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COLONIZATION OF SORGHUM BICOLOR (L.) MOENCH BY THE N2-FIXING ENDOPHYTE GLUCONACETOBACTER DIAZOTROPHICUS

Vanessa Yoon

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Recommended Citation Yoon, Vanessa, "COLONIZATION OF SORGHUM BICOLOR (L.) MOENCH BY THE N2-FIXING ENDOPHYTE GLUCONACETOBACTER DIAZOTROPHICUS" (2011). Digitized Theses. 3309. https://ir.lib.uwo.ca/digitizedtheses/3309

This Thesis is brought to you for free and open access by the Digitized Special Collections at Scholarship@Western. It has been accepted for inclusion in Digitized Theses by an authorized administrator of Scholarship@Western. For more information, please contact [email protected]. COLONIZATION OF SORGHUM BICOLOR (L.) MOENCH BY THE N2-FIXING ENDOPHYTE GLUCONACETOBACTER DIAZOTROPHICUS

(Spine Title: Colonization of Sorghum by Gluconacetobacter diazotrophicus) (Thesis Format: Integrated Article)

: b y , „

Vanessa Yoon

Graduate Program in the Department of Biology

■ j .

A thesis submitted in partial fulfillment of the requirements for the degree of Master of Science

The School of Graduate and Postdoctoral Studies The University of Western Ontario London, Ontario, Canada

©Vanessa Yoon 2011 THE UNIVERSITY OF WESTERN ONTARIO School of Graduate and Postdoctoral Studies

CERTIFICATE OF EXAMINATION

Supervisor Examiners

Dr. Lining Tian Dr. Hugh Henry

Co-Supervisor Dr. Greg Thom

Dr. Sheila Macfie Dr. Brian McGarvey Supervisory Committee

Dr. Hugh Henry

Dr. Krzysztof Szczyglowski

The thesis by

Vanessa Yoon

entitled:

Colonization of Sorghum bicolor (L.) Moench by the N2-Fixing Endophyte Gluconacetobacler diazotrophicus

is accepted in partial fulfillment o f the requirements for the degree of Master of Science

D ate______Chair of the Thesis Examination Board

li ABSTRACT

Gluconacetobacter diazotrophicus is a N2-fixing endophyte to study biological nitrogen fixation in cereal crops. The objectives of this study were to establish colonization of sorghum with G. diazotrophicus and to evaluate the effects o f the environment and sucrose on colonization. Seven

sorghum cultivars inoculated with the bacterium were grown under greenhouse and field

conditions. In the sweet and grain cultivars, G. diazotrophicus was either detected in the roots,

stems and/or leaves. Differences in colonization were observed among the different plant

cultivars and plant growth conditions. G. diazotrophicus was detected in a higher proportion of

sweet sorghum plants than grain sorghum plants. Bacterial presence was detected in a higher proportion o f field-grown plants than greenhouse-grown plants. Strong positive correlations

between sucrose concentration and the proportion o f plants showing bacterial presence and

bacterial population numbers were observed, indicating that colonization is dependent on the

sucrose concentration of the plant. i,

Keywords: colonization, endophyte, Gluconacetobacter diazotrophicus, N2 fixation, sorghum ACKNOWLEDGEMENTS

I would like to express my sincere gratitude towards my supervisor, Dr. Lining Tian, and co­

supervisor, Dr. Sheila Macfie, for their guidance and support during the entire, process o f this research. Constantly providing, an intellectually stimulating and challenging environment, I am thankful for their patience and understanding with the progress o f my studies. Dr. Lining Tian’s

and Dr. Sheila Macfie’s ideas and constructive comments have contributed substantially to the

development of this thesis and are much appreciated.

I am grateful towards the members o f my advisory committee, Dr. Hugh Henry and Dr.

Krzysztof Szczyglowski, for their helpful suggestions and remarks that allowed further progress

of my work. I would like to thank Dr. Brian McGarvey and Bob Poes for their technical

assistance and input. Thanks to Dr. Zhongmin Dong and Dr. Kevin Vessey for providing the

bacterial strains used in this project. Additionally, I would like to thank my labmates, Cindy

Wang, Nikita Eskin and Jessie Wong, for their insights, discussions and technical assistance.

Special thanks to Agriculture Environmental Renewal Canada Inc. and NSERC for their support

and funding of this research.

Lastly, and most importantly, I would like to acknowledge my family; my parents, Seong Hee

Yoon and Eun Sook Yoon, and my brother, Michael Yoon. Without my family, I would not be

where I am today. I am forever thankful for their love, patience and support through thick and

thin and will always be indebted to them.

IV TABLE OF CONTENTS

Certificate of Examination......

Abstract and Keywords...... ^...... hi

Acknowledgements...... ;...... ;...:...... ;...;.:...... :...:...... ;...... ^

Table of Contents,...... ;...... v

List of Tables...... ;...... ix

List of Figures....;.;...... :...... x

List of Abbreviations...... xii

Chapter 1. Introduction...... 1

1.1 Nitrogen in agriculture......

1.2 Costs of nitrogen additions„...... ;....;...... ;.;...... :...... ;...... ;...... 4

1:3 Biological nitrogen fixation & agriculture,...... ;;...... ;.;.;.....;...... ;...... 6

1.4 Rationale, hypothesis & objectives...... 12

References...... ;...... ;...... ;...... 14

Chapter 2. Gluconacetobacter diazotrophicus: A N 2-fixing bacterial species...,;...... ;....;.;...21

2.1 Introduction....;,...... :...... :...... :;;.....;.:..21

2.2 Materials & Methods ...... ;...... :...... 22

2.2:1 G. diazotrophicus strains...... ::...... :...... 22

2.2.2 Cultivation of G. diazotrophicus......

2.2.3 Detection of G. diazotrophicus by polymerase chain reaction,...... 25

2.2.4 Evaluation of nitrogenase activity of G.

2.2.5 Statistical analyses,...... ;.....;...... ;...... ;...... :...29

2.3 Results ......

v 2.3.1 Bacterial growth...... 32

2.3.2 Detection of G'. diazotrophicus...... 32

2.3.3 Nitrogenase activity of G. diazotrophicus ...... 36

2.4 Discussion...... ;.:...... 36

2.4.1 Laboratory cultures o f G. diazotrophicu s^ %,m..,mmm^; mm.....,..^:mi,-...... 36

2.4.2 Activity of nitrogenase in G.diazotrophicus, ...... 38

2.5 Conclusion...... 39

References...... 41

Chapter 3. Colonization o f sorghum by Gluconacetobacter diazotrophicus: A greenhouse and, field study...... 44

3.1 Introduction...... 44

3.2 Materials & Methods...... 48

3.2.1 Sorghum cultivars...... 48

3.2.2 Bacterial strains & inoculum...... I...... „48

3.2.3 Germination of sorghum...... ;.50

\ 3.2.4 Inoculation o f sorghum...... „50

3.2.5 Sample preparation & extraction for polymerase chain reaction (PCR)...... „51

3.2.6 PCR detection of G. diazotrophicus from plant tissues...... „56

3.2.7 Re-isolation & enumeration of G diazotrophicus population from plant tissues...... 53

3.2.8 Visual confirmation of colonization by.G. diazotrophicus...... „53

3.2.9 Statistical analyses...... 54

3.3 Results...... „55

3.3.1 Germination o f sorghum1..:„„:.;„.i.;;.„„.„c„:..‘...... 55

3.3.2 Detection of G. diazotrophicus from greenhouse sorghum tissues,...... 55

vi c

3.3.3 Detection of G. diazotrophicus from field sorghum tissues...... 60

3.3.4 Effect o f growth environment on G. diazotrophicusdetection...... 63

3.3.5 Population o f G. diazotrophicus in greenhouse plant tissues...... 67

3.3.6 Population o f G. diazotrophicus in field plant tissues...... 67

3.3.7 Effect o f growth environment on G diaztrophicus population,...... 71

3.3.8 Visualization of G. diazotrophicus colonization...... 76

3.4 Discussion...... „76

3.4.1 Colonization o f sorghum by G. diazotrophicus...... „76

3.4.2 Influence of plant cultivar on colonization o f sorghum by G. diazotrophicus,...... 81

3.4.3 Environmental influence on colonization o f sorghum by G. diazotrophicus ...... 8 6

3.5 Conclusion...... 90

References...... 93

Chapter 4. Endophytic association between Gluconacetobacter diazotrophicus and sorghum:

Colonization factors and inoculation effects...... 100

4.1 Introduction...... 100

4.2 Materials & Methods,...... 103

4.2.1 Sugar analysis in sorghum...... 103

4.2.2 Agronomic measurements o f sorghum in the field...... 104

4.2.3 Statistical analyses...... 105

4.3 Results...... 106

4.3.1 Sugar concentration in sorghum...... 106

4.3.2 Relationship between sorghum sugar concentration and colonization...... 106

4.3.3 Effect o f G. diazotrophicus on plant growth...... 110

4.4 Discussion...... 110

vii 4.4.1 Influence of sugar on colonization of sorghum by G. diazotrophicus.... •••■■•••••••••a 110

116

4.5 Conclusion 119

References.. taaaaaaaaaaaaa 121

Chapter 5. Conclusion...... 126

5.1 Colonization o f sorghum by Gluconacetobacter diazotrophicus-. Summary...... 126

5.2 Future directions...... 129

References...... 133

Appendix...... ;...... 135

Curriculum Vitae...... 143 LIST OF TABLES

Table 1-1 Nitrogen removal rates in grain crops...... 3

Table 2-1 G. diazotrophicus discovered from different crops and sources...... 23

Table 2-2 Differentiating characteristics for G. dUàotropM cusj.;.jJ.^m„^ ^ J^ i...... 24

Table 2-3 Species-specific PCR primers for identification/detection o f G. diaztrophicus ...... 26

Table 2-4 Nitrogenase activity of G. diazotrophicus strains PAL5 and MAd3A measured by the acetylene reduction assay...... 1...... 37

Table 3-1 Sorghum cultivars from Agriculture Environmental Renewal Canada Inc....:...... 49

Table 3-2 Growing mix components for Sun Gro Sunshine Mix 2 ...... 52

Table 3-3 Germination success of seven sorghum cultivars for greenhouse trial...... 56

Table 3-4 Germination success of seven sorghum cultivars for field trial,...... 57

Table 3-5 Proportion o f greenhouse plants showing presence of G. diazotrophicus...... 58

Table 3-6 Proportion o f field plants showing presence of G. diazotrophicus...... ,62

Table 3-7 Most probable number o f G. diazotrophicus population in greenhouse plant tissues ..6 8

Table 3-8 Most probable number of G. diazotrophicus population in field plant tissues...... 72

IX LIST OF FIGURES

Figure 1-1 Schematic structure of the nitrogenase enzyme complex;...... :...... ;...... ;;....;....;....8

Figure 2-1 Ethylene standard curve...... 30

Figure 2-2 Bovine serum albumin standard curve for bacterial protein estimation...„.„„>„...... :...31

Figure 2-3 G. diazotrophicus morphology on LGIP plates supplemented with (NIL^SC^...... 33

Figure 2-4 Nested PCR amplification for detection of G. diazotrophicus strains...... 34

Figure 2-5 Sensitivity o f detection for nested PGR amplification of PAL5 and MAd3A...... 35

Figure 3-1 Effect of tissue type on the proportion o f greenhouse plants showing presence o f G. diazotrophicus,...... 59

Figure 3-2 Effect o f cultivar type on the proportion o f greenhouse plants showing presence o f G.

Figure 3-3 Effect of tissue type on the proportion of field plants showing presence of G. rfiazo/rqp/j/cMi..;...... ;..;...... ;...... ;...... ;...... ;...... :...... 64

Figure 3-4 Effect of cultivar type on the proportion of field plants showing presence of G. diazotrophicus ...... ;...... :...... :...... :..65

\ Figure 3-5 Effect of growth environment on the proportion of plants showing presence of G. diazotrophicus ...... :...... ;...... :;;..66

Figure 3-6 Effect of tissue type on G. diazotrophicus population numbers in greenhouse plants. 69

Figure 3-7 Effect of cultivar type on G. diazotrophicus population numbers in greenhouse plants..

...... 70

Figure 3-8 Effect of tissue type on G. diazotrophicus population numbers in field plants...... 73

Figure 3-9 Effect of cultivar type on G. diazotrophicus population numbers in field plants...... 74

Figure 3-10 Effect o f growth environment on G. diazotrophicus population numbers. 75

x Figure 3-11 Colonization of N111 roots by gusA-marked G. diazotrophicus strain

UAP5541/pRGS561...... 77 ) <■ Figure 3-12 Colonization of 275 roots by gusA-marked G. diazotrophicus strain

UAP5541/pRGS561...... 78

Figure 3-16 Effect o f growth environment on G. diazotrophicus population in leaf tissues...... 8 8

Figure 3-17 Colonization of N111 roots by gusA-marked G. diazotrophicus strain

UAP5541/pRGS561...... 90

Figure 4-1 Sucrose concentration in sorghum cultivars...... 107

Figure 4-2 Sucrose concentration in sorghum tissues...... 108

Figure 4-3 Relationship between sucrose and bacterial presence and bacterial population numbers in greenhouse sorghum plants...... 109

Figure 4-4 Relationship between sucrose and bacterial presence and bacterial population numbers in field sorghum plants...... 111

Figure 4-5 Height and dry weight of cultivar N 111 two months post-inoculation with G. diazotrophicus ...... 112

\ Figure 4-6 Height and dry weight of cultivar 275 two months post-inoculation with G. diazotrophicus ...... 113

xi LIST OF ABBREVIATIONS

A AFC Agriculture and Agri-Food Canada

ADP Adenosine diphosphate

AM Arbuscular mycorrhiza

ATP Adenosine triphosphate

BLAST Basic local alignment search tool

BNF Biological nitrogen fixation

bp Basepair(s)

BSA Bovine serum albumin

CFU Colony forming unit(s)

cm Centimeter(s)

DNA Deoxyribonucleic acid

dNTPs Deoxyribonucleotide triphosphates

EG Endoglucanase

EPMG Endopolymethylpolygacturonase

EtBr Ethidium bromide

EXG Endoxyloglucanase

g Gram(s)

GC Gas chromatograph

GC-FID Gas chromatograph-flame ionization detector

GUS ^-glucuronidase

h Ilour(s)

ha Hectare

Xll IAA Indole-3-acetic acid kg kilogram m Meter(s)

MD Mean difference mg Milligram(s) min Minute(s) mL Milliliter(s) mm Millimeter(s) mM Millimolar(s)

MOX 2% me thoxy amine »HCl in pyridine

MPN Most probable number

MSTFA A-methyl-A'-(trimethylsilyl) trifluoroacetamide nm Nanometer(s)

No. Number

Nod Nodulation

PCR Polymerase chain reaction

PGPB ' Plant growth-promoting bacteria

PGPR Plant growth-promoting rhizobacteria ppm Parts per million

PVPP Polyvinyl pyrrolidone rDNA) Ribosomal deoxyribonucleic acid rpm Revolutions per minute s Second(s)

SE Standard error

X lll Mg Microgram(s)

|iL Microliter(s)

Mm Micrometer(s)

|iM Micromolar(s)

UV Ultraviolet ,

UWO University o f Western Ontario

V ; . Voit : v:v Volume: volume

XIV 1

CHAPTER 1. Introduction

1.1 Nitrogen in agriculture

Nitrogen, a primary constituent o f nucleotides and proteins, is essential for life and central to living systems; however, biologically available nitrogen is typically in short supply thus limiting plant growth and primary production (Smil, 2002). The vast majority o f nitrogen in the biosphere exists as atmospheric N2, which is bound by an exceptionally strong triple bond that can only be broken by a few processes (Galloway et al., 2004). Two natural processes that can provide the energy required to break the triple bond are lightning and N2-fixing microbes with specialized enzymes that convert N2 to NH3 (Galloway et al., 2004). However, more N2 is now fixed through industrial processes during the production of nitrogen fertilizer via the Haber-

Bosch process, in which natural gas is subjected to steam reforming to produce hydrogen that reacts with N2 under high temperature and pressure to form NH3 (Smil, 2002). The discovery and commercialization o f this process has fulfilled a substantial fraction of the world’s agricultural needs (Robertson and Vitousek, 2009).

The addition o f nitrogen is vital in agricultural systems to sustain and increase crop yields

(Robertson and Vitousek, 2009). Intensive nitrogen management permits more food to be grown on a given area of land increasing its human carrying capacity and relieving the pressure for new land clearing (Christianson and Vlek, 1991; Cassman et al., 1995; Zhu and Chen, 2002). To maximize growth and crop yields arid fully understand nitrogen in agriculture, grasping the fundamental need to match nitrogen supply to crop nitrogen demands is required. Nitrogen as much as any other nutrient that is removed during harvest must be replaced to sustain productivity in a cropping system (Robertson and Vitousek, 2009). The difference with nitrogen from other plant nutrients, including phosphorus, potassium, calcium, and to a lesser extent 2 magnesium and boron, is the minimal pool of organic and mineral nitrogen in most soils (Jarvis et al., 1996; Murphy et al., 2000). Removing nitrogen from the field through harvest means that less nitrogen enters the soil as plant residue, resulting in less nitrogen being recycled over time for future plant use (Vitousek et al., 1997). Nitrogen depletion can happen quickly under intensive cropping. In major grain cropping systems, maintaining crop nitrogen removal rates of

100-260 kg nitrogen year' 1 (Table 1-1) poses a significant challenge in restoring the levels of soil nitrogen reserves (Robertson and Vitousek, 2009). Overall, nitrogen is used inefficiently in annual crop systems. Cereal crops, including Triticum (L.) spp. (wheat), maize and B rassica napus (L.) (canola), need 35-175 kg nitrogen ha' 1 depending on the soil type (Huffman et al.,

2008). However, typically 50% or less of fertilizer nitrogen actually makes it into the crop

(Cassman et al., 1993; Raun et al., 2002; Bundy and Andraski, 2005). For replacing lost nitrogen and managing intensive cropping systems, most grain producers choose synthetic fertilizers as a source of nitrogen (Smil, 2002; Heffer, 2009). Synthetic nitrogen fertilizers are accessible and transportable options of nitrogen input, so much so that the usage has increased from approximately 4 million tonnes nitrogen year' 1 in the 1960s (Food and Agriculture Organization of the United Nations, 2009a) to roughly 105 million tonnes nitrogen year' 1 in 2009 (Food and

Agriculture Organization o f the United Nations, 2009b). The upsurge in fertilizer application has led to the consensus that intensive agriculture has the capability to meet the food needs o f 8 -1 0 billion people by increased production, while substantially decreasing the proportion o f the population who go hungry - a feat that should not be slighted (Waggoner, 1995; Tillman et al.,

2002). Table 1-1. Nitrogen removal rates in grain crops (adapted from Robertson and Vitousek, 2009)______Crop ‘ Yield (tonne ha'1) Grain Nitrogen Removed (kg ha'1)

Z ea mays L. (maize) 1 0 .0 . 2 6 0 "

Triticum aestivum L. (wheat) 5.4 ; ; : 1 08

Oryza saliva L. (rice) 7.9 ; 142 , .

Sorghum bicolor (L.) Moench 9.0 270 (sorghum)

Glycine max (L.) Merr. 2 .8 176 (soybean) 4

1.2 Costs of nitrogen additions

The addition o f fertilizer nitrogen to agricultural systems and the benefits that follow

appear to be clear-cut; however, with those advantagés come certain costs. Nitrogen added to

agricultural fields often does not reach its ultimate target and is frequently out of balance, either

negatively or positively, with the amount removed in harvested products (Vitousek et al., 2009).

While intensively cultivated maize-based systems in Kenya only receive 11 kg nitrogen ha'1.via

fertilizer and manure, as much as 59 kg nitrogen ha' 1 is removed in grain and secondary harvests

resulting in a net imbalance of -48 kg nitrogen ha' 1 (Bunemann et al., 2004; Sanchez et al.,

2007). In contrast, annual fertilizer and manure additions o f 649 kg nitrogen ha' 1 and removal of

only 361 kg nitrogen in yields cause a net imbalance of 288 kg nitrogen ha' 1 in the wheat/maize

double-cropping systems in Northern China (Ju et al., 2009). It is often difficult to synchronize

the supply o f nitrogen to biological demands and thus the balance is tipped in either direction.

Many agricultural systems in developed or developing regions typically have positive nitrogen

balances, in which nitrogen supply is often in excess o f the amount taken up by crops (Vitousek

et al., 2009). : V;", =::. L: ' \ In conjunction with agricultural harvesting, major pathways o f nitrogen loss from

cropped ecosystems include surface and groundwater leaching, denitrification to N2,

volatilization o f NH3, as well as N2O, NO and NO2 fluxes to the atmosphere (Robertson and jVitousek, 2009). Since nitrogen is highly mobile in the biosphere, losses from agricultural

systems can alter downwind and downstream ecosystems substantially (Robertson and Vitousek,

2009). Based on Rabalais et al.’s (2002) review, a result o f runoff from the agricultural hubs of

the United States, which drain via the Mississippi River, is the massive hypoxic zone in the Gulf

o f Mexico, one of the most prominent environmental costs of adding excessive nitrogen to a 5

system. This “dead zone”, so named for the low dissolved oxygen levels, resulted from the

interaction between fresh-water inputs to the Gulf and the high supply o f nutrients in that fresh­ water. Hypoxia is the result o f algal growth, due to high nutrient levels, that sink and die deep in the water column where the dissolved oxygen is consumed during their subsequent

decomposition by bacteria. Eutrophication of the Gulf waters caused by nitrogen, originating

from agricultural systems in the Mississippi basin, has increased the size of the hypoxic zone to just over 20,000 km2 (Mclsaac et al., 2002).

