Biodiversity of Diazotrophs in Rhizosphere of Potato (Solanum …prr.hec.gov.pk/jspui/bitstream/123456789/9446/1/Tahir... · 2019-12-31 · Thesis Submission Approval This is to certify

Biodiversity of Diazotrophs in Rhizosphere

of Potato (Solanum tuberosum L.)

Tahir Naqqash

2016

Department of Biotechnology

Pakistan Institute of Engineering and Applied Sciences

Nilore, Islamabad, Pakistan

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Reviewers and Examiners

Foreign Reviewers

1. Prof. Dr. Dittmar Hahn

Designation: Professor

University: Department of Biology, Texas State University, USA.

Mailing Address: 601 University Drive San Marcos, Texas 78666, USA.

Telephone: +1 (512) 245 3372

Email Address: [email protected]

2. Dr. Philippe Normad

Designation: Research Director (C.N.R.S.)

University: Microbial Ecology Laboratory, Universite Claude Bernard, Lyon I.

Mailing Address: Microbial Ecology Laboratory, UMR CNRS 5557, F-69622

Villeurbanne Cedex.

Telephone: +33 (0) 4 7243 1377

E-mail Address: [email protected]

Thesis Examiners

1. Prof. Dr. Asghari Bano


University: Department of Bio-sciences, University of Wah, Wah Cantt.

Mailing Address: Department of Bio-sciences, Faculty of Basic Sciences, University

of Wah, Wah Cantt, Pakistan

Telephone: +92 (51) 9157000


2. Prof. Dr. M. Kaleem Abbasi


University: Department of Soil & Environmental Sciences, University of Poonch.

Mailing Address: Faculty of Agriculture, Rawalakot Azad Jammu and Kashmir,

Pakistan

Telephone: +92 (5824) 960046


3. Prof. Dr. Amer Jamil


University: University of Agriculture Faisalabad

Mailing Address: Molecular Biochemistry Lab, Department of Chemistry,

University of Agriculture Faisalabad

Telephone: +92 (41) 9201104


Head of the Department (Name): Prof. Dr. Shahid Mansoor (S.I)

Signature with Date: _________________________________

mailto:[email protected]

Thesis Submission Approval

This is to certify that the work contained in this thesis entitled Biodiversity of

Diazotrophs in Rhizosphere of Potato (Solanum tuberosum L.), was carried out by

Tahir Naqqash, and in my opinion, it is fully adequate, in scope and quality, for the

degree of Ph.D. Furthermore, it is hereby approved for submission for review and thesis

defense.

Supervisor: _____________________

Name: Dr. Kauser A. Malik

Date: 16 December, 2016

Place: NIBGE, Faisalabad.

Co-supervisor:____________________

Name: Dr. Sohail Hameed


Place: NIBGE, Faisalabad

Head, Department of Biotechnology: ___________________

Name: Dr. Shahid Mansoor


Place: NIBGE, Faisalabad

Biodiversity of Diazotrophs in Rhizosphere

of Potato (Solanum tuberosum L.)

Tahir Naqqash

submitted in partial fulfillment of the requirements

for the degree of Ph.D.

2016

Department of Biotechnology

Pakistan Institute of Engineering and Applied Sciences

Nilore, Islamabad, Pakistan

ii

Dedicated To

My Father

Muhammad Gul Khan (late)

iii

Acknowledgments

All praises and thanks are ascribed to ALLAH ALMIGHTY, without HIS grace

nothing is possible in this universe. It is HIM only, who bestowed HIS countless

blessing on me and many people around me so that we were able to achieve the

objectives of this research project. Countless blessings, mercy and peace be upon The

Holy Prophet Muhammad (PBUH), the source of inspiration and guidance to whole

of humanity forever.

Though only my name appears on the cover of this dissertation, a great many

people have contributed to its production. I owe my gratitude to all those people who

have made this dissertation possible and because of whom my graduate experience has

been one that I will cherish forever. Special thanks are due to my Father (late) who

sacrificed his past for our future and my Mother who waited a long time for our

success. I am especially thankful to my Brothers and Sisters who raised me like son

and never let me feel absence of my father and provide me the spiritual and moral

support during this long period of study, research work and in thesis writing.

My deepest gratitude is to my advisor, Dr. Kauser A. Malik (SI, TI, HI). I have

been amazingly fortunate to have an advisor who gave me the freedom to explore on

my own. I am also lucky to have Dr. Sohail Hameed [Director, Academics & Coord.

(Biology)] as my co-advisor, has been always there to listen and give advice, and at the

same time the guidance to recover when my steps faltered. He taught me how to

question thoughts and express ideas. His patience and support helped me overcome

many crisis situations and finish this dissertation. I hope that one day I would become

as good an advisor to my students as he has been to me.

I am deeply grateful to Prof. Dr. Jan Dirk van Elsas for hosting me at

Microbial Ecology Group at University of Groningen, The Netherlands during IRSIP

fellowship. He gave me the chance to work with highly skilled people and to learn a lot

from them and the long discussions that helped me sort out the technical details of

metagenomics work. I am thankful to every member of Microbial Ecology Group and

among all, I am in debt to say thanks to Michele C. Pereira e Silva, who helped me a

iv

lot with my metagenomics work. Due to her time and interest, I managed to handle

large piles of data and extracted valuable information from it regarding potato

metagenome of Pakistan soils; a never explored area before.

I would also like to appreciate and acknowledge the efforts of Dr. Shahid

Mansoor (SI), Director (NIBGE) and ex-directors of NIBGE, Faisalabad, for

maintaining the honor of this institute among other research organizations of Pakistan.

