amended with 0.1mM tryptophan, Appendix 5.1; Prinsen et al. 1993; Kim et al.
1997). This starter culture was grown with shaking (~100rpm) overnight and then 200µL
was used to inoculate 50 ml of NBY* broth in a 250ml Erlenmeyer flask plugged with
cotton wool. This culture was grown with shaking (-80rpm) overnight. The culture was
collected into a 50ml tube and centrifuged at 3000rpm for 10 minutes (Pengnoo et al.
2006). The supernatant was discarded, the bacterial pellet was resuspended in 0.1 x
phosphate buffered saline (PBS, Appendix 5.1) and then the microbial suspension was
centrifuged as above (i.e. bacterial cells were washed).
The pellet was then
resuspended in 25 ml of PBS, potassium-phosphate buffer or water as indicated below
(Jetiyanon and Kloepper 2002; Dobbelaere et al. 2002; Thirup et al. 2001; Kadouri et al.
2003; Bashan et al. 2006). Bacteria were resuspended in buffer to avoid subjecting the
cells to hypo-osmotic stress which occurs when the extracellular concentration of solutes
is less than within the cell cytoplasm; under such conditions there is a tendency for water
to move into the cell (causing swelling) and for an efflux of solutes to occur (Sleator and
Hill 2001; Welsh 2000).
To prepare a bacterial suspension with a defined concentration of cells (CFU/ml), initially
a standard curve was produced, for the optical density at 600nm (OD600) of ten fold serial
dilutions of each strain of bacteria and viable numbers of cells of each dilution plated on
nutrient agar (Landa et al. 2004). Following this, measurement of OD600 was used to
determine the concentration of cells in bacterial suspensions. Combinations of bacteria
were prepared by mixing bacterial suspensions at 1:1 or 1:1:1:1 ratio as appropriate
(Dekkers et al. 2000; Landa et al. 2002).
107
5.2.b. Plants and growth conditions.
All plants were grown in peat:sand (50:50) that had been steam pasteurised at 60oC for
45 minutes. This results in the elimination of plant pathogenic organisms from the soil but
the competitive potential of the indigenous microflora is retained (Broadbent et al. 1971).
Ginger cv. Canton tissue culture plants were obtained from Sharon Hamill, Department of
Primary Industries and Fisheries, Maroochy Research Station).
These plants were
generated via shoot tip culture and grown in axenic conditions in Murashige and Skoog
medium (Murashige and Skoog 1962) for 4-5 weeks as described by Smith and Hamill
(1996). For ex vitro acclimatisation, agar was gently washed from the roots with tap
water in a shaded area; plants were transplanted into a vermiculite/perlite mix and grown
under a humidifying cover for seven weeks. Fully acclimatised plants were used in all
experiments. As first generation tissue cultured plantlets have a very small rhizome, in
order to be able to detect a growth response, plants were grown for a minimum of three
months following application of bacterial treatments as described below.
In all experiments synthetic fertilisers were applied at minimal rates, at approximately half
of the recommended rate or at the onset of symptoms of nutrient deficiency and then to
prevent nutrient deficiency.
Pale green leaf colour and yellowing of leaf tips were
considered to indicate nitrogen and potassium deficiencies respectively (Asher and Lee
1975).
At completion of trials plants were destructively harvested and measured. Plants were
dried at 60oC for approximately one week for determination of dry weights. Surface area
was measured with a Li-Cor LI-3100 Area Meter (Lincoln Nebraska, USA).
5.2.c. Experimental design.
Pot trials were set up in a randomised complete block design on a single bench, where
each block contained one replicate from each treatment group.
The software program
Statistical Package for the Social Sciences (SPSS) was used for the following statistical
108
analyses. The Levene’s test was used to assess the homogeneity of variance. Two-way
analysis of variance (ANOVA) was used to assess the effect on plant growth, of
application method for the introduction of different bacteria (α=0.05). Where a single
method of bacterial application was used one-way ANOVA was used to determine if
there were statistically significant differences between the means of different treatment
groups (α=0.05). The Least Significant Difference test as described by Fischer was used
to make multiple comparisons between the means of treatment groups (LSD, p<0.05)
(Goulden 1939).
Once levels of inherent variability were determined in initial trials with ginger tissue
culture plants, for the seed piece trial and subsequent tissue culture trial, the sample size
(n) required to detect a 10% increase in plant growth was calculated iteratively (i.e. by
testing different values of n) as described by Zar (1984), using the formula n = s2/σ2(tα,ν +
tβ(1),ν)2, where s is the standard deviation of rhizome weight (determined in previous
trials), σ is the size of the increase to be detected, α (set at 0.05) is the probability of a
type I error, β (set at 0.20) is the probability of a type II error and ν represents degrees
of freedom.
5.2.1. Wheat as a model system for testing efficacy of bacterial inoculants in
promoting plant growth.
5.2.1.a. Application of bacteria to wheat seed.
Wheat seed (Triticum aestivum cv. Janz) was surface sterilised as described by
Dobbelaere et al. (1999) and modified to include preliminary rinses in sterile milli-Q water
to reduce indigenous microbial contaminants. Briefly, the seed was rinsed three times in
sterile milli-Q (s.m.) water, immersed momentarily in seventy percent ethanol, rinsed
three times in s.m. water, then soaked in one percent sodium hypochlorite for five
minutes and rinsed four times in s.m. water.
Bacteria were applied to wheat as a seed treatment only or as a seed treatment followed
by soil drenching as described in Table 19. The use of 1% methylcellulose as a sticker
agent (Raaijmakers and Weller 1998; Rapauch and Kloepper 1998; de Souza et al.
109
2003) in the bacterial seed treatment was observed to be inhibitory to germination of this
wheat seed variety and was therefore not used further. For all applications in this trial,
bacteria were resuspended in 0.1 x PBS (Appendix 5.1; Jetiyanon et al. 2003; Thirup et
al. 2001; Dobbelaere et al. 2002; Jetiyanon and Kloepper 2002; Bashan 1986a). The
seed was soaked in either water or 0.1 x PBS (two different controls) or a bacterial
suspension (4ml per forty seeds) for 30 minutes and dried under sterile air for 30 minutes
(Thirup et al. 2001; Dekkers et al. 2000).
Seeds were germinated in Petri dishes
between sheets of moistened sterile filter paper for forty-eight hours in the dark
(Dobbelaere et al. 2002).
Table 19. Treatments and application methods used in the wheat trial.
Bacteria Applied
Seed Treatment#
(CFU/ml)
1
Water
-
1
Buffer (0.1 x PBS)
-
1
B. subtilis A13
1 x 108
2
B. subtilis A13
1 x 108
1
B. coagulans NCTC 10334
1 x 108
2
B. coagulans NCTC 10334
1 x 108
1
P. putida KT2442
1 x 108
2
P. putida KT2442
1 x 108
App.
Method*
*
#
Application Method 1: Seed treatment is the only method of bacteria application.
Application Method 2: Seed treatment plus soil drench.
Concentration of cells applied in the seed treatment (colony forming units/ml).
5.2.1.b. Application of bacterial drenches and wheat growth conditions.
Germinated seeds were planted in 7.5mm diameter pots that contained 190g of
pasteurised peat:sand.
The bacterial suspension was applied at planting (106 CFU/ml,
25ml per pot) as a soil drench and at weekly intervals as appropriate (Table 19); two
different control treatments received either water or buffer at this time (Bashan 1986a;
Kadouri et al. 2003; Jetiyanon and Kloepper 2002; Bressan and Borges 2004). Plants
110
were grown at 25oC day/15oC night with a 14 hour photoperiod and 85% humidity in a
growth chamber (Lindne and May; Kadouri et al. 2003). Fifteen plants per treatment
were used (three plants per pot, five pots per treatment). Plants were watered twice
weekly with deionized water (d. water) as required.
After three weeks plants were
sampled. Plant roots were washed in d. water to remove adhering soil and blotted dry.
Fresh weight and surface area were measured for the complete plant, shoots and roots.
Plant height was also measured.
5.2.2. Effect of application method on bacterial induced growth response in
ginger tissue culture plants (ginger tissue culture trial I).
Initial ginger tissue culture trials were undertaken in a growth cabinet, to prevent
senescence caused by reduced temperature and shortened day length at the time the
trial was undertaken. Growth cabinet settings were based on average temperatures and
humidity in regional conditions in September-October that occur during the early growth
of ginger (http://bom.gov.au). A variety of application regimes and concentrations of
bacterial cells were assessed for selected reference strains B. subtilis A13,
B. coagulans NCTC 10334 and P. putida KT2442.
Bacteria were resuspended in 0.1 X PBS (Jetiyanon et al. 2003; Thirup et al. 2001;
Dobbelaere et al. 2002; Jetiyanon and Kloepper 2002; Bashan 1986a) and were applied
by three different methods that included: 1) Plant roots were dipped into the bacterial
suspension for 30 minutes and 50 ml of bacterial suspension was also applied to the
soil at planting (root dip and drench); 2) 50 ml of bacterial suspension was applied to the
soil at planting (drench only) and; 3) Plant roots were dipped into a bacterial suspension
for 30 minutes prior to planting (root dip only) (Bressen and Borges 2004; Akkopru and
Demir 2005; Munoz-Rajas and Caballero-Mellado 2002; Suman et al. 2005; Jetiyanon
and Kloepper 2002; Zhang et al. 2004; Siddiqui and Shakat 2002).
Different
concentrations of bacteria were assessed as indicated in Table 20.1. For methods (1)
and (2) bacterial drenches were reapplied fortnightly for the first month and then every
four weeks for the remainder of the trial.
111
Seven plants of uniform size (~10cm in height) were used for each treatment group.
Plants were grown in 125mm diameter pots (planted 21/06/06) that contained one litre
of pasteurised peat:sand.
Plants were maintained in a growth cabinet (Lindne and May) at 28oC day/15oC night
with a 16-hour photoperiod and 48-70% humidity. Plants were watered three times per
week with d.water. Thrive® general purpose fertiliser (25ml, 1.0g/L) was added to each
pot at 2, 28, 50, 64, 78 days after planting (DAP). Sulphate of potash (0.4g, Searles®)
was added to each pot 5 and 50 DAP. After 12 weeks plants were sampled and fresh
weight of shoots, roots and rhizomes were determined. Surface area and dry weight of
shoots, the number of shoots per plant and the width of the stem at the base of the plant
were measured.
Figure 11. Acclimatised tissue cultured ginger plants that had been maintained for
several weeks in a growth cabinet.
112
Table 20.1. Treatments applied in ginger tissue culture trial 1.
No.
Treatment
Conc.
Bacteria
(CFU/ml)*
1
Water
-
Soil drench
2
0.1 X PBS
-
Soil drench
3
Water
-
Root dip and drench
4
0.1 X PBS
-
Root dip and drench
5
B. subtilis A13
1 x 107
Root dip and drench
6
B. subtilis A13
1 x 107
Soil drench
7
B. subtilis A13
1 x 105
Root dip and drench
8
B. subtilis A13
1 x 105
Soil drench
9
B. subtilis A13
1 x 107
Root dip
7
Application Method
10
P. putida KT2442
1 x 10
Root dip and drench
11
P. putida KT2442
1 x 107
Soil drench
12
P. putida KT2442
1 x 105
Root dip and drench
13
P. putida KT2442
1 x 105
Soil drench
14
B. coagulans NCTC 10334
1 x 107
Root dip and drench
15
B. coagulans NCTC 10334
1 x 10
7
16
B. coagulans NCTC 10334
1 x 105
Soil drench
Root dip and drench
17
B. coagulans NCTC 10334 1 x 105
Soil drench
* Concentration of bacteria applied, colony forming units/ml; application of bacterial
suspensions containing 1 x 107 CFU/ml results in 105 -106 CFU/ml of soil.
113
5.2.3. Evaluation of plant growth promoting ability of additional bacterial strains
in ginger tissue culture plants (ginger tissue culture trial II).
Based on preliminary results in the first ginger tissue culture trial, a further trial was
established where bacteria were applied as a root dip and soil drench to ginger tissue
culture plants.
Application of B. subtilis A13 and B. coagulans NCTC 10334 was
repeated to assess the consistency of the plant growth response to introduction of these
bacteria. The effect of application of A. brasilense Sp7, A. lipoferum Br-17, Bacillus F1,
Bacillus F2 (field isolates) and selected combinations of these bacteria were also
assessed.
Plant Growth conditions.
Plants had been maintained in a growth cabinet for approximately 5 weeks (15-20cm
height) prior to treatment. Bacteria were resuspended in 0.1 x PBS for application to
ginger tissue culture plants according to Table 20.2 (28/7/06). Plant roots were dipped
into a suspension of bacteria for 30 minutes. Bacterial drenches were also applied at
planting, fortnightly for the first month and then every four weeks for the remainder of
the trial. Plants were transplanted into 125mm pots that contained one litre of
pasteurised peat:sand. Ten replicate plants per treatment were used. Plants were
further maintained in a growth cabinet and fertiliser was applied as described in ginger
tissue culture trial I. Plants were sampled after 12 weeks. Surface area of shoots, the
number of shoots per plant and the width of the stem at the base of the plant were
recorded. Fresh weight of shoots, roots and rhizomes were measured.
rhizome dry weights were recorded.
Shoot and
Roots were used for assessment of bacterial
colonisation.
Assessment of root colonisation by introduced bacteria (ginger tissue culture trial II).
Culture based methods were employed to determine whether introduced bacteria could
be isolated from plant roots. Plant roots from each treatment group were combined to
produce a composite sample. Rhizoplane and rhizosphere suspensions were prepared
as described in Chapter 4. For the isolation of B. subtilis and B. coagulans, rhizoplane
and rhizosphere samples were heat treated at 80oC for twenty minutes and then plated
onto Salt V8 agar (Appendix 5.1) as described by Backman and Turner (1991).
114
Semisolid NFb agar (Dobereiner 1995; Eckert et al. 2001) was used to isolate A.
brasilense and A. lipoferum. Approximately 10ml of NFb agar was inoculated with 50µl
of rhizoplane or rhizosphere suspension and incubated at 30oC for seven days. A loop
of pellicle forming culture (typically formed by microaerophilic bacteria such as
Azospirillum species) was streaked onto solid NFb amended with yeast extract and
Congo red (Appendix 5.1). Rhizosphere and rhizoplane suspensions prepared from the
roots of non-inoculated plants and dilution buffers served as controls.
Table 20.2. Treatments used in ginger tissue culture trial II.
No.
Treatment
Concentration of Applied
Bacteria (CFU/ml)*
1
Water control
-
2
Buffer control (0.1 X PBS)
-
3
Bacillus subtilis A13
1 x 107
4
Bacillus coagulans NCTC 10334
1 x 107
5
A. brasilense Sp7
1 x 107
6
A. brasilense Sp7
1 x 105
7
A. lipoferum Br-17
1 x 107
8
Bacillus F1
1 x 107
9
Bacillus F2
1 x 107
10
A. brasilense Sp7 + B. coagulans NCTC 10334
1 x 107
11
A. brasilense Sp7 + A. lipoferum Br-17
1 x 107
12
A. lipoferum Br-17 + B. coagulans NCTC 10334 1 x 107
* Concentration of bacteria used in root dip and soil drenches; application of bacterial
suspensions containing 1 x 107 CFU/ml results in 105 -106 CFU/ml of soil. A reduced
concentration of A. brasilense Sp7 was also tested, as too high a concentration of this
bacterium may negatively impact on plant growth (Dobbelaere et al. 1999).
115
5.2.4. Evaluation of plant growth promoting activity of selected bacteria in ginger
grown from seed pieces.
Bacteria selected for the ginger seed piece trial were those that significantly promoted
growth of tissue culture plants (Trial II). Combinations of these treatments were also
assessed.
Ginger rhizomes (cv. Queensland) were produced on a site established with Fusarium
free tissue culture plants (DPI&F Research Farm, Bundaberg). Rhizomes had been cut
into seed pieces and treated with the fungicide benomyl (1.0g/L) as per standard
industry practice (Stirling 2004). Seed pieces of equivalent size (~ 50g, Figure 12) were
dipped into a bacterial suspension (106 CFU/ml in 0.1% methylcellulose) for 10 minutes
(Table 20.2, 20/10/06). Controls were also dipped in 0.1% methylcellulose (Desai et al.
2002). Seed pieces were then allowed to dry at ambient temperature for two days, as
per standard industry practice, to prevent rotting that may be associated with planting
wet seed material (Hamill, personal communication 2006).
Thirty replicates per
treatment were used. The seed pieces were planted in ten-liter plant bags that
contained seven litters of pasteurised peat:sand mix amended with fertiliser (per 100L of
peat:sand 34g ammonium sulphate, 11.5g superphosphate, 15.5g potash of sulphate,
3g magnesium sulphate, 0.9g copper sulphate, 1.2g zinc sulphate, 0.9g iron sulphate,
450g lime; pH 5.5; Sanewski 2002). Plants were maintained in a shade-house under
natural conditions, were rain-fed and watered by means of an overhead irrigation
system as required.
Bacteria were resuspended in 0.01 M potassium-phosphate buffer (Appendix 5.1; Lin et
al. 1983; Kadouri et al. 2003; Bashan et al. 2006) for application of bacterial drenches,
to avoid potential negative effects of adding salt via buffer to the plants, as was
indicated in previous ginger tissue culture trials.
Bacterial suspensions (70 ml
containing 1 x 107 CFU/ml of bacteria, or ~ 105 CFU/ml of soil) were applied at planting
and 14, 30 and 60 DAP; two control treatments received either water or buffer at this
time as appropriate.
116
Sulphate of Potash (1.2g) was applied 14, 42 and 60 DAP.
Peters Professional®
fertiliser (2g/L, 250ml) was applied every 4 weeks until January and then every 14 days.
Plants were harvested after 17 weeks, typical of early harvest. Soil was removed from
the roots by hosing.
Fresh and dry weight of roots, shoots and rhizomes were
measured. Plant height, stem width, and number of knobs were recorded.
Table 21. Treatments applied in ginger seed piece trial.
Treatment No. Bacterial Treatments
1
Water Control
2
Buffer Control (0.02M potassium-phosphate buffer)
3
A. brasilense Sp7
4
Bacillus F2
5
A. brasilense Sp7 + B. coagulans NCTC 10334
6
A. brasilense Sp7 + B. coagulans NCTC 10334 + Bacillus F2
7
A. brasilense Sp7 + Bacillus F2
Figure 12. Ginger seed pieces used for testing of bacterial treatments.
117
5.2.5. Effect of alginate beads for the delivery of A. brasilense Sp7 on the growth
response of ginger tissue culture plants (alginate bead trial).
A further trial was conducted using ginger tissue culture plantlets, where an alginate bead
formulation was compared to soil drenches for the application of the model strain A.
brasilense Sp7.
Preparation of Dried Alginate Beads
Alginate beads were prepared as described by Bashan (1986b) and van Elsas et al.
(1992) as follows.
Sterile glassware, media, solutions, filter paper, filtration units,
needles and syringes were used to prepare alginate beads in a Class II cabinet.
A.
brasilense Sp7 was cultured overnight in 25 ml of NBY* broth or TYG broth (Bashan et
al. 2002; Appendix 5.1). Cultures were centrifuged at 3000rpm and the bacterial pellet
was washed in 0.1 x PBS.
The bacterial pellet was then resuspended in 5ml of
broth:skim milk (1:1, broth:20% skim milk powder, Difco). This suspension was mixed
with 20ml of 2% alginate (low viscosity, Sigma-Aldrich) with gentle shaking for 15
minutes.
The control was prepared similarly, with the addition of broth:skim milk but
without bacteria. The alginate suspension was dripped aseptically, with the use of a 23gauge needle and syringe, into a stirred solution of 0.2M calcium chloride; the beads
were allowed to cure for at least 30 minutes. The beads were recovered on Whatman
filter paper and washed three times in 0.85% saline. The beads were then returned to a
flask containing NBY* or TYG broth as appropriate and incubated with gentle shaking
overnight at 28oC. The beads were recovered on Whatman filter, washed in saline as
described above and dried overnight on filter paper in a biological safety cabinet. The
dried beads were stored in an airtight container at room temperature.
To determine the viability of bacteria in the alginate beads, 0.1M potassium phosphate
buffer (Appendix 5.1; Bashan 1986b) was added to a weighed sample of the beads
(bacteria or control) and incubated with agitation in a 100ml Erlenmeyer flask for at least
2 hours at 28oC. For wet beads 24ml of 0.1M potassium phosphate buffer was added
per 400mg of alginate beads. In the case of dried beads 25mg of beads were added to
25ml of 0.1M potassium phosphate buffer. Serial dilutions of these suspensions were
118
plated onto nutrient agar. Plates were incubated at 28oC for up to 4 days and examined
for colony number and purity.
Application methods
Cells from an overnight culture of A. brasilense Sp7 were resuspended in water (106
CFU/ml) to avoid positive or negative buffer effects. Ten millilitres of this suspension
was applied to ginger tissue culture plants (20-25 cm height) growing in
vermiculite/perlite in seedling trays (Bashan et al. 1987; Roncato-Maccari et al. 2003;
Njoloma et al. 2006). After three days the plants were transferred to 125 mm diameter
pots that contained one litre of pasteurised peat:sand (50:50 9/2/07). A. brasilense Sp7
was then also applied to the soil as a drench or as the alginate bead formulation, Table
22 (Bashan et al. 1987).
Table 22. Treatments applied in the alginate bead trial.
No.
Treatment applied
1
Water control
2
Alginate bead control
3
A. brasilense Sp7 applied as a soil drench*
4
A. brasilense Sp7 applied to soil as alginate beads*
* A suspension of A. brasilense Sp7 was also applied to plants growing in
vermiculite/perlite prior to transplantation in soil.