, Excess soluble nitrogen, lost from agricultural systems, can be transformed into organic

or volatile forms that can leach into groundwater, thereby influencing human health (Abrahams,

2002; Powlson et al., 2008). Groundwater supplies in agricultural regions around the world tend

to have elevated NOa'-nitrogen contamination usually associated with nitrogen fertilizer use

(Zhang et al., 1996; Agrawal et al;, 1999; Townsend et al., 2003). NO3' contamination of

groundwater is a critical problem due to its persistence once contamination occurs. Potential

health effects of high NO3' levels in the drinking water include reproductive problems,

methemoglobinemia and cancer (Fan & Steinberg, 1996; Ward, 2009). It is well noted that long-

\ term consumption of water with NO3' concentrations of 6.3 ppm has been associated with a

higher risk of Non-Hodgkins Lymphoma (Townsend et al., 2003).

Rivers and oceans represent an important sink o f excess nitrogen; however, nitrogen can

also be lost to the atmosphere as gases including the environmentally benign gas N2, the

radiatively active gas N2O, the air polluting gases NO and NO2, and NH3, which transfers large

amounts o f nitrogen to downwind ecosystems (Robertson and Vitousek, 2009). In the

troposphere, N2O is not reactive, but it is able to absorb outgoing radiation from the Earth thus

allowing it to act as a powerful greenhouse gas that is 300 times more effective than CO2 6

(Robertson and Vitousek, 2009). Referring to global budgets, 5-6 million tonnes of N2 0 -nitrogen comes from an anthropogenic source, with around 80% of this source being associàted with agriculture (Galloway et al., 2004).

NO is the primary oxide that oxidizes in the atmosphere to form NO2 and other compounds, collectively known as NOx (Robertson and Vitousek, 2009). As of the mid-1990s, about 80% o f terrestrial NOx emissions were anthropogenic, with 25% coming from agricultural processes such as burning, land clearing and fluxes from agricultural soils (Galloway et al.,

2004). The elevation of NOx results in the oxidation of atmospheric hydrocarbons and CO leading to the production of O3 (Sillman et al., 1990; Sillman and West, 2009). At high concentrations, particularly in high-NOx regions, tropospheric O3 acts as a greenhouse gas and oxidant that can be harmful to human health and plant growth (Powlson et al., 2008). Although primary sources of NOx stem from urban and industrial systems, certain regions experience high

NOx and O3 concentrations as a consequence o f agriculturally based biomass burning (Thompson et al., 2001). NOx, whether emitted from energy or agricultural systems, eventually gets deposited on downwind ecosystems either as gases, such as NOx and HNO3 vapour, or as dissolved forms, such as HNO3 and oxidized organic nitrogen (Holland et al., 2005). As such, appropriate management of nitrogen in agricultural systems is critical. However, it is difficult to overcome the challenge of enhancing agricultural productivity,'while reducing the transfer of nitrogen to non-target ecosystems. ; 1 1.3 Biological nitrogen fixation & plant agriculture

■ Biological nitrogen fixation (BNF), which involves the conversion of atmospheric N2 into

NH3 by N2-fixing bacteria, plays a significant role in the environment and the agricultural world

(Dixon and Kahn, 2004). As an important part o f the nitrogen cycle, N2 fixation is one form of 7 nitrogen input that can help replenish the overall nitrogen content of the biosphere and compensate for the losses incurred from denitrification (Dixon and Kahn, 2004). During BNF,

N2 is reduced to NH3 through multiple electron transfers, which shortly undergoes protonation to

NH4+ based on the pH o f the environment (Kim and Rees, 1992). NH4+ is then used for the subsequent synthesis of biomolecules (Kneip et al., 2007). The reduction of N2 to NH4+ is an

ATP-dependent, energy consuming reaction that is catalyzed in all N2-fixing organisms via the nitrogenase enzyme complex (Kneip et al., 2007). Nitrogenase is a multifaceted metalloenzyme comprised of two main functional subunits: dinitrogenase reductase (azoferredoxin) and dinitrogenase (molybdoferredoxin) (Hageman and Burris, 1978). The structural components of these subunits are the N if (N2 fixation) proteins, NifH (homodimeric azoferredoxin), also known as the Fe protein, and NifD/K (heterotetrameric molybdoferredoxin), also known as the Mo-Fe protein (Hageman and Burris, 1978) (Figure 1-1). The smaller dimeric component functions as an ATP-dependent electron donor to the larger heterotetrameric component, which contains the enzyme catalytic site (Dixon and Kahn, 2004). Three types o f nitrogenases are known based on the composition o f their metal centres: molybdenum and iron (Mo-Fe), vanadium and iron (V- \ Fe) or iron and iron (Fe-Fe) (Bishop and Premakumar, 1992). Among all N2-fixing bacteria, the most common nitrogenase enzyme complex is the Mo-Fe type (Eady, 1996). However, the depletion o f Mo can induce the synthesis of the two alternative nitrogenases, containing the V-Fe or Fe-Fe cofactors, in several organisms e.g. Azotobacter vinelandii and Rhodobacter capsulatus

(Eady, 1996). The surface-exposed 4Fe-S cluster that bridges the two subunits of the dimer protects both the protein components of the nitrogenase enzyme (Einsle et al., 2002; Seefeldt et al., 2004). The Mo-Fe protein contains two types o f metal centres: the P cluster (8Fe-7S) and the

FeMo cofactor (MoFevS^homocitrate), which acts as the substrate reduction site (Einsle et al., Pyruvate + CoA 2 ADP + 2 P| Figure 1-1. Schematic structure of the nitrogenase enzyme complex (adapted from Kneip et al., 2007) Nitrogenase is comprised o f two main functional subunits: 1) dinitrogenase reductase composed ■ . 9

2002; Seefeldf et al., 2004). The general stoichiometry of N2 reduction under optimal conditions is shown in Equation 1-1.

Equation 1-1. Général reaction of molecular N2 fixation N2 + 8 e- + 8 H+ + 16 MgATP 2 NH3 + H2 + 16 M gAD P+16 Pr

N2 fixation is a widespread function o f prokaryotic cells that is established among various groups o f bacteria as well as some archaea (Murray and Zinder, 1985; Leigh, 2000). Within the eubacteria, N2-fixing members have been described among the proteobacteria, cyanobacteria, actinobacteria, spirochaetes, clostridiales, purple-sulfur bacteria and green-sulfur bacteria (Kneip et al., 2007). Among the numerous bacterial species that have the ability to fix N2, only a certain number of N2-fixing bacteria are known to interact with eukaryotes (Kneip et al., 2007). Inputs of fixed nitrogen into an ecosystem come from two basic BNF systems: 1) symbiotic N2 fixation in legumes, which contribute 12-25 million tonnes of fixed nitrogen annually; and 2) endophytic/associative N2 fixation in non-legumes, which contribute annual inputs of 5.5 million tonnes fixed nitrogen (Herridge et al., 2008). The mutualistic symbioses between the proteobacteria o f the order Rhizobiales, known as rhizobia, with plants of the order F a b a les, which comprises the legumes, are the most extensively studied interactions between bacteria and plants (Kneip et al., 2007).

The rhizobia-legume symbiosis is characterized by typical root-nodule ; structures of the plant host, which house the bacteria in a microaerobic environment providing the anoxic conditions for N2 fixation (Pamiske, 2000). In return for the fixed nitrogen, the bacteria are supplied with energy-rich carbon compounds within the nodulated plant roots (Kneip et al.,

2007). The highly regulated process o f nodule formation involves rhizospheric rhizobia entering the plant root epidermis and inducing nodule formation by reprogramming the root cortical cells

(Kneip et al., 2007). Flavonoids secreted by the plant host leads to the subsequent induction of ' ' ■' • 10 bacterial nodulation (nod) genes (Goethals et al., 1992; Bladergroen and Spaink, 1998). The nodulation (Nod-) factors, encoded within the n od genes, play a role in the nodulation process by initiating changes in the. host, including root hair deformation, membrane depolarization, intracellular calcium oscillations, and the initiation of cell division in the root cortex that leads to establishing the meristem and nodule primordium (Gage, 2004). Early in the symbiotic interaction, rhizobia grow and divide inside a tubule called an infection thread (Gage, 2004).

Formation o f the infection thread occurs when the bacteria become trapped between the cell walls o f the sharp bend or curl of the deformed root hairs (Callaham and Torrey, 1981).

Invagination of the plant cell wall in the curl leads to the growth of an infection thread down the inside o f the root hair toward the root interior (Robledo et al., 2008). Inside the infection thread, rhizobia grow and divide filling up the tubule (VandenBosch et al., 1989; van Workum et al.,

1998). The infection thread undergoes multiple branching as it grows through the root and enters the nodule, thereby increasing the sites of entry for the bacteria (Gage, 2004). Once inside the nodule, the bacteria reside within parenchyma cells, where they are localized in membrane- bound vesicles, synthesizing proteins for N2 fixation and proteins to maintain the symbiotic relationship (Gage, 2004).

Although the rhizobia-legume symbiosis is a major source of fixed N2 in some cropping systems, progress in stimulating root nodule formation in non-legumes has been slow. As a

Jresult, studies on endophytic N2-fixing bacteria have expanded thanks to their ability to colonize non-legumes without any specific structures. Endophytes are microorganisms that reside within plants without causing apparent harm to the host (Stone et al., 2000). N2-fixing endophytes have been consistently found in the root, stem, leaf and tuber tissues of various agricultural, horticultural, and forest species (Sturz et al., 2000). Endophytic colonization by N2-fixing 11 bacteria can support the growth and development of their plant hosts through BNF and the supply o f growth-promoting substances (Bhattacharjee et al., 2008). A well-documented example o f endophytic N2 fixation in a graminaceous plant is the occurrence of BNF in

Saccharum L. spp. (sugarcane). In 1988, a new species of theAcetobacter genus, first called

Acetobacter diazotrophicus Gillis et al. 1989 and later renamed as Gluconacetobacter diazotrophicus (Gillis et al. 1989) Yamada et al. 1997 was discovered in the interior of the sugarcane plant (Cavalcante and Dobereiner, 1988, Gillis et al., 1989, Yamada et al., 1997). G. diazotrophicus is considered to be an endophyte, as it lives inside the plant tissue either latently or actively colonizing local or systemic tissues (Reis et al., 2007). Plant-growth improvement due to colonization by the N2-fixing endophyte has also been observed in several instances.

Sevilla et al. (2001) demonstrated that, under conditions o f limited fixed nitrogen, micro- propagated sugarcane plantlets inoculated with wild-type G. diazotrophicus strain PAL5 showed significant increases in dry matter and nitrogen accumulation compared to those inoculated with the N if mutant MAd3A, which does not fix nitrogen. l

The endophyticassociation betweenG. diazotrophicus and sugarcane, as well as several other sugar-rich plants such as Pennisetum purpureum Schumach. (Cameroon grass), Ananas com osus (L.) Merr. (pineapple) and Ipomoea batatas (L.) Lam. (sweet potato), suggests a preference for a high sugar environment by the bacterium (Saravanan et al., 2008). In sugarcane, sucrose accounts for 50% of the total dry matter of the stalk (Glasziou and Gayler, 1972).

Modem sugarcane cultivars are multispecies hybrids, primarily o f Saccharum officinarum L.,

Saccharum spontaneum L. and Saccharum robustum E.W.Brandes & Jeswiet ex Grassl (Zhu et al., 1997). S. officinarum stores up to 21% sucrose in the juice o f the stem, while S', spontaneum stores less than 6 % and S. robustum stores less than 10% (Zhu et al., 1997). Reminiscent of 12 sugarcane, Sorghum bicolor {L.) Moench (sorghum) is a sugar-rich cereal crop that resembles sugarcane in its storage o f sugars within the stalks. With each developmental stage, sucrose accumulation in sorghum appears to increase after the termination o f intemodal elongation, which is quite similar to the pattern of sucrose accumulation in sugarcane (Hoffmann-Thoma et al., 1996). Particularly the sweet cultivars of sorghum can accumulate large amounts of sugar in their stem parenchyma (Hoffmann-Thoma et al., 1996). Total sugar contents in the stalks and grains of various sorghum cultivars have been reported to range from 1-40% on a dry weight basis (Subramanian et al., 1987). As such, sucrose contents are comparable between sugarcane and sorghum.

1.4 Rationale, hypothesis & objectives

. The major inputs o f nitrogen into terrestrial ecosystems are through the biological and industrial fixation of atmospheric N2 to NH3 (McNeill and Unkovich, 2007). Improving the efficiency o f fertilizer nitrogen use in world agriculture is vital to the long-term sustainability of the planet (Vitousek et al., 1997). Given that there is often low efficiency of fertilizer nitrogen use, with gaseous losses contributing to global warming and leaching and erosion losses to the degradation o f watercourses and watersheds, appropriate nitrogen management seems to be a desirable goal (Vitousek et al., 1997; McNeill and Unkovich, 2007). Equally as important would be the more effective exploitation and utilization of biologically fixed nitrogen in agricultural jsystems (Herridge et al., 2008). At the very least, complimenting fertilizer nitrogen use with

BNF may ease the long-term pressure for expanded production and in some cases particular systems within the global food production framework could become more reliant on BNF, rather than chemical fertilizers, for nitrogen inputs (Herridge et al., 2008). I examined the colonization o f sweet and grain cultivars o f sorghum by G. diazotrophicus and several, factors that influence colonization. It is hypothesized that plant cultivar type and plant growth conditions will affect colonization o f sorghum by G. diazotrophicus. From past literature research, G. diazotrophicus has been isolated from sugarcane and is primarily associated with sugar-rich plants (Cavalcante and Dobereiner, 1988; Saravanan et al., 2008).

Additionally, colonization studies ; involving other endophytes generally introduce the microorganisms to the plant hosts via the roots (Hurek et al., 1994; Bellone et al., 1997;

Reinhold-Hurek and Hurek, 1998; Bressan and Borges, 2004). As such, the major predictions for this study are: 1) sweet sorghum cultivars will show higher colonization than grain sorghum cultivars; and 2 ) colonization will be the highest in plant root tissues than stem and leaf tissues.

The main objectives to the proposed research are: 1) to assess the potential for colonization of different sorghum cultivars with G. diazotrophicus', and 2) to determine the effects of sucrose and the environment on colonization. References/"-.-'

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CHAPTER 2.Gluconacetobacter diazotrophicus: A N2-fixing bacterial species

2.1 Introduction

Gluconacetobacter diazotrophicus (Gillis et al. 1989) Yamada et al. 1997 is a

microaerobic, N2-fixing bacterium that was initially isolated from SaccharumL. spp. (sugarcane)

grown in the State of Alagoas, Brazil (Cavalcante and Dôbereiner, 1988). The key strain

responsible for N2 fixation in sugarcane was originally -named Saccharobacter nitrocaptans

(Cavalcante and Dôbereiner, 1988). However, intensive taxonomic analysis of this bacterium

placed it under a distinct rRNA branch o f Acetobacter, resulting in the novel species Acetobacter

diazotrophicus Gillis et al. 1989 (Gillis et al., 1989). Nearly a decade after, the species was

transferred to the genus Gluconacetobacter based on the analysis o f 16S rDNA sequences and

the production of a predominant type of ubiquinone (Yamada et al. 1997).

Cells o f G. diazotrophicus are straight rods about 0.7-0.9 ± 2 pm in length with rounded

ends that are motile by peritrichous flagella (Gillis et al., 1989). The bacterium belongs to the

phylum Proteobacteria in section a-Proteobacteria, order Rhodospirillales and family

Acetobacteraceae (Kersters et al., 2006). Encompassed within the same family are three N2-

fixing genera including six other species: Acetobacter nitrogenifigens Dutta and Gachhui 2006,

Gluconacetobacter kombuchae Dutta and Gachhui, 2007, Gluconacetobacter johannae Fuentes-

Ramirez et al., 2001, Gluconacetobacter azotocaptans Fuentes-Ramirez et al., 2001,

Swaminathania salitolerans Loganathan and Nair, 2004 and Acetobacter peroxydans Visser’t

Hooft 1925 (Fuentes-Ramirez et al., 2001; Loganathan and Nair, 2004; Muthukumarasamy et al.,

2005; Dutta and Gachhui, 2006; Dutta and Gachhui, 2007).

To evaluate the interactions between plants and bacteria, G. diazotrophicus is typically

the model organism to study its association with non-legumes. Initially, G. diazotrophicus was considered, to b e a plant-dwelling species that did not thrive well in soil (Kirchof et al., 1998).

However, the rhizospheric occurrence and survival o f the bacterium is apparent in a variety of agricultural crops (Table 2-1). The N2-fixing bacterium is primarily associated with sugar- and starch-rich plants suchasPennisetum purpureum Schumach. (Cameroon grass), Ananas comosus

(L.) Merr. (pineapple) and Ipomoea batatas (L.) Lam. (sweet potato) (Saravanan et al., 2008).

Unlike other N2-fixing bacteria, G. diazotrophicus possesses several unique traits that make it ideal for its consistent use in studying biological nitrogen fixation (BNF). Physiological characteristics of G. diazotrophicus include: growth and nitrogenase activity in a medium of 10-

30% sucrose; high acid tolerance; absence of nitrate reductase; and tolerance of high NO3' concentrations (Stephan et al., 1991). When grown in mixed cultures with an amylolytic yeast,

G. diazotrophicus can excrete about 40% o f its fixed nitrogen into the medium (Cojho et al.,

1993). Differentiating characteristics of G. diazotrophicus are listed in Table 2-2.

Since G. diazotrophicus appears to be an ideal candidate for BNF, phenotypic, molecular and physiological characteristics of two G. diazotrophicus strains will be examined and evaluated. Culture, polymerase chain reaction (PCR), and the acetylene reduction assay were used to ensure the validity of the bacterium for further application in field and greenhouse inoculation trials, which will be addressed in Chapter 3.

2.2 Materials & Methods

2.2.1 G. diazotrophicus strains

G. diazotrophicus strains PAL5 (ATCC 49037) and MAd3A, kindly provided by Dr.

Zhongmin Dong (Saint Mary’s University, Halifax, Nova Scotia, Canada) were used in this study. PAL5, originally isolated from the roots o f sugarcane in Alagoas, Brazil, is a wild-type strain capable of N2 fixation (Cavalcante and Dobereiner, 1988). Due to an insertional Table 2-1. G. diazotrophicus discovered from different crops and sources (adapted from Saravanan et al., 2008) Country of Crop Isolation sources Reference discovery Saccharum spp. Roots, stems, leaves, Cavalcante and Brazil (sugarcane) sugarcane juice Dobereiner, 1988

Cameroon grass, Brazil Roots, stems, tubers Dobereiner et al., 1993 sweet potato

Ashbolt and Inkerman, Australia Sugarcane Mealybugs, rhizosphere 1990; Gillis et al., 1989; Li and Macrae, 1991

Caballero-Mellado et al., Mexico Sugarcane Roots, stems, mealybugs 1995; Fuentes-Ramirez et al., 1993 . . J ■ . Coffea arabica L. Jiménez-Salgado et al., Mexico Roots, stems, rhizosphere (coffee) 1997

Camellia sinensis L. (tea), coffee, M usa Matiru and Thomson, Kenya Roots, rhizosphere 1998 acum inata Colla ♦ (banana)

Eleusine coracana Roots, stems, leaves, India (L.) Gaertn. (finger Loganathan et al., 1999 rhizosphere millet)

Tapia-Hemàndèz et al., Mexico Pineapple Roots, stems, leaves 2 0 ° 0

Daucus carota L. (carrot), Raphanus India sativus L. (radish), . Roots Madhaiyan et al., 2004 Beta vulgaris L. (beetroot), coffee

India and Muthukumarasamy et al., Oryza sativa L. Republic of Roots, stems, rhizosphere 2005; Muthukumarsamy et (rice) Korea al., 2007 24

Table 2-2. Differentiating characteristics for G. diazotrophicus (adapted from Gillis et al., 1989) Characteristics Gluconacetobacter diazotrophicus Formation of:

Water-soluble brown pigments on GYC medium '' + '

y-Pyrones from D-glucose '■ ■+'

y -Pyrones from D-fructose +

5-Ketogluconic acid from D-glucose

2,5-Diketogluconic acid from D-glucose ■ ■ + ’:>■

Ketogenesis from glycerol

Growth on the following carbon sources: Ethanol

Methanol

Sodium acetate +

Growth on the following L-amino acids in the presence of ■ ' ■.. ; y. D-mannitol as a carbon source:

L“0 ily L - lln e o n m e , L^li^ptoplian

L-Asparagine

L-Glutamine ■ ■ . .

Growth in the presence o f 10% ethanol ■■■ —

Formation of H2S —

Growth in the presence of 30% D-glucose

N2 fixation and growth on dinitrogen : +■' ■

Ubiquinone type Q10

Guanine-plus-cytosine content of DNA (mol%) 61-63 ;!y +, 90% or more o f the strains are positive; /, 11-89% o f the strains are positive; 90% or more o f the strains are negative 25 inactivation of the nifD gene o f PAL5, the resulting mutant strain MAd3A (nifD~, Kmr) is resistant to kanamycin and does not fix nitrogen (Sevilla etal., 2001).

2.2.2 Culture of G.diazotrophicus

Culturing of G. diazotrophicus strains PAL5 and MAd3A was done in LGIP medium

(quantities per litre: K2HPO4, 0.2 g; KH2PO4, 0.6 g; M gSQ ^^O , 0.2 g; CaCl2*2 H2 0 , 0.02 g;

Na2Mo0 4 »2 H2 0 , 0.002 g; FeCl3#6 H2 0 , 0.01 g; bromothymol blue in IN NaOII, 0.025 g; sucrose, 100 g; yeast extract, 0.025 g; bacto agar, 4 or 15 g; 1% acetic acid, pH 5.5) (Cavalcante and Dobereiner, 1988). PAL5 and MAd3A were cultured on solid, semisolid and/or liquid LGIP medium supplemented with 10 mM NH4(S0 4 )2 and incubated at 28°C for 2-5 days in the dark.