I am thankful to Dr. Mazher Iqbal (DCS) for helping me regarding GC-MS analysis

and Mr. Javed Iqbal who facilitated me for TEM which is an important part of my

study. Thanks are due to Dr. Fathia Mubeen (PS) my lab incharge who facilitated me

for both for my academics and research needs. I have no words to say thanks to Dr.

Asma Imran (PS) who taught me, how to do research and then write it. Due to her, I

was able to perform my CLSM studies which aid worth to my findings. I am also

grateful to every member of Microbial Physiology Group, NIBGE for being around

at both fun full and hard times of my PhD.

I am highly appreciative to my friends and NIBGE fellows especially; Dr.

Muther Mansoor Qaisrani, Dr. Rizwan Haider, Muhammad Kashif Hanif,

Muhammad Tayyib Naseem, Rizwan Subhani, Naveed Ahmad, Arshad

Mehmood, Ghulam Rasool Shehzad, Malik Zahid Rasheed, Dr. Rana Muhammad

Irfan and lastly but most important Rana Shahbaz Khan who helped me stay sane

through these difficult years. Their support and care helped me overcome setbacks and

stay focused on my studies. I greatly value their friendship and I deeply appreciate their

belief in me.

Finally, I appreciate the financial support from Higher Education

Commission, Pakistan that funded my studies and research in this dissertation.

Tahir Naqqash

v

Declaration of Originality

I hereby declare that the work contained in this thesis and the intellectual content of this

thesis are the product of my own work. This thesis has not been previously published

in any form nor does it contain any verbatim of the published resources which could be

treated as infringement of the international copyright law. I also declare that I do

understand the terms ‘copyright’ and ‘plagiarism,’ and that in case of any copyright

violation or plagiarism found in this work, I will be held fully responsible of the

consequences of any such violation.

________________

(Tahir Naqqash)

NIBGE, Faisalabad.

vi

Copyrights Statement

The entire contents of this thesis entitled Biodiversity of Diazotrophs in Rhizosphere

of Potato (Solanum tuberosum L.) by Tahir Naqqash are an intellectual property of

Pakistan Institute of Engineering & Applied Sciences (PIEAS). No portion of the thesis

should be reproduced without obtaining explicit permission from PIEAS.

vii

Table of Contents

Dedication ..................................................................................................................... ii

Acknowledgments ...................................................................................................... iii

Declaration of Originality ........................................................................................... v

Copyrights Statement ................................................................................................. vi

Table of Contents ....................................................................................................... vii

List of Figures .............................................................................................................. xi

List of Tables ............................................................................................................. xiv

Abstract ....................................................................................................................... xv

List of Publications .................................................................................................. xvii

List of Abbreviations ............................................................................................. xviii

1. Introduction .............................................................................................................. 1

1.1 The Rhizosphere .................................................................................................. 2

1.1.1 Rhizospheric Diversity .................................................................................. 4

1.2 Phosphorus (P) and P Solubilization by PGPR ................................................... 7

1.3 Phytohormones Production by PGPR .................................................................. 9

1.3.1 Auxins ........................................................................................................... 9

1.3.2 Pathways of IAA Production ...................................................................... 10

1.4 Bio-control ......................................................................................................... 11

1.5 Nitrogen: One of the Important Element of Life ............................................... 12

1.5.1 Nitrogen Cycle and Nitrogen Fixation ........................................................ 12

1.5.2 Biological Nitrogen Fixation ...................................................................... 13

1.6 The Diazotrophs ................................................................................................. 14

1.6.1 Associative and Free Living Diazotrophs ................................................... 15

1.6.2 Endophytic Diazotrophs .............................................................................. 15

1.6.3 Actinorhizal Diazotrophic Symbioses ........................................................ 16

1.6.4 General Biochemistry of Biological Nitrogen Fixation .............................. 16

1.6.5 Physiological and Phylogenetic Diversity of Diazotrophs ......................... 18

1.7 Biological Nitrogen Fixation and Human Demand ........................................... 20

1.8 Potato ................................................................................................................. 22

1.8.1 PGPR Interaction with Potato ..................................................................... 23

viii

2. Materials and Methods .......................................................................................... 25

2.1 Sample Collection .............................................................................................. 25

2.2 Physiochemical Analysis of Soil Samples ......................................................... 27

2.2.1 Soil Texture ................................................................................................. 27

2.2.2 pH of Soil (pH) ........................................................................................... 27