For plants that received alginate beads, prior to transplanting a furrow was made in the
pot, the beads (100mg per pot @ 1 x 107 CFU/mg of dried beads, or approximately 1 x
106 CFU/ml of soil) were placed in the furrow and mixed with surrounding soil, so that
the beads were approximately 2 cm below the plant (Fallik and Okon 1996). For plants
where bacteria were applied as a soil drench, 50ml of a 1 x 107 CFU/ml suspension of
A. brasilense Sp7 in water was applied 1, 3 and 7 weeks after planting (-106 CFU/ml of
soil). Two different control treatments received either 100mg of dried alginate beads
119
without bacteria per pot (bead control) or water only (water control).
Twenty-two
replicate plants per treatment were used. Plants were maintained in a greenhouse and
watered twice weekly to maintain soil moisture. Thrive® general purpose fertiliser (25
ml, 1.0 g/L) was applied fortnightly. Sulphate of potash (0.4g) was added to each pot 5,
50 and 80 DAP. After 16 weeks plants were sampled and roots were washed in tap
water. Fresh and dry weight was determined for the complete plant, rhizome, roots and
shoots. The number of shoots and plant height was recorded.
5.3. Results
5.3.1. Wheat as a model system for testing efficacy of bacterial inoculants in
promoting plant growth.
The effect of bacteria introduced by different methods on the growth of wheat is
summarised in Table 23 and Figure 13. When compared to the buffer control, bacterial
treatments applied as a soil drench and seed treatment increased plant growth as
follows: leaf surface area, leaf weight and root weight were increased 22.2%, 17.9%
and 11.8% respectively for Bacillus coagulans NCTC 10334; leaf weight was increased
5.8% and leaf surface area increased 10.3% in the B. subtilis A13 treatment; and leaf
weight and surface area were increased by approximately 12% for P. putida KT2442.
One-way ANOVA indicated that the mean differences in plant growth between treatment
groups were not significant, although high levels of variability were observed.
Two-way analysis of variances indicated that application method significantly affected
the plant growth response to bacterial treatments. When B. coagulans NCTC 10334
was applied as a seed treatment as well as soil drench, leaf weight was significantly
increased compared to the application of this bacterium as a seed treatment alone
(LSD, p<0.05). Likewise, leaf weight and leaf surface area were significantly increased
for P. putida KT2442 applied to the seed and soil compared to the seed treatment
alone. In addition, when data for the different bacterial treatments (B. subtilis A13,
P. putida KT2442 and B. coagulans NCTC 10334) was combined, the application of
120
treatments to the seed as well as to the soil significantly improved plant growth when
compared to the application of bacteria as a seed treatment alone (LSD, p<0.05).
Figure 13. Growth response of wheat plants to introduction of bacterial treatments as
either a seed treatment or seed treatment as well as soil drenches.
1
1
2
3
4
5
6
7
8
2
3 4
5 6
Water
Buffer
B. subtilis A13 (seed)
B. subtilis A13 (seed + soil)
B. coagulans NCTC 10334 (seed)
B. coagulans NCTC 10334 (seed + soil)
P. putida KT2442 (seed)
P. putida KT2442 (seed + soil)
121
7 8
Table 23. Analysis of effect of application method for different bacterial treatments on growth response in wheat by two-way
ANOVA*.
Treatment
App.
Plant Weight
Method**
(grams)
Plant SA
(cm2)
Leaf Weight
(grams)
Leaf SA
(cm2)
Root Weight
(grams)
Root SA
(cm2)
Height
(cm)
Water
SS
0.34 ± 0.05
7.7 ± 1.5
0.15 ± 0.02
5.1 ± 0.6
0.20 ± 0.05
2.8 ± 1.0
21.3 ± 1.9
Buffer
SS
0.33 ± 0.08
8.4 ± 1.5
0.16 ± 0.03
5.6 ± 0.7
0.17 ± 0.06
3.3 ± 1.5
24.9 ± 2.1
Bacillus A13
Seed
0.31 ± 0.07
7.6 ± 2.1
0.15 ± 0.04
5.3 ± 1.3
0.16 ± 0.04
2.6 ± 0.8
22.5 ± 2.8
Bacillus A13
SS
0.33 ± 0.12
8.9 ± 2.0
0.17 ± 0.04
6.2 ± 1.5
0.17 ± 0.06
2.8 ± 0.9
25.8 ± 2.0
B. coagulans 10334
Seed
0.33 ± 0.03
8.3 ± 0.3
0.16 ± 0.02
5.5 ± 0.4
0.17 ± 0.03
2.9 ± 0.4
23.5 ± 1.7
B. coagulans 10334
SS
0.38 ± 0.11
10.1 ± 3.1
0.19 ± 0.06
6.9 ± 1.9
0.19 ± 0.06
3.8 ± 1.4
25.7 ± 3.0
P. putida KT2442
Seed
0.27 ± 0.07
7.4 ± 1.1
0.13 ± 0.02
4.9 ± 0.8
0.14 ± 0.04
2.7 ± 1.1
22.6 ± 2.9
P. putida KT2442
SS
0.35 ± 0.07
9.2 ± 1.0
0.18 ± 0.04
6.3 ± 0.9
0.17 ± 0.04
3.0 ± 0.6
26.0 ± 2.5
* Differences between treatment groups were not significant according to one-way ANOVA. Two-way AVOVA indicated that application
method significantly affected the plant growth response to introduced bacteria. Mean difference between numbers highlighted in bold (in
the same column) is significant (LSD, p<0.05). Mean difference between numbers that are shaded (in the same column) is significant
(LSD, p<0.05).
** Application methods: seed treatment and soil drench (SS) or seed treatment only (seed).
122
5.3.2. Effect of application method on bacterial induced growth response in
ginger tissue culture plants (ginger tissue culture trial I).
Tissue culture plants did not senesce in the controlled conditions of the growth cabinet.
Malfunctioning of the growth cabinet in the first few weeks resulted in reduced
photoperiods that may have affected plant growth.
The response of ginger tissue
culture plants to the introduction of various bacterial strains using different application
methods is indicated in Table 24, Figure 14. One-way ANOVA indicated that there were
significant differences between treatment groups. Shoot fresh and dry weight were
increased by 21.9% and 23.5% respectively, in plants that were soaked in water for
thirty minutes prior to planting in soil (water DD), compared to plants that were planted
directly into soil without soaking in water (water D); this mean difference was significant
according to the LSD test (p<0.05). Rhizome fresh weight was increased similarly,
although the mean difference was not significant. When compared to plants that were
soaked in water (water DD), growth was reduced in plants that were soaked in buffer
prior to transplantation (buffer DD), although differences were not significant. While the
majority of differences between buffer controls and bacterial treatments were not
significant, certain application methods and rates of P. putida KT2442 and B. subtilis
A13 significantly reduced rhizome weight when compared to the buffer control (Table
24).
In comparison to water DD (where growth parameters were increased compared
to water D), bacterial treatments significantly reduced growth parameters, although
differences were not significant for rhizome weight in the B. coagulans NCTC 10334
treatment. A consistent effect of application method and/or concentration of applied
bacteria on plant growth was not observed. Two-way analysis of variance indicated that
different methods of application and concentrations of bacteria did not result in
statistically significant differences in plant growth, although a high level of variability was
observed between plants of the same treatment group.
123
Figure 14. Ginger tissue culture trial I.
a
b
c
d
a.
b.
c.
d.
Ginger tissue plants 1 week after treatment.
Ginger tissue culture plants in the growth cabinet at harvest at harvest.
Representative treatments with roots intact.
Representative treatment with roots removed, illustrating difference
between well-developed and inferior sized rhizomes.
124
Table 24. Effect of different bacterial application methods on mean (± standard deviation) growth parameters in ginger tissue
culture trial I.
Treatment
Method*,
CFU/ml
Water
DD
Plant Fresh
Weight
(grams)
Stem width
(mm)
47.6 ± 7.2ay
Rhizome
Fresh
Weight
(grams)
4.9 ± 1.2a
22.2 ± 2.7a
2.1 ± 0.3a
20.5 ± 4.0a
5.1 ± 1.1a
387 ± 67a
5.3 ± 0.7
D
39.0 ± 5.9abc
4.1 ± 1.8ab
18.2 ± 3.1bc
1.7 ± 0.3bc
16.7 ± 4.1ac
3.7 ± 1.7bc
298 ± 65bc
5.2 ± 0.4
DD
40.9 ± 6.9abc
4.6 ± 1.3a
19.2 ± 2.6ab
2.0 ± 0.4ba
17.2 ± 4.5ac
3.7 ± 0.8bc
328 ± 59ab
5.3 ± 0.9
D
39.3 ± 7.0abc
4.1 ± 1.2ab
18.0 ± 1.9bc
1.7 ± 0.3bc
17.2 ± 5.5ac
3.1 ± 1.1b
309 ± 55bc
5.4 ± 0.8a
DD,log7
33.9 ± 3.1bc
2.6 ± 0.7b
16.9 ± 2.3bc
1.6 ± 0.3bc
14.4 ± 3.6bc
4.3 ± 0.8ac
284 ± 46bc
5.0 ± 0.6a
D,log7
35.4 ± 8.5bc
3.8 ± 1.8ab
16.7 ± 4.3bc
1.6 ± 0.4bc
14.9 ± 3.7bc
3.9 ± 1.1bc
288 ± 85bc
4.7 ± 2.0a
DD,log5
31.9 ± 5.7bc
3.4 ± 1.5b
15.0 ± 2.6bc
1.6 ± 0.1bc
14.2 ± 4.6bc
3.3 ± 0.5bc
270 ± 56bc
5.0 ± 0.6a
D,log5
38.1 ± 15.0bc
3.6 ± 1.8ab
16.7 ± 6.0bc
1.6 ± 0.5bc
17.8 ± 7.6ac
3.6 ± 1.8bc
284 ± 113bc
5.3 ± 0.7a
RD,log7
38.0 ± 12.3bc
3.8 ± 1.8ab
16.7 ± 3.6bc
1.5 ± 0.7cd
17.2 ± 7.1ac
3.4 ± 1.1bc
283 ± 97bc
5.4 ± 1.0a
DD,log7
38.5 ± 9.8bc
3.8 ± 1.0ab
18.0 ± 3.6bc
1.7 ± 0.3bc
16.7 ± 6.3ac
3.9 ± 0.7bc
310 ± 76bc
5.3 ± 0.8a
D,log7
35.2 ± 9.3bc
3.1 ± 1.2b
16.7 ± 2.4bc
1.5 ± 0.2cd
15.5 ± 6.7ac
3.4 ± 1.0bc
287 ± 54bc
5.2 ± 0.8a
DD,log5
31.1 ± 7.8c
2.7 ± 1.1b
15.3 ± 5.3bc
1.4 ± 0.5c
13.0 ± 1.8bc
3.1 ± 0.7b
243 ± 91c
5.5 ± 0.5a
D,log5
35.0 ± 6.7bc
3.6 ± 1.4ab
17.6 ± 3.7bc
1.5 ± 0.3cd
13.9 ± 3.7bc
4.1 ± 1.1ac
280 ± 58bc
5.6 ± 0.8a
B. coagulans DD,log7
NCTC 10334
D,log7
38.3 ± 6.3bc
4.4 ± 0.9a
18.3 ± 2.0bc
1.8 ± 0.2ac
15.6 ± 5.7ac
3.7 ± 1.1bc
295 ± 27bc
5.8 ± 0.6a
41.1 ± 12.2ab
3.7 ± 1.4ab
19.9 ± 3.1ab
1.9 ± 0.4adb
16.6 ± 5.5ac
3.7 ± 1.1bc
328 ± 98ac
5.1 ± 0.5a
DD,log5
38.6 ± 6.7bc
4.6 ± 1.3a
18.6 ± 2.9ac
1.8 ± 0.3ac
14.7 ± 5.1bc
4.2 ± 1.2ac
308 ± 83bc
5.5 ± 0.7a
D,log5
36.9 ± 5.3bc
4.2 ± 1.2a
18.3 ± 3.2bc
1.6 ± 0.4bc
14.5 ± 1.2bc
3.7 ± 0.8bc
299 ± 77bc
5.6 ± 0.7a
Buffer
B. subtilis
A13
P. putida
KT2442
*
Y
Shoot Fresh
Weight
(grams)
Shoot Dry
Weight
(grams)
Root Fresh
Weight
(grams)
Number of
Shoots
Shoot
Surface
2
Area (cm )
Method of Application: DD root dip and soil drench; D soil drench and RD root dip only; CFU/ml of bacteria applied.
Mean difference is not significant for numbers with the same letter, in the same column (LSD, p<0.05).
125
Correlations and Multiple Regression Analysis
In order to determine the value of stem width and plant height as progressive indicators
of plant growth, correlations and multiple regression analyses were performed. The
relationship between stem width and rhizome fresh weight and was investigated by
calculating the Pearson product moment correlation co-efficient, where a value of 0.314
indicated a positive correlation, of average strength, that was statistically significant
(p=0.001). The significance of both stem width and plant height in predicting rhizome
fresh weight was determined by multiple regression analysis. In this model, stem width
was strongly correlated to rhizome weight (p=0.002), while only a small positive
correlation for height was indicated, which was not statistically significant (Figure 15).
Therefore stem width was used as an indicator of plant vigour in future trials.
Figure 15.
Regression analysis assessing stem width as a predictor of rhizome
weight.
Normal P-P Plot of Regression Standardized Residual
Dependent Variable: Rhizome Fresh Weight (grams)
Expected Cum Prob
1.0
0.8
0.6
0.4
0.2
0.0
0.0
0.2
0.4
0.6
Observed Cum Prob
126
0.8
1.0
5.3.3. Evaluation of plant growth promoting ability of additional bacterial strains
in ginger tissue culture plants (ginger tissue culture trial II).
The effect of various bacterial inoculants, applied with the root dip and drench method,
on the growth of ginger tissue culture plants is summarised in Table 25 and Figure 17 –
Figure 19.
One-way ANOVA indicated that there were statistically significant
differences in mean plant growth between treatment groups. When compared to the
buffer control, application of Bacillus F2 (field isolate), Azospirillum brasilense Sp7*
(1 x 107 CFU/ml) and A. brasilense Sp7 combined with Bacillus coagulans NCTC 10334
increased rhizome fresh weight by 40.9%, 46%, and 50% respectively. These mean
differences were significant according to LSD, p<0.05. Rhizome dry weight was
increased similarly in the aforementioned treatments, although differences were only
significant for the A. brasilense Sp7 + B. coagulans NCTC 10334 treatment compared
to the buffer control. Differences in root and shoot weight, for Bacillus F2, Azospirillum
brasilense Sp7* and A. brasilense Sp7 + B. coagulans NCTC 10334, compared to the
buffer control were much smaller and not significant, with the exception of increased
root weight in the A. brasilense Sp7 + B. coagulans NCTC 10334 (Table 25). When B.
coagulans NCTC 10334 was applied alone, plant growth parameters were significantly
reduced compared to application of the bacterium in combination with A. brasilense
Sp7.
Also, when compared to the buffer control, plant growth was also generally
reduced by the application of B. coagulans NCTC 10334. Application of A. brasilense
Sp7 at 2 x 105 CFU/ml resulted in increased rhizome fresh and weight of 24.7% and
20.3% (p>0.05), which is less than increased plant growth observed when 2 x 107
CFU/ml of this bacteria was used.
In this trial, the application of B. subtilis A13 resulted in a trend towards increased
rhizome fresh and dry weight (~32%), root fresh weight (21.18%) and shoot fresh
(6.5%) and dry (11.8%) weight, when compared to the buffer control, although
differences were not statistically significant.
Biometric parameters of the buffer control were reduced when compared to the water
control, where application of buffer (0.1 x PBS) reduced rhizome fresh weight, rhizome
127
dry weight and root fresh weight by 25.7%, 12.5% and 24.7% respectively, although
differences were not statistically significant.
In comparison to the water control, application of Azospirillum brasilense Sp7 (107
CFU/ml), Bacillus F2 and a combination of A. brasilense Sp7 and B. coagulans NCTC
10334 increased rhizome fresh weight by 16.1%, 12.1% and 19.4% respectively,
although mean differences were not statistically significant.
In the A. lipoferum Br-17 treatment increased root fresh weight (8%) and shoot dry
weight (13.7%) compared to the buffer control were not statistically significant (Table
25). The positive effect of A. lipoferum Br-17 on plant growth was augmented when the
bacterium was combined with B. coagulans NCTC 10334 (p>0.05). When A. lipoferum
Br-17 was combined with A. brasilense Sp7, plant growth was reduced compared to the
application of A. brasilense Sp7 alone (p>0.05).
In analyses of root colonisation, introduced bacteria were not distinguishable from
indigenous microflora by culture-based analyses employed.
Figure 16. Ginger tissue culture II 4 weeks after planting.
128
Figure 17. Ginger tissue culture II at harvest.
a
b
1
2
3
4
5
6
7
8
9
10
11
12
a. Tissue cultured plants in growth cabinet.
b. Tissue cultured plants with roots removed:
1. Water control
2. Buffer control
3. B. subtilis A13
4. B. coagulans NCTC 10334
7
6. A. brasilense Sp7 (105 CFU/ml)
5. A. brasilense Sp7 (10 CFU/ml)
7. A. lipoferum Br-17
8. Bacillus F1
9. Bacillus F2
10. A. brasilense Sp7 + B. coagulans NCTC 10334
11. A. brasilense Sp7+ A. lipoferum Br-17;
12. A. lipoferum Br-17 + B. coagulans NCTC 10334.
129
Table 25. Effect of a range of bacterial treatments (root dip followed by soil drenches) on mean (± standard deviation) growth
parameters of ginger tissue culture plants, ginger tissue culture trial II#.
Treatment
Plant Fresh
Weight
(grams)
Rhizome
Rhizome Dry
Fresh Weight
Weight
(grams)
(grams)
Root Fresh
Weight
(grams)
Shoot Fresh
Weight
(grams)
Shoot Dry
Weight
(grams)
Height
(cm)
Shoot Surface
area
2
(cm )
Stem width
(cm)
Water
46.7 ± 7.7ady
6.5 ± 1.6ac
0.41 ± 0.15abc 21.9 ± 4.3ac
18.3 ± 3.5ab
1.9 ± 0.5a
356.0 ± 94.0ab 30.2 ± 5.1a 0.47 ± 0.05a
Buffer
41.2 ± 9.9ac
5.2 ± 2.3ab
0.36 ± 0.18abc 17.6 ± 4.3ab
18.5 ± 4.4a
1.9 ± 0.4a
338.0 ± 71.7ab 30.8 ± 2.9a 0.48 ± 1.12a
BS
47.9 ± 4.9ad
6.9 ± 1.8ac
0.48 ± 0.14acd 21.3 ± 2.9ac
19.7 ± 2.3a
2.1 ± 0.3a
375.4 ± 61.2a
30.7 ± 2.6a 0.47 ± 0.06a
BC
36.9 ± 7.6bc
4.6 ± 1.7b
0.33 ± 0.08b
16.2 ± 2.6ab
1.7 ± 0.3a
310.0 ± 41.3b
32.6 ± 3.5a 0.47 ± 0.09a
AB Sp7*
45.4 ± 12.7ad 7.6 ± 2.3c
0.53 ± 0.13bcd 20.4 ± 6.3acb
17.6 ± 6.7ab
2.0 ± 0.3a
369.5 ± 52.6ab 31.3 ± 4.1a 0.51 ± 0.08ab
AB Sp7 **
43.2 ± 9.8acd 6.5 ± 2.8abc
0.43 ± 0.15cd 18.9 ± 5.6acb
17.9 ± 3.0ab
1.9 ± 0.3a
349.5 ± 52.8ab 31.8 ± 4.2a 0.53 ± 0.12ab
AL
41.0 ± 5.4ac
0.35 ± 0.09ab 19.0 ± 4.4acb
17.0 ± 2.5ab
2.1 ± 0.9a
350.2 ± 29.4ab 31.7 ± 3.2a 0.48 ± 0.06a
Bacillus F1
41.6 ± 10.7ac 5.5 ± 2.2ab
0.39 ± 0.12ab 22.2 ± 9.0ac
13.9 ± 10.4b
1.8 ± 0.4a
340.2 ± 91.5ab 33.3 ± 3.6a 0.46 ± 0.06a
Bacillus F2
47.6 ± 7.7ad
7.3 ± 2.3ac
0.50 ± 0.17cd 21.2 ± 4.9ac
19.1 ± 1.7a
2.1 ± 0.4a
383.7 ± 63.9a
32.5 ± 3.5a 0.52 ± 0.07ab
7.8 ± 2.9c
0.56 ± 0.21d
23.9 ± 6.5c
18.9 ± 1.6a
2.2 ± 0.6a
371.7 ± 47.5a
31.9 ± 2.8a 0.57 ± 0.09b
AB Sp7+AL 43.3 ± 11.3acd 6.0 ± 2.2abc
0.41 ± 0.14c
20.0 ± 5.2acb
17.3 ± 8.1ab
1.8 ± 0.5a
364.7 ± 81.4ab 30.9 ± 4.1a 0.52 ± 0.09ab
AL+BC
0.42 ± 0.19c
19.9 ± 7.5acb
18.6 ± 5.0a
2.0 ± 0.4a
378.9 ± 96.7a
AB Sp7+ BC 50.5 ± 8.9d
5.0 ± 1.3ab
44.4 ± 9.8acd 5.8 ± 2.3ab
16.2 ± 4.9b
30.4 ± 1.8a 0.49 ± 0.03a
# BS: B. subtilis A13; BC: B. coagulans NCTC 10334; AB Sp7: A. brasilense Sp7; AL: A. lipoferum Br-17.
* A. brasilense Sp7 used at 2 x 107 CFU/ml.