Liquid cultures o f PAL5 and MAd3A were agitated at 180 rpm and incubated for 2 days in the dark. Solid LGIP (bacto agar, 15g) plate cultures were prepared using either the streak plate method with a glycerol bacterial stock or by means of the spread plate method with a diluted bacterial suspension. Semi-solid LGIP medium (bacto agar, 4g) with no added inorganic nitrogen was used for the acetylene reduction assay. Kanamycin was added to the LGIP medium, at a concentration of 100 pg mL'1, for MAd3A cultures.

2.2.3 Detection of G. diazotrophicus by polymerase chain reaction

Primers used to detect both strains of G. diazotrophicus via PCR are listed in Table 2-3.

Detection of G. diazotrophicus from the pink sugarcane mealybug Saccharicoccus sacchari

Cockerell (Homiptera: Pseudococcidae) was observed by Franke-Whittle et al. (2005) using the

16S rDNA-based primers GDI39F/GDI916R. These primers were species-specific and produced a single product from strains of G. diazotrophicus only (Franke-Whittle et al., 2005). Tian et al.

(2009) desgined 16S rDNA-based primers GDI25F/GDI923R for detection of G. diazotrophicus in maize tissues by nested PCR. Producing an 876 bp amplicon, primers GDI39F and GDI916R 26

Table 2-3. Species-specific PCR primers for detection of G. diazotrophicus Primer Sequence Reference ~ GDI39F 5 ’ -TGAGTA ACGCGTAGGG ATCTG-3 ’ Franke-Whittle et al., 2005 GDI916R 5’-GGAAACAGCCATCTCTGACTG-3’

GDI25F TAGTGGCGGACGGGTGAGTAACG Tian et al., 2009 GDI923R CCTTGCGGGAAACAGCCATCTC 27 are located inside the GDI25F/GDI923R fragment. As such, primers GDI25F and GDI923R were used to amplify an 899 bp amplicon in the first round of nested PCR. Primers GDI39F and

GDI916R were used in the second round o f nested PCR.

Standard nested PCR was performed with each 20 pL PCR mastermix consisting of: 10X o f PCR buffer, 0.2 pM o f each of the forward and reverse primers, 0.2 mM of all four dNTPs,

Taq-polymerase (5 U p L 1), and sterile milli-Q II2O. For the first round of nested PCR, 1 pL of three-day old colonies diluted in 100 pL sterilized milli-Q H2O was transferred to 200 pL PCR microtubes containing the reaction mix. The subsequent temperature profile was carried out for

35 cycles for round one of nested PCR: 45 s o f dénaturation at 95°C, 45 s of annealing at 63°C, and 60 s o f extension at 72°C. Immediately afterwards, 1 pL aliquots of the PCR products from the first round were used as templates in the second round. Modifications to the temperature profile for the second round included: 30 s of dénaturation at 95°C, 45 s of annealing at 62°C, 30 s o f extension at 72°C, and decreasing the number of cycles to 30. Prior to each temperature profile, an initial dénaturation step at 95°C was carried out for 10 min. Upon the completion of cycles o f each temperature profile, a final extension step o f 10 min at 72°C was performed and the samples were maintained at 4°C. All PCR amplifications were executed either on the

Eppendorf Mastercycler Pro S Vapo.Protect thermal cycler or the Eppendorf Mastercycler

EpGradient thermal cycler. ?

The amplification products were analyzed via gel electrophoresis. The PCR products were loaded onto 1% agarose gels (agarose, lg; TAE buffer, 100 mL) stained with 10 pL EtBr at

concentration o f 500 pg m L'1. After a running time o f 40 min at 100V, the gels were visualized with the BioRad Quantity One Gel Doc software (Version 4.4.1) using the Foto/Prep UV transilluminator. (

28

For identification of G. diazotrophicus, the PCR products should be confirmed through sequencing and basic local alignment search tool (BLAST) analysis.

2.2.4 Evaluation of nitrogenase activity of G. diazotrophicus

Nitrogenase activity in G. diazotrophicus strains PAL5 and MAd3A were tested using the acetylene reduction assay (Reis et al., 1994; Hardy et al., 1968). The acetylene reduction assay is built on the correlation between the reduction of C2H2 into C2H4 and the reduction of N2 to NH3 in the process of N2 fixation (Dilworth, 1966). According to Hardy et al. (1968), the reduction of

C2H2 serves no useful purpose to the organism, but it does provide a simple and rapid method of measuring the activity o f N2-fixing systems. Sensitivity of C2H4 detection by gas chromatography and flame ionization is the key advantage o f the acetylene reduction assay. This sensitivity makes it possible to detect low levels of N2-fixing activity in biosphere samples, bacterial cultures and nitrogenase preparations.

Both strains were cultured for 48 h in LGIP liquid medium containing lOmM (N H ^SO^

The liquid culture was centrifuged at 10,000 rpm for 15 min and the supernatant was removed.

The remaining bacterial pellet was washed twice and re-suspended in sterilized milli-Q H2O. In a

40 mL GC vial, 100 pL of the bacterial suspension was grown in 5 mL o f semisolid LGIP medium under five different conditions: 1) no added inorganic nitrogen; 2) 1 mM o f NO3'; 3) 10 mM o f NO3'; 4) 1 mM of Nil/; and 5) 10 mM of NII^. After 48 h growth at 28°C, 10% (v/v) of air was removed from the culture vial and an equal volume o f C2H2 was'injected into the vial using a syringe. Following another 48 h o f incubation at 28°C, C2H4 production was measured on an HP 5890 Series II plus gas chromatograph (GC) (Agilent Technologies), fitted with a

Carboxen 1006 plot column (30m x 0.32 mm, Supelco Canada, Oakville, Ontario, Canada), using flame ionization detection (FID) with a carrier gas o f helium. From the headspace o f each 29 vial, 10 jxL was manually sampled using a gas-tight syringe (Hamilton, Reno, NV, USA) and injected into the GC, with an injection and oven temperature o f 120°C and a detector temperature of 250°C. Negative controls consisted o f un-inoculated media that were subjected to the same protocols as the PAL5 and MAd3A cultures.

A standard curve for the quantification o f C2H4 was developed using C2H4 in helium (984 ppm) standard (Alltech Associates, Inc., Deerfield, Illinois, USA) (Figure 2-1). Overall nitrogenase activity was measured in units of nmol o f C2H4 per h per mg of bacterial protein.

Bacterial protein estimation was carried out using the Bradford method for protein quantitation (Kruger, 2002). After the acetylene reduction assay, 5 mL of sterilized milli-Q H2O was added to each vial then vortexed at max speed for 30 s .T o a new 1.5 mL microfuge tube, 1 mL o f the diluted bacterial suspension was transferred along with 0.5 mL of 1 N NaOH. The mixture was then incubated at 100°C in a heating block for 10 min. Upon cooling to room temperature, the sample was centrifuged at 10,000 rpm for 10 min. For the Bradford microassay,

800 pL o f supernatant from the sample and 200 pL o f Bradford dye (refer to Appendix for components) were transferred into a plastic cuvette and analyzed with the Eppendorf

BioPhotometer at 595 nm. To quantify the amount of bacterial protein, a calibration curve was generated using bovine serum albumin (BSA) as the protein standard (100 pg mL'1) (Figure 2-2).

2.2.5 Statistical analyses

Statistical analysis of nitrogenase activity data was conducted with SPSS 18.0 (SPSS

Inc., Chicago, Illinois, USA). C2H4 productionUnder five different nitrogen conditions was compared by one-way analysis of variance. All multiple comparisons were performed by the

Least Significant Difference post-hoc test. All statistical tests were performed at the P=0.05 level. 18000 *

16000 -

14000 *

12000 *

E 1 0 0 0 0 * m 2 8000 *

6000 *

4000 -

2000 -

o -- — ,

0 50 100 150 200 250 300 350 40 Ethylene (nmol)

Figure 2-1. Ethylene standard curve Standard curve obtained using a C2H4 in helium (984 ppm) standard for the quantification of C2H4 from the acetylene reduction assay. 31

ì

: BSA Concentration (pg/ml)

Figure 2-2. Bovine serum albumin standard curve for bacterial protein estimation Standard curve obtained via the Bradford microassay using the protein standard bovine serum albumin for the estimation of protein in the bacterial cultures used for the acetylene reduction assay. ' . ■ 32

2.3 Results

2.3.1 Bacterial growth

On solid LGIP plates supplemented with (N H ^SO ^ G. diazotrophicus strains PAL5

(Figure 2-3A) and MAd3A (Figure 2-3 B) formed smooth, circular colonies that started off semi­ transparent with no color then later became opaque and orange after 5 days o f incubation. Both

PAL5 and MAd3A produced an under-surface pellicle during early growth, which concentrated on the surface and became yellow after incubating for 5 days in semi-solid LGIP medium without added inorganic nitrogen. Similar bacterial growth patterns were observed in semi-solid

LGIP media supplemented with either (NH4)2S0 4 or KNO3,. However, the key difference detected was that the pellicle appeared to be denser and thicker with the addition of (NH4)2S0 4 or KNO3. Finally, growth in liquid LGIP medium was observed 2 days post-incubation by the presence o f turbidity in the medium. Cultures incubated in liquid LGIP with no added inorganic nitrogen showed no growth unless a nitrogen starter dose was used. During bacterial growth, both PAL5 and MAd3 A produced acid causing the LGIP medium to change in color from orange to yellow at first, then gradually leading to no color.

2.3.2 Detection of G. diazotrophicus

Strains PAL5 and MAd3A were both detected by standard nested PCR analysis. Both strains presented an 899 bp amplicon in the first round (Figure 2-4A) and an 876 bp amplicon in the second round (Figure 2-4B), using 16s rDNA-based primers GDI25F/GDI923R and

GDI39F/GDI916R, respectively. Amplification products for 10' 1 to 10"4 dilutions of bacterial

DNA were observed in the first round of nested PCR (Figure 2-5A and Figure 2-5B), while the remaining dilutions, from 1 0 '5 to 1 0 ‘7, presented amplification products during the second round

(Figure 2-5C and Figure 2-5D). Serial dilution of PAL5 (Figure 2-5A and Figure 2-5C) and 33

Figure 2-3. G. diazotrophicus morphology on LGIP plates supplemented with (NH^SC^ A, Plating o f PAL5 at 10' 4 dilution B, Plating o f MAd3A at 10’4 dilution. Both strains formed smooth colonies with regular edges. Colonies were transparent during early growth and displayed a gradual change in color to dark orange. Acid produced by the bacteria lowered the pH of the medium causing a change in color from orange to yellow to no color. Figure 2-4. Nested PCR amplification for detection of G. diaztrophicus strains A, Nested PCR round 1 B, Nested PCR round 2 * L - 100 bp marker; P - PAL5; M - MAd3A 35

Figure 2-5. Sensitivity of detection for nested PCR amplification of PAL5 and MAd3A A, Nested PCR round 1 for PAL 5 B, Nested PCR round 1 for MAd3A C, Nested PCR round 2 for PAL 5 D, Nested PCR round 2 for MAd3A * L - 100 bp marker; NC - negative control 36

MAd3A (Figure 2-5B and Figure 2-5D) samples confirmed the sensitivity o f detection for the nested PCR procedure.

2.3.3 Nitrogenase activity of G. diazotrophicus

Nitrogenase activity of G. diazotrophicus strains PAL5 and MAd3A was tested under different culture conditions. A significant difference in C2H4 production was observed among the various cultures after the addition of different nitrogen sources (F=208.33, df=3, P<0.01) (Table

2-4). C2H4 was detected in the PAL5 culture with no added inorganic nitrogen at 296 nmol h' 1 mg' 1 protein, while C2H4 was not detected in the MAd3A culture. Nitrogenase activity o f PAL5 was observed even after the addition of both concentrations of N 0 3\ Nitrogenase activity was significantly lower after the addition of 1 mM NIL^ (192.44; Cl, 175.46-209.43; P<0.01). 1 mM

N03' (27.02; Cl, 10.03-44.00; P<0.01) and 10 mM N03' (68.00; Cl, 24.00-58.00; P<0.01) compared to the cultures with no added inorganic nitrogen. Adding 1 mM NFL}+ to the PAL5 culture caused a 35% decline in nitrogenase activity, while the addition o f 10 mM N R^ resulted in no N2 fixation. MAd3A and the control cultures did not exhibit any signs o f nitrogenase activity in any of the five culture conditions.

2.4 Discussion

2.4.1 Laboratory cultures of G. diazotrophicus

Successful cultivation of G. diazotrophicus strains PAL5 and MAd3A was achieved in laboratory conditions. As observed by Cavalcante and Dobereiner (1988), G. diazotrophicus, known to occur in high numbers in sugarcane roots and stems, demonstrates optimal growth with

10% sugar and pH 5.5. LGIP medium provided the conditions that emulate sugarcane juice for the cultivation of both strains. PAL5 and MAd3A isolates in pure culture have been shown to 37

Table 2-4. Nitrogenase activity of G. diazotrophicus strains PAL5 and MAd3A measured by the acetylene reduction assay Nitrogen Nitrogenase activity (nmol C2H4 h' 1 mg’1 protein) sources PAL5 Control MAd3A Control No inorganic 296± 16A 0 0 0 nitrogen

1 m M N O f 269±15 B 0 0 0

lOmM NOs' 228± 21c 0 0 0

1 mM NH4+ 103±19 D ' ; 0 0

10 mMNH4+ 0 0 0 . * Results are ± SD averages o f 3 replicates for each treatment. A significant difference in nitrogenase activity was observed among the bacterial cultures after the addition of different nitrogen sources (F=208.329, df=3, P<0.001). Different letters represent f multiple comparisons o f nitrogenase activity that are significantly different. 38

initially form small, white colonies that continue to grow and gradually turn yellow and orange

on LGIP plates. Formation of a veil-like pellicle in semisolid LGIP medium with no added

inorganic nitrogen was similar to that of azospirilla in semisolid malate medium. Addition of

(NH4)2S 0 4 and KNO3 to liquid and semisolid LGIP medium, respectively, resulted in turbidity and formation o f a thicker pellicle, indicating nitrogen-dependent growth by G. diazotrophicus.

Both strains o f G. diaztrophicus only require a nitrogen starter dose in liquid LGIP medium due to the aerobic conditions for growth. Lastly, acid production did not interfere with bacterial

growth, indicating tolerance of low pH environments and suggesting the existence o f effective protection mechanisms o f the cells (Stephan et al., 1991). Both PAL5 and MAd3A strains

characteristically fit the description of the acid-tolerant, N2-fixing bacterium discovered by

Cavalcante and Dobereiner (1988). Necessary for further work in introducing the bacterium into plants, both PAL5 and MAd3A were successfully established in a laboratory culture.

2.4.2 Activity of nitrogcnase in G. diazotrophicus

Nitrogenase activity of G. diazotrophicus strain PAL5 in semisolid culture was confirmed via the acetylene reduction assay method. Variation in the level of nitrogenase activity was

observed based on the type of nitrogen source available to the bacterium. The addition of N Ri+

caused a decline in nitrogenase activity, with the most prominent change in nitrogenase activity

observed under high concentrations o f NR*"*-. The influence of a nitrogen source on nitrogenase

activity is apparent in several studies, all of which demonstrated either partial or full inhibition of nitrogenase (Sorger 1969; Manhart and Wong, 1980; Hartmann et al., 1986). As reviewed by

Madigan et al. (2003), nitrogenase is not inhibited initially since NI-I3 that is excreted by the bacterium is incorporated into an organic form and used in biosynthesis. When NH3 starts to

accumulate and becomes in excess, the NtrC protein then becomes repressed, which inhibits the 39 expression of NifA, the N2 fixation activator protein. For certain In fix in g bacteria, nitrogenase activity can be regulated by the presence of NH3 via the “switch-off’ effect. Excess NH3 causes a covalent modification o f the nitrogenase enzyme, which leads to the loss of enzyme activity.

When NH3 becomes limited, the modified protein converts back to the active form and resumes fixing nitrogen. -

The capacity of the nitrogenase enzyme of G. diazotrophicus to function in the presence o f low concentrations o f NH*4- may have considerable ecological importance, especially since the bacterium lacks a nitrate reductase (Cavalcante and Dobereiner, 1988). Two other known I n ­ fixing bacteria also lacking nitrate reductase are Azotobacter paspali, specifically associated with

Paspalum notatum, and Bacillus azotofixans (later renamed Paenibacillus azotofixans)(Se\dm et al., 1984; Cavalcante and Dobereiner, 1988)! The presence of NO3' has been shown to inhibit nitrogenase in Azotobacter vinelandii, which expresses the nitrate reductase enzyme (Sorger,

1969). It was suggested that NO3' itself did not repress nitrogenase, but instead NH3, the metabolic product of NO3' reduction, was the main cause o f nitrogenase inactivity (Sorger,

1969). The lack o f nitrate reductase prevents additional accumulation o f NH3 since denitrification cannot occur (Madigan et al., 2003). Thus, the ability to fix nitrogen in the presence of NO3' and low concentrations of NH4+ suggests that G. diazotrophicus could be ideally employed in the complementation o f fertilizer application with N2 fixation in the field.

2.5 Conclusion

PAL5 and MAd3A were phenotypically and molecularly confirmed to be strains of G. diazotrophicus. Both strains were able to withstand low pH environments and high sugar concentrations. Optimal growth was observed in 10% sucrose and pH 5.5. Growth in liquid 40

LGIP medium was entirely dependent on the addition of a nitrogen starter dose. Nitrogen- dependent growth was also demonstrated by the growth o f a thicker pellicle in semisolid medium supplemented with KNO3 or (N H ^SQ*. Nested PCR analysis further validated the identity of

PAL5 and MAd3A by amplifying the 899 bp and 876 bp products. Additionally, nested PCR seemed to be a sensitive method for detecting bacteria at various dilutions.

The capacity of PAL5 to fix nitrogen was tested and established by the acetylene reduction assay. Nitrogenase activity was not only observed in conditions with no added inorganic nitrogen, but also in conditions with added NO3'. The addition of ImM N H r caused a decrease in nitrogenase activity; however, the addition of lOmM NH4+ resulted in the complete inhibition o f N2 fixation. The influence of nitrogen conditions on the activity of nitrogenase indicates that there is a limiting factor for N2 fixation.

Taken together, these results and past literature reviews suggest that G. diazotrophicus may be a promising candidate for the possibility o f N2 fixation within graminaceous plants. N2- fixing bacteria associated with N2 fixation in graminaceous plants also include the rhizospheric bacteria Azospirillum Tarrand et al. 1979 spp. and Azotobacter Beijerinck 1901 spp. (Dobereiner,

1988; Stephan et al., 1991; Bashan, 1999); however, G. diazotrophicus stands out from the others due to its endophytic disposition. Culturing and characterizing both strains of G. diazotrophicus was necessary to verify that these strains were useable for further applications.

Inoculation of Sorghum bicolor (L.) Moench with G. diazotrophicus as well as factors influencing the endophytic assocation are explored in the following chapters. 41

References

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3.1 Introduction

The interior and exterior parts o f the plant remain forbidden territories for a vast majority o f microorganisms due to the production of antimicrobial compounds by the plant, including terpenoids, benzoxazinone, flavonoids, and isoflavanoids (Bais et al., 2006). However, plants are capable o f associating with an array o f microbial organisms and can occasionally require the presence o f associated bacteria for their own growth and establishment in diverse ecosystems

(Hardoim et al., 2008). Microbes often profit from the readily available nutrients in plants, while plants receive benefits from the bacterial associates by growth enhancement or stress reduction

(Hardoim et al., 2008). These mutualistic interactions could have emerged as a result of the clear positive selection exerted on the associations between plants and microbes (Thrall et al., 2007).

Microorganisms, such as bacteria, that occur within plants are classified as endophytes

(Hardoim et al., 2008). The term endophyte, meaning “in the plant” (endon Gr. = within, phyton

= plant), has been broadly used, from interactions o f mycorrhizal or mycorrhizal-like beneficial fimgi, to interactions o f saprophytic or pathogenic fungi, as well as bacteria (Rothballer, et al.,

2009). Endophytes are most commonly defined as organisms that cause “infections o f plant parts which are inconspicuous, the infected host is at least transiently symptom-less, and microbial colonization can be demonstrated internally...” (Stone et al., 2000). Despite the fact that this definition was used to characterize fungal endophytic interactions, it is equally applicable to endophytic bacteria-plant interactions (Rothballer et al., 2009). Stone et al.’s (2000) definition furthermore encompasses avirulent microorganisms and latent pathogens, stressing that endophytic bacteria do not cause apparent harm and are characterized by the lack of macroscopically visible pathologic symptoms. In order to recognize “true” endophytes, bacteria 45 must be isolated from surface-sterilized tissues and there must be microscopic evidence to visualize “tagged” bacteria inside the plant (Reinhold-Hurek and Hurek, 1998). For endophytic bacteria-plant associations that do not meet, the latter criterion, use of the term “putative” endophytes has been recommended instead (Rosenblueth and Martinez-Romero, 2006).