2.2.3 Electrical Conductivity (EC) of Soil ........................................................... 27

2.2.4 Organic Matter ............................................................................................ 27

2.2.5 Total Nitrogen in Soil ................................................................................. 27

2.2.6 Extractable Phosphorus ............................................................................... 28

2.2.7 Extractable Potassium ................................................................................. 28

2.3 Determination of Indigenous Bacterial Population ........................................... 28

2.4 Isolation of Rhizospheric Bacteria ..................................................................... 28

2.4.1 Isolation of Nitrogen Fixing Bacteria from Rhizosphere ........................... 29

2.4.2 Preservation of Bacteria .............................................................................. 29

2.5 Morphological Characterization ........................................................................ 29

2.6 Biochemical Characterization ............................................................................ 29

2.6.1 Reference Strain .......................................................................................... 29

2.6.2 Nitrogen Fixation ........................................................................................ 29

2.6.3 Analysis of Indole Acetic Acid (IAA) Production by Bacterial Isolates .... 30

2.6.4 Phenotypic Microarrays .............................................................................. 30

2.7 Molecular Characterization ................................................................................ 31

2.7.1 Extraction of Genomic DNA from Bacterial Isolates ................................. 31

2.7.2 Molecular Marker ....................................................................................... 31

2.7.3 Analysis of Bacterial Fingerprints using BOX and ERIC PCR .................. 31

2.7.4 Identification of Bacterial Isolates using 16S rRNA Sequence Analysis .... 32

2.7.5 Amplification of NifH Gene ........................................................................ 32

2.7.6 DNA Sequencing and Sequence Analysis .................................................. 33

2.7.7 Phylogenetic Analysis ................................................................................. 33

2.8 Plant Inoculation Studies ................................................................................... 35

2.8.1 Preparation of Seeds ................................................................................... 35

2.8.2 Preparation of Bacterial Inoculum .............................................................. 35

2.8.3 Growth Room Experiments ........................................................................ 35

2.8.4 Experiment 1: Potato Plant Inoculation in Sand Culture ............................ 35

2.8.5 Experiment 2: Potato Plants Inoculated in Soil Culture ............................. 36

ix

2.9 Analysis of Plant Samples ................................................................................. 36

2.9.1 Estimation of Total N in Plant Samples ...................................................... 36

2.9.2 Estimation of 15N Abundance in Potato Plants ........................................... 37

2.10 Field Experiments ............................................................................................ 37

2.10.1 Soil Analysis and Bacterial Population ..................................................... 37

2.10.2 Locations of Field Experiments ................................................................ 37

2.10.3 Layout Plan of Experiments ...................................................................... 38

2.10.4 Agronomy of Crop .................................................................................... 38

2.10.5 Parameters Studied .................................................................................... 38

2.11 Root Colonization Studies ............................................................................... 39

2.11.1 Ultra-structure Studies using Transmission Electron Microscopy (TEM) 39

2.11.2 Root Colonization Studies using Confocal Laser Scanning Microscope

(CLSM) ................................................................................................................ 40

2.12 Metagenomics Studies ..................................................................................... 41

2.12.1 Rhizospheric Soil DNA Extraction ........................................................... 42

2.12.2 DGGE Analysis for Microbial Diversity .................................................. 42

2.12.3 Quantitative PCR ...................................................................................... 43

2.12.4 Pyrosequencing of NifH Gene .................................................................. 44

2.12.5 NifH Gene Metagenomics Data Analysis ................................................. 45

3. Results ..................................................................................................................... 47

Part 3.1 Bacterial Isolation and Characterization .................................................... 47

3.1.1 Physico-chemical Characteristics and Bacterial Population Analysis of Soil

Samples ................................................................................................................ 47

3.1.2 Bacterial Isolation and Morphological Characterization ............................ 47

3.1.3 Analysis of Plant Growth Promoting (PGP) Traits In vitro ........................ 50

3.1.4 Phenotypic Microarray Analysis ................................................................. 53

3.1.5 Molecular Characterization ......................................................................... 56

Part 3.2 Plant Inoculation and Colonization Studies ............................................... 73

3.2.1 Pot Experiment (Sand Culture) ................................................................... 73

3.2.2 Root Colonization Studies through Transmission Electron Microscopy

(TEM) .................................................................................................................. 79

3.2.3 Root Colonization Studies through Confocal Laser Scanning Microscopy

(CLSM) ................................................................................................................ 86

3.2.4 In vivo N2 Fixation Analysis using 15N Analysis (Soil Culture) ................. 91

x

3.2.5 Field Experiments ....................................................................................... 93

Part 3.3 Metagenomics Studies of Potato Rhizosphere ......................................... 104

3.3.1 Extraction of Soil DNA ............................................................................ 104

3.3.2 Quantification of the Total Bacterial Community .................................... 104

3.3.3 Quantification of the N-fixing Community .............................................. 104

3.3.4 Changes in the Structural Community based on 16S rRNA ..................... 106

3.3.5 Changes in the Diazotrophic Community based on NifH Gene................ 106

3.3.6 Pyrosequencing (NifH Gene) .................................................................... 109

4. Discussion.............................................................................................................. 112

4.1 Bacterial Isolation and Characterization .......................................................... 112

4.2 Plant Inoculation and Colonization .................................................................. 115

4.3 Metagenomics Analysis of Potato Rhizosphere .............................................. 119

5. References ............................................................................................................. 123

xi

List of Figures

Figure 1-1 Plant microbe interaction in the rhizosphere of plant root [29]. ............ 3

Figure 1-2 The diversity and richness of different microbial communities

associated with Arabidopsis thaliana and bulk soil [43]. ...................... 5

Figure 1-3 Plant microbe interaction showing beneficial and harmful effects of

rhizobacteria [42]. .................................................................................. 7

Figure 1-4 Phosphate solubilization by plant growth promoting rhizobacteria [60].