** A. brasilense Sp7 used at 2 x 105 CFU/ml; All other bacteria were used at 2 x 107 CFU/ml.
Y Mean difference is not significant for numbers with the same letter, in the same column (LSD, p<0.05).
130
Figure 18. Effect of bacterial treatments on the fresh weight of ginger tissue culture
plants (Ginger Tissue Culture Trial II)*.
A. Plant Fresh Weight
B. Rhizome Fresh Weight
12.0
Mean Rhizome Fresh Weight (grams)
Mean Plant Fresh Weight (grams)
60.0
50.0
40.0
30.0
20.0
10.0
10.0
8.0
6.0
4.0
2.0
0.0
0.0
Water Buffer BS
BC AB7* AB7 ** AL
F1
F2 AB7+ AB7+ AL+
BC AL BC
Water Buffer BS
Treatment
BC AB7* AB7 ** AL
F1
F2 AB7+ AB7+ AL+
BC AL BC
F1
F2
Treatment
Error bars: +/- 1 SD
Error bars: +/- 1 SD
D. Root Fresh Weight
C. Shoot Fresh Weight
30.0
25.0
Mean Root Fresh Weight (grams)
Mean Shoot Fresh Weight (gram s)
30.0
20.0
15.0
10.0
20.0
10.0
5.0
0.0
0.0
Water Buffer BS
BC AB7* AB7 ** AL
F1
F2
Water Buffer BS
AB7+ AB7+ AL+
BC AL BC
BC AB7* AB7 ** AL
AB7+ AB7+ AL+
BC AL BC
Treatment
Treatment
Error bars: +/- 1 SD
Error bars: +/- 1 SD
*BS: B. subtilis A13; BC: B. coagulans NCTC 10334; AB Sp7: A. brasilense Sp7 (*107
CFU/ml, ** 105 CFU/ml); AL: A. lipoferum Br-17; F1: Bacillus F1; F2: Bacillus F2.
131
Figure 19. Effect of bacterial treatments on the dry weight and growth parameters of
ginger tissue culture plants (Ginger Tissue Culture Trial II).
A. Rhizome Dry Weight
B. Shoot Dry Weight
3.00
Mean Shoot Dry Weight (grams)
Mean Rhizome Dry Weight (gram s)
0.80
0.60
0.40
0.20
2.50
2.00
1.50
1.00
0.50
0.00
0.00
Water Buffer BS
BC AB7* AB7 ** AL
F1
F2
Water Buffer BS
AB7+ AB7+ AL+
BC AL BC
BC AB7* AB7 ** AL
F1
F2
AB7+ AB7+ AL+
BC AL BC
Treatment
Treatment
Error bars: +/- 1 SD
Error bars: +/- 1 SD
C. Plant Surface Area
D. Stem Width
0.60
400.0
Mean Stem Width (cm)
Mean Plant Surface Area (square cm)
500.0
300.0
200.0
0.40
0.20
100.0
0.0
0.00
Water Buffer BS
BC AB7* AB7 ** AL
F1
F2 AB7+ AB7+ AL+
BC AL BC
Water Buffer BS
Treatment
BC AB7* AB7 ** AL
F1
F2
AB7+ AB7+ AL+
BC AL BC
Treatment
Error bars: +/- 1 SD
Error bars: +/- 1 SD
BS: B. subtilis A13; BC: B. coagulans NCTC 10334; AB Sp7: A. brasilense Sp7 (*107
CFU/ml, ** 105 CFU/ml); AL: A. lipoferum Br-17; F1: Bacillus F1; F2: Bacillus F2.
132
5.3.4.
Evaluation of plant growth promoting activity of selected bacteria in
ginger grown from seed pieces.
The effect of bacterial treatments on the growth of ginger grown from seed pieces is
summarised in Table 26, Figure 21 - Figure 23. When compared to the water control,
rhizome fresh weight was increased by 10.5% in plants that received the potassiumphosphate buffer. Rhizome fresh weight was similarly increased in the A. brasilense
Sp7 treatment.
Root dry weight was increased by 15% in the A. brasilense Sp7
treatment compared to the buffer control. One-way analysis of variance indicated that
differences between treatment groups were not statistically significant (alpha = 0.05;
Table 26) and data was normally distributed.
Figure 20. Ginger seed piece trial 9 weeks after planting.
133
Figure 21. Ginger seed piece trial at harvest (17 weeks after planting).
a
b
c. 1
2
3
4
5
6
7
a. Ginger plants in shade house at harvest.
b. Rhizome with seed piece (white arrow) attached.
c. Ginger plants after hosing of roots:
1. Water control
2. Buffer control
3. A. brasilense Sp7
4. Bacillus F2
5. A. brasilense Sp7 + B. coagulans NCTC 10334
6. A. brasilense Sp7 + B. coagulans NCTC 10334 + Bacillus F2
7. A. brasilense Sp7 + Bacillus F2
134
Table 26. Effect of bacterial treatments on the mean growth parameters of ginger plants grown from seed pieces (± standard
deviation).*
Treatment
Plant Fresh
Weight
(grams)
Plant Dry
Weight
(grams)
Rhizome
Rhizome Shoot Fresh Shoot Dry
Fresh Weight Dry Weight
Weight
Weight
(grams)
(grams)
(grams)
(grams)
Root Fresh
Weight
(grams)
Root Dry Number
Weight of Shoots
(grams)
Stem
Width
(mm)
Height
(cm)
Number of
Knobs
Water
582.6 ± 106.6
44.0 ± 8.0
205.0 ± 47.7
12.6 ± 4.0
278.1 ± 51.3
26.0 ± 4.1
99.5 ± 45.6
5.1 ± 2.4
6.8 ± 1.8
11.1 ± 1.1
102.6 ± 8.0
14.2 ± 3.6
Buffer
608.1 ± 102.6
44.6 ± 8.5
226.6 ± 42.0
14.3 ± 3.9
285.3 ± 47.5
26.6 ± 4.2
96.3 ± 35.2
4.7 ± 1.8
6.6 ± 1.7
11.3 ± 1.2
105.1 ± 8.0
14.4 ± 3.5
A. brasilense Sp7
618.3 ± 86.3
(AB)
45.4 ± 7.2
219.3 ± 39.7
13.7 ± 3.5
296.9 ± 43.7
27.1 ± 3.5
102.1 ± 53.0
5.4 ± 4.6
7.2 ± 2.1
11.2 ± 1.2
102.9 ± 10.5 14.7 ± 3.0
Bacillus F2
579.2 ± 71.1
42.9 ± 5.8
203.2 ± 40.8
12.2 ± 3.2
276.2 ± 38.8
26.4 ± 3.2
100.1 ± 41.2
4.3 ± 1.9
6.6 ± 1.1
11.0 ± 1.3
103.2 ± 7.3
14.3 ± 2.9
AB + B. coagulans
583.5 ± 95.4
(BC)
44.2 ± 7.8
211.5 ± 46.8
12.8 ± 3.3
274.3 ± 44.1
25.9 ± 3.8
94.9 ± 36.1 5.0 ± 3.8
6.4 ± 1.5
11.0 ± 1.1
104.0 ± 6.1
14.2 ± 3.1
AB+BC+F2
595.6 ± 85.7
44.1 ± 7.0
214.4 ± 44.8
13.7 ± 3.5
279.4 ± 49.1
26.6 ± 3.9
101.8 ± 47.2
4.3 ± 2.1
7.0 ± 1.3
11.2 ± 1.5
102.6 ± 10.2 14.4 ± 2.3
AB+F2
622.6 ± 107.8
46.2 ± 7.5
222.8 ± 44.7
13.5 ± 4.0
291.2 ± 58.9
27.6 ± 5.0
108.7 ± 56.3
4.6 ± 1.5
6.9 ± 1.2
11.1 ± 1.3
101.8 ± 9.1
*Mean differences were not significant according to one-way ANOVA.
135
15.2 ± 3.5
Figure 22. Effect of bacterial treatments on fresh weight of ginger grown from seed
pieces.
A. MeanPlant FreshWeight
B. MeanRhizomeFreshWeight
300.0
Rhizome Fresh Weight (grams)
Plant Fresh Weight (grams)
800.00
600.00
400.00
200.00
250.0
200.0
150.0
100.0
50.0
0.0
0.00
Water Buffer
AB
F2
Water Buffer
AB+BC AB+BC+ AB+F2
F2
Treatment
F2
AB+BC AB+BC AB+F2
+F2
Treatment
C. MeanShoot FreshWeight
D. MeanRoot FreshWeight
200.00
Mean Root Fresh Weight (grams)
400.0
Shoot Fresh Weight (grams)
AB
300.0
200.0
100.0
150.00
100.00
50.00
0.00
0.0
Water Buffer
AB
F2
Water Buffer
AB+BC AB+BC AB+F2
+F2
AB
F2 AB+BC AB+BC AB+F2
+F2
Treatment
Treatment
Buffer: 0.02M potassium phosphate buffer; AB: A. brasilense Sp7; F2:
Bacillus coagulans NCTC 10334. Bars represent +/- SD.
136
Bacillus F2; BC:
Figure 23. Effect of bacterial treatments on the dry weight of ginger grown from seed
pieces.
A. MeanPlant DryWeight
B. Mean Rhizome DryWeight
20.0
50.00
Rhizome Dry Weight (grams)
Plant Dry Weight (grams)
60.00
40.00
30.00
20.00
10.00
0.00
15.0
10.0
5.0
0.0
Water Buffer
AB
F2
AB+BC AB+BC AB+F2
+F2
Water Buffer
Treatment
F2
AB+BC AB+BC AB+F2
+F2
Treatment
C. Mean Shoot Dry Weight
D. Mean Root Dry Weight
12.00
Root Dry Weight (grams)
40.00
Shoot Dry Weight (grams)
AB
30.00
20.00
10.00
10.00
8.00
6.00
4.00
2.00
0.00
0.00
Water Buffer
AB
F2
Water Buffer
AB+BC AB+BC AB+F2
+F2
AB
F2
AB+BC AB+BC AB+F2
+F2
Treatment
Treatment
Buffer: 0.02M potassium phosphate buffer; AB: A. brasilense Sp7; F2: Bacillus F2; BC:
Bacillus coagulans NCTC 10334. Bars represent +/- SD.
137
5.3.5. Effect of alginate beads for the delivery of A. brasilense Sp7 on the growth
response of ginger tissue culture plants (alginate bead trial).
An increase in the number of viable cells per gram by approximately of one order of
magnitude was obtained when alginate beads were prepared from A. brasilense Sp7
cells grown in NBY broth amended with tryptophan (NBY*) compared to TYG broth
(Table 27).
Results of the alginate bead trial are summarised in Table 28, Figure 24 – Figure 25.
When compared to the application of A. brasilense Sp7 as a soil drench, delivery of this
bacterium in alginate beads resulted in significantly increased plant weight, shoot
weight, root fresh weight and number of shoots. In comparison to the water control, the
application of the alginate bead formulation of A. brasilense Sp7 resulted in augmented
plant fresh weight (28%), plant dry weight (32.6%, rhizome fresh weight (25%), rhizome
dry weight (21%), shoot fresh weight (45%), root fresh weight (23%), root dry weight
(39%) and number of shoots (63%) (mean differences were significant, with the
exception of rhizome dry weight LSD, p<0.05). When compared to the bead control,
application of A. brasilense Sp7 in alginate beads increased rhizome fresh weight,
rhizome dry weight and shoot dry weight by 23.3%, 17.7% and 12% respectively,
although mean differences were not significant. Fresh and dry weight of the complete
plant, shoots and roots were significantly higher in the alginate bead control compared
to the water control (p<0.05; Table 28).
Even though for A. brasilense Sp7 applied as a drench (aqueous suspension) plant dry
weight, rhizome fresh weight, root dry weight and shoot fresh weight were increased by
12.5%, 10.5%, 25.85% and 5.3% respectively, when compared to the water control,
mean differences were not significant. Generally plants that received A. brasilense Sp7
as a drench had increased yellowing/senescence of shoots that was not observed when
the bacterium was applied as alginate beads.
138
Figure 24. Alginate beads and their effects on the growth of ginger tissue culture
plants.
a
Wet Beads
Dried Beads
b
1
2
3
4
a. Wet and dry alginate beads.
b. Effect of alginate bead formulation on the growth of tissue culture plants
1. Water control.
2. Bead control.
3. A. brasilense Sp7 applied as soil drenches.
4. Alginate bead formulation of A. brasilense Sp7.
139
Table 27. Viable numbers of cells of A. brasilense Sp7 in alginate beads.
Bead Type
TYG Brothx
NBY* Brothx
NBY* Broth#
Wet
1.5 x 107
2.9 x 108
…
Dried
1.5 x 108
2.0 x 109
1 x 1010
* Numbers represent CFU per gram of beads.
# Beads were returned to broth for overnight incubation.
X Beads were not returned to broth after preparation.
Table 28. Effect of alginate carrier material on the mean growth response (± standard deviation) of ginger tissue culture plants to
introduction of A. brasilense Sp7.*
Treatment
Water
Plant Fresh
Weight (grams)
Plant Dry
Weight
(grams)
Rhizome
Rhizome Shoot Fresh Shoot Dry Root Fresh
Fresh Weight Dry Weight
Weight
Weight
Weight
(grams)
(grams)
(grams)
(grams)
(grams)
Root Dry
Weight
(grams)
Plant Height
(cm)
Number of
Shoots
85.7 ± 23.5a
9.2 ± 2.7a
23.0 ± 7.8a
4.3 ± 1.5a
16.7 ± 6.9a 1.9 ± 0.7a 46.0 ± 13.9a
3.1 ± 1.3a
366.4 ± 50.6a
3.5 ± 2.3a
Bead Control
105.9 ± 21.5b
11.2 ± 2.4bc
23.2 ± 6.9a
4.4 ± 1.5a
23.1 ± 6.6b 2.5 ± 0.6b 59.6 ± 13.9b
4.3 ± 1.3bc
382.6 ± 47.2a
5.2 ± 2.0b
AB7 Drench
88.9 ± 27.0a
10.3 ± 3.0ac
25.4 ± 7.5ac
4.5 ± 1.3a
16.2 ± 6.5a 1.9 ± 0.7a 47.3 ± 17.3a
3.9 ± 2.1ac
376.0 ± 50.0a
3.8 ± 1.4a
AB7 Beads
109.5 ± 28.0b
12.2 ± 3.4b
28.7 ± 11.5bc
5.2 ± 2.1a
24.2 ± 7.b 2.8 ± 0.7b 56.6 ± 14.9b
4.3 ± 1.5bc 379.2 ± 110.2a
5.7 ± 2.7b
* Mean difference is not significant for numbers with the same letter in the same column (LSD, p<0.05).
140
Figure 25. Effect of an alginate bead formulation of A. brasilense Sp7 on the fresh
weight of ginger*.
B. Rhizome Fresh Weight
95% CI Rhizome Fresh Weight (grams)
95% CI Plant Fresh Weight (grams)
A. Plant Fresh Weight
125.00
100.00
75.00
30.0
25.0
20.0
Water
Water
Bead Control
AB7 Drench
Bead Control
AB7 Drench
AB7 Beads
AB7 Beads
Treatment
Treatment
C. Shoot Fresh Weight
D. Root Fresh Weight
95% CI Root Fresh Weight (grams)
95% CI Shoot Fresh Weight (grams)
65.00
25.0
20.0
15.0
60.00
55.00
50.00
45.00
40.00
Water
Bead Control
AB7 Drench
Treatment
AB7 Beads
Water
Bead Control
AB7 Drench
Treatment
* Error bars represent standard deviation; AB7 A. brasilense Sp7.
141
AB7 Beads
Figure 26. Effect of an alginate bead formulation of A. brasilense Sp7 on the dry weight
of ginger.
A. Plant Dry Weight
B. Rhizome Dry Weight
6.50
95% CI Rhizome Dry Weight (grams)
95% CI Plant Dry Weight (grams)
14.00
13.00
12.00
11.00
10.00
9.00
8.00
6.00
5.50
5.00
4.50
4.00
3.50
Water
Bead Control
AB7 Drench
AB7 Beads
Water
Treatment
Bead Control
AB7 Drench
AB7 Beads
Treatment
C. Shoot Dry Weight
D. Root Dry Weight
3.25
5.00
95% CI Root Dry Weight (grams)
95% CI Shoot Dry Weight (grams)
3.00
2.75
2.50
2.25
2.00
4.50
4.00
3.50
3.00
2.50
1.75
1.50
2.00
Water
Bead Control
AB7 Drench
Treatment
AB7 Beads
Water
Bead Control
AB7 Drench
Treatment
* Error bars represent standard deviation; AB7 A. brasilense Sp7.
142
AB7 Beads
5.4. Discussion
Results of this study indicated that in wheat, the application of bacteria (B. subtilis A13,
B. coagulans NCTC 10334 and P. putida KT2442) as a seed treatment and soil drench
produced significant increases in leaf weight and surface area compared to the bacterial
treatment of seed alone (Table 23, Table 29).
This may have indicated that the
application of bacteria to the soil enabled establishment of threshold populations of
bacteria required for an improved plant growth response. As populations of introduced
bacteria may decline with time and distance from the point of inoculation, the application
of bacteria to the soil has been proposed as a means to improve the performance of
inoculants in field conditions (Kloepper et al. 2004; Martin and Bull 2002). A consistent
response to Bacillus inoculants applied as a soil treatment (with or without seed
application) has been demonstrated in tomato, cucumber and pepper in greenhouse
and field conditions (Kokalis-Burelle et al. 2006; Zehnder et al. 2000a; Jetiyanon et al.
2003; Jetiyanon and Kloepper 2002).
In the current study, the application of B. subtilis A13 increased leaf weight (5.8%) and
leaf surface area (10.3%), although not significantly (Table 23, Table 29).
This is
comparable to the response documented by Merriman and colleagues (1974), where
foliage dry weight was increased by 11.9% (and tiller number was increased by 32.7%)
in wheat inoculated with B. subtilis A13 and grown in fertilised plots in field conditions.
Broadbent and colleagues (1977) reported a striking increase in the growth of wheat
following the introduction of B. subtilis A13 in greenhouse conditions without additional
fertiliser, although such a response was not observed in field conditions
Additional
fertiliser was not used in the present study, but as plants were not grown to maturity
further comparison with previous trials was not possible.
When evaluating effects of application methods (including root dip followed by soil
drenches and soil drenches alone) on the response of micropropagated ginger plants to
the introduction of B. subtilis A13, B. coagulans NCTC 10334 and P. putida KT2442,
mean differences in growth parameters were not significant in comparison to the buffer
control (ginger tissue culture trial I: Table 24, Table 30). This may have been due to a
143
high degree of inherent variability between plants of the same treatment group.
Significantly reduced rhizome weight and shoot fresh weight resulted from the
application of P. putida KT2442 when data from the different application methods was
combined.
Different growth parameters were also significantly reduced by the
application of B. subtilis A13, depending on the application method used (Table 24,
Table 30). P. putida KT2440 (the parent strain of KT2442) is known for its ability to
degrade aromatic compounds and is reported to be an efficient coloniser of the
rhizosphere in broad-bean, corn, pea and barley, although the growth response of
plants to the bacterium was not reported in these studies (Molina et al. 2000; EpsinosaUrgel et al. 2002; Molbank et al. 2007). B. subtilis A13 was previously shown promote
the growth of wheat, a variety of nursery plants, cotton, carrots and peanuts but be
inhibitory to the growth of other nursery plants (Turner and Backman 1991; Broadbent
et al. 1977; Brannen and Kenney 1997; Merriman et al. 1994). Reasons why bacteria
may have a deleterious effect on plant growth include the production of metabolites or
phytotoxins that are inhibitory to root and shoot growth/functioning (Brimecombe et al.
2001; Jagadeesh et al. 2006).
Alternatively, a negative effect of bacteria on plant
growth may result from production of inhibitory concentrations of plant hormones such
as indole acetic acid, if too high a concentration of bacteria is applied (Dobbelaere et al.
1999).
In ginger tissue culture trial I, statistically significant (p<0.05) increased plant growth
resulted from soaking acclimatised tissue culture plant roots in water for 30 minutes
prior to planting in soil (Water DD compared to Water D, Table 24, Table 30). The fresh
weight to dry weight ratio was not different between plants soaked in water to those
transplanted directly into soil, indicating that improved plant growth resulted from
increased plant biomass and not just increased water content. Results suggested that a
negative effect of the buffer on plant growth occurred when plant roots were dipped in
phosphate buffered saline (0.1 X PBS), although a negative effect of applying the buffer
to the soil was not evident (discussed later).
As effects of the buffer on plant growth
were not known at the time the second ginger tissue culture trial commenced, the
method of root dip followed by soil drench was chosen (bacteria suspended in 0.1 X
PBS, ginger tissue culture trial II), as this may potentially enable beneficial bacteria
144
access to root colonisation sites prior to transplanting into a soil that has a competitive
indigenous microflora.