Bacterial endophytes can be classified as either obligate or facultative (Bacon and

Hinton, 2006; Schulz and Boyle, 2006). Obligate endophytes are strictly dependent on the host plant for their growth and survival (Hardoim et al., 2008). Facultative endophytes have a life cycle stage in which they exist outside host plants and live in other habitats, primarily soil

(Hardoim et al., 2008). It is believed that the vast majority o f endophytic microorganisms have a tendency towards the biphasic lifestyle, alternating between plants and the environment

(Rosenblueth and Martinez-Romero, 2006). Endophytes that infect plants from soil must be competent root colonizers (Sturz et al., 2000). Successful endophytic associations can be determined by the chance of emerging roots coming into contact with effective numbers of bacteria that possess dedicated genetic systems involved in bacterial-plant crosstalk (Ryan et al.,

2008). In Hardoim et al.’s (2008) review, it was proposed that facultative endophytes could be further categorized as competent, opportunistic, or passenger endophytes. Competent endophytes include those bacteria that possess key genetic machinery required for colonizing the plant and persisting in it. Opportunistic endophytes are proficient rhizosphere colonizers that colonize the narrow region o f the soil surrounding the roots, which become endophytic by inadvertently entering the root tissues even though they lack the key genes to reside in the plant. Finally, passenger endophytes enter plants purely by chance due to absence o f any machinery for efficient root colonization or entry. • • A 46

The best-studied interactions o f N2-fixing bacteria with plants leading to efficient

biological nitrogen fixation (BNF) are the root nodule-forming rhizobia-legume symbioses

(Rothballer et al., 2009). However, most agricultural crops such as Triticum L. spp. (wheat),

Oryza sativa L. (rice), Z ea mays L. (maize), and Saccharum L. spp. (sugarcane) are not able to

form the root nodules in which the bacteria grow and perform N2 fixation. Because of this much

of the continuing research on N2 fixation with non-legumes has focused on N2-fixing bacterial

endophytes (Rothballer et al., 2009). Gluconacetobacter diazotrophicus (Gillis et al. 1989)

Yamada et al. 1997 is considered to be a true facultative competent endophyte that serves as a

major source of BNF to sugarcane (James et al., 1994; Sevilla et al., 2001). The crop can

accumulate between 100-200 kg nitrogen ha"1 per season (Reis et al., 2007). However, nearly all

o f this fixed nitrogen is removed at harvest through the burning of 25% o f the senescent leaves,

which leaves less than 10% of the fixed nitrogen in the field (Graham et al., 2002; de Resende et

al., 2006). As a result, continuous cropping of sugarcane should deplete soil nitrogen reserves

and cane yields should gradually decline (Rothballer et al., 2009). This decline is not observed

even after decades or centuries of cane cropping suggesting that sugarcane must benefit

significantly from fixed nitrogen inputs through BNF (Rothballer et al., 2009). BNF inputs have

been shown to range from 25-60% o f the plant nitrogen, suggesting that N2-fixing bacteria need

to fix N2 effectively in sugarcane to generate this amount of nitrogen nutrition (Boddey et al.,

■ 2001). : - v - V .

Sorghum bicolor (L.) Moench (sorghum) is a cereal crop originated in northern Africa

that is now widely cultivated in tropical and subtropical regions (Dicko et al., 2006). According

to the Food and Agriculture Organization of the United Nations, the top five countries for

sorghum production, as of 2009, are: India, United States o f America, Nigeria, Sudan and ' 47

Ethiopia. Additionally, sorghum is the fifth most important cereal crop after wheat, rice, maize

and Hordeum vulgare L. (barley) (Food and Agriculture Organization of the United Nations,

2009). Sorghum belongs to the family Poaceae, subfamily Panicoideae and tribe Andropogoneae,

which consists of some of the most efficient biomass accumulators (Dicko et al., 2006).

Important for productivity, sorghum carries out C4 photosynthesis, due to biochemical and

morphological specializations that increase net carbon assimilation at high temperatures (Hatch

and Slack, 1966). Sorghum also has the potential to adapt itself to the natural environment and is

often called a “nature-cared crop” since it requires minimal anthropogenic care such as irrigation

and insect removal (Dicko et al., 2006). The drought-resistant crop can adapt to hot, semi-arid

environments that are too dry for other cereals, as well as to heavy soils o f the tropics where

tolerance to waterlogging is often a requirement (Dicko et a!., 2006).

As in sugarcane, sorghum shows high sucrose accumulation and increased biomass particularly in the stalks (Reddy and Reddy, 2003). However unlike sugarcane, sorghum is

known for its wide adaptability, drought resistance, waterlogging tolerance, and saline-alkali tolerance (Reddy and Reddy, 2003). As such, the interest in sorghum production has increased

due to these characteristics as well as its importance and use for human consumption, animal

feed, alcohol production and industrial products (Dicko et al., 2006). Cultivation of sugarcane

could become difficult given that water availability may potentially become a . constraint to

agricultural production (Reddy et al., 2005). As such, sorghum, particularly the sweet cultivars,

would be a logical crop alternative, since sorghum can be grown with less irrigation and rainfall

and decreased cost inputs (Reddy et al., 2005).

Inoculation of cereals with N2-fixing bacteria seems to be the next plausible step for the

enhancement o f crop yields and is now the focal point of numerous studies in agronomy, biology ' ■ 48'' and environmental science. By inoculating different sorghum cultivars with G. diazotrophicus grown under greenhouse and field conditions, the capacity of different sorghum cultivars for colonization will be examined in this chapter. It is hypothesized that plant cultivar and growth conditions will influence the capacity for colonization o f sorghum by G. diazotrophicus. Using various sorghum cultivars in two different growth environments, two separate trials were carried out to determine the factors influencing the endophytic association, which will be addressed further in Chapter 4.

3.2 Materials & Methods

3.2.1 Sorghum cultivars

Seven different sorghum cultivars were used to evaluate the colonization capacity of G. diazotrophicus under greenhouse and field conditions. Five o f the seven cultivars were grain sorghum and the remaining two cultivars were sweet sorghum (Table 3-1). All seeds were provided by Dr. Om P. Dangi and Dr. K. Anand Kumar of Agriculture Environmental Renewal

Canada Inc. (Delhi, Ontario, Canada).

3.2.2 Bacterial strains & inoculum

Using liquid LGIP medium (refer to Chapter 2, Section 2.2.2 for components), bacterial suspensions o f G. diazotrophicus strains PAL5 and MAd3A (refer to Chapter 2, Section 2.2.1 for details of the strains) were cultured and prepared for inoculation. After 48 hrs growth, cultures were collected at 10,000 rpm for 15 min and re-suspended in 0.9% NaCl solution to a concentration of ~108 colony forming unit (CFU)/ml. Concentration was determined by plating o f serial dilutions of the bacterial suspension. A ten-fold dilution series consisting of 1 0 '1 to 1 0 '8 dilutions was performed for both strains. For each dilution, 100 pi aliquots were plated on solid f ì 49

ì

I. '

Table 3-1. Sorghum cultivars from Agriculture Environmental Renewal Canada Inc. Cultivar Background 98W Grain, inbred parental

Nutriplus BM R Silage, tall, large biomass

275 Grain, parental

CG SH 27 :;'V Grain, hybrid, commercial

' CGSH 8 Grain, hybrid, commercial

n u i ; Sweet, inbred

CSSH 45 -Sweet'■■■ ■" : LGIP medium and incubated for five days at 28°C in the dark. Following incubation, GFUs were

counted from each plate and the bacterial concentration (per ml) was calculated using Equation

3-1

Equation 3-1. Concentration of bacteria Number of CFU Bacterial concentration = Plated volume (ml) x Plated dilution

For visualization purposes, the gw^-marked G. diazotrophicus strain

UAP5541/pRGS561, generously provided by Dr. Kevin Vessey (Saint Mary’s University,

Halifax, Nova Scotia, Canada), was additionally cultivated for inoculation. The wild-type G.

diazotrophicus strain UAP5541 was conjugated with the plasmid pRGS561 (Smr, Spr), allowing

for the constitutive expression of gusA -nptll (Fuentez-Ramirez et al., 1999). As.with PAL5 and

MAd3A, a suspension of UAP5541/pRGS561 was cultured in liquid LGIP medium for 48 hrs

and collected at 10,000 rpm for 15 min. The bacterial pellet was then resuspended in 0.9% NaCl o solution to a concentration of ~10 CFU/ml.

3.2.3 Germination of sorghum

Prior to germination, all seeds were thoroughly washed with tap water, surface-sterilized

with 1% NaCIO for 15 min, and rinsed with sterilized milli-Q H2O. For each cultivar, 72 seeds

were planted in 1:3 (v:v) sand:vermiculite mix in plastic germination trays with 1 seed/hole. All jcultivars were germinated under greenhouse conditions with daily watering until the seedlings

reached the 2- to 3-leaf stage (15-20 days after germination).

3.2.4 Inoculation of sorghum

Two colonization studies, one conducted in the greenhouse and the other conducted in the

field, were carried out with all plants of each cultivar within each trial subjected to the same

procedures and methods. Both trials were executed at the facilities of the Southern Crop ■ 51

Protection, and Food Research Centre, Agriculture and Agri-Food Canada, London, Ontario,

Canada. Seedlings at the 2- to 3-leaf stage were removed from the germination trays and washed with tap water to get rid of any adhering sand/vermiculite. Inoculation o f sorghum with PAL5

and MAd3A was performed via the pruned-root dip method. For each cultivar, 10% of the root

lengths were cut then inoculated by submerging the roots in the bacterial inoculum for 30 min.

Control plants were inoculated with 0.9% NaCl solution without the bacterial suspension.

Immediately after inoculation, seedlings for both trials were transferred into individual pots

containing 1:3 (v:v) sand:Sun Gro Sunshine Mix 2 (Table 3-2). Field seedlings were initially left

in the greenhouse for one week to allow for acclimatization and ensure inoculation. A total of 30 plants per cultivar for each trial (10 plants inoculated with PAL5, 10 plants inoculated with

MAd3A, and 10 un-inoculated plants) were grown under greenhouse and field conditions for 60

days. Plants grown in the greenhouse were watered daily.

3.2.5 Sample preparation & extraction for polymerase chain reaction (PCR)

Plants from both trials were harvested and collected 60 days post-inoculation. To remove the adhering soil, debris and external microorganisms, samples were first washed with tap water then surface-sterilized for 15 min with 1% NaClO. Each plant was separated into roots, stems and leaves then cut into 5 cm pieces and each tissue type stored separately in plastic 2 lb bags at -

80°C prior to analysis. j Using a mortar and pestle, bacteria were extracted from greenhouse plant tissues by

grinding 2 g o f tissue in 4 ml of sterilized milli-Q H2O. Simultaneously, 50 mg PVPP was added to absorb polyphenols and humic substances that are known to chelate Mg , which is an

important element in PCR (Möller et al., 1994). Extraction of bacteria from field plant tissues was performed by grinding 4 g o f tissue in 8 ml of sterilized milli-Q H2O with 100 mg PVPP. I 52 }■

'5

Table 3-2. Growing mix components for Sun Gro Sunshine Mix 2 Sun Gro Sunshine Mix 2 / LB2 (Basic Professional Growing Mix)

Components:

• Canadian Sphagnum peat moss

■ ■ • Coarse perlite • -Gypsum;:'

• Dolomitic limestone The ground tissue samples were then filtered through a moist Qualitative 415 9 cm filter paper, collecting 1.5 ml aliquots. After centrifuging at 10,000 rpm for 15 min, the pellets were re­ suspended in 100 pi o f autoclaved milli-Q ffeO.

For nested PCR, 1 pi o f the extracted plant samples were used to detect G. diazotrophicus within the plant tissues in the first round. In the second round, 1 pi o f the PCR products from the first round were successively used as templates. For details on the PCR protocol, refer to Chapter

2, Section2.2.3.

3.2.6 Re-isolation & enumeration of G. diazotrophicus population from plant tissues

The most probable number (MPN) method (Sutton, 2010) was used to verify bacterial population numbers within the plant tissues. The bacterial population was quantified in three plants o f each cultivar that tested positive during PCR screening. All plants were separated into roots, stems and leaves and surface-sterilized with 1% NaCIO for 15 min. Using a mortar and pestle, 2 g o f tissue were ground up in 4 ml of sterilized milli-Q ^ O .: After serially diluting the ground tissue samples to 10'1 to 10'4 dilutions, 100 pi aliquots were collected and inoculated into three replicate 40 ml vials containing 5 ml of nitrogen-free semisolid LGIP medium. After five days o f incubation at 28°C in the dark, population size was estimated by establishing the MPN.

Using the McCrady table (de Man, 1983) (refer to Appendix), MPN was determined based on the pellicle growth o f N2-fixing bacteria in three replicate vials for each dilution. Through phenotypic characteristics, the identity of G. diazotrophicus was then confirmed by isolating the bacteria on solid LGIP plates supplemented with 10 mM N H ^SO ^.

3.2.7 Visual confirmation of colonization by G. diazotrophicus

Colonization of cultivars Nlll and 275 by G. diazotrophicus was confirmed visually via the histochemical/?-glucuronidase (GUS) assay. Thirty-six seeds of each cultivar were prepared 54 for germination as mentioned in Section 3.2.3. At the 2- to 3-leaf stage, the pruned-root dip method (refer to Section 3.2.4) was performed using the gusA-marked strain of G. diazotrophicus. Seedlings were transplanted and grown under greenhouse conditions in separate pots containing 1:3 (v:v) sand: Sun Gro Sunshine Mix 2. For each cultivar, 20 plants were inoculated with UAP5541/pRGS561 and five plants served as un-inoculated controls.

One week after inoculation, two plants of each cultivar were harvested every four days to test for colonization by the -marked strain o f G. diazotrophicus. Plants were first washed under running tap water and rinsed with sterilized milli-Q H2O. Separating the tissues into roots, stems and leaves, all samples were cut into 2 cm pieces and placed into individual 1.5 mL microcentrifuge tubes. To each tube, 500 pi of the IX histochemical staining solution (refer to

Appendix for components) was added and left to incubate overnight at 37°C to allow the degradation of X-Gluc by /2-glucuronidase. Following incubation, the staining solution was removed and replaced with 70% ethanol to clear any pigmentation from the plant. For the stems and leaves, repeated changes with 95% ethanol were necessary to adequately remove the green pigmentation from the tissues. All tissues were stored in 70-95% ethanol at 4°C. Bacterial colonization was visualized and photographed using the Nikon SMZ1500 stereomicroscope fitted with a Nikon Digital Eclipse DXM1200 digital camera.

3.2.8 Statistical analyses j All statistical analyses were performed with SPSS 18.0 (SPSS Inc., Chicago, Illinois,

USA). Three-factor univariate analysis o f variance was conducted to compare proportion of plants showing bacterial presence and bacterial population numbers among the different cultivars, tissues and growth environments. All multiple comparisons were evaluated with the 55

Least Significant Difference post-hoc test. All statistical tests were performed at the P=0.05 level.

3.3 Results

3.3.1 Germination of sorghum

Germination success varied among the different sorghum cultivars (Table 3-3 and Table

3-4). Cultivars 98W, Nutriplus BM R, 275, CGSH 27, CGSII 8 and N 111 showed greater than

80% germination success allowing 30 plants per cultivar for each trial for inoculation.

Germination success for CSSH 45, one o f the two sweet cultivars, was 55%, thus allowing only

18 plants per trial for inoculation.

3.3.2 Detection of G. diazotrophicus from greenhouse sorghum tissues

All plants, inoculated and grown in the greenhouse, were screened by nested PCR to detect G. diazotrophicus within plant tissues. G. diazotrophicus was detected in the roots, stems and leaves o f the inoculated plants signified by the amplified 876 bp product indicator (refer to

Appendix for PCR data). Although G. diazotrophicus was found in both grain and sweet cultivars, bacterial detection was not observed in all inoculated tissue samples. As such the proportion o f plants showing bacterial presence was calculated, using Equation 3-2.

Equation 3-2. Proportion of plants showing bacterial presence Number o f PCR positive samples Proportion o f plants showing bacterial presence = — —------— ------—— ------——— Number o f inoculated plants

The proportion of plants showing bacterial presence in the roots, stems and leaves o f each cultivar ranged from 50-92%, 20-75%, and 0-67%, respectively (Table 3-5). A significant difference in the proportion of plants showing bacterial presence was noted among the root, stem and leaf tissues, indicating an effect of tissue type (F=12.79, df=2, P<0.01) (Figure 3-1). There Table 3-3. Germination success of seven sorghum cultivars for greenhouse trial Cultivar Number o f plants emerged Germination success (%)

CG SH 27 V 36 100

98W • 35 97 ;

CGSH 8 32 91

Nutriplus BM R 36 : ' 100

275 30 . : 83

N 111 36 100

C SSH 45 . 20 : \ : : 56 * Total number o f seeds planted for each cultivar: 36 Table 3-4. Germination success of seven sorghum cultivars for field trial Cultivar Number of plants emerged Germination success (%) CG SH 27 36/ 100

98W V . 33'.'', ;■ 92 ■

CGSII 8 36 l o o ;

Nutriplus BM R 35 9 i ■ v '.

275 32.'',- 89

N i l l 33 . 92 .

CSSH 45 !9 ; 53

* Total number o f seeds planted for each cultivar: 36 Table 3-5. Proportion of greenhouse plants showing presence of G. diazotrophicus Proportion of Plants Showing Bacterial Presence (%) Cultivar Root Stem Leaf CG SH 27 55 20 ; 0

98W 50 40 10

CGSIT 8 70 60 30

Nutriplus BM R 80 60 25

275 60 ■; 5 5 45 :

N i l i 85 ■ 75 . 65

C SSH 45 92 75 ■ 67

* Proportion calculated from PCR screening results v Root . -'Stem ■ ■ ■, Leaf . , Sorghum Tissue

Figure 3-1. Effect of tissue type on the proportion of greenhouse plants showing presence of G. diazotrophicus ■ A significant difference in the proportion o f plants showing bacterial presence was observed among the root, stem and leaf tissues, indicating an effect of tissue type (F=12.79, df=2, P<0.01). Among the tissues, different letters represent multiple comparisons that are significantly different. Error bars represent SE. .¡ ^ . . 60 was a higher proportion of plants showing bacterial presence in the roots compared to the stems

(15.24; Cl, 0.50-29.98; P=0.04; where Cl represents confidence interval) and leaves (35.71; Cl,

20.98-50.45; P<0.01). A higher proportion o f plants showed bacterial presence in the stems compared to the leaves (20.48; Cl, 5.74-35.21; P=0.01).

Among the seven different sorghum cultivars, the proportion of plants showing bacterial presence varied significantly and the effect o f cultivar type was observed (F=6.46, df=6, P<0.01)

(Figure 3-2). The sweet cultivar N il 1 had a higher proportion of plants showing bacterial presence than the grain cultivars CGSH 27 (50.00; Cl, 27.49-72.51; PO.01) and 98W (41.67;

Cl, 19.15-64.18; P<0.01). The sweet cultivar CSSH 45 had a higher proportion o f plants showing bacterial presence than all five grain cultivars (P<0.05).

During nested PCR screening, G. diazotrophicus, was not detected in the root, stem and leaf tissues o f all control plants.

3.3.3 Detection of G. diazotrophicus from field sorghum tissues

PCR screening was performed to confirm that G. diazotrophicus was detectable within the root, stem and leaf tissues of all field plants. In the roots, stems and leaves of the inoculated plants, the bacterium was detected by testing positive for the 876 bp product after the second round o f amplification (refer to Appendix for PCR data). Although G. diazotrophicus was present in both the grain and sweet cultivars, the bacterium was not detected in all the inoculated jissue samples; as a result, the proportion o f plants showing bacterial presence was calculated using Equation 3-2 (refer to Chapter 3, Section 3.3.2).

The proportion o f plants showing bacterial presence in the roots, stems and leaves o f each cultivar ranged from 45-100%, 40-100%, and 5-85%, respectively (Table 3-6). Among the root, stem and leaf tissues, a significant difference in the proportion of plants showing bacterial 61

CGSH27 98W CGSH8 Nutriplus 275 N i l i CSSH 45 BMR Sorghum Cultivar

Figure 3-2. Effect of cultivar type on the proportion of greenhouse plants showing presence of G.diazotrophicus A significant difference in the proportion o f plants showing bacterial presence was observed among the cultivars, indicating an effect o f cultivar type (F=6.46, df=6, P<0.01). Among the cultivars, different letters represent multiple comparisons that are significantly different. Error bars represent SE. ' " ' • ■ 62

Table 3-6. Proportion of field plants showing presence of G. diazotrophicus Proportion of Plants Showing Bacterial Presence (%) Cultivar Root Stem Leaf CG SH 27 5° '':V. 50 : ■■ v,^5;'

. 98W 55 ; 45 r 30

C G S II8 45 ■.V. 5'

Nutriplus BM R 75 75 65

275 85 85 70

N i l i 100 100 85

C SSH 45 83 83 67

* Proportion calculated from PCR screening results

t 63 presence was observed indicating the effect of tissue type (F=9.01, df=2, PO.Ol) (Figure 3-3).

There was a higher proportion of plants showing bacterial presence in the roots (23.81; Cl,

10.90-36.72; P O .O l) and stems (21.67; Cl, 8.76-34.57; PO .O l) compared to the leaves.

Among the various sorghum cultivars, significant differences in the proportion o f plants

showing bacterial presence was observed, indicating an effect of cultivar type (F=14.31, df=6,

P<0.01) (Figure 3-4). The sweet cultivar N il 1 had a higher proportion o f plants showing bacterial presence than the grain cultivars CGSH 27 (60.00; Cl, 40.29-79.71; PO.Ol), 98W

(51.67; Cl, 31.95-71.38; PO.Ol), CGSH 8 (65.00; Cl, 45.29-84.71; PO.Ol), and Nutriplus BMR

(23.33; Cl, 3.62-43.05; P=0.02). The sweet cultivar CSSH 45 had a higher proportion o f plants showing bacterial presence than the grain cultivars CGSH 27 (42.78; Cl, 23.06-62.49; PO .O l),

98W (34.44; Cl, 14.73-54.16; PO.Ol), and CGSH 8 (47.78; Cl, 28.06-67.49; PO.Ol).

During nested PCR screening, G. diazotrophicus was not detected in the root, stem and leaf tissues o f all control plants.

3.3.4 Effect of growth environment on G. diazotrophicus detection

The proportion of field plants and greenhouse plants showing bacterial presence was compared to determine the overall effect o f the growth environment on colonization. An overall significant difference in the proportion o f plants showing bacterial presence was observed between the greenhouse and field trials among the seven cultivars (F=9.49, df=13, PO.Ol)

¡(Figure 3-5). For cultivar CGSH 8, a significantly higher proportion of greenhouse plants than field plants showed bacterial presence (23.33; Cl, 2.80-43.87; P=0.03). For cultivar 275, a significantly higher proportion o f field plants than greenhouse plants showed bacterial presence

(26.67; Cl, 6.13-47.20; P=0.01). 64

?•

100

* 90 - 01u § 80 A « £ 7; 70 •c

i 60

50

40

30

20 ii 10 i li 0 Root Stem Leaf Sorghum Tissue

Figure 3-3. Effect of tissue type on the proportion of field plants showing presence of G. diazotrophicus A significant difference in the proportion of plants showing bacterial presence was observed among the tissues, indicating an effect of tissue type (F=9.01, df=2, P<0.01). Among the tissues, different letters represent multiple comparisons that are significantly different. Error bars represent SE.