................................................................................................................ 9

Figure 1-5 Tryptophan dependent pathways for the production of IAA in microbes

and plants ............................................................................................. 11

Figure 1-6 The nitrogen cycle [94] ........................................................................ 14

Figure 1-7 Protein model of molybdenum nitrogenase. ........................................ 17

Figure 1-8 Cluster of nif genes involved in atmospheric nitrogen fixation. .......... 18

Figure 1-9 Phylogenetic tree of diazotrophs based on 16S rRNA [132]. ............... 19

Figure 1-10 The cascade of reactive nitrogen (Nr) forms and associated

environmental problems [133]. ............................................................ 21

Figure 1-11 Potato consumption per capita per year in Pakistan. ........................... 23

Figure 2-1 Geological information of sampling sites. ........................................... 25

Figure 2-2 1kb DNA ladder used to measure the size of the bands. ..................... 31

Figure 2-3 Restriction map of vector pBBR1MCS-4. ........................................... 41

Figure 3-1 Total bacterial population (CFU) of bacteria and diazotrophs (MPN) in

the rhizospheric soil samples from potato. .......................................... 49

Figure 3-2 BOX-PCR fingerprinting patterns of different isolates from potato

rhizosphere. .......................................................................................... 57

Figure 3-3 ERIC-PCR fingerprinting patterns of different isolates from potato

rhizosphere. .......................................................................................... 58

Figure 3-4 Composite dendrogram generated from the data for BOX-PCR

fingerprints of bacterial isolates from potato rhizosphere. .................. 59

Figure 3-5 Composite dendrogram generated from the data for ERIC-PCR

fingerprints of bacterial isolates from potato rhizosphere. .................. 60

Figure 3-6 Phylogenetic tree base on 16S rRNA sequences (1014 bp) of

Azospirillum sp. isolated from potato rhizosphere ............................... 65

Figure 3-7 Phylogenetic tree base on 16S rRNA sequences (603 bp) of Rhizobium

sp. and Agrobacterium sp. isolated from potato rhizosphere ............... 66

Figure 3-8 Phylogenetic tree base on 16S rRNA sequences (979 bp) of Bacillus sp.

isolated from potato rhizosphere .......................................................... 67

xii


Brevundimonas sp. isolated from potato rhizosphere .......................... 68


Stenotrophomonas sp. isolated from potato rhizosphere ..................... 69

Figure 3-11 Agarose gel photograph showing amplified nifH gene from bacterial

strains from potato rhizosphere. ........................................................... 71

Figure 3-12 Phylogenetic tree based on nifH sequences (211 bp) of the bacterial

strains isolated from potato rhizosphere .............................................. 72

Figure 3-13 Effect of bacterial inoculation on potato plants. .................................. 76

Figure 3-14 Effect of different bacterial strains on potato shoot and root dry weight.

.............................................................................................................. 77

Figure 3-15 Effect of different bacterial strains on potato shoot and root N contents.

.............................................................................................................. 77

Figure 3-16 Population dynamics of inoculated bacterial strains in rhizosphere of

potato at different time intervals after sowing. .................................... 78

Figure 3-17 Electron micrographs of ultra-thin sections of potato root inoculated

with Azospirillum sp. TN03 ................................................................. 80


with Rhizobium sp. TN04 .................................................................... 81






with Brevundimonas sp. TN37 ............................................................ 84

Figure 3-22 Electron micrographs of un-inoculated potato root control ................. 85

Figure 3-23 Confocal image potato (variety Kuroda) root inoculated with YFP-

labelled Azospirillum sp. TN03............................................................ 87


labelled Rhizobium sp. TN04 ............................................................... 88


labelled Azospirillum sp. TN09............................................................ 89


labelled Brevundimonas sp. TN37 ....................................................... 90

Figure 3-27 Principal component analysis showing the relationship between

different experimental locations and treatments. ............................... 101


different experimental locations and treatments. ............................... 102

Figure 3-29 Field view of inoculated potato (variety Kuroda) plants ................... 103

Figure 3-30 Abundance of total bacterial community based on 16S rRNA in potato

rhizosphere. ........................................................................................ 105

xiii

Figure 3-31 Abundance of diazotrophs based on nifH gene in potato rhizosphere.

............................................................................................................ 105

Figure 3-32 Comparison of total bacterial population to the diazotrophic community

in rhizosphere of potato across different regions. .............................. 106

Figure 3-33 Denaturing gradient gel of amplified 16S rRNA from potato rhizosphere.

............................................................................................................ 107

Figure 3-34 Non-metric multidimensional scaling analysis based on fingerprints

generated by PCR-DGGE of 16S rRNA from potato rhizosphere. .... 107

Figure 3-35 Denaturing gradient gel of amplified nifH gene from potato rhizosphere.

............................................................................................................ 108

Figure 3-36 Analysis based on fingerprints generated by PCR-DGGE of nifH gene

from potato rhizosphere. .................................................................... 108

Figure 3-37 Refraction indexes of Shannon, chao1, abundance and diversity of

diazotrophs in rhizosphere of potato from different regions. ............ 110

Figure 3-38 Relative abundance of diazotrophs in rhizosphere of potato from

different regions. ................................................................................ 111

xiv

List of Tables

Table 1.1 Important diazotrophic genera reported for their nitrogen fixation ability

with different crop plants. .................................................................... 22

Table 2.1 Geographical information of sampling sites ........................................ 26

Table 2.2 PCR and thermal conditions for BOX-PCR and ERIC-PCR analysis,

16S rRNA and nifH genes. ................................................................... 34

Table 2.3 PCR and thermal conditions for PCR-DGGE analysis and qPCR for 16S

rRNA and nifH genes and pyrosequencing for nifH. ............................ 46

Table 3.1 Physico-chemical properties of soil samples ....................................... 48

Table 3.2 Morphological characteristics of bacterial isolates purified from

rhizosphere of potato. ........................................................................... 49

Table 3.3 Nitrogenase activity of bacterial isolates purified from potato growing

areas of Punjab. .................................................................................... 51