While in ginger tissue culture trial I the introduction of B. subtilis A13 resulted in a trend
toward reduced growth of ginger tissue culture plants, in ginger tissue culture trial II this
bacterium induced a positive growth response. In this second trial, the B. subtilis A13
treatment increased rhizome fresh and dry weight (~32%), root fresh weight (21.18%)
and shoot fresh (6.5%) and dry weight (11.8%) when compared to the buffer control,
although mean differences were not statistically significant (Table 25, Table 31). The
difference in the response of tissue cultured ginger plants to the introduction of B.
subtilis A13 in the two trials may have been a result of the variable nature of the plant
response to this bacterium, as has been reported in nursery plants, wheat and peanuts
(Broadbent et al. 1977; Backman and Turner 1991). It has been previously shown that
the time of planting may also influence the plant growth response to B. subtilis A13
(Backman and Turner 1991), therefore it is possible that in the present study reduced
photoperiods, due to growth cabinet malfunctioning in the first trial, may have
contributed to this differential response of ginger plants to the introduction of B. subtilis
A13 (Table 32).
In ginger tissue culture trial II, the application of Bacillus F2 (field isolate), A. brasilense
Sp7 (107 CFU/ml) and a combination of A. brasilense Sp7 and B. coagulans NCTC
10334 to micropropagated ginger plants significantly increased rhizome fresh weight by
40.9% to 50% when compared to the buffer control. When compared to the water
control, plants in the buffer control had a reduced rhizome fresh weight of approximately
20% (p>0.05; Table 25, Table 31). In contrast, results from ginger tissue culture trial I
suggested a negative effect of dipping of plant roots into the buffer, but not from the
application of the buffer to the soil. As the buffer contained 13.6 mM NaCl, trends
toward reduced rhizome weight may have indicated that tissue culture plants were
sensitive to salt (hyper-osmotic) stress. Under conditions of salt stress, an efflux of
water and an accumulation of Na+ in the cytosol may result from osmotic gradients
across the cell membrane (Sleator and Hill 2001; Welsh 2000). In salt tolerant plants
and bacteria the preferential uptake of potassium rather than sodium ions and the
145
accumulation of cytosolic osmolytes may prevent the occurrence of detrimental
concentrations of cytoplasmic Na+. Similarly bacterial inoculation may confer resistance
to hyper-osmotic stress in plants; for example under conditions of salt stress, in maize
plants inoculated with Azospirillum spp. potassium was increased whereas in noninoculated plants Na+ was increased (Hamdia et al. 2004). Amelioration of salt stress
following inoculation of lettuce with A. brasilense Sp7, and by a range of other plant
beneficial bacteria in wheat, tomato, lettuce, squash, chickpea and faba bean has
previously been shown (Mayak et al. 2004; Bacilio et al. 2004; Hamaoui et al. 2001;
Barassi et al. 2006; Yildirim et al. 2006). Improved tolerance of plants to increased
salinity (caused by irrigation) is important for the maintenance of productivity in
agricultural systems (Powers and McSorley 2000).
Azospirillum inoculants may also enhance plant growth by improving water status of
plants, which may be of particular value for improving the growth of micropropagated
plants, as water stress following ex vitro transplantation is a typical cause of reduced
productivity of tissue cultured plantlets (Okon and Labandera-Gonzalez 1994; Nowak
1998).
As these plants are cultured in enclosed vessels in vitro, conditions of high
relative humidity are produced. This may contribute to inefficient transfer of water from
roots to shoots and poorly regulated leaf transpiration, causing water stress of plants
following ex vitro transplantation. In addition lack of development of a waxy cuticle
during in vitro growth is further implicated in water stress of plants during acclimatisation
(Nowak 1998; Posposilova et al. 1999; Nowak and Shulaev 2003; Krishna et al. 2005).
Sensitivity of tissue cultured plants to biotic and abiotic stresses following
transplantation may also be caused by impaired photosynthetic capacity due to
supplementation of in vitro culture media with carbohydrates (Nowak 1998). Reduced
water stress of transplanted micropropagated tomato and potato plants following
bacterization has had much success and has the potential for increasing productivity of
tissue cultured plants in commercial applications (Nowak et al.1997; Pillay and Nowak
1997). Enhanced growth and survival of bacterized micropropagated plants may also
result from competitive displacement of pathogenic and deleterious organisms (Pandey
et al. 2000).
Mia and colleagues (2005) used A. brasilense Sp7 to inoculate
micropropagated banana plants, grown under hydroponic conditions with 33% N146
fertiliser, and reported increased nutrient content, earlier flowering and enhanced yield
and fruit quality. Inoculation of micropropagated photinia plants (an ornamental shrub)
with A. brasilense Sp7 resulted in increased root fresh weight and surface area, as well
as increased survival rate of plants (Larrabu et al. 2007). In the present study, the
application of A. brasilense Sp7 to micropropagated ginger plantlets resulted in
significant increases in rhizome fresh weight (46%), but increases in root fresh weight
(16%) were not significant (Table 25, Table 31). Although when A. brasilense Sp7 was
co-inoculated with B. coagulans NCTC 10334, both rhizome and root fresh weight were
increased significantly compared to the buffer control (55% and 36% respectively).
Accordingly, results of this study suggested that bacterial treatments Bacillus F2,
A. brasilense Sp7 and A. brasilense Sp7 combined with B. coagulans NCTC 10334
enabled ginger tissue culture plants to overcome sensitivity to hyper-osmotic stress
imposed by the presence of salt in the buffer used to apply the bacteria (where reduced
colonisation of neutral or deleterious rhizobacteria may have been involved). It is
possible that the A. brasilense Sp7 improved sensitivity to salt stress by inducing the
preferential uptake of potassium, rather than sodium as increased rhizome growth was
not always accompanied by increased root weight. The improved growth of ginger
tissue culture plants that were soaked in water prior to planting in soil (following
acclimatisation) may have resulted from improved water status of ginger tissue culture
plants or other unidentified factors. Increased rhizome fresh weight is of significance,
as this part of the ginger plant has commercial value.
Phosphate buffered saline is used in the preparation of bacterial suspensions to avoid
subjecting the bacteria to osmotic shock (Bashan et al. 1993).
Often such bacterial
suspensions (in PBS) are applied as seed treatment rather than as a soil drench,
although application to soil and plant roots are also reported (Jetiyanon and Kloepper
2002; Dobbelaere et al. 1999; Jetiyanon et al. 2003; Mia et al. 2005). In most studies,
only one control is used, either buffer control or water and bacteria resuspended in
buffer have been compared to application of water only (Njoloma et al. 2006). This may
alter the interpretation of effects of bacteria on plant growth.
For example, had the
water control not been included in the ginger tissue culture trials, a negative effect of the
buffer would not have been identified and only the positive effect of the bacteria on plant
147
growth would have been evident, perhaps leading to an over-estimation of the effect of
the bacteria if the plants were not subject to salt stress.
Therefore the value of using
appropriate controls, in this case water as well as buffer controls, is extremely important
for accurate interpretation of effects of treatments on plant growth.
In order to avoid potential complicating effects of salt on plant growth, a potassium
phosphate buffer was selected for application of bacteria to ginger plants grown from
seed pieces. A positive effect of the buffer on plant growth was suggested (p>0.05),
where rhizome fresh weight was increased by 11.7% in plants that received the buffer
compared to the water control (Table 26, Table 34). This is likely to have been due to
the presence of soluble potassium and orthophosphate in the buffer, which may have
been used as nutrients by the ginger plants (supplementary potassium, as sulphate of
potash and phosphate are required for optimal growth of ginger in commercial
production). Differences in growth parameters between bacterial inoculated plants and
the buffer control were of a small size and not statistically significant. Root dry weight
was increased in the A. brasilense Sp7 treatment by 15.5% when compared to the
buffer, although the mean difference was also not significant.
Differences may not
have been significant due to the high amount of inherent variability observed between
plants of the same treatment group, which was greater than levels observed in ginger
tissue culture trials. The ability of A. brasilense Sp7 to enhance root growth in wheat,
via the production of indole-acetic acid has been previously shown (Dobbelaere et al.
1999; Dobbelaere et al. 2003). Enhanced root growth may enable the increased uptake
of water and nutrients, resulting in augmented plant growth. Lin and colleagues (1983)
demonstrated that inoculation of maize and corn with A. brasilense Sp7 resulted in
significant increases in plant nutrients, including potassium. However, inoculation with
Azospirillum spp. may not result in improved plant growth in fertile or heavily fertilised
soil (Okon and Labandera-Gonzalez 1994).
As improved potassium uptake may be a
mechanism involved in the plant growth response to Azospirillum spp., the application of
potassium phosphate buffer might have reduced potential effects of the bacteria, as
levels of soil fertilisers may affect the activity of PGPB (Dobbelaere et al. 2001). This
again demonstrated the value of buffer, as well as water controls in the trial.
148
Another possible reason for a marginal response of ginger grown from seed to
introduced A. brasilense Sp7 may have been due to competing indigenous microflora
(ginger seed pieces were not surface sterilised as this is damaging to cells on the
rhizome surface).
Furthermore, ginger grown from seed pieces may not be as
responsive to the introduction of bacteria as tissue cultured plants that are transplanted
from a sterile environment. This hypothesis is consistent with the findings of Nowak and
Sharma (1998), where the in vitro bacterization of tomato plants resulted in resistance
to Verticillum dahliae, while such a response was not observed when the bacterial
inoculant was applied following transplantation into soil.
Bashan (1986b) also
demonstrated that inoculation with A. brasilense Cd was more effective when applied at
sowing, rather than 20 days after planting, and suggested that by this later stage the
majority of root colonisation sites may have already been occupied.
In addition,
bacterial treatments were applied to the roots of micropropagated ginger plants,
whereas ginger plants grown from seed pieces remain in an inactive state for several
weeks after planting, before germination and roots begin to emerge. The presence of a
root mass in micropropagated plants and lack thereof in initial stages of growth in seedgrown plants may be an important difference in colonisation and establishment of
bacterial inoculants in these plants; bacterization of tissue culture plants prior to planting
may have enabled introduced bacteria access to root colonisation sites, while in ginger
grown from seed bacteria must establish on the seed surface, where competing
bacteria may be present (as the mother rhizome from which the seed is cut is dug from
soil), or there may be little effect of seed treatment.
Alternative application methods that have been used to improve the effectiveness of
PGPB include the use of a carrier material such as alginate (Zohar-Perez et al. 2005;
van Elsas et al. 1992). In the current study, use of use of alginate beads for the delivery
of A. brasilense Sp7 resulted in significantly increased growth of ginger tissue cultured
plants compared to the application of this bacterium as a soil drench (Table 28, Table
35).
Bashan and colleagues (1987) used ELISA based methods to demonstrate
increased levels of root colonisation by A. brasilense Cd occurred in wheat when the
bacterium was applied as alginate beads compared to a liquid suspension. This may be
attributed to the continued release of bacteria at rates of up to 1 x 106 CFU/gram per
149
day as the beads degrade and protection of bacteria under competitive and adverse soil
conditions (Fages 1992; Bashan 1986b). Therefore it is possible that in the current
study, use of alginate beads to deliver A. brasilense Sp7 resulted in improved levels of
root colonisation by the bacterium.
As culture based methods employed in this
research did not enable discrimination of the introduced bacteria from indigenous
microflora, levels of root colonisation were not able to be determined.
When the combined effects of the alginate beads and A. brasilense Sp7 were
considered, rhizome weight was significantly increased (24.8%) compared to the water
control (Table 28, Table 35). Considering that the rhizome of the ginger plant has
commercial value, it is noteworthy that only the alginate bead formulation of A.
brasilense Sp7 significantly increased rhizome weight.
Results suggested that
increased rhizome weight by A. brasilense Sp7 was augmented with the use of the
alginate bead carrier and the alginate bead material enhanced shoot and root growth.
Sodium alginate, derived from brown marine kelp alga, is composed of mannuronic and
guluronic acids. Such oligosaccharides are also found in plant and fungal cell walls and
may modulate plant growth and activate plant defence responses at low concentrations
(John et al. 1997; Etzler 1998; Fry et al. 1993; Farmer et al. 1991). Xu and associates
(2003) demonstrated that enzymatically digested polyguluronate promoted root
elongation of carrot (dicotyledon) and rice (monocotyledon). Growth promotion of rice
and peanuts was demonstrated following the root or foliar application of depolymerised
alginate (Hien et al. 1999). In tobacco, plant height and weight were increased and
defense against tobacco mosaic virus was induced following the foliar application of
depolymerised algal oligosaccharides (Laporte et al. 2007). Phenylalanine lyase and
total peroxidase defense responses were induced in wheat following the application of
alginic acids extracted from brown kelp (Chandia et al. 2004).
In previous studies where alginate bead formulations of bacteria have been used,
frequently a positive effect of the alginate bead material on plant growth may not be
detected when application of alginate beads without bacteria is the only control used or
when application of the bacteria in alginate beads is compared to the application of free
cell suspensions (Young et al. 2006; Trivedi et al. 2005; van Elsas et al. 1992). Thus
150
the value of water control as well as carrier (or buffer) controls was again reiterated, so
that effects of carrier materials or buffers on plant growth can be identified.
5.5. Conclusion
In conclusion, buffers and carrier materials used to prepare bacterial inoculants were
observed to have marked effects on plant growth. These effects could be identified due
to the inclusion of buffer/carrier controls as well as water controls in the trials. A dilute
phosphate buffered saline solution negatively impacted on the growth of ginger tissue
culture plants, which was overcome by application of Bacillus F2 (field isolate),
A. brasilense Sp7 and a combination of A. brasilense Sp7 and B. coagulans NCTC
10334. Conversely a potassium phosphate buffer had a positive effect on the growth of
ginger grown from seed pieces and may have masked the effects of introduced
bacteria, which may have otherwise stimulated uptake of ions similar to those present in
the buffer or the bacteria may have exhibited reduced activity under higher levels of
available nutrients. Alternatively ginger plants grown from seed may be less responsive
than tissue cultured plants to the introduction of bacteria or application methods were
not optimal in the seed-piece trial.
Amongst several strains of bacteria tested under reduced levels of fertiliser, the trials
undertaken demonstrated the enhanced growth of ginger tissue culture plants was most
pronounced following the introduction of A. brasilense Sp7, an extensively characterised
type strain known to produce the phytohormone IAA (Dobbelaere et al. 1999; Kadouri et
al. 2003; Tarrand et al. 1978). A mechanism involved in the growth promotion of ginger
tissue culture plants following the introduction of A. brasilense Sp7 and Bacillus F2 may
have included improved resistance to hyper-osmotic stress. The positive response of
micropropagated ginger plants to the introduction of A. brasilense Sp7 was augmented
by the use of alginate as a carrier material and alginate also had a phytostimulatory
effect.
Results suggested that the soaking of plant roots in water following
acclimatisation and prior to planting in soil might reduce water stress and improve
growth of micropropagated ginger tissue culture plants.
151
Tissue cultured ginger is used to establish mother blocks that supply commercial
growers with planting material free from Foz infestation. Tissue cultured ginger plants
often don’t produce rhizomes of commercial size that are much smaller than those from
plants vegetatively propagated plants. Improved rhizome development in first
generation ginger tissue culture plants may reduce high levels of wastage encountered
due to formation of inferior sized rhizomes that are not suitable for use as seed pieces
(Smith and Hamill 1996). This may facilitate increased usage and reduced production
costs of this source of clean planting material in the ginger industry.
152
Appendix 5.1. Solutions and Media.
0.1 X Phosphate Buffered Saline (Sambrook et al. 1989).
Per liter of 0.1 X PBS contains: NaCl 0.8g, 20mg KCl, 144 mg Na2HPO4, 24 mg
KH2PO4, pH 7.4. The buffer was autoclaved before use.
0.1 M Potassium Phosphate Buffer (Sambrook et al. 1989).
Per liter 0.1M potassium phosphate buffer contains 49.7 ml 1M K2HPO4 and 50.3 ml 1M
KH2PO4, pH 6.8. The 0.02M potassium phosphate buffer was prepared by performing a
1:50 dilution of the stock solution in deionised water. Buffers were autoclaved before
use.
Salt V8 Agar (Turner and Backman 1991).
Per liter the medium contains 400ml V-8 juice, NaCl 40g, dextrose 1g, agar 20g, pH 5.2.
NFb Media (Dobereiner 1995; Eckert et al. 2001).
Per litre this media contains D,L-Malic acid 5g (Fluka), K2HPO4 0.5g, MgSO4.7H2O 0.2g,
NaCl 0.1g, CaCl2.2H2O 0.02g, minor element solution 2ml, bromothymol blue 2ml (0.5%
solution in 0.2M KOH; 50mg/10ml), FeCl2 10mg, vitamin solution 1ml, agar 15g (plates)
or 1.8g (semisolid media), yeast extract (Fluka)+/- 50mg (+plates). So that iron and
salts did not precipitate, ingredients were added in the sequence listed. Per 10ml, the
minor element solution contained CuSO4.H2O 4mg, ZnSo4.7H2O 1.2mg, Na2MoO4.2H2O
10mg and MnSO4.H2O 15mg. Per 10ml the Vitamin Solution contained biotin 1mg and
pyridoxol-HCl 2mg; this solution was filter sterilised and added to the media after
autoclaving. For the preparation of agar plates, following sterilisation of the media, 37.5
µg/ml of sterile aqueous Congo red was added (Bastarrachea et al. 1988; Katupitiya et
al. 1995).
153
TYG Broth (Bashan et al. 2002).
Per litre the medium contains tryptone 5g, yeast extract 5g, D-glucose 5g, KOH 4.8g,
NaCl 1.2g, MgSO4 0.25g, K2HPO4 0.13g, CaCl2 0.22g, 0.17g K2SO4, Na2SO4 2.4g,
NaHCO3 0.5g, Na2CO3 0.09g, FeIII-EDTA 0.07g, pH 7.0.
NBY (Nutrient Broth-Yeast Extract Medium: Vidaver 1967; Kim et al. 1997).
Overnight cultures of bacteria were prepared in sterile NBY, that contains per litre:
nutrient broth 8g (Sigma-Aldrich), yeast extract 2g (Fluka), K2HPO4 2g, KH2PO4 0.5g,
glucose 5g and MgSO4.7H2O 0.25g, 15g of agar was added for the preparation of
plates.
Glucose (Sigma-Aldrich) was added as a 10% filter sterilised solution after
autoclaving and cooling of the media. In order to promote bacterial indole acetic acid
production, filter sterilised tryptophan (Sigma-Aldrich) was added to the medium (final
concentration 0.1 mM tryptophan; Kadouri et al. 2003; Prinsen et al. 1993).
154
Appendix 5.2. Supplementary data for wheat trial.
Table 29. Percentage difference in growth parameters of bacterial treatments
compared to the buffer control for wheat.
Treatment
Application Plant
Leaf
Root Plant SA Leaf SA Root SA Height
Method
Weight Weight Weight
Water
Seed + Soil
3.0
-4.6
15.2
-8.2
-9.4
-15.6
-14.6
Buffer
Seed + Soil
…
…
…
…
…
…
…
B. subtilis A13 Seed
-7.3
-8.5
-6.9
-8.6
-6.0
-19.6
-9.7
B. subtilis A13 Seed + Soil
-1.4
5.8
0.4
5.9
10.3
-13.9
3.4
1.2
1.3
1.2
-0.9
-2.1
-12.8
-5.5
14.8
17.9
11.8
21.0
22.2
14.0
3.2
-17.4
-18.8
-17.7
-12.1
-14.0
-18.6
-9.5
5.9
12.5
0.4
9.6
12.2
-9.3
4.6
B. coagulans
Seed
10334
B. coagulans
Seed + Soil
10334
P. putida
Seed
KT2442
P. putida
Seed + Soil
KT2442
SA: surface area
155
Appendix 5.3. Supplementary data for ginger tissue culture trial 1
Table 30. Percentage difference in growth parameters for bacterial treatments
compared to the buffer control in ginger tissue culture trial I.
Plant Rhizome Shoot Root No. of Shoot Stem
Fresh
Fresh
Fresh Fresh Shoots Surface width
Weight Weight Weight Weight
Area
Treatment
Application
Method*,
Concentration
(CFU/ml)
Water
Dip+drench
16.3
7.2
15.5
19
38.9
17.9
0
Buffer
Dip+drench
…
…
…
…
…
…
…
B. subtilis A13 Dip+drench,
log7
Dip+drench,
log5
Root dip only,
log7
Dip+drench,
P. putida
KT2442
log7
Dip+drench,
log5
B. coagulans Dip+drench,
10334
log7
Dip+drench,
log5
-17.2
-43.5
-12.2
-16.3
17
-13.5
-5.3
-21.9
-25.7
-22
-17.3
-10
-17.5
-6
-7
-17.2
-11.1
-0.2
-7.3
-13.8
1.7
-5.8
-16.7
-6.4
-2.7
4.3
-5.4
0.8
-24
-41.1
-20.2
-24.2
-15.1
-26
4.3
-6.4
-3.5
-4.7
-9.4
0.3
-10
8.7
-5.7
0
-3.4
-14.4
12.7
-6
4.3
Water
Drench only
-0.8
0
0.6
-0.1
19.7
-3.5
-3.9
Buffer
Drench only
…
…
…
…
…
…
…
-9.9
-6.6
-7.2
-0.4
24.5
-6.6
-12.2
-3
-11.2
-7.2
0.1
15.2
-8.2
-1.3
-10.3
-24.1
-7.4
-0.3
10.6
-7
-3.5
-10.8
-12.4
-2.2
-0.6
33.5
-9.3
4.4
-3.6
-9.8
7.8
-0.1
19.7
-3.4
-5.7
-6
1.5
1.7
-0.5
19.7
-3.3
3.5
B. subtilis A13 Drench only,
log7
Drench only,
log5
Drench only,
P. putida
KT2442
log7
Drench only,
log5
B. coagulans Drench only,
log7
10334
Drench only,
log5
156
Appendix 5.4. Supplementary data for ginger tissue culture trial II
Table 31. Percentage difference in plant growth parameters for bacterial treatments
compared to the buffer control for ginger tissue culture trial II.