J. c 100

£ 90 0 1 u S 80 s & 70 ‘C

60 flO 0 0 c 1 50 o J Z ' i A U) 40 c 5 CL 30 o c' o 20

R 8 10

0 CGSH27 98W CGSH8 Nutriplus 275 Nlll CSSH45 , ''BMR Sorghum Cultivar

Figure 3-4. Effect of cultivar type on the proportion of field plants showing presence of G. diazotrophicus A significant difference in the proportion of plants showing bacterial presence was observed among the different sorghum cultivars, indicating the effect o f cultivar type (F= 14.31, df=6, P|<0.01). Among the cultivars, different letters represent multiple comparisons that are significantly different. Error bars represent SE. 66

CGSH27 98W CGSH8 Nutriplus 275 Nlll CSSH 45 BMR ■' Sorghum CuMvar

Figure 3-5. Effect of growth environment on the proportion of plants showing presence of G. diazotrophicus A significant difference in the proportion of plants showing bacterial presence was observed between the greenhouse and field trials among the seven cultivars (F=9.49, df=13, P<0.01). Within a cultivar, different letters represent multiple comparisons that are significantly different. Error bars represent SE.

f 67

3.3.5 Population of G. diazotrophicus in greenhouse plant tissues

For quantification and confirmation of bacterial colonization, the MPN method was used to enumerate the population of G. diazotrophicus within plant tissues that screened positive for bacterial detection by PCR. Bacterial population numbers in the roots ranged from 8.43x102 to

7.16x10 MPN g tissue, while the population numbers in the stems varied from 3.74x10 to

1.68xl03 MPN g'1 tissue. Leaves of the inoculated plants had population numbers ranging from low as 1.08xl02 to as high as 1.28xl03 MPN g'1 tissue (Table 3-7). Among the root, stem and leaf tissues, there was a significant difference in bacterial population numbers, indicating the effect o f tissue type (F=18.19, df=2, P<0.01) (Figure 3-6). Higher bacterial population numbers were observed in the roots compared to the stems (2.18x103; Cl, 1.23xl03-3.13xl03; P<0.01) and leaves (2.74X103; Cl, 1.79x 103-3.69x 103;P<0.01).

The effect of cultivar type was observed with significant differences in bacterial population numbers among the different sorghum cultivars (F=3.73, df=6, P<0.01) (Figure 3-7).

The sweet cultivar N il 1 had higher bacterial population numbers than all the other cultivars

(P=0.04). ; :

During PCR screening, bacteria were not detected in the leaves of CGSH 27. As a result,

MPN was not determined for the leaf tissues o f CGSH 27. From the roots, stems and leaves of the control plants, re-isolation of G. diazotrophicus was not observed.

3.3.6 Population of G. diazotrophicus in field plant tissues

Using the MPN method, the population o f G. diazotrophicus from field sorghum tissues was enumerated to confirm bacterial colonization. In the root tissues, bacterial population numbers ranged from 1.15xl03 to 1.86x104 MPN g'1 fresh weight. Population numbers in the stems varied from 5.67xl02 to 6.88xl03 MPN g'1 freshweight. Lastly, the leaves of the Table 3-7. Most probable number of G. diazotrophicus population in greenhouse plant tissues Bacterial Population (MPN g'1 tissue) Cultivar Root Stem Leaf CGSH 27 1.69X103. 6.97x102 . n/a

98W 8.45 xlO2 4.56xl02 1.08xl02

CGSII 8 9.47x102 3.74xl02 1 72x102

NutriplusBMR 3.61 xlO3 1.68xl03 1.88xl02

2 7 5 3.04x103 4.43 xlO2 2.46 xlO2

N i l i 7.16X103 1.58xl03 1.28xl03

C SSH 45 4 .12x l03 8.87xl02 2.21 xlO2

* Results are averages o f three PCR positive replicates/cultivar. 69

4000 -

A 3500 *

| 3000 v *4 ' ' ' bo €LZ 2500 s o 2000 15

1500 B

1000

500

Root Stem Leaf Sorghum Tissues

Figure 3-6. Effect of tissue type on G. diazotrophicus population numbers in greenhouse plants A significant difference in bacterial population numbers was observed among the root, stem and leaf tissues, indicating the effect of tissue type (F=18.19, df=2, PO.Ol). Among the tissues, different letters represent multiple comparisons that are significantly different. Error bars represent SE. j l pa t/ .;/ ,'■/■• -plants/,.,;'/ Figure 3-7. Effect of cultivar type on on type cultivar of Effect 3-7. Figure A significant difference in bacterial population numbers was observed among the cultivars, cultivars, the among observed was numbers population bacterial in difference significant A letters represent multiple comparisons that are significantly different. Error bars represent SE. represent bars Error different different. cultivars, the Among significantly are that comparisons P<0.01). multiple df=6, represent letters (F=3.73, type f cultivar o effect the indicating

Bacterial Population (MPN g*1 tissue) 4000 3500 -1 4500 G. diazotrophicus G. population numbers in greenhouse in numbers population 70 71 inoculated plants had population numbers ranging from 1.53x10 to 7.97x10 MPN g' fresh weight (Table 3-8). Indicating an effect of tissue type, a significant difference in bacterial population numbers was observed among the root, stem and leaf tissues o f the various sorghum cultivars (F=40.19, df=2, P<0.01) (Figure 3-8). Higher bacterial population numbers were observed in the roots compared to the stems (5.44xl03; Cl, 3.43xl03-7.46xl03; P<0.01) and leaves (9.07xl03; Cl, 7.05xlO3-1.11 xlO4; P<0.01). There were higher bacterial population numbers in the stems compared to the leaves (3.63xl03; Cl, 1.61xl03-5.64xl03; P<0.01).

Among the sorghum cultivars, bacterial population numbers within plant tissues significantly differed, indicating an effect of cultivar type (F=9.52, df=6, P<0.01) (Figure 3-9).

The sweet cultivars N il 1 and CSSH 45 had higher bacterial population numbers than four o f the grain cultivars (P<0.01). Bacterial population numbers in the grain cultivars CGSH 27, 98W,

CGSH 8, and Nutriplus BM R were not significantly different (P>0.06).

From the roots, stems and leaves o f the control plants, re-isolation o f G. diazotrophicus was not observed.

3.3.7 Effect of growth environment on G. diazotrophicus population

Population numbers o f G. diazotrophicus within the roots, stems and leaves o f each cultivar from each colonization trial were compared to assess the potential effect o f the environment in which the plants were grown. Among the seven cultivars, an overall significant

in bacterial population numbers was observed between the greenhouse and field trials

(F=11.46, df=13, P<0.01) (Figure 3-10). Higher bacterial population numbers were observed in field plants than greenhouse plants for cultivars 275 (6.03x103; Cl, 3.63xl03-8 43xl03; PO.01),

N il! (4.91 xlO3; Cl, 2.51xl03-7.30xl03; P<0.01) and CSSH 45 (6.79x103; Cl, 4.40x103-

9.19xl03; P<0.01). Table 3-8. Most probable number of G. diazotrophicus population in field plant tissues Bacterial Population (MPN g'1 tissue) Cultivar Root Stem Leaf CG SH 27 3 20x103 5.67xl02 2.20x102

98W 2.46x103 9.07x102 2.28 xlO2

CG SH 8 1.15X103 3.04x103 1.86xl02

Nutriplus BM R 7.68x103 4.41x10s 3.72x102

2 7 5 ■; 1.55X104 6.17xl03 1.53xl02

N i l i 1.72X104 6.88xl03 6.90xl02

C S S II45 1.86xl04 6.23x10s 7.97x102

*"U Results are averages of three PCR positive replicates/cultivar. 73

12000 I

A

10000 □ tfk(A 8000

a. 2 c o 6000

a a.o Ti 4000 •c

2000 C

Root Stem Leaf Sorghum Tissue

Figure 3-8. Effect of tissue type on G. diazotrophicus population numbers in field plants A significant difference in bacterial population numbers was observed among the root, stem and leaf tissues, indicating an effect of tissue type (F=40.19, df=2, P<0.01). Among the tissues, different letters represent multiple comparisons that are significantly different. Error bars represent SE. 12000 I

^ 10000 - «i

CGSH27 98W CGSH 8 Nutriplus 275 N l l l CSSH45 BMR Sorghum Cultivar

Figure 3-9. Effect of cultivar type on G. diazotrophicus population numbers in field plants A significant difference in bacterial population numbers was observed among the cultivars, indicating the effect of cultivar type (F=9;52, df=6 , P<0.01). Among the cultivars, different letters represent multiple comparisons that are significantly different. Error bars represent SE. 75 \i. /

12000 n

_ 10000

* 8000 z • CL 2

| 6000 jj 3 ■Greenhouse Ol o □ Field £

Figure 3-10. Effect of growth environment on G. diazotrophicus population numbers A significant difference in bacterial population numbers was observed between the greenhouse and field trials among the seven cultivars (F=l 1.46, df=13, P<0.01). Within a cultivar, different letters represent multiple comparisons that are significantly different. Error bars represent SE. 76

3.3.8 Visualization of G. diazotrophicus colonization

Colonization of N il 1 and 275 by UAP5541/pRGS561 was confirmed visually through

the expression o f the dark blue precipitate resulting from the degradation of X-Gluc by /?-

glucuronidase. GUS activity in the roots o f both cultivars was observed, indicating colonization

by the gws/4-marked strain o f G. diazotrophicus. Colonization was observed one week after

inoculation; however, the presence of GUS activity was minimal. As time post-inoculation

increased, the presence o f GUS activity progressed throughout the root tissues. Control plants

did not present any GUS activity at any time. (Figure 3-11 and Figure 3-12).

3.4 Discussion

3.4.1 Colonization of sorghum by G. diazotrophicus

The endophytic nature o f G. diazotrophicus has been primarily associated with sucrose-

rich plants, such as sugarcane, Ipomoea batatas (L.) Lam (sweet potato) and Pennisetum purpureum Schumach. (Cameroon grass) (Saravanan et al., 2008). Although the bacterium has

been isolated from a number o f graminaceous plant species, previous reports have shown that G.

diazotrophicus does not naturally occur in sorghum. Dobereiner et al. (1988) were not able to

isolate G. diazotrophicus from grain and sugar cultivars of sorghum. Likewise, isolation o f the

bacterium was not achieved in samples representing six different forage grasses, sorghum, maize

and 11 weed species from sugarcane fields (Boddey et al., 1991; Reis et al., 1994). Although

sorghum may not be a natural endophytic habitat o f G. diazotrophicus, there have been several

attempts to introduce the bacterium to the plant. Colonization of S. bicolor and Sorghum vulgare

Pers. by G. diazotrophicus in conjunction with arbuscular mycorrhizal (AM) fungi was observed 77

Figure 3-11. Colonization of N lll roots by gw.v/1-marked G. diazotrophicus strain UAP5541/pRGS561 A-C, Colonization one week post-inoculation. D-F, Colonization two weeks post-inoculation. G-I, Colonization 3 weeks post-inoculation. J-L, Colonization 4 weeks post-inoculation. M-O, Colonization 5 weeks post-inoculation. C, F, I, L, O, Un-inoculated controls. * Plants were harvested every 4 days from 1 week post-inoculation 78

Figure 3-12. Colonization of 275 roots by gM&4-marked G. diazotrophicus strain UAP554l/pRGS561 A-C, Colonization one week post-inoculation. D-F, Colonization two weeks post-inoculation. G-I, Colonization 3 weeks post-inoculation. J-L, Colonization 4 weeks post-inoculation. M-O, Colonization 5 weeks post-inoculation. C, F, I, L, O, Un-inoculated controls. * Plants were harvested every 4 days from 1 week post-inoculation 79 under gnotobiotic conditions (Paula et al., 1991; Adriano-Anaya et al., 2006;

Meenakshisundaram and Santhaguru, 2009).

Colonization o f sorghum tissues by G. diazotrophicus was attained in both the greenhouse and field trials. The bacterium was detected in each of the cultivars with the proportion of plants showing bacterial presence ranging from as low as 0 % to as high as 1 0 0 % depending oh the tissue. Among the sorghum cultivars, there was a tendency of higher bacterial population numbers and higher proportion of plants showing bacterial presence in the root tissues than in the stem and leaf tissues. Variability in colonization among the roots, stems and leaves was also observed in sugarcane. Li and McRae (1992) and Reis et al. (1994) isolated G. diazotrophicus from the roots, stems and aerial parts of several Australian and Brazilian sugarcane cultivars. The initial site of infection and the pattern o f colonization of the plant interior may explain the preference' of G. diazotrophicus to specific tissue parts. James et al.

(1994) demonstrated that G. diazotrophicus in sugarcane seedlings, axenically grown in a modified plant growth medium containing sucrose, began to colonize the root surfaces by 4 days after inoculation. By 15 days after inoculation, root apoplasts and xylem vessels of sugarcane seedlings were positive for the bacterium. Localization of the bacterium in the apoplasts and xylem vessels o f inoculated sugarcane seedlings was further confirmed by Fuentes-Ramirez et al.

(1999) and Sevilla et al. (2001). James et al. (1994) suggested that G. diazotrophicus rapidly colonizes the sugarcane roots through loose cells o f the root tips and cracks at the lateral root junctions, subsequently entering the vascular system. Presence o f the bacteria in xylem vessels proposed a means o f systemic spread, via the transpiration stream, into the shoots as the seedling continues to grow. In the stem, G. diazotrophicus particularly concentrates in the nodes, since the xylem is convoluted at these points and the flow o f solutes, as well as bacteria, through the 80 xylem into the next intemode is impeded. Despite the low sugar levels, the xylem has been

suggested as a suitable location for G. diazotrophicus and other endophytic In fix in g bacteria

due to the low pC>2 that allows for nitrogenase expression (James and Olivares, 1997).

Initial entry o f G. diazotrophicus into sorghum was facilitated by the pruned-root dip method. Pruned-root dip has been shown to be the most effective method of introducing endophytic bacteria into plants (Bressan and Borges, 2004). The mechanism by which G. diazotrophicus initially penetrates the plant root is still not known. Cell wall degrading enzymes,

such as cellulase, hemicellulase and pectinase, have been implicated in overcoming the plant barrier by beneficial plant microorganisms such as Rhizobium Frank 1889 sp p F ran kia

Brunchorst 1886 spp., Azospirillum Tarrand et al. 1979 spp. and A zoarcus Reinhold-IIurek et al.

1993 spp. (Adriano-Anaya et al., 2005). In Adriano-Anaya et al.’s (2005) study, G. diazotrophicus was induced to produce several plant cell wall degrading hydrolytic enzymes when grown in sucrose as the sole carbon source. Strains PAL5 and UAP5541 were capable of producing endoglucanase (EG), endopolymethylpolygacturonase (EPMG) and endoxyloglucanase (EXG ), which were detected, in pure culture, at concentrations o f 2 x 10 ,

<2 ■ *2 1 10x10 and 5x10 units mg protein, respectively. This suggests that the penetration of roots by

G. diazotrophicu s and its subsequent mobility inside the plant may be the result of hydrolytic activity o f the bacterium. . '

^ Introduction of G. diazotrophicus to sorghum has been previously documented in a few

studies. Luna et al. (2010) studied the colonization behavior o f G. diazotrophicus in sorghum and wheat via colony counting of homogenized tissues and microscopy of organ sections. Although

Luna et al. (2010) demonstrated that colonization was achieved in the roots, stems and leaves of

sorghum and wheat, a molecular-based technique to detect G. diazotrophicus was not performed. 81

Other studies focusing on the endophytic association between sorghum and G. diazotrophicus primarily involves the use o f AM fungi in conjunction with G. diazotrophicus. Overall positive effects on sorghum after dual inoculation with AM fungi and G. diazotrophicus include: 1) increased bacterial and fungal population numbers re-isolated from the plant (Paula et al., 1991);

2) enhancement o f drought tolerance (Meenakshisundaram and Santhaguru, 2009); and 3) increased fresh and dry matter yield (Meenakshisundaram and Santhaguru, 2011). In the present work, colonization o f sorghum by G. diazotrophicus alone was achieved, as indicated by the proportion of plants showing: bacterial presence and the re-isolation o f the bacterium from inoculated tissues. Screening of the plant samples by PCR permitted a molecular-based approach to demonstrate the colonization o f sorghum with G. diazotrophicus. Colonization was further confirmed using a microbiological method via re-isolation of the bacterium from the colonized tissues. The re-growth o f the bacterium on LGIP plates was characterized based on phenotypic characteristics; however, detection of G. diazotrophicus by PCR from the re-isolation samples could provide a molecular-based method to complement the microbiological method of characterization. Although colonization of the plant was fully established, further work is needed to understand the mechanisms of entry and systemic spread of G. diazotrophicus. ;

3.4.2 Influence of plant cultivar on colonization of sorghum by G. diazotrophicus

Consistency in colonization of plant tissue by G. diazotrophicus is rarely observed even within the same plant species. Different sugarcane cultivars, whether grown in the same region or all over the world, show different population rates by the bacterium. The number o f isolated

G. diazotrophicus strains from root and stem samples, collected from traditional and newly cultivated areas in Northeast and Southeast Brazil, varied greatly among numerous sugarcane cultivars (Cavalcante and Dobereiner, 1988). Specifically in the Rio de Janeiro State, occurrence . ' . .. ' .82 of the bacterium in four different cultivars (CB45-3, NA56-79, RB73-9735 and RB73-9359) varied from -0.01x10s to ~100xl0 5 cells /g fresh weight (Reis et al., 1994). In several cultivated regions o f Mexico, the percentage o f plants from which G. diazotrophicus was isolated ranged between 0-65% from the roots and stems o f an assortment of sugarcane cultivars(Fuentes-

Ramirez et al., 1993).

: Although G. diazotrophicus is readily capable o f colonizing sorghum, both the rates of bacterial detection and bacterial population numbers varied among the different cultivars in each o f the greenhouse and field colonization trials. With the proportion of plants showing bacterial presence and bacterial population numbers ranging broadly, certain cultivars were more predisposed to colonization over others. The overall proportion o f plants showing bacterial presence and bacterial population numbers for cultivars Nlll and CSSH 45 were higher than those observed for 98W, CGSH 27 and CGSH 8 . Cultivars showing no significant difference in the proportion o f plants showing bacterial presence were clustered into two groups: group 1 consisted of N111, CSSH 45, 275, and Nutriplus BM R; while group 2 consisted of 98W, CGSH

27 and CGSH 8 .

Differences in bacterial endophytic communities between cultivars o f the same species have been previously recognized in a wide range o f plants, van Overbeek and van Elsas (2008) noted the effects of plant cultivar on plant-associated bacterial community structures among three different Solarium tuberosum L. (potato) cultivars (Achirana Inta, Desiree and Merkur).

Adams and Kloepper (2002) documented significant cultivar effects for threshold communities o f endophytic bacteria recovered from within seeds and internal tissues o f 10 different field- grown Gossypium L. spp. (cotton) cultivars. Finally, Muños-Rojas and Caballero-Mellado (2003) analyzed the population dynamics of G. diazotrophicus, in association with sugarcane, revealing 83 that cultivar M EX 57-473 was able to harbor the bacterium in greater numbers than four other sugarcane cultivars that were assessed. Not only did the sugarcane cultivar influence the persistence o f the plant-bacteria interaction, but also it could potentially clarify the discrepancies in the frequencies and bacterial numbers o f G. diazotrophicus recovered from sugarcane plants analyzed in various studies (Cavalcante and Doberiener, 1988; Fuentes-Ramirez et al., 1993;

Bueno dos Reis et al., 2000; Reis et al., 1994).

The variability observed in colonization among the different sorghum cultivars may be due to how the plant communicates and responds to the bacteria. Communication between the plant and bacteria, whether commensalistic, mutualistic, symbiotic, or pathogenic, is critical in the entire colonization process, which often begins through compounds that are exuded from the roots of plants (Bais et al., 2006). Simultaneously, bacterial recognition of these specific compounds must occur for plant-microbe associations to be compatible (Bais et al., 2006).

Chemotaxis to root exudates is one factor that strongly contributes to the competitiveness and specificity in bacterial colonization (Lugtenberg et al., 2001). Different bacterial strains have been described to be positively chemotactic to sugars, amino acids, various dicarboxylic acids and aromatic compounds (Brencic : and Winans, 2005). Plants communicate to attract microorganisms for their own ecological and evolutionary benefit; however, the intricacy of the interaction and the detailed mechanisms involved in the putative selection process is difficult to understand (Hardoim et al., 2008). Referring to the well-studied Rhizobium -plant interaction, which encompasses highly evolved species-specific communication, may give light to the selective associations of other plant-microbe systems (Bais et al., 2006). According Smith and

Goodman’s (1999) review, within a plant species, there exists cultivar-strain specificity that can be qualitatively described as either compatible or incompatible. Compatible interactions lead to 84 successful infection and formation of N2-fixing nodules in the plant, while incompatible interactions may show signs o f early infection, nodule formation does not occur. The key factor in nodule formation is the bacterial recognition of polycyclic aromatic compounds called flavonoids released by the plant and the plant detection of nodulation (Nod) factors (also known as lipo-chito-oligosaccharides) that are synthesized and excreted by the bacteria (Stougaard,

2000; Cullimore et al., 2001; Brencic and Winans, 2005). Nod factors are synthesized and exported from the bacteria by the products o f n od genes,: which are present in all Rhizobium

(Geurts et al., 2005). Each Rhizobium sp. possesses species-specific n od genes, which direct species-specific modifications of the basic Nod factor (Cullimore et al., 2001). Therefore, the specific set o f Nod factors, produced by each Rhizobium sp., plays a crucial role in host specificity (Cullimore et al., 2001). One observation of cultivar-strain specific interactions was in peas nodulated by R. legum inosarum bv. v iciae (Hogg et al., 2002). The pea cultivar Afghanistan could not be nodulated by several strains of Rhizobium that nodulate other cultivars (Hogg et al.,

2002). The resistance to certain Rhizobium sixains, was isolated as a single recessive gene, known as sym-2, in the host (Lie, 1984; Davis et al., 1988). This gene interacts with a specific nodulation gene, nodX, present only in strains that nodulate primitive pea cultivars (Davis et al.,

1988). The product of the cultivar-specific nodX gene acetylates a nodulation factor to instigate a compatible interaction with the cultivar Afghanistan (Finnin et al., 1993). Although quantitative genetic mechanisms of the host may affect the relative ability of different strains to occupy root nodules, the mode o f action that results in cultivar-strain specificity is not fully understood

(Smith and Goodman, 1999).