Table 3.4 Quantification of IAA produced by bacterial isolates in LB medium . 52

Table 3.5 Differential metabolic profiling of bacterial isolates associated with

potato roots (Biolog PM2A Microplate analysis) ................................ 53

Table 3.6 Identification of bacterial isolates from rhizosphere of potato based on

16S rRNA gene sequence analysis. ..................................................... 62

Table 3.7 NifH gene sequence similarity of nitrogen fixing bacteria from

rhizosphere of potato. ........................................................................... 71

Table 3.8 Effect of inoculation of different nitrogen fixing strains on shoot growth

parameters of potato plant grown in sterilized sand. ........................... 74

Table 3.9 Effect of inoculation of different nitrogen fixing strains on root growth

parameters of potato plant grown in sterilized sand. ........................... 75

Table 3.10 Effect of inoculation of nitrogen fixing strains on growth parameters, N

contents and 15N abundance of potato plant grown in sterilized soil and

estimates of biologically fixed nitrogen. .............................................. 93

Table 3.11 Physico-chemical analysis and bacterial population of soil from

experimental sites. ................................................................................ 94

Table 3.12 Comparison of treatment means and location means of potato inoculated

with bacterial isolates under field conditions....................................... 99

Table 3.13 Comparison of treatment means and location means of potato inoculated

with bacterial isolates under field conditions....................................... 99

Table 3.14 Potato growth parameters as affected by interaction between bacterial

treatments and locations under field conditions................................. 100

Table 3.15 Potato growth parameters as affected by interaction between bacterial

treatments and locations under field conditions................................. 100

xv

Abstract

Potato is the third most important food crop which requires high fertilizer that leads to

environmental pollution. The use of biofertilizer is environment friendly and cost

effective hence is suitable for sustainable agriculture. In this study, 44 bacterial isolates

including mostly Azospirillum, Bacillus, Brevundimonas, Enterobacter, Pseudomonas

and Rhizobium spp. were isolated from potato rhizosphere of which 32 were able to fix

atmospheric N2 and 29 showed indole acetic acid (IAA) production. Strains showed

high metabolic and genetic diversity based on BioLog, ERIC and BOX-PCR analysis.

Out of potential plant growth promoting rhizobacteria (PGPR) tested for inoculation

response in potato, it was observed that the N2 fixation (%Ndfa) decreases with the

increase in applied N for Rhizobium sp. TN04 and Azospirillum sp. TN09 whereas no

effect was observed for Azospirillum sp. TN03. Azospirillum sp. TN09 showed the

highest %Ndfa among three tested isolates. Under field conditions, Azospirillum spp.

TN03, TN09 and Rhizobium sp. TN04, Rhizobium sp. TN04 increased plant height

(18%), dry weight (18.5%) tuber dry weight (14%) and tuber yield (19%). Azospirillum,

Brevundimonas and Rhizobium spp. also maintain substantial population with potato

roots. Metagenomics were employed to study total bacterial diversity in the rhizosphere

of potato from major potato growing areas of Punjab, Pakistan showed that structural

community (based on 16S rRNA) is almost similar across all the regions whereas

functional diversity (based on nifH) of diazotrophs is mostly variable. Abundance of

diazotrophs (108-1010 copies of nifH g-1 of soil) changed among regions.

Pyrosequencing of nifH validated maximum diversity in Sheikhupura (446 species)

whereas minimum in Gujranwala area (291 species). Overall in the potato rhizosphere,

most abundant class was α-proteobacteria followed by β-proteobacteria. The study

concludes that abundance of different genera varies across different regions due to

many biotic and abiotic factors. Azospirillum spp. TN03, TN09 and Rhizobium sp.

TN04 omnipresent and have potential to improve potato yield up to 15-19%

subsequently saving 50% nitrogenous fertilizer, are recommended as potential

candidates for biofertilizer production for potato crop. The study also shows that potato

xvi

harbor a diverse range of novel diazotrophs (not known earlier) which opens new

horizons for future prospects.

xvii

List of Publications

1. T. Naqqash, S. Hameed, A. Imran, M. K. Hanif, A. Majeed, and J. D. van Elsas,

"Differential growth stimulation response of potato towards inoculation with

taxonomically diverse plant growth promoting rhizobacteria," Frontiers in Plant

Science, vol. 7, pp. 144, 2016.

xviii

List of Abbreviations

°C Degree centigrade

µL Micro litre

µm Micro meter

10X 10 times

15N Isotope of nitrogen with atomic

ABA Abscisic acid

ANOVA Analysis of variance

ARA Acetylene reduction assay

ATP Adenosine triphosphate

BLAST Basic local alignment search tool

BNF Biological nitrogen fixation

cfu Colony forming units

CLSM Confocal laser scanning microscope

cm Centimeter

CRD Completely randomized design

DAS Days after sowing

DGGE Denaturing gradient gel electrophoresis

HPLC High performance liquid chromatography

IAA Indole-3-acetic acid

LB Luria Bertani

MPN Most probable number

N Nitrogen

N2 Atmospheric nitrogen

Nr Reactive nitrogen

NCBI National Center for Biotechnology Information

NFM Nitrogen free malate

P Phosphorus

PCA Principal component analysis

PCR Polymerase chain reaction

xix

PGPR Plant growth promoting rhizobacteria

pH Hydrogen ion concentration

ppm Parts per million

PSB Phosphate solubilizing bacteria

qPCR Quantitative PCR

RCBD Randomized complete block design

rpm Revolution per minute

TEM Transmission electron microscope

YFP Yellow fluorescent protein

1. Introduction

Soil is the main source which provides fundamental ecosystem services i.e. water

regulation, nutrient cycling, toxic compounds and transformation of organic materials