Treatment *
Plant Rhizome Rhizome Root Shoot Shoot Shoot Height
Fresh Fresh
Dry
Fresh Fresh
Dry Surface
Weight Weight Weight Weight Weight Weight area
Stem
width
Water
13.1
25.7
12.5
24.7
-1.3
1.3
5.4
-1.8
-2.1
Buffer
…
…
…
…
…
…
…
…
…
BS
16.0
32.6
32.8
21.2
6.5
11.8
11.1
-0.2
-2.1
BC
-10.4
-12.2
-7.8
-7.7
-12.5
-10.5
-8.3
5.9
-3.1
AB Sp7*
10.2
46.0
46.4
16.0
-5.2
8.0
9.4
1.7
5.2
AB Sp7 **
4.8
24.7
20.3
7.5
-3.4
0.7
3.4
3.4
9.4
AL
-0.7
-3.1
-2.2
8.0
-8.2
13.7
3.6
3.1
-1.0
F1
0.9
6.2
9.4
26.3
-24.8
-4.7
0.7
8.3
-4.2
F2
15.3
40.9
38.9
20.8
3.0
15.8
13.5
5.7
8.3
22.3
50.0
54.7
35.9
1.7
20.4
10.0
3.7
17.7
4.9
15.8
13.6
14.0
-6.9
-1.0
7.9
0.4
7.3
7.7
11.6
16.9
13.2
0.5
6.8
12.1
-1.4
1.0
AB Sp7 +
BC
AB Sp7 +
AL
AL + BC
*BS: B. subtilis A13; BC: B. coagulans 10334; AB Sp7: A. brasilense Sp7;
AL: A. lipoferum Br-17; F1: Bacillus F1 (field isolate); F2: Bacillus F2 (field isolate).
Table 32.
cabinets.
ment
Comparison of plant growth in two ginger tissue culture trials in growth
Plant Fresh
Weight
(grams)
Rhizome Fresh
Weight
(grams)
Root Fresh
Weight
(grams)
Shoot Fresh
Weight
(grams)
Shoot Surface
area
(grams)
Stem width
(mm)
GT1
GT2
GT1
GT2
GT1
GT2
GT1
GT2
GT1
GT2
GT1
GT2
Water
47.6
46.7
4.9
6.5
20.5
21.9
22.2
18.3
387
356
5.3
4.7
Buffer
40.9
41.3
4.6
5.2
17.2
17.6
19.2
18.5
329
338
5.3
4.8
BS
33.9
47.9
2.6
6.9
14.4
21.3
16.9
19.7
284
375
5
4.7
BC
38.3
37
3.4
4.6
14.2
16.2
15
16.2
271
310
5
4.7
*BS: B. subtilis A13; BC: B. coagulans 10334.
GT1: Ginger tissue culture trial I; GT2: Ginger tissue culture trial II.
157
Appendix 5.5. Supplementary data for ginger seed piece trial.
Table 33. Percentage difference in growth parameters of bacterial treatments compared
to buffer control in ginger seed piece trial.
Treatment#
Plant
FW
Plant Rhizome Rhizome Shoot Shoot Root
DW
FW
DW
FW
DW
FW
Root No. of
DW Shoots
1. Water
-4.2
-1.5
-9.5
-11.7
-2.6
-2.2
3.4
7.7
2.6
2. Buffer
…
…
…
…
…
…
…
…
…
3. AB7
1.7
2
-3.2
-3.9
4.1
1.8
6.1
15.5
8.6
4. F2
-4.8
-3.7
-10.4
-15
-3.2
-0.9
3.9
-9.4
0.5
5. AB7 + BC
-4.1
-0.8
-6.6
-9.9
-3.9
-2.5
-1.5
6.2
-2.6
6. AB7 + BC + F2 -2.1
-1.1
-5.4
-4.2
-2.1
-0.2
5.7
-7.7
6.1
7. AB7 + F2
3.6
-1.7
-5.9
2
3.7
12.8
-3.2
4.1
2.4
* FW: Fresh Weight; DW: Dry Weight.
# AB Sp7: A. brasilense Sp7; BC: B. coagulans 10334; F2: Bacillus F2 (field
isolate).
Table 34. Percentage difference in growth parameters of bacterial treatments
compared to the water control in ginger seed piece trial*.
atment#
Plant
FW
Plant
DW
Rhizome Rhizome Shoot
FW
DW
FW
Shoot
DW
Root
FW
Root
DW
No. of
Shoots
1. Water
…
…
…
…
…
…
…
…
…
2. Buffer
4.4
1.3
10.5
13.5
2.6
2.3
-3.2
-8
-2.9
3. AB7
6.1
3.3
7
9
6.8
4.2
2.6
6.5
5.4
4. F2
-0.6
-2.4
-1
-3.6
-0.7
1.4
0.6
-16.5
-2.5
5. AB7 + BC
0.2
0.5
3.2
2.3
-1.4
-0.2
-4.7
-2.2
-5.4
6. AB7 + BC + F2
2.2
0.2
4.6
8.7
0.5
2.2
2.3
-14.9
2.9
7. AB7 + F2
6.9
5
8.7
6.7
4.7
6.1
9.2
-10.8
1
* FW: Fresh Weight; DW: Dry Weight.
# AB Sp7: A. brasilense Sp7; BC: B. coagulans 10334; F2: Bacillus F2 (field
isolate).
158
Appendix 5.6. Supplementary data for alginate bead trial.
Table 35. Percentage difference in growth parameters of bacterial treatments and
controls in the alginate bead trial*.
Treatment
Plant
FW
Plant
DW
Rhizome Rhizome Shoot
FW
DW
FW
Shoot
DW
Root
FW
Root
DW
No. of
Shoots
Bead control cf water control
23.7
21.7
1.3
2.3
38.3
31.6
29.6
38.7
48.6
AB7 Drench cf water control
3.8
12.5
10.5
3.7
-3.1
5.3
2.8
25.8
8.6
AB7 Beads cf bead control
3.3
9.1
23.3
17.7
4.8
12
-5
0
9.6
AB7 beads cf AB7 drench
23.1
32.6
13
15.6
49.4
40
19.7
10.3
50
AB7 Beads cf water control
27.8
32.6
24.8
20.9
44.9
47.4
23
38.7
62.9
* FW: Fresh weight; DW: Dry Weight; cf: compared to; AB7: A. brasilense Sp7.
159
Chapter 6. In vitro and in vivo analysis of interactions between bacterial isolates
and Fusarium oxysporum f. sp. zingiberi.
6.1. Introduction
In addition to improving plant growth, certain plant growth promoting bacteria (PGPB)
may also increase resistance against seed- and soil-borne phytopathogens (Lucy et al.
2004; Fravel 2005; Haas and Defago 2005; Ryan et al. 2008). Increased resistance to
disease may result from improved plant vigour as a consequence of increased root
growth and enhanced nutrient uptake and water status following the application of
PGPB (Vessey 2003). Induced systemic resistance, production of antibiotics, volatiles,
lytic enzymes and sideophores, degradation of pathogen virulence factors and
competitive exclusion are further mechanisms by which PGPB may reduce the
incidence of disease in agronomically important crops (Whipps 2001; Compant et al.
2005; Kloepper et al. 2004; Mercado-Blanc and Bakker 2007; Gnanamanickam et al.
2002; Chin-A-Woeng et al. 2003). Bacteria that reduce disease via direct antagonism of
plant pathogens or by induction of systemic resistance have been referred to as
biocontrol PGPB or biopesticides (Bashan and Holgiun 1998; Haas et al. 2002).
Fluorescent Pseudomonas and Bacillus species have been reported to improve
resistance against diseases caused by Fusarium oxysporum, which cause severe
losses in crop production worldwide (Cazorla et al. 2007; Koumoutsi et al. 2004;
Domenech et al. 2006; Landa et al. 2004; Bakker et al. 2007; de Boer et al. 1999; Bapat
and Shah 2000; Benhamou et al. 1998; Larkin and Fravel 1998). The forma specialis of
this disease that infects ginger, Fusarium oxysporum f. sp. zingiberi (Foz), has caused
devastating losses in regional production (Stirling 2004).
While Foz has typically
affected ginger rhizomes left in the ground until late harvest, high incidences of failed
emergence and poor establishment in the late 1990s were associated with this disease.
Factors linked with the earlier onset of this disease included planting of infected seed
pieces, build up of inoculum levels in soil caused by leaving rhizomes in the ground for
longer periods (due to increased demand for fresh ginger), seasonal conditions, poor
rotational practices and mechanization of the industry (Stirling 2004).
In order to
reduce such losses caused by Foz the use of clean planting material has been eminent.
160
This has included discarding of infected seed pieces and strict hygiene procedures
during seed preparation or the use of disease free planting material. Planting material
free from Foz infection has been produced in sites established with tissue cultured
ginger plants, which have not previously grown Foz affected ginger (Foz is not found in
virgin soil but is introduced via contaminated seed). While dipping of planting material in
a fungicide prior to planting may reduce infection of the seed piece by Foz, this is not
expected to prevent infection of the new rhizome or roots (that grows out from the seed
piece) by soil-borne propagules of this pathogen. Once introduced into the soil,
chlamydospores of Foz may persist for many years, and measures to reduce infection
by these soil borne propagules are not known (Pegg et al. 1974; Stirling 2004).
While biocontrol activity of Bacillus and fluorescent Pseudomonas strains against many
Fusarium oxysporum form species has been demonstrated, only one account of the use
of plant growth promoting bacteria against Foz was found in searches of literature;
Sharma and Jain (1979) reported antagonism and reduced incidence of Foz in ginger
following the application of a B. subtilis strain to ginger seed pieces and soil, although a
detailed account of this study was not described. Therefore in the present study, firstly
the nature of the in vitro interaction between Foz and bacterial isolates that promoted
growth of ginger tissue culture plants as described earlier (A. brasilense Sp7, B. subtilis
A13 and Bacillus F2) and further isolates listed in Chapter 4 (P. fluorescens,
B. megaterium NCTC 10342, B. subtilis DAR26659, B. subtilis ATCC 6633 and several
field isolates) was assessed. Initially the in vitro antagonistic ability of bacterial isolates
was investigated in dual culture assays with Foz on PDA and Waksman agar plates
(Berg et al. 2002). Different types of culture media were used as in vitro antagonism
may occur on one type of media and not another, which may be related to the effects of
media composition on metabolite secretion.
To examine the nature of the in vitro
interaction in further detail, dual culture assays were performed on agar films on
microscope slides, so that contact lysis or aberrant fungal growth could be observed
(Shankar et al. 1994). The demonstration of in vitro antagonism by a bacterial isolate
does not always translate into biocontrol activity in the natural environment (SharifiTehrani et al. 1998; Fravel 2005). This may be due to factors such as differences in the
production of antifungal metabolites on laboratory media and soil (as a result of nutrient
161
availability, population density and quorum sensing) or antagonism of introduced strains
by indigenous microflora (Duffy and Defago 1999; Zhang and Dong 2004).
Thus
bacteria that inhibited the growth of Foz in vitro were also tested for their ability to
reduce the incidence of disease in ginger plants grown in soil that was inoculated with
this pathogen.
6.2. Materials and Methods
6.2.1. Fusarium oxysporum cultures.
Two isolates of Fusarium oxysporum f. sp. zingiberi, BRIP 44987 and BRIP 44963
(provided by Hamill, Maroochy DPI&F) with demonstrated pathogenicity in ginger
(Stirling 2001) were maintained on potato dextrose agar at 24oC.
6.2.2. Dual culture assays on agar plates.
Bacterial cultures listed in Table 13 and selected field isolates were maintained as
described earlier. Bacteria were streaked across the centre of potato dextrose agar
(Oxoid) and Waksman agar (Berg et al. 2002; Appendix 6.1) plates with a sterile
inoculating loop. The plates were then incubated at 28oC overnight. The following day
an agar plug from the leading edge of the two different Foz isolates was placed on
either side of the bacterial streak.
The plates were then incubated at 24oC and
inspected after 7, 10 and 14 days for evidence of inhibition of fungal growth. The
assays were repeated three times.
162
6.2.3. Dual culture assays on microscope slides.
Dual culture assays were performed on microscope slide agar films as described by
Shankar and colleagues (1994). Briefly, potato dextrose agar was poured onto sterile
microscope slides to produce an agar film. The bacteria were streaked onto the agar at
one end of the slide and incubated at 28oC overnight. A plug from the leading edge of
Foz BRIP 44963 was then inoculated onto the agar film at the other end to the bacteria,
at a distance of approximately 4cm.
An agar film inoculated with Foz but without
bacteria served as the control. The slides were then incubated at 24oC for 10 days.
Fungal growth on the slides was examined under a stereo microscope (Nikon SMZ800).
A segment of agar that included the leading edge of the fungus was then transferred to
a clean microscope slide and stained with lactophenol cotton blue for 15 minutes and
then examined under a light microscope (Olympus BH-2).
6.2.4. Effect of bacterial treatments on incidence of Foz infection in ginger tissue
culture plants.
A bioassay to demonstrate the infection of ginger plants by Foz in greenhouse
conditions
has
not
been
developed.
Levels
of
seed
based
inocula
of
Fusarium oxysporum pathogens used to infect subterranean clover, banana and
cyclamen have ranged between 0.5% and 5% w/w or v/v (Elmer 2002; Barbetti and
Sivasithamparam 1987; Smith and Smith 2003).
In the current study soil was
inoculated with low levels of Foz to approximate numbers of infective propagules
estimated in heavily infested field soil, in order to examine the efficacy of bacterial
inoculants under conditions that may be encountered in the field. This inoculum was
produced on rye grass seeds (described next), which may induce the production of
resistant chlamydospores and provide a nutrient base to aid in the establishment of the
fungus in the soil (Dewan and Sivasithamparam 1988; Burgess et al. 1988). Given lack
of disease development, a suspension of spores was later applied to the soil.
163
Rye Grass Seed Foz Inoculum.
Deionised water (125ml) was added to 100g of rye grass seed in a conical flask, the
flasks were plugged with cotton wool and autoclaved for 30 minutes on three
consecutive days (El-Tarabilly et al. 1997). Ten plugs from the leading edge of Foz
BRIP 44963 (growing on PDA) were used to inoculate each flask. The flasks were
incubated at 24oC for ten days and stirred regularly.
Viability of the fungus was
confirmed by placing colonised seeds on PDA.
Plant inoculation and plant growth conditions.
Fully acclimatised ginger tissue cultured plantlets (cv. Canton, 20-25cm height), growing
in vermiculite/perlite in seedling trays, were treated with 10 ml of a suspension of
bacteria (106 CFU/ml in water) prepared as described earlier and as listed in Table 36
(Bashan et al. 1987; Roncato-Maccari et al. 2003; Njoloma et al. 2006). Control
treatments received water at this time. After 3 days plants were transplanted into 1L of
pasteurised peat: sand in 125mm diameter pots. At time of transplantation three rye
grass seeds, heavily colonised with Foz, were added to each pot approximately 3cm
below the plant (~0.01% v/v; approximating levels detected in preliminary analysis of
heavily infested field soil: Hamill, personal communication 2006).
Three controls
treatments received either water only, sterile RG seeds or seeds that had been
inoculated with Foz (bacterial treatments were not applied in these control treatments).
Bacterial inoculants (50ml, 2 x 107 CFU/ml in water) were also applied to the soil 1, 4
and 7 weeks after planting as appropriate (Jetiyanon and Kloepper 2002; Zhang et al.
2004).
Twenty-two replicate plants per treatment were used. Eight extra control plants that
received only the Foz inoculum were included for sampling during the course of the trial
to monitor disease progression. Plants were maintained in a greenhouse and watered
twice weekly to maintain soil moisture. Thrive® general purpose fertiliser (25 ml, 1.0
g/L) was applied fortnightly. Sulphate of potash (0.4g) was added to each pot 5, 50 and
80 days after planting (DAP).
164
After 7 weeks symptoms of Foz infection were not apparent, as determined by visual
examination of cut roots and rhizomes in the control treatment that was inoculated with
Foz.
Therefore a spore suspension of Foz was applied to the pots (10ml @ 105
spores/ml). This suspension was prepared by adding sterile milli-Q water to 10-day-old
plates of Foz BRIP 44963 and using a sterile plate spreader to agitate/remove spores
from the mycelium; this was repeated three times (Omar et al. 2006). The resultant
suspension was filtered through sterile gauze. The number of spores per millilitre was
determined with a Neubauer hemocytometer and the suspension was diluted in sterile
deionised water to a concentration of 105 spores/ml (Omar et al. 2006). Ten millilitres of
the spore suspension was applied to each pot (103 spores/ml of soil).
A 2cm layer of
vermiculite was then used to cover the soil to prevent cross contamination of pots by
splashing and aerosols when watering (Bashan 1986b).
After 16 weeks plants were sampled and roots were washed in tap water. Plants were
rated for above ground (leaf yellowing) and below ground symptoms (rhizome
discoloration) of Foz infection (Table 37). Isolations were performed to determine if Foz
could be recovered from rhizomes. Rhizomes were surface sterilised by momentarily
dipping into 1% sodium hypochlorite, rinsing in sterile distilled water and flaming in
100% ethanol (Stirling 2004). Pieces of rhizome (from the interior) were cut with a
sterile scalpel blade from five surface sterilised rhizomes from each treatment group
and placed onto PDA (Oxoid) amended with streptomycin (50ppm, Sigma-Aldrich).
Plates were incubated at 28oC for ten days.
Fresh and dry weight was measured for the complete plant, rhizome, roots and shoots.
The number of shoots and plant height was recorded.
6.2.5. Experimental design.
The sample size (n) required to detect a 10% increase in plant growth was calculated
iteratively (i.e. by testing different values of n) as described by Zar (1984), using the
formula n = s2/σ2(tα,ν + tβ(1),ν)2, where s is the standard deviation of rhizome weight
(determined in previous trials), σ is the increase to be detected, α (set at 0.05) is the
165
probability of a type I error, β (set at 0.20) is the probability of a type II error and ν
represents degrees of freedom.
The greenhouse trial was set up in a randomised complete block design on a single
bench.
Each block contained one replicate from each treatment group. The software
program Statistical Package for the Social Sciences (SPSS) was used for the following
statistical analyses. The Levene’s test was used to assess the homogeneity of variance
for each variable measured. One-way ANOVA was used to determine if statistically
significant differences were present between the means of treatment groups
(alpha = 0.05). The least significant difference (LSD) test as described by Fisher, was
used to compare the means of different treatment groups (p<0.05).
Table 36. Bacterial treatments used in the in planta Foz bioassay.
Treatment
No.
Treatment
1
Water control
3
Sterile RG Seed
6
Foz infested RG Seed
7
A. brasilense Sp7 + Foz
8
B. megaterium NCTC 10342 + Foz
9
P. fluorescens + Foz
10
B. subtilis DAR26659 + Foz
11
B. subtilis A13 + Foz
12
B. subtilis ATTC 6633 + Foz
13
Above Bacillus isolates + A. brasilense Sp7 + Foz
166
Table 37. Rating of plants for symptoms of Foz infection.
Rating of Shoot Yellowing
Rating of Rhizome discoloration
Rating
Shoot Yellowing
Rating
Rhizome Discoloration
1
No yellow shoots
1
No rhizome discoloration
2
< 25% of shoots yellow
2
3
25% to 50% of shoots yellow
3
4
50% to 75% of shoots yellow
4
5
> 75% of shoots yellow
5
< 25% of rhizome discoloured
25% to 50% of rhizome
discoloured
50% to 75% of rhizome
discoloured
> 75% of rhizome discoloured
6
All shoots wilted, plant dead
6
100% of rhizome discoloured
6.3. Results
6.3.1. Dual culture assays on agar plates.
Results of the dual culture assays, where bacterial isolates and Foz were grown on agar
plates, is summarised in Table 38 and Figure 27.
An altered colour of Foz (red)
occurred in the presence of certain bacteria. Results were very similar for assays
conducted on both Waksman agar and PDA.
167
Table 38. Nature of interaction between bacterial isolates and Foz on agar plates.
Interaction
Bacterial Isolate
+++
B. subtilis DAR26659, B. megaterium NCTC 10342, P. fluorescens,
Pseudomonas Dz5 (field isolate)
++
B. subtilis ATCC 6633, Acidovorax N1 field isolate
+
B. subtilis A13, P. putida KT2442
-
A. brasilense Sp7, Bacillus F2, B. coagulans 10334
+++ Zone of inhibition of greater than 1.5 cm.
++
Zone of inhibition of approximately 0.5cm.
+
After 7 days the zone of inhibition was greater than 1cm, after 10 days Foz was
able to grow adjacent to the bacteria and after 14 days increased proliferation of Foz
occurred along the streak of bacteria.
No inhibition of the growth of Foz and increased proliferation of the fungus
occurred along the streak of bacteria.
168
Figure 27. Effect of bacteria on the in vitro growth of Foz on potato dextrose agar
(PDA) and Waksman agar (WA) plates.
a
b
Foz
44987
Foz
44987
Foz
44963
Foz
44963
PDA
WA
B. subtilis DAR26659
c
PDA
WA
B. megaterium NCTC 10342
d
Foz
44987
Foz
44987
Foz
44963
Foz
44963
PDA
WA
B. subtilis ATCC 6633
a.
b.
c.
d.
PDA
WA
B. subtilis ATCC 6633
Foz and B. subtilis DAR26659.
Foz and B. megaterium NCTC 10342.
Foz and B. subtilis ATCC 6633.
Foz and B. subtilis A13.