Beyond the scope of Rhizobium -legume interactions, various plants have evolved different responses to colonization by endophytic bacteria. Differential expression o f numerous ■■85 genes with unknown functions was observed during sugarcane-bacterial associations (Rocha et al., 2007). Vinagre et al.’s (2006) research demonstrated that the shr5 gene was differentially expressed in sugarcane aftér inoculation with specific N2-fixing bacteria. The shr5 gene encodes a signal transduction protein that is likely to be involved in the plant host defense response. The expression of shr5 decreased when exclusively inoculated with G. diazotrophicus,

Herbaspirillum seropedicae Baldani et al. 1986 emend. Baldani et al. 1996 and/or Azospirillum brasilen se Tarrand et al. 1979. In response to inoculation with pathogenic bacteria, there was no change in the expression of shr5.

Colonization may also induce different physiological responses by different cultivars.

Muftoz-Rojas and Caballero-Mellado (2003) suggested that certain physiological and metabolic changes could conceivably modify the establishment and endophytic permanence o f the microorganism in the plant. Changes in tissue water relations, concentration of sucrose and enzymatic activities are known to occur during plant growth and have the potential to influence the residing population o f G. diazotrophicus in sugarcane. A study by Funnell-IIarris et al.

(2010) examined the effect of sorghum cultivars on populations of root-associated and soil fluorescent Pseudomonas Migula 1894 spp.. Although not ^ -fixin g , populations o f

Pseudomonas spp. were differentially affected by the two sorghum: cultivars, Redlan and

RTx433. When cycling the two cultivars, numbers of Pseudomonas spp. had increased jsignificantly only in the presence o f cultivar RTx433, suggesting that changes in root populations may have been affected by plant cultivar type. The biochemical difference between the roots of

Redlan and RTx433 sorghum seedlings was the differential production of the allelopathic compound, sorgoleone. Redlan produced relatively large amounts of sorgoleone while RTx433 86 produced the lowest, which coincides with the higher populations of Pseudomonas spp. during the cycling of RTx433.

Variation in colonization capacities o f endophytic bacterial communities seems to be unpredictable, even within individuals o f the same plant species. Factors that drive the endophytic associations, from the early stages o f root colonization of young plants to the established communities seen in mature plants, are not well understood. Despite the limited research into host variation for interactions with beneficial plant-associated microbes, evidence from well-studied systems, such as • plant-pathogen and Rhizobium -legume interactions, may provide some insight. Preceding evidence has shown that root exudate patterns, including the amount and type o f compounds, gene expression during plant-bacterial associations, and plant physiological stage all may influence root colonization by microorganisms. Together, this may well suggest that the variability in colonization o f different sorghum cultivars may be attributed to factors o f similar grounds.

3.4.3 Environmental influence on colonization of sorghum by G. diazotrophicus

Not only are the mechanisms involved in plant-microbe interactions quite complex, biotic and abiotic factors that influence microbial colonization further complicate the understanding of the entire process. In the present work, the effect o f the growth environment on colonization of seven different sorghum cultivars was evaluated. Variability in the proportion of plants showing jbacterial presence and bacterial population numbers was observed among the seven cultivars that were grown under greenhouse and field conditions. When comparing the data between the two colonization trials, a higher proportion of field-grown plants than greenhouse-grown plants showed bacterial presence with higher bacterial population numbers. Differences in the 87

conditions o f the environment could have influenced plant growth and development that in turn

could have affected the ability of the endophytes to persist within the plant.

■ Functions of the rhizosphere are o f central importance for plant nutrition, health and

quality (Berg and Smalla, 2009). Plant-microorganism interactions in the rhizosphere specifically

contribute to carbon sequestration, ecosystem functioning, and nutrient cycling in various natural

ecosystems as well as in agricultural and forest systems (Singh et al., 2004). The structural and O- . . ' ... , .

functional diversity of rhizospheric bacterial communities can be influenced by a number of

factors including climate and season, grazers and animals, pesticide treatments, soil typé and

structure, and plant health and developmental stage (Lemanceau et al., 1995; Siciliano et al.,

2001; Graner et al., 2003; Jousset et al., 2006; Rasche et al., 2006). The extent of these factors

influencing the rhizospheric populations and thus affecting plant growth and development and

indirectly determining colonization is not fully understood.

Different soil types not only harbor specific populations of microorganisms, but can also

influence the microbial community structure and function of the rhizosphere (Fierer and Jackson,

2006). As investigated by Da Silva et al. (2003), soil type was the overriding determinative

factor that influenced the community structures o f Paenibacillus Ash et al. 1994 emend.

Behrendt et al. 2010 populations in the rhizosphere o f maize. Dalmastri. et al. (1999) noted a

tore prominent influence by soil type on the genetic diversity o f maize root-associated

urkholderia cepacia (Palleroni and Holmes 1981) Yabuuchi et al. 1993 populations. Chiarini et al. (1998) collected samples from the rhizosphere of maize cultivar Airone and recognized a clear soil-type effect, with microbial densities and community structures varying significantly among the five different field sites. Lastly, Marschner et al. (2001) observed variation in the rhizosphere communities o f chickpea and rape host plants grown in different soils. It is evident ' 88 that the type o f soil and its properties contribute to the numerous nuances o f microbial communities. Edaphic variables including texture, temperature, aeration, physico-chemical characteristics and pH can result in distinct microbial communities and spatial variability

(Cavigelli et al., 1995; Gelsomin et al., 1999; Carelli et al., 2000). Analyzing the distribution of microbial isolates from two uncultivated soils, Latour et al. (1996) reported a difference in populations of fluorescent pseudomonads from one uncultivated soil to the other. Differing in texture and in their natural response towards fusarium wilts, Chateaurenard soil with pH 8.05 has a suppressive response, while Dijon soil with pH 6.91 has a conducive response. These differences may not only have affected the survival of the fluorescent pseudomonads, but also its activity levels within the rhizosphere. Soil temperature also appears to have an affect on rhizospheric microbial populations. Bowers and Parke’s (1993) study evaluated the effect of soil temperature on rhizosphere colonization o f pea taproots by Pseudomonas fluorescens Migula

1895 at a constant soil-water matric potential. Higher bacterial populations at low temperatures

(16°C) and a linear decrease as temperatures increased indicated the effect o f soil temperature on colonization and taproot length. Soil influences the size and composition of the rhizosphere microflora in different ways (Wieland et al., 2001; Buyer et al., 2002). The differences in the rhizosphere could lead to differences in the growth and development o f each plant, which could either interfere or enhance colonization o f the plant.

J The environment in which plants are grown in largely determines the growth and development o f the plant. As such, differences in the environmental conditions could influence the physiological state o f the plant, which could influence bacterial community structures and colonization (Van Overbeek and van Elsas, 2008; Hallmann and Berg, 2006). Fuentes-Ramirez et al. (1999) observed that sugarcane plants grown under a high nitrogen fertilization regime as 89 opposed to low nitrogen fertilization showed reduced colonization by G. diazotrophicus. Under high nitrogen concentrations, synthesis o f sucrose by the plant is reduced. As such, the supply of nitrogen to the, plants altered its physiology by causing a decrease in sucrose needed for endophytic growth. Other fluctuating environmental factors could be the reason for a wide variation in bacterial population numbers within sugarcane plants. Mufioz-Rojas and Caballero-

Mellado (2003) recorded a range of bacterial numbers of G. diazotrophicus in sugarcane, from

• 104..to 10s CFU g'1 fresh weight. Further studies reported cell numbers ranging from 105-107 i i , CFU g fresh weight and even as low as 10 to 10 CFU g' fresh weight (Cavalcante and

Dobereiner, 1988; Reis et al., 1994). Varying environmental conditions, particularly in the amounts o f rainfall, have been proposed as the rationalization for the discrepancies in bacterial numbers o f G. diazotrophicus (Mufioz-Rojas and Caballero-Mellado, 2003).

Due to the complexity and interactive nature of numerous environmental determinants, the task of revealing the direct effect o f any one variable on any aspect of microbial activity, such as colonization, can be quite convoluted (Bowers and Parke, 1993). The relative importance o f the rhizosphere, plant growth and development, and plant physiology in the colonization process is still difficult to determine as these factors may either act independently or together.

Root colonization by bacteria largely depends on the occurrence of an effective plant root- bacterium interaction (Rosenbleuth and Martinez-Romero, 2006). The probability of this interaction occurring initially depends on the abundance, diversity, physiological status and distribution of the endophytes (Berg and Smalla, 2009). Thus, the effect of the environment, whether indirect or direct, on the dynamics of endophytic colonization warrants further research. 90

3.5 Conclusion

In the present work, sorghum was used as a model host for G. diazotrophicus to look

further into the complexities of the interactions between plants and endophytic microorganisms.

Colonization o f sorghum tissues by G. diazotrophicus was attained under both greenhouse and field conditions. The bacterium was detected in each of the cultivars with the proportion o f plants showing bacterial presence ranging from as low as 0% to. as high as 100% depending on the tissue. As observed in sugarcane, the variability was mostly noted among the different tissues, with detection in the roots, stems and leaves o f the sorghum plant. Among the seven sorghum cultivars, there was a tendency o f higher population numbers and higher proportion o f plants

showing bacterial presence in the root tissues than in the stem and leaf tissues. The pruned-root dip method o f inoculation, thus making the roots as the initial site of entry, may be the plausible explanation for the observed colonization patterns in the root tissues. In sugarcane, the initial site o f entry and the location o f the bacterium have been narrowed down to the root tissues and the xylem vessels, respectively. However, the exact mechanism of entry has yet to be determined, though there has been a strong indication of the involvement of cell wall degrading hydrolytic enzymes. -

Among the different cultivars, the proportion of plants showing bacterial presence and bacterial population numbers varied significantly in each o f the greenhouse and field colonization trials. Certain cultivars were more susceptible to colonization than were others. The overall proportion o f plants showing bacterial presence and bacterial population numbers for cultivars N 111 and CSSH 45 were higher than those observed for 98W, CGSH 27 and CGSH 8.

Consistency in colonization of plant tissue by G. diazotrophicus is rarely observed even within the same plant species. Different population rates by the bacterium have been observed within 91

the same , plant species, indicating the possibility of a plant cultivar effect on microbial

colonization. Potato, sugarcane and cotton cultivars all have shown variation in microbial

colonization rates and population numbers amongst its own species. Although plant cultivar

seems to specify the success of bacterial colonization, the exact mechanisms involved in the

specificity has not yet been fully disclosed. Specific communication, whether by physiological,

chemical or genetic mechanisms, between the plant and bacterium may play a role in the

specificity o f the endophytic association. Chemotaxis to root exudates or differential expression

of certain genes involved in the plant-endophyte establishment could provide the basis for

explaining the specificity observed between G. diazotrophicus and the different sorghum cultivars/'-':’... .V--/'■

Additional peripheral factors that interactively influence microbial colonization further complicates the understanding o f the processes involved. Variability in the proportion o f plants showing bacterial presence and bacterial population numbers was observed among the seven cultivars that were grown under greenhouse and field conditions. With significant differences mainly observed in the roots and stems, a higher proportion of plants showing bacterial presence and higher bacterial population numbers were observed under field conditions than greenhouse conditions. Rationalization for the observed variability between the two colonization trials may be the different environmental factors that influence growth. Climate and season, soil type and structure and plant health and developmental stage are thought to influence the community and structure o f the rhizosphere. Colonization o f roots by introduced microorganisms has been tied to several factors one o f which includes general rhizosphere dynamics. It is very possible that the rhizosphere dynamics in the field could have provided a beneficial environment for both sorghum and G. diazotrophicus to thrive well together. The environmental conditions in the field which could have influenced the colonization success of G. diazotrophicus. However, the extent to which these elements sway microbial populations and affect the rhizosphere is not yet understood; ■ -

The variation observed in the proportion of plants showing bacterial presence and bacterial population numbers may be attributed to a number of interactive factors based on the environment, the bacterium and the plant itself. Determining the appropriate conditions for bacterial colonization could result in increasing the numbers o f the bacterial population residing within the plant. Increased bacterial colonization would be ideal in the efficient application of endophytic bacteria, such as G. diazotrophicus, for BN F. G. diazotrophicus is predominantly known as an endophyte that prefers to colonize sugar-rich plant hosts. Although colonization of sorghum by G. diazotrophicus was achieved, bacterial detection was more pronounced in the sweet cultivars than the grain cultivars. Whether sugar content in the sweet sorghum types 93

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CHAPTER 4. Endophytic association between Gluconacetobacter diazotrophicus and sorghum: Colonization factors and inoculation effects

4.1 Introduction

Sucrose, the principal carbohydrate translocated from source to sink tissues, is a fundamental end product of photosynthesis in most plants (Stitt et al., 1987). During the day, sucrose accumulates in the leaves preceding dissemination to the rest of the plant (Lunn and

Hatch, 1995). At night, the stored sucrose is remobilized to maintain the export of sucrose to sink tissues and to support respiration in the leaves (Lunn and Hatch, 1995). Saccharum L. spp.

(sugarcane) is often a suitable candidate for the study of sugar storage since sucrose accounts for

50% o f the total dry matter of the stalk (Glasziou and Gayler, 1972). Sucrose contents o f about

20% fresh weight are attained in mature intemodal tissues, while young intemodes from the top portion of the stalk often contain much lower levels of sucrose, glucose and fructose - each roughly at 5% fresh weight (Glasziou and Gayler, 1972). Sucrose is synthesized in the photosynthetic tissue of the leaf mesophyll, from there it is loaded into the phloem then moves out of the leaves and towards the sink tissues, which include developing shoot and root apices and storage organs (Lalonde et al., 2003; Rae et al., 2005). Sucrose accumulation often begins in the intemodes during the stage o f stalk elongation and continues long after elongation ceases, indicating the maturation o f sugarcane (Rose and Botha, 2000).

Sorghum bicolor (L.) Moench (sorghum) is a tropical self-pollinating plant that is native to Africa, but is now produced throughout the tropical, semi-tropical and arid regions o f the world (Dicko et al., 2006). Sorghum can be classified into four groups based on its intended purpose: 1) grain sorghum, which is primarily used as food in tropical areas and as raw materials for alcoholic beverages, sweets and glucose; 2) sweet sorghum, which is used as material for sweetener syrups; 3) broom sorghum, which is used as material to make brooms; and 4) grass sorghum, which is mainly grown for feed and forage use. All types of sorghum contain sugar in the stalks (Bian et a l, 2006). Higher sugar contents are typically found in sweet sorghum, which

is currently processed for production o f table syrups and livestock feed (Reddy et al., 2005).

Sweet sorghum cultivars have been revealed to accumulate considerable amounts o f sugars in their stem parenchyma, with sucrose (92%) as the predominant type o f sugar trailed by glucose

(4.5%) (McBee and Miller, 1982; Bian et al., 2006). Sugar content in the juice extracted from sweet sorghum can vary from 16-23% B rix and the quality of sugar prepared from sweet sorghum is comparable to that of sugarcane (Reddy et al., 2005; Channappagoudar et al., 2007).

The pattern of sucrose storage and accumulation in sorghum is similar to that observed in

sugarcane. Specifically, the sweet cultivars o f sorghum accumulate considerable amounts of sucrose in their stem parenchyma (Hoffmann-Thoma et al., 1996). Sucrose is the key storage carbohydrate in sorghum stems, while starch is the major storage carbohydrate in the seeds

(Tarpley et al., 1994). Carbohydrate reserves and sucrose accumulation can vary among cultivars and are influenced by a multitude of factors such as stem elongation, leaf number, and the number o f nodes and intemodes formed (Hoffmann-Thoma et al., 1996). The amount o f solutes retained in the stem tissue may additionally be affected by the competition between grain and stem, the two sink organs, during plant development (Hoffmann-Thoma et al., 1996). In one sweet sorghum cultivar, Rio, the onset of sucrose accumulation coincided with the extension of the panicle into the sheath of the flag leaf (Lingle, 1987). In conjunction with plant developmental factors, sorghum carbohydrate contents are also influenced by temperature, time o f day, maturity, cultivar, stem section, spacing and fertilization (McBee and Miller, 1982;

Amaducci et al., 2004). Sorghum cultivars experienced diurnal fluctuations of carbohydrate contents with sucrose varying widely in the upper leaves (McBee and Miller, 1982). Sugars are 102 first synthesized during seed germination and then transported to sites where they are required for growth (Amuti and Pollard, 1977). Soluble sugars seem to play an important role in the osmotic regulation of cells and the expression of certain genes during germination (Reynolds and

Smith, 1995; Yu et al., 1996).

Sucrose is not only a key component for plant growth and development, but also a primary carbon source for certain microbial endophytes. Gluconacetobacter diazotrophicus

(Gillis et al. 1989) Yamada etal. 1998 is believed to utilize sucrose in its endophytic association with sugarcane (Tejera et al., 2004). Under laboratory conditions, G. diaztrophicus is capable of pellicle formation in N-free semisolid medium with 10% added sucrose and can grow in media containing up to 30% sucrose (Cavalcante and Dobereiner, 1988; Reis et al., 1994). Growth of the bacterium is observed at pH values ranging from 2.5 to 7.0 (Stephan et al., 1991). Thus, G. diazotrophicus seems well adapted to sugarcane tissues as it shows optimum growth with 10% sucrose and pH 5.5 (Cavalcante and Dobereiner, 1988; Reis et al., 1994). Since the bacterium can subsist under different sucrose concentrations, it is possible that sugar is an important factor in colonization. .

Successful colonization of plants with N2-fixing bacterial endophytes, such as G. diazotrophicus, could result in plant growth promotion, mediated by the production and excretion of phytohormones (Rothballer et al., 2009). These bacteria can be categorized as plant growth-promoting bacteria (PGPB), which are defined as “free-living soil, rhizosphere, rhizoplane, endophytic and phyllosphere bacteria that under certain conditions are beneficial for plants” (Bashan and de Bashan, 2005). Primarily involved in root growth promotion are indole acetic acids (IAA) and other auxins, as demonstrated by Loper and Schroth (1986), Barzani and

Friedman (1999), and Patten and Glick (2002). The production of phytohormonal substances such as auxins by different PGPB has been proposed as one of the mechanisms to explain plant growth-promotion. It has been previously 103

demonstrated that G. diazotrophicus is capable of producing the phytohormone, indole-3-acetic acid, and, thus, there IS potential for plant growth promotion after inoculation of plants with thebacterium(Fuentes-

Ramirez et al., 1993).

Sorghum cultivars showing a gradient of sucrose concentrations were inoculated with G. diazotrophicus in Chapter 3. This chapter will establish the effect of sucrose on colonization and determine the significance of sugars in the endophytic interaction between the plant host and the bacterium. It is hypothesized that there will be an increased bacterial presence in sorghum plants with higher sucrose concentrations. Additionally, the relationship between promotion of plant growth, if present, and inoculation with G. diazotrophicus, will be investigated.

4.2 Materials & Methods

4.2.1 Sugar analysis in sorghum

Sugar analysis was performed on all seven sorghum cultivars (refer to Chapter 3, Section

3.2.1 for details of the cultivars). Three plants o f each sorghum cultivar were analyzed for sucrose concentration via gas chromatography-flame ionization detection (GC-FID). All plants were separated into roots, stems and leaves, which were freeze-dried and individually ground into powder using a Wiley mill fixed with a No. 20 mesh screen. For the analysis of sucrose, 200 mg o f the powdered plant samples were collected for extraction. All samples were subjected to

Jtwo rounds of extraction by vortexing with 5 mL of milli-Q H2O for 20 s then centrifuging at

2500 rpm for 15 min at 24°C in each round. Combining both the 5 mL aliquots, 100 pL of the extracts were transferred to 2 mL target DP™ glass reaction vials for centrifugal evaporation using the Savant SVC 100H SpeedVac Concentrator. Prior to analysis via GC-FID, the dried sugar extracts were first subjected to methoximation derivatization for 90 min at 30°C with the ' . . 104 methoxyamine HC1 (MOX™) reagent (ThermoFisher Scientific). Subsequently, each sample was then submitted to trimethylsilyl derivatization for 30 min at 37°C with jV-methyl-JV-

(trimethylsilyl) trifluoroacetamide (MSTFA) (Sigma-Aldrich®). Two pL samples were

introduced, using helium as the carrier gas, by split-less injection with an injection temperature of 280°C on the Hewlett Packard 5890 II Plus GC equipped with a DB-5 capillary column (30 m x 0.25 mm, 0.25 pm thickness, J& W Scientific). The column temperature gradient was programmed at an initial 70°C, followed by a ramp at 5°C min'1 up to 330°C, held for six min.

Sucrose standards of known concentrations were analyzed to generate a calibration curve for the quantitative calculation o f sucrose from each sample based on the relative peak area distinguished from the gas chromatogram. ' ^

4.2.2 Agronomic measurements of sorghum in the field

Agronomic measurements for the sweet cultivar N i l 1 and grain cultivar 275 (chosen for their relatively high proportion of plants showing bacterial presence) were collected to determine the effect o f colonization on plant growth.