as well as control of diseases and pests [1, 2]. It offers an extremely heterogeneous and

vibrant environment for its microbiota in the form of a range of microhabitats formed

by different combinations of solid fractions of soil [3]. These microhabitats help and

promote the development and maintenance of an extremely large number of niches [4]

which have a direct effect on the living fraction of soil. The microbes plays an important

role in almost all soil processes [5] and are significant drivers for almost all

biogeochemical cycles in terrestrial ecosystems such as nitrogen and carbon cycle [6]

and ultimately shaping soil structure [7]. So soil microbial communities are very

important in maintaining the quality of both natural and agriculturally managed soil

systems. These microorganisms are highly responsive to environmental changes, such

as abiotic (pH, temperature, soil structural or textural type and soil moisture) and biotic

factors (the diversity and composition of the microbial communities) [8-10].

All the forms of life are assumed to be ascended over 3.5 billion years ago from

a single common ancestor. Bacteria are the most widely distributed and flexible of all

different organisms. They have been recognized in every single distinctive kind of

environments on Earth. These are found deep inside the earth's crust upto the oceans

and are discovered from acid mine seepage and even living inside of mammalian guts.

Bacteria are thought to be the main contributors of the biomass of the world due to their

adaptability to diverse and broad range of environmental conditions. Until the discovery

of microscope in 1683 by Anton von Leeuwenhoek, the existence of bacteria was

unknown however these were used for plant growth promotion since ancient times as

unknown however these were used for plant growth promotion since ancient times as

Theophrastus (372-287 BC) suggested mixing of different soils in order to improve soil

productivity [11].

Around 80-90% of soil processes are controlled and mediated by soil microbes

1. Introduction

2

so it must be true to say that these are indeed an important part of soil [12]. They are

essential part of soil ecosystems, involved in all biotic activities in order to make it

beneficial in terms of nutritive values and ultimately for sustainable crop production

[13]. Bacterial activity, abundance and composition will mainly regulate sustainable

yield of agricultural land [14].

Different bacteria perform different functions like some are involved in nutrient

cycling other are effecting water dynamics. Few resides in root zone of plants as

pathogen while other produce a range of antibiotics [15] to suppress these pathogens.

For the better use of these microbes for sustainable agriculture, it is necessary to

understand the rhizosphere and rhizospheric diversity of bacteria.

1.1 The Rhizosphere

Plant life is the primary energy source that drives the terrestrial soil ecosystem. The

micro-ecosystem of plants is very complex and it harbors many different types of

bacteria which colonize almost all parts of plants i.e. roots, stem, leaves, flowers, fruits

and even seeds [16-18]. Due to the scarcity of biologically available nutrients in the

soil, plant roots must explore large volumes of soil to obtain the nutrients which they

require for their growth and survival. As a result, plant roots provide a supply of

biologically available nutrients to a large range of microorganisms in their immediate

vicinity. The plant roots can be divided into three zones of influence (Figure 1-1);

a) The rhizosphere; the volume of soil surrounding the root that is subject to

influence by the root [19]

b) The rhizoplane; the root surface and tightly adhering soil particles.

c) The root interior; bacteria capable of colonizing the root interior are known as

endophytes [18].

Biomass and composition of plant communities can be affected directly or

indirectly by rhizospheric microbiota [20]. Numerous organisms play an important role

to these processes, leading to a large number of interactions between antagonists,

mutualistic symbionts and plants, both above ground and below ground [21].

However many of the microbes can cross the endodermis barrier, moving from the

root cortex to the vascular system, and subsequently thrive as endophytes in leaves,

1. Introduction

3

tubers, stem and other organs [22]. The magnitude of endophytic colonization of host

plant tissues and organs reflects the capability of bacteria to adjust to these particular

ecological niches [23]. Consequently, intimate associations between host plants and

bacteria can be formed [24] without harming the plant [22, 25].

Plant roots excrete various organic substances into the rhizosphere such as sugars,

amino acids, vitamins etc. that are collectively referred to as exudates. The composition

of the exudates varies with plant species, physiological condition of the plant such as

age and nutritional status, and abiotic conditions [26-28]. Root exudates within the

rhizosphere support the growth and metabolism of diverse microbial populations

inhabiting this soil zone. The importance of the rhizosphere arises from the release of

exudates from the root and the resulting influence of increased microbial activity on

nutrient cycling and plant growth (beneficial, neutral, harmful or variable).

Figure 1-1 Plant microbe interaction in the rhizosphere of plant root [29].

1. Introduction

4

1.1.1 Rhizospheric Diversity

Biodiversity or biological diversity is defined as the set of species and their genetic

material in a specific ecosystem. It includes examination at the ecosystem, genetic

diversity levels and species [30]. The influence of microbial diversity on the endurance

of ecosystem functioning has been studied intensely with relationships often observed

between soil, microbial diversity and ecosystem sustainability and plant quality [9, 31-

33]. The rhizosphere microbial communities are very diverse both genotypic and

phenotypically so characterization of rhizospheric communities is quite difficult

(Figure 1-2). Bacteria are the most numerous inhabitants of the rhizosphere, with

population numbers typically ranging between 106-109 g-1 of rhizosphere soil [34]. The

number of bacteria associated with plant roots g-1 of soil is 10 to 100 folds greater than

the bacterial density associated with bulk soil [35]. Plant rhizosphere has harbor very

divers bacterial communities like Proteobacteria (Azospirillum, Enterobacter,

Pseudomonas, Rhizobium, Serratia etc.), the Gram-positive bacteria Firmicutes

(Bacillus) and the Actinobacteria (Streptomyces).