169
Figure 27. continued…
e
Foz
44987
f
Foz
44987
Foz
44963
PDA
Foz
44963
WA
PDA
WA
Acidovorax N1
Dz2
g
Foz
44987
h
Foz
44987
Foz
44963
Foz
44963
PDA
WA
Pseudomonas Dz5
e.
f.
g.
h.
PDA
WA
P. fluorescens
Foz and Dz2 field isolate (unidentified).
Foz and Acidovorax N1 (field isolate).
Foz and Pseudomonas Dz5 (field isolate).
Foz and P. fluorescens (wild type isolate)
170
Figure 27. continued…
i
Foz
44987
j
Foz
44987
Foz
44963
PDA
WA
A. lipoferum
PDA
WA
A. brasilense Sp7
k
Foz
44963
PDA
WA
Bacillus F2
Foz
44987
PDA
WA
Bacillus F1
l
Foz
44963
PDA
WA
B. coagulans 10334
PDA
WA
B. subtilis 1184
Foz 44963, PDA
i.
Foz and A. lipoferum Br-17 (2 plates on left), A. brasilense Sp7 (2 plates
on right).
j. Foz and Bacillus F2 and Bacillus F1 (field isolates)
k. Foz and B. coagulans 10334, B. subtilis 1184
l. F. oxysporum f. sp. zingiberi BRIP 44963
171
6.3.2. Dual culture assays on microscope slides.
The interaction between bacterial isolates and Foz on microscope slide agar films is
shown in Figure 28 – Figure 39.
.
Figure 28. Microscope slide agar film culture of Foz alone or with bacterial isolates.
Foz 44963
B. subtilis A13 B. subtilis 26659
Foz 44963 Pseudomonas Dz5
Dz11
Foz 44963
B. megaterium B. subtilis 6633
Foz 44963 A. brasilense Sp7
Dz2
a. Foz 44963 and B. subtilis A13, B. subtilis 26659 on a microscope slide agar film.
b. Foz 44963 and B. megaterium NCTC 10342, B. subtilis ATCC 6633 on a
microscope slide agar film.
c. Foz 44963 and Pseudomonas Dz5, Dz11 (field isolates) on a microscope slide
agar film.
d. Foz 44963 and A. brasilense Sp7, Dz2 (field isolate) on a microscope slide agar
film.
172
Figure 29. Culture of Foz on agar film on a microscope slide agar film under
magnification. Arrows indicate chlamydospore like structures at hyphal tips. Hyphal
growth is straight and radially oriented, with infrequent branching near hyphal tips.
a
b
cc
a.
b.
c.
d.
d
Stereomicroscope microscope image.
Stained with lactophenol cotton blue at 40X magnification.
Stained with lactophenol cotton blue at 400X magnification.
Stained with lactophenol cotton blue at 400X magnification.
173
Figure 30. Microscope slide agar film culture of Foz and B. subtilis DAR26659 under
magnification. White arrow indicates inhibition of radially oriented hyphal growth and
compacted growth of hyphae; green arrows indicate lysis of hyphae; red arrows indicate
abnormal hyphal branching; orange arrow indicates altered directionality of growth and
increased looping growth of hyphae.
a
b
c
d
a.
b.
c.
d.
Stereomicroscope microscope image.
Stained with lactophenol cotton blue at 40X magnification.
Stained with lactophenol cotton blue at 100X magnification.
Stained with lactophenol cotton blue at 400X magnification.
.
174
Figure 31. Microscope slide agar film culture of Foz and B. subtilis A13 under
magnification. White arrow indicates inhibition of radially oriented hyphal growth and
compacted growth of hyphae; green arrows indicate chlamydospore – like structures
within the hyphae (compared to the hyphal tip in the control); orange arrow indicates
altered directionality of growth.
a
b
c
d
a.
b.
c.
d.
Stereomicroscope microscope image.
Stained with lactophenol cotton blue at 40X magnification.
Stained with lactophenol cotton blue at 100X magnification.
Stained with lactophenol cotton blue at 400X magnification.
175
Figure 32. Microscope slide agar film culture of Foz and P. fluorescens under
magnification. White arrow indicates inhibition of radially oriented hyphal growth and
compacted growth of hyphae; red arrow indicates increased hyphal branching; orange
arrow indicates altered directionality of growth and increased looping growth of hyphae,
green arrow indicates increased coiling of hyphae.
a
b
c
d
a.
b.
c.
d.
Stereomicroscope microscope image.
Stained with lactophenol cotton blue at 40X magnification.
Stained with lactophenol cotton blue at 100X magnification.
Stained with lactophenol cotton blue at 400X magnification.
176
Figure 33. Microscope slide agar film culture of Foz and A. brasilense Sp7 under
magnification. Red arrow indicates growth of Foz into the cells of A. brasilense Sp7;
green arrow may indicate products from lysis of bacteria.
]
a
b
c
d
a.
b.
c.
d.
Stereomicroscope microscope image.
Stained with lactophenol cotton blue at 40X magnification.
Stained with lactophenol cotton blue at 100X magnification.
Stained with lactophenol cotton blue at 400X magnification.
177
Figure 34. Microscope slide agar film culture of Foz and B. subtilis ATCC 6633 under
magnification. Orange arrow indicates inhibition of radially oriented hyphal growth and
compacted growth of hyphae; green arrows indicate fragmentation of hyphae, red
arrows indicate altered directionality of growth.
a
b
c
d
a.
b.
c.
d.
Stereomicroscope microscope image.
Stained with lactophenol cotton blue at 40X magnification.
Stained with lactophenol cotton blue at 100X magnification.
Stained with lactophenol cotton blue at 400X magnification.
178
Figure 35. Microscope slide agar film culture of Foz and B. megaterium NCTC 10342
under magnification. White arrow indicates inhibition of radially oriented hyphal growth
and compacted growth of hyphae; orange arrow indicates altered directionality of
growth.
a
b
c
d
a.
b.
c.
d.
Stereomicroscope microscope image.
Stained with lactophenol cotton blue at 40X magnification.
Stained with lactophenol cotton blue at 100X magnification.
Stained with lactophenol cotton blue at 400X magnification.
179
Figure 36. Microscope slide agar film culture of Foz and Pseudomonas Dz5 under
magnification. White arrow indicates inhibition of radially oriented hyphal growth and
compacted growth of hyphae; green arrows indicate increased branching near the
hyphal tip; red arrows indicate increased looping growth and coiling of hyphae.
a.
b.
c.
d.
Stereomicroscope microscope image.
Stained with lactophenol cotton blue at 40X magnification.
Stained with lactophenol cotton blue at 100X magnification.
Stained with lactophenol cotton blue at 400X magnification.
180
Figure 37. Microscope slide agar film culture of Foz and Dz11 under magnification.
White arrow indicates inhibition of radially oriented hyphal growth and compacted
growth of hyphae; green arrows indicate abnormal/increased hyphal branching.
a.
b.
c.
d.
Stereomicroscope microscope image.
Stained with lactophenol cotton blue at 40X magnification.
Stained with lactophenol cotton blue at 100X magnification.
Stained with lactophenol blue at 400X magnification.
181
6.3.3. Effect of bacterial treatments on incidence of Foz infection in ginger tissue
culture plants.
Results of the Fusarium trial are summarised in Table 39 and Figure 37 to Figure 40.
Inoculation with low levels of Foz resulted in inconsistent infection of ginger plants.
While yellowing of above ground plant material was evident amongst many plants,
brown discoloration of rhizomes was observed less frequently (~20% of plants in the
inoculated control, Table 39, Table 40). Fungi morphologically typical of Foz were only
obtained from rhizome isolations in approximately 5% of the inoculated controls. Thus
yellowing of above ground plant material may have been in part due to leaf senescence,
as a result of shortened day length nearing completion of the trial. However, plant dry
weight, rhizome dry weight and plant height were significantly reduced in plants
inoculated with Foz compared to the seed control, indicating the pathogen negatively
impacted on plant growth.
While the incidence of disease symptoms was slightly
reduced in plants inoculated with B. subtilis A13 and B. subtilis DAR26659, differences
were not significant, which may have been associated with inconsistent infection by
Foz. A marginal effect of B. subtilis A13 on the growth of ginger tissue cultured plants
was observed.
Growth promotion that resulted from the introduction of B. subtilis
DAR26659 was highly significant, where all plant growth parameters were increased, by
up to 64%, when compared to the inoculated control.
Following the application of
P. fluorescens shoot dry weight, rhizome fresh weight and rhizome dry weight were
increased by 19.1%, 9.9% and 9.1% respectively, although differences were not
significant. In this treatment group several plants displayed extensive wilting and plant
death; given that Foz was not isolated from rhizomes, it is unclear whether plant growth
was negatively affected as a result of the bacterial treatment or whether this was due to
unrelated factors. Growth was significantly reduced in plants that were inoculated with
B. subtilis ATTC 6633. Statistically significant increased growth was observed in the
seed control compared to the water control, which indicated a positive effect of the rye
grass seed material on the growth of ginger plants.
182
Figure 38. Glasshouse trial assessing effect of bacterial treatments on growth and
infection of ginger tissue culture plants by Foz.
a
b
b
a
c
1 2 3 4 5 6 7 8 9 10
d
a. 4 weeks after planting.
b. 16 weeks after planting.
c. Orange arrow indicates healthy rhizome, green arrow indicates discoloured
rhizome.
d.
1. Water control
2. Seed control
3. Foz inoculated control
4. A. brasilense Sp7
5. B. megatarium,
6. P. fluorescens
7. B. subtilis 22659
8. B. subtilis A13
9. B. subtilis ATCC 6633
10. Combination of strains
(treatments (4-10 were also inoculated with Foz)
183
Table 39. Effect of bacterial treatments on mean growth parameters and incidence of Foz infection (± standard deviation)
in tissue cultured ginger plants (Fusarium trial).*
Treatment
Plant Fresh
Weight (grams)
Plant Dry
Weight (grams)
Rhizome Fresh
Weight (grams)
Rhizome Dry
Weight (grams)
Shoot Fresh
Weight (grams)
Shoot Dry
Weight (grams)
Water
85.7 ± 23.5ad
9.2 ± 2.7ae
23.0 ± 2.8a
4.3 ± 1.5ab
16.7 ± 6.9ae
1.9 ± 0.7a
Seed Control
98.1 ± 19.8b
11.2 ± 2.2b
25.2 ± 7.8a
4.9 ± 1.1a
20.2 ± 6.6abd
2.2 ± 0.6bcd
Foz
91.4 ± 18.4ab
9.7 ± 1.7ac
23.2 ± 5.9a
4.1 ± 1.2be
19.6 ± 6.0abd
2.1 ± 0.4abc
A. brasilense Sp7
95.6 ± 15.8ab
11.0 ± 2.3bc
25.2 ± 5.9a
4.5 ± 1.0ab
17.8 ± 4.2ade
2.1 ± 0.4abc
B. megaterium NCTC 10342
99.5 ± 18.3b
10.1 ± 1.9abc
21.6 ± 6.9a
3.9 ± 1.1be
20.1 ± 4.5abd
2.3 ± 0.4cd
P. fluorescens
101.5 ± 23.8b
10.8 ± 2.9bc
25.5 ± 4.9a
4.5 ± 1.7ab
22.8 ± 4.0b
2.5 ± 0.3d
B. subtilis DAR26659
133.2 ± 19.8c
15.1 ± 2.3d
37.2 ± 8.1b
6.7 ± 1.5c
30.3 ± 6.9c
3.2 ± 0.7e
B. subtilis A13
91.5 ± 19.6ab
9.7 ± 2.0ac
23.9 ± 7.3a
4.4 ± 1.3ab
17.0 ± 4.5ae
1.9 ± 0.4abc
B. subtilis ATCC 6633
78.7 ± 22.3d
8.2 ± 2.3e
21.5 ± 6.6a
3.4 ± 1.2de
15.2 ± 4.6e
1.9 ± 0.5abc
All Bacillus and A. brasilense Sp7
101.4 ± 18.8b
10.6 ± 2.4bc
24.3 ± 7.5a
4.1 ± 1.4be
21.4 ± 11.5bd
2.3 ± 0.6bd
Table 39 continued …
Treatment
Root Fresh
Weight (grams)
Root Dry
Weight (grams)
Number of
Shoots
Height
(cm)
Rating of Shoot
Yellowing
Discoloured
Rhizomes
Water
46.0 ± 13.9ae
3.1 ± 1.3ae
3.5 ± 2.3a
366.4 ± 50.6ab
1.5 ± 1.1ac
1.00 ± 0.00a
Seed Control
52.7 ± 12.2ad
4.0 ± 1.3bd
4.3 ± 2.0ac
386.2 ± 51.1a
1.2 ± 0.4a
1.00 ± 0.00a
Foz
48.6 ± 15.1ae
3.4 ± 1.2ade
4.9 ± 1.8bc
335.8 ± 89.0b
1.5 ± 1.0ac
1.27 ± 0.55a
A. brasilense Sp7
52.6 ± 12.4ad
4.5 ± 2.2bc
4.0 ± 1.7ac
372.4 ± 51.0a
1.5 ± 0.6ac
1.36 ± 0.90a
B. megaterium NCTC 10342
57.8 ± 13.8bcd
3.9 ± 1.0bd
5.0 ± 1.4bc
355.0 ± 40.5ab
1.5 ± 1.0ac
1.32 ± 0.78a
P. fluorescens
54.2 ± 12.8ad
3.9 ± 1.4bd
5.4 ± 1.4b
364.8 ± 44.7ab
1.5 ± 1.1ac
1.32 ± 0.72a
B. subtilis DAR26659
65.6 ± 15.4c
5.1 ± 1.7c
5.9 ± 2.4b
433.2 ± 81.8c
1.2 ± 0.5ac
1.14 ± 0.47a
B. subtilis A13
50.6 ± 13.8ad
3.4 ± 1.0ade
3.6 ± 1.3a
370.9 ± 61.8a
1.3 ± 0.7ac
1.05 ± 0.21a
B. subtilis ATCC 6633
41.9 ± 14.8e
3.0 ± 1.5e
3.6 ± 1.4a
367.9 ± 49.4ab
2.1 ± 1.7bc
1.32 ± 0.65a
All Bacillus and A. brasilense Sp7
55.6 ± 16.0ad
4.1 ± 1.3bd
4.4 ± 1.3ac
366.2 ± 36.9ab
1.6 ± 1.1ac
1.27 ± 0.63a
* Mean difference between numbers with the same letter is not significant (LSD, p<0.05).
184
Figure 39. Effect of bacterial treatments on the fresh weight of tissue cultured ginger
plants that were inoculated with Foz.
Bars
represent
±
standard
deviation.
AB7:
A.
brasilense
Sp7;
BM: B. megaterium NCTC 10342;
PF: P. fluorescens ;
BS 59: B. subtilis
DAR26659; BS6633: B. subtilis ATCC 6633; Bac + AB: Combination of Bacillus
strains with A. brasilense Sp7.
185
Figure 40. Effect of bacterial treatments on the dry weight of ginger tissue cultured
plants that had been inoculated with Foz.
Bars
represent
±
standard
deviation.
AB7:
A.
brasilense
Sp7;
BM: B. megaterium NCTC 10342;
PF: P. fluorescens ;
BS 59: B. subtilis
DAR26659; BS6633: B. subtilis ATCC 6633; Bac + AB: Combination of Bacillus
strains with A. brasilense Sp7.
186
Figure 41. Effect of bacterial treatments on growth parameters and development of
symptoms of Foz infection (number of yellow shoots and rhizome discoloration).
D. Mean Rhizome Discolouration
Rh i zo m e d i sc o u r a ti o n
2.5
2
1.5
1
0.5
0
Water Seed FOZ AB7
BM
PF BS59 BS13 BS66 Bac/A
33
B7
Treatment
Bars
represent
±
standard
deviation.
AB7:
A.
brasilense
Sp7;
BM: B. megaterium NCTC 10342;
PF: P. fluorescens ;
BS 59: B. subtilis
DAR26659; BS6633: B. subtilis ATCC 6633; Bac + AB: Combination of Bacillus
strains with A. brasilense Sp7.
187
6.4. Discussion
Dual culture assays indicated that several bacterial isolates strongly antagonised the
growth of Foz; other isolates moderately inhibited the growth of Foz and; certain
isolates did not inhibit the growth of Foz at all and increased proliferation of the fungus
occurred along the streak of bacteria (Table 38).
Further analyses of these Foz-
bacteria in vitro interactions on microscope slide agar films demonstrated that
inhibition of the radially oriented hyphal growth at the leading edge of the Foz was
often associated with compacted growth and increased branching and looping growth
of hyphae.
Bolwerk et al. (2003) reported analogous anomalies in the growth of
Fusarium oxysporum f. sp. lycopersici (Fol) during in vitro culture with a biocontrol
strain of P. chlororaphis. These researchers also observed abnormal chlamydospore
like structures were formed within the hyphae of Fol when co-cultured with
P. chlororaphis. Similarly, in the present study chlamydospore-like structures were
present in hyphal tips when Foz was cultured alone, but were not observed when Foz
was co-cultured with antagonistic bacteria, with the exception of B. subtilis A13 where
abnormal chlamydospore-like structures were observed within the hyphae. Inhibition
of chlamydospore spore formation is significant as these structures are associated
with the long-term survival of Foz in the soil. Thus, antagonistic bacteria such as
those described might have the potential for reducing persistent populations of Foz in
contaminated soils.
B. subtilis A13, is the parent strain of the well-characterised commercial strain
B. subtilis GB03, which is known to produce fungitoxic iturin lipopeptides (Kloepper et
al. 2004b). The iturin lipopeptide mycosubtilin, as well as surfactin are produced by B.
subtilis ATCC 6633 (Leenders et al. 1999), which caused fragmentation of hyphae
when co-cultured with Foz in this study. Fengycins are a further type of extracellular
antifungal lipopeptides excreted by certain B. subtilis strains (Romero et al. 2007;
Koumoutsi et al. 2004; Ongena et al. 2007). Iturins, surfactins and fengycins, may
insert into cell membranes forming ion-conducting pores; this results in increased
permeability to K+ and other ions and membrane destabilisation which may cause cell
188
death (Maget-Dana et al. 1992; Maget-Dana and Peypoux 1994; Heerklotz and Seelig
2001; Deleu et al. 2005; Grau et al. 2000; Sheppard et al. 1991; Montesino 2007;
Mizumoto et al. 2006; Vanittanakom et al. 1986). In the present study, it is possible
that antifungal lipopeptides produced by B. subtilis DAR26659 contributed to the lysis
of Foz hyphae by this bacterium. The production of extracellular bacterial enzymes
(chitanase or glucanase) that degrade components of fungal cell walls (Whipps 2001)
may have been a further mechanism involved in lysis of hyphae by B. subtilis
DAR26659. In certain instances both lytic enzymes and lipopeptides may be required
to antagonise the growth of fungal pathogens (Harish et al. 1998). The production of
fungitoxic metabolites or lytic enzymes may enable bacteria to use fungal products as
a substrate following hyphal lysis, referred to as bacterial mycophagy (Ahn et al. 2006;
Kamilova et al. 2007).
Antifungal lipopetides and metabolites have also been implicated in biocontrol of
fungal plant pathogens by certain Pseudomonas spp. (Koumoutsi et al. 2004;
Raajmakers et al. 2006; de Souza et al. 2003). Bolwerk et al. 2003 demonstrated the
antifungal metabolite phenazine-1-carboxamide (PCN) produced by the biocontrol
strain P. chlororaphis PCL1391 caused anomalies in the in vitro growth of
Fusarium oxysporum f. sp. lycopesici. The authors proposed that inhibition of radially
oriented hyphal growth, increased formation of hyphal branching and “looping growth”
may have been a result of the influence of the phenazine compounds on hyphal
electrical currents that may be involved in polarised growth. When examining the
interaction of fluorescently labelled Fol and biocontrol Pseudomonas strains in the
tomato rhizosphere, it was suggested that the increased branching and altered
directionality of hyphal growth may have indicated the fungus was attempting to find
penetration sites not colonised by the antagonistic bacteria (Bolwerk et al. 2003).
Similarly in the current study, increased formation of hyphal branches (that often did
not undergo extension) and looping growth may have indicated that Foz was
attempting to avert fungitoxic substances and grow away from the antagonistic
bacteria.
189
Further mechanisms that may have been involved in the inhibition of the in vitro
growth of Foz by P. fluorescens may have included sideophore production, evidenced
by the presence of yellow-green pigments. These sideophores sequester iron and
make it unavailable to other microorganisms (O’Sullivan and O’Gara 1992).
Production of volatile substances is a further mechanism by which antagonistic
bacteria may inhibit the in vitro growth of pathogenic fungi (Desai et al. 2002).
It is
interesting that in this study an altered colour of Foz (red instead of purple) was
observed in dual culture with Pseudomonas spp. It is known that increased formation
of sclerotia (compacted, detached mycelial mass that may undergo dormancy) may
result in Fusarium oxysporum cultures having a blue colour (Nelson et al. 1983). Thus
it is tempting to speculate that the red colour of Foz in certain dual culture assays may
have been due inhibition of sclerotia formation (blue colour).
The increased proliferation of Foz along the streak of other bacteria such as
A. brasilense Sp7 and Bacillus F2 suggested that the fungus utilised products from
these bacteria as a growth substrate. Many fungi feed as saphrophytes, obtaining
nutrients from organisms killed by antibiosis (Black 1999).
Microscope slide assays
indicated the directional hyphae of Foz grew toward and into the cells of A. brasilense
Sp7; darker staining of the medium at the zone of contact possibly indicated lysis of
the bacterium.
Similarly, Barron (1988) demonstrated that Pseudomonas and
Agrobacterium isolates stimulated the formation of unbranched, directional hyphae by
several types of fungi. These hyphae grew into bacterial colonies causing lysis of the
bacterial cells and presumably used bacterial products as a nutrient source as
increased proliferation of the fungus occurred on the bacterial colony on water agar.