For each cultivar, 72 seeds were germinated and seedlings were then inoculated and grown as described in Chapter 3, Section 3.2.3 and Section 3.2.4. Bacterial suspensions o f PAL5 and MAd3A were cultured as described in Chapter 3, Section 3.2.2. To determine whether plant growth promotion is solely due to N2 fixation or due to the presence o f plant growth regulators, plants were divided into two inoculation treatments: treatment with PAL5 and treatment with

MAd3A (refer to Chapter 2, Section 2.2.1 for details of the strains). Control plants were inoculated with 0.9% NaCl solution without the bacterial suspension. Upon inoculation, seedlings were transferred into individual pots containing 1:3 (v:v) sand:growing mix #1 (refer to

Chapter 3, Table 3-2 for components) and left in the greenhouse to acclimatize and ensure 105 inoculation. After 1 week in the greenhouse, seedlings were then transplanted into the field. A total o f 60 plants per cultivar (20 plants inoculated with PAL5, 20 plants inoculated with

MAd3 A, 20 un-inoculated plants) were grown in the field for 60 days.

Sixty days after inoculation, plant height was evaluated by measuring the distance from the soil surface to the tip o f the sorghum head. Plant weight was assessed by collecting the stems and leaves in paper bags for air-drying at 28°C in a warm storage room for 2 weeks, prior to weighing. Root samples were not weighed since there was too much inconsistency in collecting all the tissues and removing all the adhering soil.

4.2.3 Statistical analyses

All statistical analyses were performed with SPSS 18.0 (SPSS Inc., Chicago, Illinois,

USA). Two-factor univariate analysis of variance was conducted to compare sucrose concentration among the different cultivars and tissues. To determine the correlation between sucrose concentration and the proportion o f plants showing bacterial presence and between sucrose concentration and population numbers, Pearson’s correlation analyses were performed.

Finally, one-factor analysis o f variance was used to compare plant height and plant weight among the different inoculation treatments. All multiple comparisons were evaluated with the

Least Significant Difference post-hoc test. All analysis of variance statistical tests were performed at the P=0.05 level, while all Pearson’s correlation bivariate tests were performed at the P=0.01 level. 106

4.3 Results

4.3.1 Sucrose concentration in sorghum

Sucrose was detected and analyzed in the roots, stems and leaves o f all seven sorghum cultivars via GC-FID. Among the cultivars, sucrose concentrations ranged from 0.02-44.26 mg'1 g dry weight in the roots, 92.68-172.70 mg'1 g dry weight in the stems, 0.14-10.17 mg'1 g dry weight in the leaves. Plant sucrose concentration significantly varied among the different sorghum cultivars (F=6.684, df=6, P=<0.001) (Figure 4-1). The sweet cultivar Ni l 1 had higher sucrose concentrations than the grain cultivars CGSH 27 (35.88; Cl, 17.78-53.99; P<0.01; where

Cl represents confidence interval), 98W (35.29; Cl, 17.19-53.40; PO.01), and CGSH 8 (28.11;

Cl, 10.00-46.21; P<0.01). The sweet cultivar CSSH 45 had higher sucrose concentrations than four of the five grain cultivars (P<0.04).

Significant differences in sucrose concentrations were also noticed among the roots, stems and leaves (F=293.55, df=2, P<0.01) (Figure 4-2). The stems had the highest sucrose concentrations compared to the roots (116.95; Cl, 105.10-128.80; P<0.01) and leaves (128.69;

Cl, 116.84-140.55; PO.01).

4.3.2 Relationship between sorghum sucrose concentration and colonization

To determine whether a relationship between colonization and the concentration of sucrose in plants exist, Pearson’s correlation bivariate tests were conducted using the colonization trial data from Chapter 3 and the sugar analysis data from the current chapter. For the greenhouse trial, a positive linear relationship was observed between the proportion o f plants showing bacterial presence and plant sucrose concentration (r=0.95, P<0.01, Figure 4-3A) as well as between bacterial population numbers and sucrose concentration (r=0.82, PO .01, Figure

4-3B). Similarly, a positive linear relationship was observed between the proportion o f plants 107

CGSH27 98W CGSH 8 Nutriplus 275 N l l l CSSH45 ' BMR Sorghum Cuitivar

Figure 4-1. Sucrose concentration in sorghum cultivars Plant sucrose concentrations significantly varied among the different cultivars (F=6.68, df=6, P=<0.01). Among the cultivars, different letters represent multiple comparisons that are significantly different. Error bars represent SE. 108

160 -

B 140 *

120 *

5

i A i

Root Stem Leaf Sorghum Tissue

Plant sucrose concentrations significantly varied among the roots, stems and leaves (F=293.55,

\

!

t I' 109

Figure 4-3. Relationship between sucrose and bacterial presence and bacterial population numbers in greenhouse sorghum plants A, A positive linear correlation was observed between the proportion of plants showing bacterial presence and plant sucrose concentration (r=0.95, P<0.01). B, A positive linear correlation was observed between bacterial population numbers and plant sucrose concentration (r=0.81, P<0.01). showing bacterial presence and sucrose (r=0.89, P<0.01, Figure 4-4A) and bacterial population numbers and sucrose (r=0.96, P<0.01, Figure 4-4B) for the field trial. Both trials indicated a strong correlation between sucrose and colonization. As sucrose concentrations increased, the proportion o f plants showing bacterial presence as well as bacterial population numbers correspondingly increased. Fittingly, greater sucrose concentrations were observed in the sweet cultivars Nlll and CSSH 45, which were more susceptible to colonization, with a greater proportion o f plants showing bacterial presence and larger bacterial population numbers than those observed for the grain cultivars.

4.3.3 Effect of G. diazotrophicus on plant growth

Agronomic measurements were collected for both N il 1 and 275 to establish any plant growth promotion effects as a result of colonization by G. diazotrophicus. Between all three treatments, the sweet cultivar Nlll did not show significant differences in plant height'(F=l.75, df=2, P=0 18, Figure 4-5A) and weight (F=0.14, df=2, P=0.87,Figure 4-5B). Similarly, the grain cultivar 275 did not show significant differences in plant height (F=0.49,'df=2, P=0.62, Figure 4-

6A) and weight (F=1.46, df=2, P=0.24, Figure 4-6B).

Agronomic measurements were not compared between the two cultivars since each cultivar is characteristically different in height and weight.

4.4 Discussion

4.4.1 Influence of sugar on colonization of sorghum by G. diazotrophicus

As anticipatedj both the sweet cultivars used in the present work (N111 and CSSH 45) contained greater amounts of sucrose in the roots, stems and leaves than those measured for the grain cultivars. The highest amounts of sucrose were observed in the stalks o f N111 and CSSH I l l

Figure 4-4. Relationship between sucrose and bacterial presence and bacterial population numbers in field sorghum plants A, A positive linear correlation was observed between the proportion o f plants showing bacterial presence and plant sucrose concentration (r=0.89, P<0.01). 2?, A positive linear correlation was i ■ observed between bacterial population numbers and plant sucrose concentration (r=0.96, PO .O l). 112

225 i

MAd3A Control Treatment A

180 i

PAL5 MAd3A Control Treatment

Figure 4-5. Height and dry weight of cultivar N il 1.two months post-inoculation with G. diazotrophicus A, No significant difference in height was observed between plants colonized with PAL5, plants colonized with MAd3A and un-inoculated control plants (F = l.75, df=2, P=0.18). B, No significant difference in dry weight was observed between plants colonized with PAL5, plants colonized with MAd3A and un-inoculated control plants (F=0.14, df=2, P=0.87). 113

140

PAL5 MAd3A Control Treatment A

50 1

Figure 4-6. Height and dry weight of cultivar 275 two months post-inoculation with G. diazotrophicus A, No significant difference was observed between plants colonized with PAL5, plants colonized with MAd3A and un-inoculated control plants (F=0.49, df=2, P=0.62). B, No significant difference in weight was observed between plants colonized with PAL5, plants colonized with MAd3 A and un-inoculated control plants (F= 1.46, df=2,Pr=0.24). 45. Additionally, the grain cultivars varied in sucrose concentration, with CGSH 27 having the lowest sucrose concentration to 275 having the highest. The primary sugars present in sorghum have been determined to be fructose, glucose, raffinose, sucrose and maltose (Anglani, 1998).

Sweet sorghum stalks are composed o f pith and bark, which are present in roughly equivalent amounts on a dry matter basis: on a fresh matter basis, pith can account for 65% o f the stem

(Billa et al., 1997). Sucrose and glucose are primarily found in the pith (71% of dry weight), which exhibits twice the amount o f sugars in relation to the bark (34.6% o f dry weight) (Billa et al.,1997). Sucrose is believed to be the key sugar in sorghum. Subramanianetal. (1980) reported that sucrose could comprise between 68.7-82.7% o f the total soluble sugars in various sorghum cultivars. Boyer and Liu (1983) also reported sucrose to be the predominant sugar and that sugary sorghums are known to contain twice the quantity o f sugars as grain sorghums.

Though plenty of studies investigated the association between G. diazotrophicus and sugar-rich plants, colonization studies comparing low- and high-sugar plants are lacking. One study looked at the colonization o f various Canadian maize cultivars, ranging in sucrose concentrations, by G. diazotrophicus (Tian et al., 2009). It was confirmed that the sweet maize cultivars had higher efficiency o f bacterial colonization than the grain maize cultivars, indicating that sucrose concentration has a positive effect on colonization and bacterial growth (Tian et al.,

2009). In the present study, sweet sorghum types generally demonstrated greater colonization success than grain sorghum types, which could be attributed to the sucrose concentrations of the sorghum cultivars.

Colonization success, in terms o f bacterial presence and bacterial population numbers, was correlated to the sucrose concentration o f the plant. Higher plant sucrose concentrations were consistently associated with higher proportion of plants showing bacterial presence and 115 greater bacterial population numbers. This could indicate that colonization may be dependent on the sucrose concentration o f the plant. The correlation observed confirms the tendency o f G. diazotrophicus to associate with plant species that are rich in sugars. The endophytic association between sugarcane and G. diazotrophicus is the primary exemplar of the bacterium’s preference for high sucrose environments (Cavalcante and Dobereiner, 1988). G. diazotrophicus is capable o f growing in media ranging in sucrose from 10-30% (Reis et al., 1994; Cavalcante and

Dobereiner, 1988). The bacterium can subsist on different carbon sources, such as glucose, fructose, galactose and glycerol, but sucrose seems to influence various metabolic functions and characterizations of G. diazotrophicus. Tejera et al.’s (2004) study demonstrated that glucose dehydrogenase and alcohol dehydrogenase activities within G. diazotrophicus significantly increased when sucrose was supplied to the growth medium. It was suggested that the heightened enzyme activities indicated a high osmotic potential and the possible activation of fermentative pathways within the bacterium that may coincide with its ability to localize within the sugarcane stem. Sugarcane stem juice has shown total soluble sugar concentrations of 15% and 22% in apoplastic and symplastic sap, respectively. As such, sucrose is thought to be the main carbon source used by G. diazotrophicus in its endophytic association with sugarcane.

For N2 fixation activity, G. diazotrophicus has been shown to have osmotolerance to 20%

-30% sucrose (Cavalcante and Dobereiner, 1988). However, the mechanism by which the bacterium tolerates high levels of sucrose is unknown. At such high sucrose concentrations, , levan is thought to accumulate and act as an omsoprotectant (Velazquez-Hemandez et al., 2011).

Levan, a linear fructose polymer, is one o f many bacterial exopolysaccharides that have been indicated in adherence and colonization o f inert surfaces, biofilm formation, protection against abiotic factors, resistance to extreme temperatures and osmotic conditions, nutrient acquisition 116 and as adherence or virulence factors (Danese et al., 2000; Laue et al., 2006; Russo et al., 2006;

Lemer et al., 2009; Dunne, 2002). Levansucrase, an extracellular enzyme with saccharolytic activity, is used by G. diazotrophicus to produce levan and break sucrose down into glucose and fructose (Alvarez and Martinez-Drets, 1995; Velâzquez-Hernandez et al., 2011).

Correspondingly, Velâzquez-Hemândez et al. (2011) showed that disruption of the IsdA gene in

PAL5, which codes for levansucrase, altered the bacterium’s osmotolerance to sucrose as well as its ability to tolerate high NaCl concentrations and dessication. The consequential effects of disrupting the IsdA gene in G. diazotrophicus suggested that levansucrase might partake in the adaptation of the bacterium to its high sugar environment.

Sugary sorghums may provide a selective environment for certain bacterial endophytes to grow in the absence of competition from other microorganisms. High sucrose concentrations can lead to osmotic stress and in turn suppress growth of microorganisms that are incompatible to the sucrose solute (Attfield and Kletsas, 2000; Csonka, 1989). Intrinsically, the prevalence of G. diazotrophicus in sugary plants can be attributed to the bacterium’s osmotolerance to elevated amounts o f sucrose. This osmotic compatibility may explain the observed positive correlation between sucrose concentration and the proportion of plants showing bacterial presence and bacterial population numbers among the different sorghum cultivars.

4.4.2 Plant growth promotion by G. diazotrophicus

For the benefit of their plant hosts, bacteria that fix nitrogen, such as G. diazotrophicus, have the proficiency to use alternative plant growth-promotion mechanisms, such as phytohormone production, phosphate solubilization, siderophore production and biological control (Mehnaz, 2011). Riggs et al.’s (2001) greenhouse and field trials collectively demonstrated the positive effect of G. diazotrophicus on maize productivity. For certain maize cultivars, .inoculation with PAL5 showed increases in dry weight in the greenhouse and yield increases in the field (Riggs et al., 2001). The effects o f plant growth-promotion by G. diazotrophicus have been more extensively noted in its association with sugarcane. Muftoz-Rojas and Caballero-Mellado (2003) observed a 26% increase in plant dry weight of micropropagated sugarcane plants in the greenhouse, Suman et al. (2005) measured an increase in plant biomass o f almost 19-50% in a pot trial, and finally Govindarajan et al. (2006) saw a 13-16% yield increase in field trials. In this study, plant growth-promotion was not observed with N 111 and

275. In comparison to the un-inoculated controls, treatment of both sorghum cultivars with PAL5 and MAD3 A resulted in no difference in plant height and weight measurements.

The performance o f PGPB to colonize the plant and induce plant growth promotion can be influenced by certain factors including the environment and the plant cultivar. Environmental factors, such as soil hydric stress and seasonal changes, can contribute to the efficiency o f PGPB

(Mehnaz, 2011). In Oliveira et al.’s (2006) study, the effects of soil type, sugarcane cultivars, and nitrogen rates, was observed on the productivity of sugarcane when inoculated with N2- fixing bacteria. Improved growth-promoting effects were observed for the inoculants in soils with lower and medium fertility and without nitrogen fertilizers compared to those in soils with higher fertility. Moutia et al.’s (2010) study demonstrated that plant cultivar and drought stress can influence plant growth promotion o f sugarcane by Azospirillum Tarrand et al. 1979

(Approved Lists 1980) emend. Falk et al. 1985 spp.. Two sugarcane cultivars R570 and

M l 176/77 adapted to different agro-climatic zones were inoculated with Azospirllum sp. with and without stress. When subjected to drought stress, cultivar R570 responded negatively, while

M l 176/77 responded positively with 15% improved shoot growth and 75% more root dry mass. 118

Microbial competition could also be a factor influencing colonization by PGPB and thus the level o f plant growth promotion induced in the plant. PGPB can be used independently or as a mixture for inoculating plants in pots or fields (Mehnaz, 2011). Whether a mixture of bacterial isolates or inoculation with a single bacterial species is used, the effects of plant growth- promotion can vary. Oliveira et al.’s (2002) study focused on the effect o f inoculating micropropagated sugarcane with five different strains of Infixing bacteria (G. diazotrophicus,

Herbaspirillum seropedicae Baldani et al. 1986 emend. Baldani et al. 1996, Herbaspirilium rubrisubalbicans (Christopher and Edgerton 1930) Baldani et al. 1996, Azospirillum amazonense

Magalhaes et al. 1984 and Burkholderia Yabuuchi et al. 1993 emend. Gillis et al. 1995 spp.) in various combinations. During the acclimatization period, total endophytic population and plant survival was influenced negatively after inoculation with all five N2-fixing strains. Potentially due to the competition for nutrients between plant host and an endophytic N2-fixing bacterial community, sugarcane seedlings inoculated with fewer strains (inoculated with either G. diazotrophicus or Herbaspirillum spp.) showed significantly increased dry matter than those inoculated with all five strains. Overall, a maximum rise o f 39% in total biomass was documented in the inoculated plants over the uninoculated controls, emphasizing the importance o f strain selection and inoculum combination for obtaining higher performance in the plant.

The abilities o f PGPB can vary due to the influence o f extraneous variables.

Environmental stresses and microbial competition could have adverse effects on the bacterial population numbers for colonization. Lower population numbers could in turn account for the lack o f plant growth-promotion observed in this study. The use of PGPB either discretely or as a mixture could be another factor involved in shaping the bacterial population numbers o f the plant-microbe relationship. Despite the fact that G. diazotrophicus strains PAL5 and MAd3A were used independently, the competition from prospective bacterial isolates in the field could

possibly explain the lack o f plant growth-promotion observed with cultivars 275 and N 111.

4.5 Conclusion

Both the sweet cultivars NI 1 l and CSSH 45 contained greater amounts o f sucrose in the

roots, stems and leaves than those of the grain cultivars. As expected, the stalks of NI 11 and

CSSH 45 contained the highest amounts o f sucrose. O f the grain cultivars, CGSH 27 had the

lowest sucrose concentration while 275 had the highest. The proportion of sweet sorghum plants

showing bacterial presence was notably higher than the proportion of grain sorghum plants,

which could be attributed to the sucrose concentrations o f the cultivars. A positive linear

correlation was observed between sucrose measurements and the proportion of plants showing

bacterial presence. This indicates that colonization of the plant, in terms o f bacterial presence and

bacterial population numbers, is dependent on the sucrose concentration of the plant. Higher

plant sucrose concentrations were consistently associated with higher proportion o f plants

showing bacterial presence and greater bacterial population numbers. Sucrose is believed to be

the main source of carbon for G. diazotrophicus in its endophytic association with sugarcane.

Sweet sorghum cultivars are comparable to sugarcane in their storage and accumulation of

sucrose, which is the predominant type o f sugar in both crops. Additionally, a high sucrose

Environment may be selective, due to osmotic stress, for certain bacterial endophytes to grow in the absence o f competition from other microorganisms. The osmotolerance to elevated amounts

of sucrose could explain the prevalence of G. diazotrophicus in sugary plants. As such, the

adaptation o f G. diazotrophicus to sugar-rich environments mayexplain the correlation . . .. ■ ■ . 120

observed. Sucrose appears to be a key factor for G. diazotrophicus, which could be used as a

stipulation in determining potential endophytic associations.

Alternative to biological nitrogen fixation (BNF), In fix in g endophytes have been

capable o f using mechanisms for plant growth-promotion, including phytohormone production,

phosphate solubilization, siderophore production and biological control. There has been

indication that G. diazotrophicus can induce plant growth-promotion in its hosts, particularly

sugarcane. However, plant growth-promotion was not observed in the present work with the

sorghum cultivars N111 and 275. A number of factors that influence bacterial colonization and

| bacterial population numbers, such as the conditions o f the environment, plant cultivar, and j microbial competition, could very well explain for the lack o f growth-promotion observed. The

1 factors involved in the promotion of plant growth are numerous such that speculations as to why j plant growth promotion was not observed necessitate further investigation.

The endophytic association between plants and microorganisms is a multifaceted process

in nature. Due to the various elements involved, it is still difficult to fully comprehend the

mechanisms behind the relationship between plants and endophytic diazotrophs. Plant sucrose

; concentration is only one aspect involved in the interaction between G. diazotrophicus and

sorghum. To folly benefit from bacterial endophytes and potentially reach BNF in non-legumes,

ongoing research must continue to understand further the precise mechanisms and factors that

jare involved. The following and final chapter will discuss recent advances and future work in

BNF with non-leguminous plants. 121

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CHAPTER 5. Conclusion

5.1 Colonization of sorghum by Gluconacetobacter diazotrophicus

In this study, colonization of different Sorghum bicolor (L.) Moench (sorghum) cultivars by Gluconacetobacter diazotrophicus (Gillis et al. 1989) Yamada et al. 1998 and the effects of cultivar type (sweet and grain) and growth conditions (greenhouse and field) were investigated.

It was concluded that plant sucrose and plant growth conditions both influenced the colonization o f sorghum by G. diazotrophicus. The effect o f sucrose was indicated by: 1) a higher proportion o f sweet sorghum cultivars showing bacterial presence than grain sorghum cultivars; 2 ) bacterial population numbers were higher within plant tissues o f the sweet cultivars than the grain cultivars; and 3) a strong positive correlation between sucrose concentration and the proportion o f plants showing bacterial presence. The inclination of G. diazotrophicus to associate with sugar-rich plant species may explain the correlation observed. G. diazotrophicus is a N2-fixing endophyte that was originally isolated from Saccharum (L.) spp. (sugarcane), a plant species that is recognized for its sugar storage since sucrose accounts for 50% of the total dry matter o f the stalk (Glasziou and Gayler, 1972). Like sugarcane, sweet cultivars of sorghum demonstrate a similar pattern in its storage and accumulation of sucrose, with certain cultivars accumulating considerable amounts of sucrose (McBee and Miller, 1982; Bian et al., 2006) Sucrose is believed to be the main source of carbon for G. diazotrophicus in its endophytic association with sugarcane. As such, the plant host supplies the necessary provisions for the bacterial endophyte to subsist. Additionally, high sucrose levels could lead to osmotic stress for certain microorganisms suggesting that the osmotolerance to elevated amounts of sucrose could explain the prevalence o f G. diazotrophicus in sugary plants. Sucrose appears to be a key factor for G. , !27

diazotrophicus, which could be used as a stipulation in determining potential endophytic

associations.