Over the past few decades, a number of molecular techniques like microbe

tagging by florescent genes, stable isotopes and mutagenesis helped to get the better

understanding of processes and players that work in the rhizosphere [36-38]. The

microbial ecology of the rhizosphere of plant species in their natural ecosystems [39]

and phenomenon of microorganisms impact on biodiversity, resource allocation even

the above-ground interactions with herbivores and their common enemies [40, 41] were

explored in detail.

The interaction between plants and microorganisms can be harmful, beneficial,

or neutral for the plants where microbe may have direct or indirect effects as sometimes

the soil conditions also changes due to the effect of microorganisms [42].

1. Introduction

5

Figure 1-2 The diversity and richness of different microbial communities

associated with Arabidopsis thaliana and bulk soil [43].

Beneficial

Rhizobacteria which are beneficial to plant are known as plant growth promoting

rhizobacteria (PGPR). The term PGPR is used from three decades for rhizobacteria

which are nonpathogenic, strongly colonize surface of plant’s roots and increase its

yield by one or more mechanisms [44, 45]. A potential PGPR is designated as PGPR

when it has capability to effect on the plant positively upon inoculation, hence

indicating that good competitive skills over the existing rhizospheric communities is

also a very important character.

1. Introduction

6

PGPR affect plant growth by direct and indirect ways [46], where the direct ways

includes:

Lowering of ethylene concentration

Atmospheric nitrogen fixation

Mineral phosphate solubilization

Phytohormone production

Zinc mobilization

Indirect mechanisms are;

Release metabolites that have antifungal activity

Production of antibiotics

Compete with pathogenic microbes for the sites on the roots

Siderophore production

Induce systemic resistance

Synthesis of antifungal cell wall lysing enzymes

Interactions between PGPR and plants in the rhizosphere clearly affect growth and

development of the crop plants and yield [13, 47, 48]. PGPR have a very wide range of

genera among which common examples include Acinetobacter, Arthrobacter,

Azospirillum, Bacillus, Bradyrhizobium, Burkholderia, Cellulomonas, Frankia,

Pantoea, Pseudomonas, Rhizobium, Serratia, Streptomyces and Thiobacillus.

Harmful

As root exudates attract beneficial microorganism, these may have equal attraction for

pathogenic population that can have adverse effects on plant growth [42, 49]. There are

many factors which defines the number and diversity of pathogenic communities,

however, quantity and quality of the rhizodeposits and the microbial interaction that

age going on in the soil are of prime importance. As the beneficial microorganisms,

phtyto-pathogens can also grow in the bulk soil, however in rhizosphere their activity

is increased and where the infection occur, due to the availability of root exudates [50].

These microbes also affect plant growth both by direct and indirect ways. These

adverse effects are caused by;

Production of type III effectors and toxins

Production of phytotoxins e.g. coronatine, syringomycin, pectatelyases.

1. Introduction

7

Auxin production leading to gal formation

Competition for nutrients

However, bacteria can behave as pathogens or symbionts depending on the

environmental conditions [51] such as light, nutrient, water or temperature stress, size

of inoculums, host developmental signals. Similar bacterial strain can be beneficial and

harmful depending upon the set of agricultural conditions and host species.

Figure 1-3 Plant microbe interaction showing beneficial and harmful effects of

rhizobacteria [42].

1.2 Phosphorus (P) and P Solubilization by PGPR

Phosphorus (P) is considered as vital nutrient for plants. It is present in soil as mineral

salts or incorporated into organic compounds. However this P is non-available, despite

agriculture soil are rich in it [52]. Even application efficiency of applied P rarely

exceeds 30% due problems like fixation in soil [53], run-off and leaching [54] losses.

Therefore, the unavailability of P in soils has been reported as an important growth

limiting factor for plants. Moreover P is not a renewable resource and it is obtained

from phosphate rocks, which may be depleted in 50-100 years [55]. Therefore,

1. Introduction

8

exploring alternative forms of agriculture, where nutrient conservation is key, is of vital

importance.

The soil microbes have the ability to transform unavailable soil P to the forms

available to plants. Microbial communities acclimatizes dissolvable P and hinder it

from fixation or adsorption [48, 56] as shown in figure 1-4. Among the soil microbes,

credible groups of phosphate solubilizing bacteria belong to genera Bacillus,

Enterobacter, Pseudomonas and Rhizobium [37, 48, 57].

These phosphate-solubilizing microorganisms use diverse mechanism(s) to

solubilize the insoluble types of the phosphate. The principal mechanism of phosphate

solubilization depends on organic acid discharge by microorganisms as a result of sugar

metabolism. Rhizobacteria use root exudates specially sugar compounds for the

production of organic acids [37, 58], which act as great chelators of Ca2+ associated

with discharge of phosphates from insoluble phosphates [59]. Huge numbers of the

phosphate-solubilizing microorganisms bring down the pH of the medium by

production of organic acids, for example lactic, acetic, succinic, malic, tartaric, oxalic,

gluconic, 2-ketogluconic and citrus extracts [37, 48]. Phosphate solubilization

mechanism can be divided in to below two main ways;

I. Mineral solubilization of phosphate

II. Organic solubilization of phosphate

1. Introduction

9

Figure 1-4 Phosphate solubilization by plant growth promoting rhizobacteria

[60].