Such activity was not demonstrated by F. oxysporum, although this may have been
due to the type of bacteria and fungal strains tested in that study (Baron 1988).
In
addition Lasik and colleagues (1979) demonstrated that the wheat pathogen
Gaeumannomyces graminis var. tritici preferentially used polysaccharides of bacterial
origin rather than those derived from the wheat mucigel. Thus the hypothesis that
roots colonised with a bacterium that may act as a substrate or attractant for
pathogenic fungi may result in increased incidence of disease was considered, as
190
inoculation with certain bacteria may result in increased incidence of disease.
Few
studies have been reported that have assessed the effect of inoculation with
A. brasilense on disease development in plants. Bashan and Bashan (2002) reported
a reduced incidence of P. syringae infection of tomato inoculated with A. brasilense
Cd and proposed mechanisms involved may have been displacement of the plant
pathogen or reduced disease as a result of increased plant vigour. In contrast,
Romero et al. (2003) demonstrated a two fold increase in the severity of bacterial spot
(Xanthomonas campestris) in cherry tomato but not fresh market tomato following
inoculation with A. brasilense Sp7, and proposed this may have been attributed to
undefined molecules secreted by the bacterium that influenced plant signalling. In the
current study, infection of ginger tissue cultured plantlets by Foz, as determined by
rhizome discoloration, was slightly increased in plants treated with A. brasilense Sp7
compared to control plants inoculated with Foz, although differences were not
statistically significant. In general, inconsistent infection by Foz was observed in this
trial, therefore effects of bacterial treatments on disease progression could not be
substantiated.
Symptoms of Foz infection were slightly reduced in B. subtilis A13, B. subtilis
DAR26659 and P. fluorescens treatments compared to the inoculated control,
although again differences were not significant.
A reason for lack of disease
development (typically assessed by rhizome discoloration) may have included
insufficient levels of Foz inoculum used.
It is also possible that conditions were not
amenable to the growth of Foz, as water potential and temperature have been
previously shown to affect disease incidence/severity incited by fungal pathogens
(Landa et al. 2004; Harveson 1998). Even though infection by Foz was inconsistent,
significant reductions in plant dry weight, rhizome dry weight and height in plants
inoculated with Foz compared to the seed control indicated that the fungus negatively
impacted on plant growth. Given that rhizome discoloration and isolation of Foz from
rhizomes was infrequent, it is possible that infection of plants had occurred in roots
and had not progressed to the rhizome in many plants (due to time course of
191
infection).
However, Pegg and colleagues (1974) reported that entry of Foz is
primarily via the rhizome.
With respect to growth promoting activities of bacteria observed in the current trial,
treatment of plants with A. brasilense Sp7 increased rhizome fresh weight and root dry
weight by 8.5% and 31.3% respectively, although differences were not significant.
This is consistent with results of earlier trials, where suspensions of this bacterium
increased rhizome growth by 10% to 16% (p>0.05).
Plant growth was marginally
affected by the introduction of B. subtilis A13 (increased rhizome weight of 6%,
p>0.05) when compared to the inoculated control in this trial, while in previous trials in
one instance a trend toward improved growth and in another a trend toward reduced
growth was documented. Thus a variable growth response of ginger plants to the
introduction of B. subtilis A13 has been observed.
A commercial formulation of this
strain was the first successfully registered biopesticide in the USA; this product has
been superseded by a preparation containing a derivative of this strain, B. subtilis
GB03 that has superior root colonising ability. GB03 is used extensively in the USA in
cotton for reduced infection by F. oxysporum (Brannen and Kenny 1997; Jacobsen et
al. 2004).
B. subtilis GB03 has also been combined with Bacillus amyloliquefaciens
IN937a, which acts by inducing systemic resistance, in a chitin based preparation, for
growth promotion and resistance to Fusarium oxysporum mediated diseases in cotton
and a variety of vegetable crops (Kloepper et al. 2004b). These commercial examples
have demonstrated the feasibility of the use of bacterial inoculants for growth
promotion and reduced disease in agricultural production systems.
In this study, marked improvement in growth of tissue culture plants resulted from
introduction of
B. subtilis DAR26659 (all growth parameters were increased, by ~
60% for rhizome fresh and dry weight, ~50% for shoot fresh weight, shoot dry weight
and root dry weight. p<0.0001). This B. subtilis strain, which also caused lysis and
abnormal hyphal branching of Foz in vitro, was isolated from diseased wheat seed
(Noble, personal communication 2007). Many efficient biocontrol agents have been
isolated from diseased plants (Mew et al. 1994). The manifestation of plant disease
192
may alter microbial populations detected in the rhizosphere, for example populations
of antibiotic producing fluorescent Pseudomonas spp. may be either increased or
depressed in the rhizosphere of diseased wheat plants, depending on the infecting
phytopathogen (Mazzola and Cook 1991).
The development of take all decline
following the monoculture of wheat and suppressiveness to Fusarium wilt following
monoculture of peas has been associated with an accumulation of antibiotic producing
fluorescent
pseudomonads (McSpadden-Gardener and Weller 2001; Landa et al.
2002). In addition, the colonisation of hyphae of plant pathogenic fungi by certain
Pseudomonas spp. is involved in the biocontrol activity of these bacteria (de Weert et
al. 2004; Yang et al. 1994). The ability of certain biocontrol bacteria to then cause
lysis of hyphae and use fungal products as a substrate (Ahn et al. 2006; Kamilova et
al. 2007) may explain their increased incidence in diseased plants. In general, the use
of biocontrol PGPB is generally effective as a preventative measure, and not when the
plant has already succumbed to infection. Thus when a plant is colonised by such
bicontrol PGBP, these bacteria may prevent infection by phytopathogens. Given the
observed in vitro antagonism toward the growth of Foz and a marked increase in
growth of micropropagated plants, B. subtilis DAR26659 may have potential for use as
a PGPB in ginger.
6.5. Conclusion
In conclusion, a number of bacterial strains that were able to antagonise the in vitro
growth of Foz were identified in this study. Of these bacteria, B. subtilis DAR26659
caused antagonism of radially oriented growth and hyphal lysis of Foz in vitro. This
strain of B. subtilis produced striking increases (~60%) in the growth of ginger tissue
cultured plants and this is the first report of growth promoting activity of this bacterium.
While the incidence of Foz infection in tissue culture plants was slightly reduced
following introduction of B. subtilis DAR26659, assessment of the biocontrol activity of
this bacterium in planta was limited by inconsistent infection of plants by the pathogen.
193
Thus B. subtilis DAR26659 is considered a candidate microorganism for further
investigation of the ability to induce growth promotion and resistance to Foz in ginger.
194
Appendix 6.1. Media
Waksman Agar contained per liter: 5g proteose peptone, 10g glucose, 3g meat extract,
5g NaCl, 20g agar pH 6.8.
Appendix 6.2. Supplementary data for Fusarium trial.
Table 40. Percentage difference between treatment means, demonstrating the effect of
bacterial treatments on the growth of ginger tissue cultured plants inoculated with Foz.
Treatment
Plant
FW
Plant Rhizome Rhizome Shoot
DW
FW
DW
FW
Shoot
DW
Root
FW
Root
DW
No. of
Shoots
Height
(cm)
Water
-6.2
-5.2
-1
3.9
-15
-10.8
-5.3
-10
-28.6
9.1
Seed Control
7.3
15.2
8.5
20
2.8
5.5
8.5
18.7
-12.8
15
Foz
…
…
…
…
…
…
…
…
…
…
A. brasilense Sp7
4.6
13.5
8.5
9
-9.1
-1
8.2
31.3
-17.4
10.9
B. megaterium
NCTC 10342
8.8
3.7
-7
-5.5
2.5
8.6
18.9
14.9
1.1
5.7
P. fluorescens
11
11.1
9.9
9.1
16.2
19.1
11.5
15.1
10.8
8.6
B. subtilis 26659
45.7
54.9
60.4
64.1
54.8
53.6
35
49.1
19.7
29
B. subtilis A13
0.1
-0.4
3.2
6.4
-13
-7.9
4
-1.1
-25.8
10.5
B. subtilis 6633
-13.9
-15.9
-7.2
-16.9
-22.4
-10.7
-13.7
-12.9
-25.8
9.6
9.2
4.7
2
9.3
9.8
14.5
20.6
-10.9
9.1
Bacillus + A.
brasilense Sp7
195
Chapter 7. General Discussion and Conclusions
Discussion
There is an increasingly recognised need to implement sustainable practices in
agricultural production systems, to maintain and improve productivity of soils and also to
address human health and environmental concerns associated with intensive farming
(Wolf and Synder 2003; Garbeva et al. 2004).
International research has targeted
integrated approaches to sustainable agriculture, where inoculants containing plant
growth promoting bacteria (PGPB) may play a role (Jacobsen et al. 2004; MercadoBlanco and Bakker 2007). In addition, the development of bacterial inoculants that
promote resistance to seed- and soil-borne plant pathogens would be of significant
value where no other means of disease control exists or where chemical controls are
limited (for example in organic production) (Fravel 2005; Lucy et al. 2004; Compant et
al. 2005).
In addition the ability of PGPB to reduce required application rates of
inorganic fertilisers, enhance nutrient uptake and increase yield in crop production are
further reasons why research in this area is valuable (Dobbelaere et al. 2004; Vessey
2003; Okon and Labandera-Gonzalez 1994; Muthukumarasamy et al. 2006).
Mechanisms by which PGPB may improve plant growth and resistance to disease have
been extensively investigated (Bashan et al. 2004; Kloepper et al. 2004; O'Sullivan and
O'Gara 1992; Cook et al. 1995; Whipps 2001; van Loon et al. 2006; Lutgenberg et al.
2002). PGPB have been isolated from rhizosphere soils, plant surfaces/tissues and
composts (Hallmann et al. 1997; Krause et al. 2003; Fisher et al. 2006; Cazorla et al.
2007).
The non-selective culture of beneficial microorganisms that are present in
compost, to produce “compost tea”, which is applied to crops as a source of
microorganisms that may provide benefits for plant growth and soil health, is an
increasingly popular agricultural practice (Scheuerell and Mahaffee 2002; Ingram and
Milner 2007). As stated by Noble and Roberts (2004) concerns about the presence of
potentially harmful organisms (plant and human pathogens) are a major limitation to the
increased uptake of composted waste by potential end users in the horticultural and
agricultural sectors. A number of studies demonstrated that human pathogen indicator
organisms were able to grow in compost tea produced with the soluble carbon additive
molasses (Duffy et al. 2004; Ingram and Milner 2005). The USA National Organic
196
Standards Board (NOSB) recommended both quality assurance testing to demonstrate
that a particular system produces compost tea with low levels of indicator organisms
and withholding periods for compost teas made with additives (CTTFR 2004). In
Australia guidelines advising on the use of compost tea have not been established. In
addition reports that document the types of beneficial organisms present in compost
teas are lacking.
Accordingly, in the current study, prior to testing the effects for
compost tea on the growth of ginger, microbiological analyses were performed to
determine if human pathogenic organisms could be detected. Results indicated that
faecal coliforms could be present at high levels in aerated teas (approximating levels
found in raw sewerage) despite the absence of “foul smells” (a common method for the
assessment of the potential for compost tea to contain human pathogenic organisms).
Analyses suggested that enteric contaminants that were detected at high levels in the
liquefied compost prior to fermentation persisted in aerated cultures produced with a
variety of growth substrates including those that did not contain soluble carbon based
additives. This is in agreement with findings of Sturz et al. (2006) who found Klebsiella
pneumoniae and Escherichia spp.
preparation.
were present in a commercial compost tea
In addition, these bacteria were detected in the potato phylloplane
following spray application of the compost tea (Sturz et al. 2006). A recent study by
Ingram and Milner (2007) also demonstrated that pathogenic indicator organisms were
able to grow in compost tea produced with additives such as kelp, even when soluble
carbon additives were not used. The growth of pathogenic indicator organisms was
also shown to in the additives alone (Ingram and Milner 2007). Similarly, in the current
study microbial contaminants were isolated from additives used in the production of
compost teas/microbial cultures, which might result in the end product of fermentation
not containing the desired microorganisms.
Ingram and Milner (2007) also reported that when enteric bacteria were not detected in
the compost source material and the compost tea was produced without additives, the
growth of enteric bacteria in the tea did not occur (Ingram and Milner 2007). However,
the efficacy of teas produced without additives and whether other compost resistant
pathogens, such as Legionella spp., Clostridium spp. and Pseudomonas aeruginosa,
are able to grow in these preparations remains to be assessed. Biofilm formation and
197
sanitation of culture and fertigation (application) equipment is also an area for further
consideration (Sadovski et al. 1978; Martin and Bull 2002).
In attempting to avert the culture of pathogenic microorganisms introduced from the
compost source material, two commercially available mixed microbial inoculants,
purported to contain non-pathogenic beneficial microorganisms (Bacillus spp.,
Pseudomonas spp., Rhizobium spp., Trichoderma spp. and others) were used to
produce microbial cultures by methods similar to those used for compost tea (open air
containers, non-sterile conditions and active aeration).
Culture based analyses
indicated that pathogenic organisms were present following fermentation of these
inoculants, despite the use of growth substrates that did not contain pathogenic
contaminants. Testing of the untreated microbial inoculants suggested that they were a
source of contaminants.
Phylogenetic analysis (16S rDNA sequencing) and
biochemical testing (by independent commercial laboratories and performed in this
study) indicated that these contaminants included Enterobacteriaceae, Klebsiella
pneumoniae,
Pseudomonas
aeruginosa,
Bacillus
cereus,
Stenotrophomonas
maltophilia and Photobacterium damsela. The detection of these human pathogenic
bacteria in commercial preparations supports the view of Kennedy et al. (2004), that
regulating agencies need to develop and administer quality control standards in the
Australian biofertiliser industry. Such quality control standards, administered by the
Australian Legume Inoculant Research Unit, have been associated with the success of
Rhizobium inoculants in the legume industry (Bullard et al. 2005; Deaker et al. 2004).
Long-term exposure to bioaerosols, similar to those that may be generated during the
production and spray application of compost teas/microbial cultures, has been
associated with diseases such as hypersensitivity pneumonitis, allergic alveolitis and
chronic obstructive pulmonary disease (Millner et al. 1994; Hansen et al. 2003; Bunger
et al. 2005; Ivens et al. 1999). Exposure to bioaerosols is of particular concern where
pathogens are present that may be transmitted via the respiratory route, such as
Klebsiella pneumoniae. In this study, risk analysis indicated that the use of personal
protective equipment (PPE) only reduced risks associated with exposure to bioaerosols
generated during the production of microbial cultures containing human pathogenic
organisms to a moderate level, which still required correction. The use of a bioreactor
198
(enclosed vessel) to contain bioaerosols reduced the risk associated with exposure to
bioaerosols generated during production of contaminated cultures to a low (acceptable)
level. In accordance with these risk reduction measures (use of PPE and bioreactors),
the common practice of active sniffing of compost teas, recommended as a way to
assess their pathogenic potential (Ingham 2004), should be strongly discouraged.
In order to reduce risks associated with the application of microbial cultures, it was
suggested that drip irrigation would not be expected to produce the same type of
bioaerosols as spray application.
Further studies are required to assess the nature of
bioaerosols produced by drip-application of microbial cultures/compost tea. Application
to the soil may also reduce the risk of contamination of fresh produce by food-borne
pathogens, in a similar manner to drip irrigation compared to spray irrigation (Sadovski
et al. 1978). As suggested by Canadian governments, compost tea should not be
applied to edible portions of plants and withholding periods recommended by the NOSB
should be observed to reduce the risk of contamination of fresh produce by food-borne
pathogens following compost tea application (Ministry of Agriculture and Lands MOAL
2005; CTTFR 2004).
However, drip irrigation is not suitable for all broad acre
applications, including many ginger production systems, due to the high density of
plants cultivated per hectare and the need for spray irrigation to prevent sunburn of
leaves during the early stages of the growth.
Risk analysis indicated that generation of bioaerosols during spray application of
microbial brews containing Enterobacteriaceae or K. pneumoniae was associated with a
high level of risk, even when skin, respiratory and eye protection were used. Further,
these risk control measures do not address the potential for compost teas to increase
pathogen loads in the environment, which might result in increased incidence of human
disease via runoff into waterways or dispersal of bioaerosols (Cabelli et al. 1979; Entry
and Farmer 2001). The distance that bioaerosols may be carried downwind from the
point of application of compost teas is not known.
Recer and colleagues (2001)
demonstrated that bioaerosol levels 500m downwind from a composting facility were
significantly increased above background levels.
It has also been shown that
incidences of communicable diseases were increased in communities that surrounded
agricultural sites where wastewater was applied via spray irrigation (Katzenelson et al.
199
1976). As such, further research is required to determine the distance travelled by
bioaerosols generated during the spray application of compost tea/microbial cultures, as
the dispersal of pathogenic organisms that may be present in these preparations may
constitute a public health risk.
In addition, the safe disposal of large volumes of
contaminated cultures (hundreds to thousands of litres) should be addressed, in order
to prevent run-off or leaching into waterways and groundwater, which might add to
problems of environmental microbial pollution.
Furthermore, the importation and
subsequent mass production and field application of unlegislated microbial products
might also become a threat to national biosecurity, particularly in farm environments,
exemplified by the detection of Photobacterium damsela, a fish pathogen and human
flesh eating bacteria, in a commercial inoculant in this study. Given that risks were not
mitigated even when personal protective equipment was employed during production
and application of microbial cultures containing Enterobacteriaceae and K. pneumoniae,
the non-selective culture of compost/microbial inoculant microorganisms could not be
recommended and such preparations were precluded from further research. Therefore,
this study was limited to the analysis of a relatively small number of cultures and
analysis of beneficial organisms present was not performed. As a result of this study,
there has been an increased awareness of the importance of using personal protective
equipment in the production and application of microbial brews (even though all risks
are not controlled) amongst local industries.
Further, concerns for the potential of
uncontrolled microbial culturing to result in the growth of human pathogenic organism
and associated risks to human health have been relayed to local industries.
The (small scale) application of bacterial inoculants containing Class 1 bacteria, that are
not likely to cause human disease, was determined to be associated with a low-level
risk when cultures were produced under controlled laboratory conditions and skin, eye
and respiratory protection (P2 respirator) were used during application.
Thus, in the
current study, pure cultures of laboratory produced, non-pathogenic bacterial stains
were selected for further research that targeted the use of microorganisms to improve
the growth of ginger. These bacterial strains were obtained from roots of locally grown
ginger or sourced from reference culture collections. The isolation of plant-adapted
bacteria from the rhizosphere and rhizoplane, suited to regional environmental and
seasonal conditions, also provided information on the type of bacteria that could be
200
naturally associated with the ginger root. Isolations performed during the early stages of
growth of seed grown ginger indicated that fluorescent Pseudomonas spp. were
dominant in the rhizosphere in non-fumigated soil, while Bacillus simplex/macroides
were dominant in the rhizosphere in fumigated soil (on King’s B agar). Analyses later in
the season indicated that fluorescent Pseudomonas populations in the ginger
rhizosphere in non-fumigated soil had declined. This is in agreement with previous
research that has shown that rhizosphere bacteria may be plant specific and may vary
over the growth cycle of a particular plant type (Smit et al. 2001; Mew et al. 1994; Wong
1994).
Identification of representative bacteria isolated on nitrogen-free media at this later
stage of plant growth, indicated that Pseudomonas spp., Rhizobium-Agrobacterium spp.
and Aneurinibacillus spp. were present in rhizoplane samples in non-fumigated soil,
while Acidovorax spp. and Pseudomonas spp. were isolated from rhizoplane samples in
fumigated soils.
Acidovorax spp. have previously been isolated from plant surfaces
and have been investigated for bioremediation purposes due to their ability to degrade
aromatic compounds (Andrade et al. 1997; Sun et al. 2007; Monferran et al. 2005;
Nestler et al. 2007). Thus it is possible Acidovorax spp were detected in fumigated soil
due to the biodegradative capacity of this bacterium. Rhizobium spp. have previously
been isolated in association with non-legumes, and may enhance the growth of such
plants via the production of phytostimulatory hormones or by inducing systemic
resistance (Hallman et al. 2001; Reddy et al. 1997; Kennedy et al. 1997).
While the
present study was limited to a brief analysis of ginger root associated bacteria, further
research that provides a more detailed description of the types of bacteria naturally
present in the ginger rhizosphere, rhizoplane and plant tissues, in different soil types
and at different stages of growth, and effects of introduced bacteria on these indigenous
microbial populations, may provide a basis for the development of more effective
inoculation strategies for the augmentation of PGPB populations in crop production
systems (Watt et al. 2006; Whipps 2001). The optimisation of application methods may
also improve the reliability of PGPB in field conditions (Kennedy et al. 2004; Bressan
and Borges 2004).
201
Many different of methods of application of plant growth promoting bacteria have been
reported, that vary for example in the concentration of bacterial applied, frequency or
method of application (soil/root/seed) and formulation of bacteria (use of buffers and
methylcellulose for resuspending bacteria, formulation into alginate beads, dusts and
powders). Therefore initial trials in the present study evaluated the growth response of
plants to the introduction of selected reference strains of bacteria by various methods
(soil or soil as well as seed/plant inoculation). Wheat and then ginger tissue culture
plants were used as indicator plants for assessment of growth promoting activities of
selected bacteria prior to the study of ginger grown from seed pieces (the conventional
planting material), the use of which is limited by a short, specific planting time
(September/October) and an annual growing season.