The effect o f the growth environment was indicated by the presence of G. diazotrophicus

observed in a higher proportion of field-grown plants than greenhouse-grown plants and,

correspondingly, field-grown plants showing higher bacterial population numbers. Colonization

by introduced microorganisms has been tied to several factors one o f which includes general

rhizosphere dynamics (van Veen et al., 1997). It is possible that the rhizosphere dynamics in the

field could have provided the necessary conditions for the endophytic association between

sorghum and G. diazotrophicus. The environmental conditions in the field could have provided

the appropriate provisions for healthier growth o f the sorghum plant itself, which could have

influenced bacterial colonization. However, the extent to which these elements sway microbial

populations and affect the rhizosphere is not fully understood.

The association between endophytic N2-fixing bacteria and sugarcane has been

investigated either under greenhouse or field growth conditions (Munoz-Rojas and Gaballero-

Mellado, 2003; Govindarajan et al., 2006; Oliveira et al., 2006). Most studies involving

sugarcane, however, have not provided a comparison between the two growth environments. A

recent study by Mehnaz et al. (2010) investigated the growth promoting effects of bacterial

isolates from Z eam ays L. (maize) under both greenhouse and field conditions. Although it was jobserved that different bacterial isolates had different plant growth promotion effects between

the two growth environments, colonization success was not considered. Further research

providing comparisons o f different growth conditions may be beneficial to comprehending the

processes involved in the endophytic interactions and relationships. ■ '128

In contrast to studies involving sugarcane and G. diazotrophicus, plant growth promotion

was not observed in this study with the sorghum cultivars inoculated with the bacterium. Plant

height and weight among the treated plants were not significantly different than those that were

not treated with G. diazotrophicus. Several possibilities exist as to why plant growth promotion

was not observed in this study. High nitrogen content in the soil could have reduced colonization

o f sorghum, resulting in low numbers of G. diazotrophicus. Several studies have shown a

decrease in colonization by G. diazotrophicus in sugarcane after the application of high amounts

o f nitrogen fertilizers (Fuentes-Ramirez et al., 1999; Muthukumarasamy et al., 1999;

Muthukumarasamy et al., 2002; Medeiros et al., 2006). In addition, the life stage of the plant at

harvest could have influenced the numbers and persistence of G. diazotrophicus within sorghum.

A decrease in bacterial population related to the age of the plant was observed in certain

sugarcane cultivars (Munoz-Rojas and Caballero-Mellado, 2003). Competition from other

microorganisms in the soil could also have affected the degree to which G. diazotrophicus could

promote or stimulate growth of the sorghum plant. Oliveira etal. (2002) demonstrated an

increase in dry matter in sugarcane seedlings inoculated with fewer strains, either G.

diazotrophicus or Herbaspirillum spp., compared to those inoculated with five different N2-

fixing strains (G. diazotrophicus, Herbaspirillum seropedicae, Herbaspirillum rubrisubalbicans,

' Azospirillum amazonense and Burkholderia sp.). The number o f speculations as to why plant

growth promotion was not observed is abundant and continued research is required. j

Understanding the complicated mechanisms and factors involved in the endophytic association

between G. diazotrophicus and sorghum is fundamental to moving forward. 129

5.2 Future directions

Sucrose appears to be a consistent factor among the endophytic interactions between G. diazotrophicus and many plant species. The likelihood o f sucrose influencing colonization is apparent from this study as well as from the fact that G. diazotrophicus primarily associates with sugar-rich plants including Pennisetum purpureum Schumach. (Cameroon grass), A nanas com osus (L.) Merr. (pineapple) mdIpomoea batatas (L.) Lam. (sweet potato) (Saravanan et al.,

2008). However, colonization and ultimately N2 fixation in graminaceous plants is a complicated process that is affected by a multitude o f independent and interconnected factors. Following the success in inoculating sorghum with G. diazotrophicus in this study, the research may be extended to focus on manipulating certain biotic and abiotic factors that could potentially affect colonization.

Colonization is influenced to some extent by external variables such as nitrogen fertilization. Previous studies with sugarcane indicated that the presence o f high supplied nitrogen inhibits the colonization o f sugarcane by G. diazotrophicus (Fuentes-Ramirez et al.,

1999; Muthukumarasamy et al., 1999; Muthukumarasamy et al., 2002; Medeiros et al., 2006). As such, adjusting the fertilizer usage in field trials could deliver insight into providing the optimal conditions for colonization. By varying the exact amount o f nitrogen fertilizer used in the field, such as applying a gradient of concentrations ranging from low to high, and enumerating the bacterial population within the plant could determine under which condition the bacterial numbers would be increased. Additionally, measuring the amount of nitrogen being leached from the system, under a controlled laboratory environment, in the presence and absence o f the bacteria could provide insight into how much fertilizer should be applied. ■ . 130

Rhizospheric populations can also influence colonization through competition for plant

resources. Controlling the microbial communities and the bacterial species used for inoculation could establish a fitting combination required for enhanced colonization. Growing the plants in an axenic culture and inoculating the seedlings with a different combination of bacterial species could establish whether there is an effect of competition or co-existence on colonization.

Moreover, other factors implicated in colonization, such as soil type, time o f inoculation, supply of plant nutrients, etc., must also be manipulated individually and collectively since these factors do interact with one another and the combination that enhances bacterial colonization is not yet known.

To further study the colonization process and the distribution of the bacterium within the plant, microscopic confirmation with in situ identification o f thé microorganism would further reinforce the plant-microbe association. Isolation from surface-sterilized plant tissues is regarded only as a single step towards the characterization o f colonization. As such, labeling the bacterium through immunological, molecular biological (fluorescence in situ hybridization), or gene marker

(green fluorescence protein) techniques would provide direct evidence of endophytic localization. Additionally, the use o f reporter gene tagging could be a promising approach to achieve a better understanding of the activities and functions o f endophytes inside the host as well as the conditions o f their microenvironment.

| Specifying the bacterial strain and matching plant partner down to the cultivar level is equally important to further clarify the complexities of the endophytic association. The plant host plays a major role in allowing or prohibiting the entry and distribution of the bacterium. In the' case o f plant pathogens, the mechanisms involved are quite well understood (Chisholm et al.,

2006). It is possible that similar key determinants such as the lipopolysaccharide and .131 exopolysaccharide structures, the flagellin type, and the type o f signaling molecules produced by the bacteria are potential elicitors o f the plant’s innate immune response (Bakker etal., 2007).

Issues that still need to be explored include: 1) increasing endophyte numbers in plant tissues without stimulating a strong defense response; 2 ) determining common signaling pathways in plant hosts to accommodate endophytes; and 3) clarifying characteristics unique to endophytic bacteria as opposed to soil or rhizospheric bacteria or pathogens. Fully understanding the specificity between the plants and endophytes and the essential components for host entry could lead to optimization of the colonization process.

The focal point of this study was to assess the capacity for colonization o f sorghum by G. diazotrophicus. Since G. diazotrophicus is an endophytic N2-fixing bacterium, the next conceivable step would be to evaluate the potential for N2 fixation in fresh plant tissues via the acetylene reduction assay. The acetylene reduction assay is based on the correlation between the reduction of C2H2 into C2H4 and the reduction of N2 to NH3 (Hardy et al., 1968). It provides a simple, rapid and sensitive method o f measuring the activity o f N2-fixing systems via gas chromatography and flame ionization. Since the reduction o f C2H2 into C2H4 occurs simultaneously as N2 is reduced to NH3, the measurement of ethylene production indirectly measures N2 fixation that is occurring in the system.

Endophytic associations, especially the interactions between N2-fixing microorganisms and plant hosts, need to be further exploited for the promotion of plant health and low-input sustainable agricultural applications. Understanding the mechanisms that enable endophytic bacteria to interact with plants could lead to achieving plant-bacterial partnerships for a range of biotechnological functions. Without a doubt, BNF does improve the nitrogen status of soils and the use of BNF in a farming system could represent a profitable approach to prevent the decline arise, 133

References -

Bakker, P. aH. M., Pieterse, C. M. J., & van Loon, L. G. (2007). Induced systemic resistance by fluorescent Pseudomonas spp. Phytopathology, 97(2), 239-243.

Bian, Y. L., Yazaki, S., Inoue, M., & Cai, H. W. (2006). QTLs for sugar content of stalk in sweet sorghum (Sorghum 'bico lo r T,. Moench). Agricultural Sciences in China, 5(10), 736-744.

Chisholm, S. T., Coaker, G., Day, B., & Staskawicz, B. J. (2006). Host-microbe interactions: shaping the evolution of the plant immune response. Cell, 124(4), 803-814.

Fuentes-Ramírez, L. E., Caballero-Mellado, J., Sepúlveda, J., & Martínez-Romero, E. (1999). Colonization o f sugarcane by Acetobacter diazotrophicus is inhibited by high N- fertilization. FEMS Microbiology Ecology, 29(2), 117-128.

Glasziou, K., & Gayler, K. (1972). Storage of sugars in stalks o f sugar cane. The Botanical Review, 38(4), 471-489. . ■V'V:V /

Govindarajan, M., Balandreau, J., Muthukumarasamy, R., Revathi, G., & Lakshminarasimhan, C. (2006). Improved yield o f micropropagated sugarcane following inoculation by endophytic Burkholderia vietnamiensis. Plant and Soil, 280(1-2), 239-252.

Hardy, R. W. F., Holsten, R. D., Jackson, E. K., & Bums, R. C. (1968). The acetylene-ethylene assay for N2 fixation: laboratory and field evaluation. Plant Physiology, 43(8), 1185-1207.

McBee, G. & Miller, F. R. (1982). Carbohydrates in sorghum culms as influenced by cultivar, spacing, and maturity over a diurnal period. Crop Science, 22(2), 381-385.

Medeiros, A., Polidoro, J. C., & Reis, V. M. (2006). Nitrogen source effect on Gluconacetobacter diazotrophicus colonization of sugarcane (Saccharum spp.). P lan t an d Soil, 279(1-2), 141-152.

Mehnaz, S., Kowalik, T., Reynolds, B., & Lazarovits, G. (2010). Growth promoting effects of com (Z ea m ays) bacterial isolates under greenhouse and field conditions. Soil Biology and Biochemistry, 42(10), 1848-1856.

Muñoz-Rojas, J., & Caballero-Mellado, J. (2003). Population dynamics of Gluconacetobacter diazotrophicus in sugarcane cultivars and its effect on plant growth. M icrobial E cology, 46(4), 454-64.

Muthukumarasamy, R., Revathi, G., & Lakshminarasimhan, C. (1999). Influence of N fertilisation on the isolation o f Acetobacter diazotrophicus and Herbaspirillum spp. from Indian sugarcane varieties. Biology and Fertility o fSoils, 29(2), 157-164. 134

Muthukumarasamy, R., Revathi, G., & Loganathan, P. (2002). Effect of inorganic N on the population, in vitro colonization and morphology o f Acetobacter diazotrophicus { syn. Gluconacetobacter diazotrophicus). Plant and Soil, 243(\), 91-102.

Oliveira, A. L. M., Canuto, E. L., Urquiaga, S., Reis, V. M., & Baldani, J. I. (2006). Yield of micropropagated sugarcane varieties in different soil types following inoculation with diazotrophic bacteria. Plant and Soil, 284(\-2), 23-32.

Oliveira, A. L. M., Urquiaga, S., Dobereiner, J., & Baldani, J. I. (2002). The effect of inoculating endophytic N2-fixing bacteria on micropropagated sugarcane plants. Plant and Soil, 242(2), 205-215.

Saravanan, V. S., Madhaiyan, M., Osborne, J., Thangaraju, M., & Sa, T. M. (2008). Ecological occurrence of Gluconacetobacter diazotrophicus and nitrogen-fixing Acetobacteraceae members: their possible role in plant growth promotion. M icrobial Ecology, 55(1), 130-40. van Veen, J. A., van Overbeek, L. S., & van Elsas, J. D. (1997). Fate and activity of microorganisms introduced into soil. Microbiology and Molecular Biology Reviews, 61(2), 121-135.

Vitousek, P., Naylor, R., Crews, T., David, M., Drinkwater, L., Holland, E., Johnes, P., Katzenberger, J., Martinelli, L. A., Matson, P. A., Nziguheba, G., Ojima, D., Palm, C. A., Robertson, G. P., Sanchez, P. A., Townsend, A. R., Zhang, F. S. (2009). Nutrient imbalances in agricultural development. Science, 324(5934), 1519-1520. APPENDIX

Bradford dye reagent Reagent Instructions

1. Dissolve 100 mg Coomassie blue G250 in 50 ml 95% ethanol

2. Mix solution with 100 ml 85% phosphoric acid

Bradford dye 3. Make up to 1 L with distilled water

4. Filter through Whatman No. 1 filter paper

5. Store in amber bottle at room temperature

* Stable for weeks, however dye may precipitate, so should be filtered before use.

Histochemical GUS staining reagents and solutions X-Gluc IX staining solution Components Final volume: 50 mL 1 part X-Gluc 5X stock solution 10 mL

4 parts incubation buffer 40 mL

X-Gluc 5X stock solution Components Final volume: 32 mL X-glucuronide (M.W. 522.0) 0.1256 g

DMSO (dissolve X-glucouronide in DMSO 8.0 mL first)

Incubation buffer 24.0 mL

X-Gluc Incubation buffer Components Final volume: 200.0 mL 0.2M N aP04, pH 7.0 100.0 mL

0 .1M K3[Fe(CN)6] 1.0 mL

0.1M K4F e(CN) 6 • 3 H20 1.0 mL

0.5MNa2EDTA 4.0 mL

Triton X -100 0.2 mL

milli-Q 1120 93.8 mL

* Reagents are light sensitive and must be wrapped in foil and stored at 4°C. 136

MPN table for 3 x 1 , 3x0.1, and 3*0.01 g (ml) Number of positive results MPN 95% Confidence limits 0 0 0 <0.30 0.00-0.94 .... j 0 0 0.30 0.01-0.95 0 1 0 0.30 0 .0 1 - 1 .0 0 0 1 1 0.61 0.12-1.70 0 2 0 0.62 0.12-1.70 0 3 0 0.94 0.35-3.50 1 0 0 0.36 0.02-1.70 1 0 1 . 0.72 0.12-1.70 1 0 2 , 1.1 0.4-3.5 1 1 0 0.74 0.13-2.00 1 1 1 1.1 0.4-3.5 1 2 0 1.1 0.4-3.5 1 2 1 1.5 0.5-3.8 1 3 o 1 .6 0.5-3.8 2 : 0 0 0.92 0.15-3.50 2 0 1 1.4 0.4-3.5 2 0 ; 2 2 .0 0.5-3.8 2 1 0 1.5 0.4-3.8 2 : 1 1 : 2 .0 0.5-3.8 2 1 ■■ 2 2.7 0.9-9.4 2 2 0 2.1 0.5-4.0 2 2 1 ; 2 .8 0.9-9.4 2 2 '. ' 2 3.5 0.9-9.4 . : 2 3 0 2.9 0.9-9.4 2 3 1 3.6 0.9-9.4 3 , 0 0 2.3 0.5-9.4 : '3" 0 1 3.8 0.9-10.4 3 0 .; ' 2 6.4 1.6-18.1 ' 3 : 1 0 4.3 0.9-18.1 3 l 7.5 1.7-19.9 3 ■ 1 ' 2 . ' 12 3-36

' V - ■ 3. --. 1 3 16 3-38 2 o 9.3 1.8-36 3 : > 2 1 . ; 15 3-38 3 2 2 21 3-40 3 ■ 2 3 29 ' ' ■ ' 9-99 3 v- 3 0 24 4-99 : 3 3 1 ■ 46 9-198 3 3 2 1 1 0 20-400 : 3 3 ■ .. 3 > 1 1 0 137

1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19

A B

1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19

C D

1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 F

1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 G H

1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 I J

1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19

K L

1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 M N Nested PCR screening of greenhouse root samples A, Détection of PAL5 in CGSH 27. B, Détection of MAd3A in CGSH 27. C, Détection of PAL5 in 98W. D, Détection of MAd3A in 98W. E, Détection of PAL5 in CGSH 8 . F, Détection of MAd3A in CGSH 8 . G, Détection o f PAL5 in Nutriplus BMR. H, Détection o f MAd3A in Nutriplus BMR. I, Détection of PAL5 in 275. J, Détection of MAd3A in 275. K, Détection of PAL5 in N111. L, Détection of MAd3 A in N111. M, Détection of PAL5 in CSSH 45. N, Détection of MAd3A in CSSH 45.

* Gel layout: lane 1 - 100 bp marker; lanes 2-5 - positive controls (bacterial colonies); lanes 6-15 - inoculated tissue samples; lanes 16-19 - negative controls (un-inoculated plants). 138

1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 A B

1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19

C D

1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19

E F

1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 G H

1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19

1 J

1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 K L

1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 1 2 3 4 5 6 7 8 9 10 11 12 13 14 15

M N Nested PCR screening of greenhouse stem samples A, Détection of PAL5 in CGSH 27. B, Détection of MAd3A in CGSH 27. C, Détection o f PAL5 in 98W. D, Détection of MAd3A in 98W. E, Détection of PAL5 in CGSH 8 . F, Détection of MAd3A in CGSH 8 . G, Détection of PAL5 in Nutriplus BMR. H, Détection of MAd3A in Nutriplus BMR. I, Détection o f PAL5 in 275. J, Détection of MAd3A in 275. K, Détection o f PAL5 in N111. L, Détection o f MAd3A in NI 11. M, Détection of PAL5 in CSSH 45. N, Détection of MAd3A in CSSH 45.

* Gel layout: lane 1 - 100 bp marker; lanes 2-5 - positive controls (bacterial colonies); lanes 6-15 - inoculated tissue samples; lanes 16-19 - negative controls (un-inoculated plants). 139

1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19

A B

12 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 C D

1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19

E F

1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19

G H

1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19

I J

1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19

K L

1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 1 2 3 4 5 6 7 8 9 10 11 12 13 14 15

M N Nested PCR screening of greenhouse leaf samples A, Detection of PAL5 in CGSH 27. B, Detection of MAd3A in CGSH 27. C, Detection of PAL5 in 98W. D, Detection of MAd3A in 98W. E, Detection o f PAL5 in CGSH 8 . F, Detection of MAd3A in CGSH 8 . G, Detection of PAL5 in Nutriplus BMR. H, Detection of MAd3A in Nutriplus BMR. /, Detection o f PAL5 in 275. J, Detection of MAd3A in 275. K, Detection o f PAL5 in N111. L, Detection of MAd3A in NI 11. M, Detection of PAL5 in CSSH 45. N, Detection of MAd3A in CSSH 45.

* Gel layout: lane 1 - 100 bp marker; lanes 2-5 - positive controls (bacterial colonies); lanes 6-15 - inoculated tissue samples; lanes 16-19 - negative controls (un-inoculated plants). 140

1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 A B

1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19

C D

1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19

E F

1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19

G H

1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19

I J

1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19

K L

1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 1 2 3 4 5 6 7 8 9 10 11 12 13 14 15

M N Nested PCR screening of field root samples A, Détection of PAL5 in CGSH 27. B, Détection of MAd3A in CGSH 27. C, Détection of PAL5 in 98W. D, Détection of MAd3A in 98W. E, Détection of PAL5 in CGSH 8 . F, Détection of MAd3A in CGSH 8 . G, Détection of PAL5 in Nutriplus BMR. H, Détection of MAd3A in Nutriplus BMR. I, Détection of PAL5 in 275. J, Détection of MAd3A in 275. K, Détection of PAL5 in N111. L, Détection of MAd3A in NI 11. M, Détection o f PAL5 in CSSH 45. N, Détection of MAd3A in CSSH 45.

* Gel layout: lane 1 - 100 bp marker; lanes 2-5 - positive controls (bacterial colonies); lanes 6-15 - inoculated tissue samples; lanes 16-19 - negative controls (un-inoculated plants). 141

1 2 .3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 A B

1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19

C D

1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19

E F

1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 1 2 3 4 5 6 7 8 9 10 11 12 1314 15 1617 18 19

G H

1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19

I J

1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19

K L

1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 1 2 3 4 5 6 7 8 9 10 11 12 13 14 15

M N Nested PCR screening of field stem samples A, Detection of PAL5 in CGSH 27. B, Detection of MAd3A in CGSH 27. C, Detection of PAL5 in 98W. D, Detection o f MAd3 A in 98W. E, Detection of PAL5 in CGSH 8 . F, Detection of MAd3A in CGSH 8 . G, Detection of PAL5 in Nutriplus BMR. H, Detection o f MAd3 A in Nutriplus BMR. I, Detection of PALS in 275. J, Detection o f MAd3A in 275. K, Detection of PAL5 in N111. L, Detection of MAd3A in N111. M, Detection of PAL5 in CSSH 45. N, Detection of MAd3A in CSSH 45.

* Gel layout: lane 1 - 100 bp marker; lanes 2-5 - positive controls (bacterial colonies); lanes 6-15 - inoculated tissue samples; lanes 16-19 - negative controls (un-inoculated plants). 142

1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 A B

1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19

C D

1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19

E F

1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19

G H

1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19

I J

1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 K L

1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 1 2 3 4 5 6 7 8 9 10 11 12 13 14 15

M N Nested PCR screening of fïeld leaf samples A, Détection of PAL5 in CGSH 27. B, Détection of MAd3A in CGSH 27. C, Détection o f PAL5 in 98W. D, Détection of MAd3A in 98W. E, Détection of PAL5 in CGSH 8. F, Détection of MAd3A in CGSH 8. G, Détection of PAL5 in Nutriplus BMR. H, Détection of MAd3 A in Nutriplus BMR. I, Détection of PAL5 in 275. J, Détection of MAd3A in 275. K, Détection of PAL5 in N111. L, Détection of MAd3A in NI 11. M, Détection o f PAL5 in CSSH 45. N, Détection of MAd3A in CSSH 45.

* Gel layout: lane 1 - 100 bp marker; lanes 2-5 - positive controls (bacterial colonies); lanes 6-15 - inoculated tissue samples; lanes 16-19 - negative controls (un-inoculated plants).