In all greenhouse trials,
fertilisers were applied at approximately half of the recommended rate. Therefore the
activity of bacteria was tested under low levels of fertiliser application.
The growth response of the indicator plant wheat, to the introduction of Bacillus subtilis
A13, Pseudomonas putida KT2442 and Bacillus coagulans NCTC 10334, was
significantly improved via application of the bacteria to soil as well as seed, compared to
application of the bacteria to seed alone. This is in agreement with other studies that
have demonstrated that application of bacteria to the soil may result in a consistent
performance of inoculants (Zehnder et al. 2001; Bressan and Borges 2004; Kloeppper
et al. 2004; Jetiyanon and Kloepper 2002).
Evaluation of selected bacterial treatments in micropropagated ginger plants indicated
that differences were not significant when comparing different application methods (soil
drench only or root dip followed by soil drench) for the introduction of
B. subtilis A13,
P. putida KT2442 and B. coagulans NCTC 10334. A high level of inherent variability
was observed in plants of the same treatment group, which may have made differences
difficult to establish.
A negative effect of soaking plant roots in 0.1 X phosphate
buffered saline (PBS) was indicated. While the growth of tissue culture plants was not
improved by bacterial treatments in this trial, a positive growth response from soaking
roots of acclimatised plants in water prior to planting in soil was suggested. Ginger
tissue culture plants, have been used to establish sites that supply industry with planting
material free from Foz. First generation tissue culture plants have a much smaller
202
rhizome than ginger plants grown from seed pieces, and produce relatively more roots
and shoots.
The identification of measures that improve rhizome growth in first
generation ginger tissue culture plants is of significance for reducing levels of wastage
due to a high incidence of plants with inferior-sized rhizomes (unsuitable for further use)
and improving productivity from this source of disease free planting material (Smith and
Hamill 1996). As discussed by Cook (2000), the use of disease free planting material
has been one of the most successful strategies in plant health management in modern
agriculture.
In a second trial (ginger tissue culture II) application of field isolate Bacillus F2,
A. brasilense Sp7 and a combination of A. brasilense Sp7 and B. coagulans NCTC
10334 (resuspended in 0.1 X PBS) resulted in increased rhizome weights of 40% to
56% compared to the buffer control. In addition, rhizome weight was reduced in the
buffer control compared to the water control. The negative effect of this buffer on plant
growth may have been due the presence of low levels of salt in the buffer, indicating a
sensitivity of micropropagated ginger plants to hyper-osmotic stress. It is known that
many types of micropropagated plants may undergo water stress during ex-vitro
acclimatisation (Nowak and Shulaev 2003). Previous reports have demonstrated that
PGPB, including A. brasilense Sp7, may improve plant growth via enhancing tolerance
to water and salt stress, by inducing the preferential uptake of potassium and enhancing
root growth (Dobbelaere et al. 1999; Okon and Labandera-Gonzalez 1994; Barassi et
al. 2006; Mayak et al. 2004; Hamdia et al. 2004). Therefore in the current study, results
indicated that the application of A. brasilense Sp7 and Bacillus F2 enabled ginger tissue
culture plants to significantly overcome hyper-osmotic stress, that may have been
caused by salt in the buffer used to apply the bacteria. Inclusion of water, as well as
buffer controls in the trial enabled these effects to be detected. The use of buffers, to
avoid subjecting bacteria to osmotic shock (Bashan et al. 1993) is not uncommon in
research that has evaluated effects of the application of bacteria to seed, plant roots
and/or soil (Jetiyanon et al. 2003; Jetiyanon and Kloepper 2002; Dobbelaere et al. 1999;
Mia et al. 2005; Njoloma et al. 2006). Most often only one control is used, either buffer
or water. Had the water control not been included in the ginger tissue culture trials, a
negative effect of the buffer would not have been identified and an over-estimation of
the positive effect of the bacteria on plant growth if the plants had not been subject to
203
salt stress may have resulted. Thus the importance of buffer as well as water controls
in greenhouse trials where bacteria are resuspended in buffer was demonstrated.
Culture based analyses were used to determine whether introduced bacteria could be
isolated from plant roots, in order to fulfil Koch’s postulate and determine levels of
bacterial colonisation. Isolates could not be distinguished from indigenous microflora by
the cultivation-dependant methods employed. Thus more specific methods may be
required to monitor introduced bacteria (Bashan et al. 1987).
For example ELISA,
employing bacteria-specific antibodies, was used to assess root colonisation by
introduced Azospirillum species (Bashan et al. 1987). Analyses using arbitrarily primed
PCR enabled the discrimination of introduced isolates of B. thuringiensis from
indigenous microflora (Brousseau et al. 1993). Antibiotic resistance, green fluorescent
protein or lacZ markers have also facilitated the tracking of introduced bacteria (Bolwerk
et al. 2003; Landa et al. 2002; Raaijmakers and Weller 2001; Notz et al. 2001).
Cultivation independent methods such as real-time PCR may also have the potential to
quantitate levels of root colonisation by introduced bacteria (Gamalero et al. 2003).
However, this method is time intensive, especially where several strains are to be
monitored, and relies on the use of specific primers, which may not be available for
bacteria such as Bacillus spp. that are ubiquitous in environmental samples.
Given
time limitations, further assessment of root colonisation was not pursued in this study,
although such analyses may provide important information for future research, for
example determining the effects of different variables (such as fertiliser, irrigation,
temperature, soil type) on levels of root colonisation may enable improved efficacy of
bacterial inoculants to be achieved.
Given the observation of high levels of inherent variability among plants of the same
treatment group, the number of replicates used per treatment was increased in
subsequent trials.
As such, a reduced number of different treatments could be
evaluated and testing was predominantly limited to assessment of effects of reference
strains of bacteria on the growth of ginger. Bacteria that promoted growth of ginger
tissue culture plants did not induce significant differences in growth parameters of
ginger grown from seed pieces (following application to seed and soil).
In this ginger
seed piece trial, a positive effect of the buffer (0.02M potassium phosphate buffer) used
204
to apply the bacteria was suggested. As the buffer contained soluble phosphate and
potassium, this may have masked effects of bacteria, as increasing potassium uptake is
a mechanism by which A. brasilense Sp7 may affect plant growth and growth promoting
activity of bacteria may not be observed under high levels of fertiliser (Lin et al.1983;
Mayak et al. 2004; Hamdia et al. 2004; Bertrand et al. 2000; Egamberdiyeva 2007;
Gunarto et al. 1999; Okon and Labandera-Gonzalez 1994). The value of buffer control
as well as water controls in this trial was again demonstrated.
The lack of a growth response of ginger grown from seed pieces may have also
resulted from failure of introduced bacteria to establish in the root environment.
Differential responsiveness of tissue cultured and seed grown ginger plants to bacterial
inoculants may result from inherent differences between these plants. More specifically,
tissue culture plants that are produced in sterile conditions do not have indigenous
microbial populations present at the time of inoculation and also have a root mass,
which may provide colonisation sites for introduced bacteria. In contrast, ginger seed
pieces do not have a root mass at the time of planting/seed-inoculation and may carry
indigenous microorganisms on the seed surface/interior that may potentially outcompete introduced bacteria for colonisation sites. The extent to which indigenous
seed piece-borne bacteria contribute to endophytic, rhizoplane and rhizosphere
populations in ginger is an area for further research.
Ginger seed pieces used in the
current study had been allowed to suberise (heal at the cut surface) for at least ten days
prior to planting, which is a common industry practice (Sanewski 2002). Increasingly
freshly cut rhizomes are used for planting.
Further research may consider the
introduction of plant beneficial bacteria to freshly cut seed pieces, which are highly
absorptive and may enable the transport of bacteria to the seed interior, where bacteria
would be protected from biotic and abiotic stresses and may have a greater prospect for
influencing plant growth (Smith, personal communication 2007). Optimisation of ginger
seed piece inoculation methods could be of significant value, as such a method could
be easily be incorporated into current seed preparation procedures. After seed pieces
are cut (from the mother rhizome) they are transported along a conveyer belt, then
immersed into a fungicide dip for several minutes before being deposited into crates for
transfer to the field. Certain bacteria, such as B. subtilis A13 are tolerant to fungicides,
and can be applied together with the fungicide during seed treatment (Backman et al.
205
1994), such that additional steps are not required for inoculation. Ideally, dipping of
ginger seed pieces into a bacterial inoculant that provides protection against Foz would
replace the fungicide dip. The development of application methods for PGPB that are
easily incorporated into current production practices is important for enabling uptake of
the technology by growers (Bashan 1998).
In the present study, a further alternative application method investigated for improved
efficacy of introduced PGPB involved the use of alginate as a carrier material. The use
of dried alginate beads for delivering A. brasilense Sp7 increased rhizome weight of
ginger tissue culture plants by 13% (p>0.05) and 25% (p<0.05) when compared to
application of this bacterium as an aqueous suspension and the water control
respectively.
Significant increases in shoot and root weight were also observed in
comparison of alginate bead formulation of A. brasilense Sp7 and the water control or
the bacterium applied as a drench. Results indicated that increased rhizome weight by
A. brasilense Sp7 was augmented by the use of the alginate bead carrier, while the
alginate bead material enhanced shoot and root growth.
Alginate contains
polysaccharides that have been shown to modulate plant growth and activate plant
defence responses at low levels (John et al. 1997; Etzler 1998; Laporte et al. 2007).
Positive effects of alginate beads on plant growth would not have been evident without
inclusion of a water control; frequently the effects of the alginate beads with bacteria is
only compared to the alginate beads without bacteria or to a liquid suspension of
bacteria (van Elsas et al. 1992; El Komy 2005; Trivedi et al. 2005; Zohar-Perez et al.
2005). Thus the value of including necessary controls in this type of trial was repeatedly
demonstrated. As well as improving performance of inoculants by protecting bacteria
from biotic and abiotic stress, alginate preparations have long-term viability, slowly
release the bacteria (thereby reducing required application rates of inoculants) and
provide a measure for containment of microorganisms (Bashan et al. 1998; van Elsas et
al. 1992). It is significant that the alginate bead formulation of A. brasilense Sp7 was
the only treatment to significantly increase rhizome weight in this trial, as the ginger
rhizome has commercial value. Therefore the use of an alginate carrier may be valuable
in further research on the use of PGPB for improved growth of ginger.
206
Lastly, the interaction of bacterial isolates with the ginger pathogen Fusarium
oxysporum forma specialis zingiberi (Foz) was investigated.
In vitro plate assays
demonstrated antagonism of the growth of Foz by a number of Bacillus and
Pseudomonas species.
Dual culture assays also indicated that proliferation of Foz
increased along the streak of other bacteria, including A. brasilense Sp7 and Bacillus
F2, suggesting that the fungus used bacterial products as a substrate. Similarly Barron
(1988) demonstrated that isolates of Pseudomonas spp. and Agrobacterium spp.
stimulated the growth of hyphae of several fungi toward colonies of these bacteria; lysis
of the bacterial cells and increased growth of the fungi suggested the bacterial products
were used as a nutrient source by the fungi.
In addition Lasik and colleagues (1979)
demonstrated that the wheat pathogen Gaeumannomyces graminis var. tritici
preferentially used polysaccharides of bacterial origin rather than those derived from the
wheat mucigel. Therefore it was hypothesized that colonisation of plant roots by a
bacterium that attracts a fungal pathogen, may increase the susceptibility of the plant to
infection, as certain bacteria may increase the incidence or severity of plant disease.
For example, Romero et al. (2003) demonstrated that bacterial spot, caused by
Xanthomonas campestri, was increased in cherry tomato following inoculation with A.
brasilense Sp7.
Symptoms of Foz infection in ginger tissue culture plants were slightly increased
following inoculation with A. brasilense Sp7 and slightly decreased by application of
B. subtilis DAR26659, B. subtilis A13 and P. fluorescens. These differences in disease
incidence between treatments and controls were not significant, which may have been
due to inconsistent infection of plants by Foz, as seen in control treatment inoculated
with the pathogen. Thus the sporadic infection of ginger plants by Foz limited the
evaluation of effects of the bacteria on disease incidence. Therefore further research is
required to establish a robust bioassay to demonstrate consistent infection of ginger
plants by Foz. This may include testing a range of Foz inoculum levels and effects of
water potential and temperature on disease progression (Landa et al. 2004; Harveson
1998).
Marked improvement in the growth of tissue culture plants resulted from introduction of
B. subtilis DAR26659 in this trial, where rhizome fresh and dry weight were increased
207
by ~ 60% and shoot fresh weight, shoot dry weight and root dry weight were increased
by ~50%. This strain also strongly inhibited the growth of Foz in vitro and caused
abnormal branching and lysis of hyphae. Therefore this strain is recommended for
further testing, to determine reproducibility of growth promoting effects, to determine
whether an even greater effect on plant growth may occur with the use of an alginate
carrier and for assessment of effects on the growth of ginger grown from seed pieces.
In three different trials a negative, positive (30% increase in rhizome weight, p>0.05) or
marginal growth response occurred following the introduction of B. subtilis A13. This
may have indicated an inherently variable response of plants to this bacterial strain or
differences may have been caused by planting time/growth conditions, previously
reported to influence the response of peanuts to the introduction of B. subtilis A13
(Turner and Backman 1991). This bacterium also antagonised the growth and caused
the formation of abnormal chlamydospore-like structures within the hyphae of Foz in
vitro. Strain A13, isolated in Australia, was one of the first commercialised biopesticides
in USA, marketed under the trade name Quantum 400®, where it was used extensively
in the cotton industry for improved resistance against Fusarium oxysporum and
Rhizoctonia soil borne pathogens (Broadbent et al. 1971; Turner and Backman 1991;
Backman et al. 1994). In peanuts, tomato, pepper and wheat, the activity of B. subtilis
A13 was also dependant on variables such as fertiliser application and water stress
(Turner and Backman 1991; Broadbent et al. 1977; Broadbent et al. 1971). Thus further
testing of B. subtilis A13, as well as A. brasilense Sp7 and B. subtilis DAR26659, under
a range of fertiliser levels and irrigation regimes may determine conditions that result in
optimal performance of these bacteria.
Additional research is also required to test the
efficacy of these bacterial strains in field soils, firstly in greenhouse conditions and then
in the field. Further studies might also determine whether introduced strains may be
transmitted via vegetative propagation of ginger, which could reduce required
application rates of bacteria.
Other areas for research, particularly for bacterial strains with commercial potential may
also include i) confirmation of the absence of human pathogenicity/virulence factors and
ii) assessment of the effect of microbial inoculants on the indigenous microflora. Conn
and Franco (2004) used terminal restriction-fragment length polymorphism with
208
actinobacteria group specific primers, to demonstrate that a commercially available
mixed microbial inoculant reduced the diversity of endophytic actinobacteria detected in
wheat roots. In contrast, no effect on indigenous actinobacterial diversity was observed
following the introduction of single strains of native actinobacteria, which also promoted
plant growth (Conn and Franco 2004). In general, it has proven difficult to establish
introduced bacterial strains in soil, and effects on indigenous microflora have typically
been minor and transient (Cook et al. 1996; Girlanda et al. 2001; Thirup et al. 2001;
Castro-Sowinski et al. 2007; Herschkovitz et al. 2005).
Difficulties in establishing
introduced bacterial populations in soil environments may result from competitive
interactions with the indigenous microflora (Turner and Backman 1991). Compant and
colleagues (2005) suggested that future research targeting the development of
inoculants of endophytic bacteria, which are comparatively well protected from biotic
and abiotic stresses might more reliably promote growth and/or resistance to disease in
field conditions.
While the use of combinations strains may improve the performance of bacterial
inoculants, in the current study combinations of strains generally reduced the growth
promoting activities of beneficial bacteria. This is in agreement with findings of Bora et
al. (2004), McSpadden-Gardener et al. (2000), and Akköprü and Demir (2005), where a
combination of P. putida strains was less effective than individual strain for the
suppression of Fusarium oxysporum. This may be due to antagonism between
combined strains, for example due to antibiosis or quorum sensing. When using
combinations of fluorescent pseudomonads Pierson and Weller (1994) also found that
the most effective combinations could vary depending on the crop. Many other
examples have demonstrated a synergistic effect of combination of strains (Guetsky et
al. 2001); Jetiyanon 2003; Rapauch and Kloepper 1998; Domenench et al. 2006; Kim et
al. 1997). Hence the development of PGPB inoculants that have a number of strains
with complementary modes of action may improve the performance and consistency of
inoculants in field conditions, although careful testing of the compatibility of strains is
required.
It is noteworthy that in the present study, the growth promoting activity of A. brasilense
Sp7 and B. subtilis DAR26659 in ginger was observed under reduced levels of fertiliser
209
application (approximately half of the recommended rate). Given that inorganic nitrogen
and super-phosphate are applied at rates of up to 750kg and 1 tonne per hectare
respectively over the growing season in commercial production (Broadley 2005), the
use of inoculants of PGPB may have the potential to alleviate the reliance of industry on
applied fertilisers. There are both significant economic benefits to growers and benefits
to the environment by reducing inputs of nitrogen fertilisers in agriculture.
More
specifically, by decreasing levels of applied nitrogenous fertilisers in agricultural
production systems leaching of nitrogen into waterways (and subsequent algal and
weed infestations) and environmental enrichment of nitrogen may be reduced (Woods
1995). Furthermore, denitrification and volatilisation of nitrogen fertilisers, produces the
greenhouse gas N2O that has an extremely high global warming potential. Thus
reduced inputs of nitrogen in agricultural production systems may contribute to
addressing the pressing issue of global warming (Kennedy et al. 2004; Venterea et al.
2005; IPCC 1996).
In summary, reduced nitrogen inputs, as well as increased yield may potentially be
achieved by the use of inoculants of A. brasilense Sp7 and B. subtilis DAR26659 in
ginger cultivation.
In addition to promoting plant under low levels of fertiliser
application, B. subtilis DAR26659 also antagonised the growth and caused lysis of
hyphae of Foz in vitro. Therefore further investigation of the biocontrol activity of this
bacterium may be of significant value, as currently no other means for control of soil
borne Foz in ginger production exits.
210
Conclusion
Outcomes of this study included increased awareness of local industries of:
•
the potential for the non-selective culture of microorganisms to result in the
growth of human pathogenic bacteria;
•
the requirement to use personal protective equipment in production and
application of such cultures whether they contain pathogenic or non-pathogenic
organisms;
•
consideration for the safe disposal of microbial brews to prevent leaching and
run off into waterways and groundwater and;
•
requirements for quality control testing to demonstrate low levels of indicator
pathogens in commercial inoculants and compost tea production systems.
Further measures that may reduce risks associated with the use of compost teas
include:
•
containment of bioaerosols during production;
•
preparation of teas without additives;
•
application of additives, such as kelp separately to the microbial culture and
without fermentation;
•
soil rather than spray application where practicable;
•
not applying to edible portions of plants and;
•
observation of withholding periods.
Further consideration should also be given to the potential for the non-selective culture
and application of microorganisms to increase and disperse populations of human
pathogenic organisms in the environment, which may constitute a public health risk. The
importation of unlegislated microbial products may pose a threat to national (and in
particular farm) biosecurity and this threat should be managed. Given that risks
associated with the production and application of large volumes of liquid that may
contain human pathogenic organisms were not ameliorated by the use of personal
protective equipment, the uncontrolled culture of microorganisms could not be
recommended and such preparations were precluded from further study under
University of the Sunshine Coast Occupational Health and Safety policies, that are in
accordance with the Queensland Workplace Health and Safety Act 1995.
211
The present study demonstrated that improved growth of ginger plants could be
achieved by using pure cultures of Class-1 bacteria in a manner associated with a low
level of risk. The importance of the inclusion of appropriate controls in greenhouse
trials, to identify effects of buffers or carrier material on plant growth was repeatedly
shown.
The extensively characterised type strain A. brasilense Sp7 improved the
growth of ginger tissue culture plants in greenhouse conditions, may enhance the
tolerance of tissue culture plants to hyper-osmotic stress and an improved growth
response may result from using alginate as a carrier material for the bacterium. The
strain B. subtilis DAR26659 induced striking improvements in the growth of
micropropagated ginger plants, which is the first report of growth promoting activity of
this strain. This is also the known first study to describe the use of bacterial inoculants
to promote the growth of micropropagated ginger plants.
In addition to promoting growth of ginger tissue culture plants, B. subtilis DAR26659
antagonised the growth and caused lysis of hyphae of F. oxysporum f. sp. zingiberi in in
vitro assays. While B. subtilis DAR26659 slightly reduced symptoms of Foz infection in
planta, inconsistent infection by the pathogen limited the evaluation of biocontrol activity
of this bacterium. Further research is required to:
•
Establish a robust bioassay to study the effects of bacterial inoculants on
infection of ginger by Foz;
•
Determine the consistency of growth promotion induced by B. subtilis
DAR26659, B. subtilis A13 and A. brasilense Sp7 in pasteurised soil and field
soils;
•
Assess the effect of alginate formulations on growth promoting and biocontrol
activity of these bacteria;
•
Investigate the activity of the bacteria under a range of fertiliser levels and;
•
Determine minimal application rates of bacteria required to induce a consistent
plant growth response.
A further understanding of the influence of these biotic, abiotic and environmental
factors on the activity of introduced bacteria may enable the implementation of
inoculants with improved reliability in field conditions. Research on the use of PGPB in
agriculture is considered to be in its infancy and the technology emergent (De Boer et
212
al. 1999; Fravel 2005). The further development of inoculants of plant growth promoting
bacteria may contribute to the development of sustainable practices in agriculture, that
enable effective plant health management with reduced inputs of inorganic fertilisers
and pesticides.
213
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