1.3 Phytohormones Production by PGPR

The phytohormones play key role in growth and yield of plants. Their importance and

positive effects have been discussed over the last 30 years by many scientists [37, 47,

61-63]. Phytohormones include auxins, gibberellins, cytokinins, ethylene and abscisic

acids (ABA), among which auxin i.e. indole-3-acetic acid (IAA) is a key controller of

many features of plant development and improvement, including cell elongation and

division, apical dominance, tropisms, flowering, separation, abscission and senescence

[64-66]. Its importance was known for plant growth promotion [67] even before its

proper identification [68, 69]. These phytohormones are also produced by majority of

the PGPR where IAA is commonly produced, although some also produce other

phytohormones e.g. gibberellins and cytokinins [70, 71].

1.3.1 Auxins

Auxins act as master control, affecting many plant processes directly or indirectly, as

well as interactively influence the synthesis and action of other phytohormones. Auxins

secreted by bacteria have been found to act as signaling molecules for communication

between bacteria to coordinate their activities [72]. One of the most prominent features

1. Introduction

10

of plants inoculated with auxin producing PGPR is the modification of root morphology

and its development. Thus, PGPR promote root growth by increasing root surface area

which ultimately promotes water and nutrient uptake, thereby indirectly stimulating

plant growth positively [47]. Auxins also control many plant mechanisms by regulating

the expression of certain genes [73]. About 80% of rhizobacteria from different crops

have ability to produce IAA as secondary metabolites. Bacteria belonging to genera

Acetobacter, Alcaligenes, Azospirillum, Bacillus, Bradyrhizobium, Enterobacter,

Pseudomonas, Rhizobium and Xanthomonas are well documented to produce IAA

which improved plant growth [37, 47, 63, 74].

1.3.2 Pathways of IAA Production

The amount and pathway for the production of IAA varies among microorganisms.

Different reports suggested the increase in the IAA production in the presence of

supplemented Tryptophan as precursor because most of the rhizobacteria have

tryptophan dependent pathways of IAA production. Below are the metabolic pathways

adopted by different microbes;

Indole-3-acetamide (IAM) pathway

Indole-3-pyruvate (IPA) pathway

Tryptophan side chain oxidase (TSA) pathway

Tryptamine pathway

Indole-3-acetonitrile (IAN) pathway

There are many factors which define production of IAA among rhizobacteria like

biosynthetic pathways adopted, location of the genes involved, regulatory sequences,

and the presence of enzymes to convert active free IAA into conjugated forms and also

the environmental conditions [63]. The pathway used for the production of IAA by

bacteria may play an important role as pathogenic bacteria use indole-3-acetamide

pathway whereas beneficial bacteria use indole-3-pyruvate pathway for IAA production

[63]. Tryptophan dependent pathways used by major IAA producing genera is shown

in figure 1-5.

1. Introduction

11

Figure 1-5 Tryptophan dependent pathways for the production of IAA in

microbes and plants

1.4 Bio-control

Including all the beneficial attributes discussed earlier, PGPR can also control a variety

of pathogens like bacteria, fungus, nematodes and are even effective against viruses.

Rhizobacteria use these mechanisms to compete, acquire and sustain favorable niches

in the rhizosphere over the other partners, which indirectly help plant to stay healthy

and disease free. They have the ability to suppress soil borne pathogens naturally [75].

These PGPR are inoculated to plants to control variety of diseases all over the world.

There are many reports about the suppression of plant pathogen by PGPR both in vitro

1. Introduction

12

and in vivo conditions [76]. The bacterial genera Agrobacterium, Alcaligenes, Bacillus,

Pseudomonas, Micromonospora, Streptomyces, Streptosporangium and Thermobifida

are known bio-control agents [15, 77, 78], among which Pseudomonas is considered as

largest group to have bio-control activity [79, 80]. A successful bio-control agent have

the following characteristic [15, 81];

Fast growing to produce mass culture

Potential to utilize diverse metabolites (root exudates)

Potential and good colonizer of roots

Capability to produce a wide range of metabolites i.e. antibiotics, HCN,

siderophores, volatiles substances.

Good competitive ability with other microbes and adaptability to

environmental stresses

Development of induced systemic resistance in plants

1.5 Nitrogen: One of the Important Element of Life

Nitrogen is one of the most essential as well as common element present in many bio-

molecules necessary for sustainable life [82, 83]. Though N was not recognized as an

element until 18th century but saltpeter (potassium nitrate) was used in the formation of

gunpowder, soup and also used as a fertilizer. It show that N was used as an important

component of many manufactured substance for centuries, even before its recognition.

Microbial communities are important part of the bio-geochemical processes in N cycle

as it is one of the most important limiting nutrient for crop production in most

developing countries [84]

1.5.1 Nitrogen Cycle and Nitrogen Fixation

Ample nitrogen is available in Earth’s atmosphere but due to inert nature (N2 gas), it is

unavailable to most of the organisms [85]. These organisms fulfill their N requirements

by utilizing fixed form of N e.g. ammonia (NH3) or nitrate (NO3−) [86]. Since fixed N

is constantly confiscated into sediments becoming unavailable for metabolism and is

also regularly converted to N2 through different processes like nitrification and

denitrification, the process of conversion of N2 to NH3 is critical step in N cycle for

sustainable life [87]. This process is known as N2 fixation [84]. Nitrogen fixation is an

important part of the nitrogen cycle as it replenishes the overall nitrogen content of the

1. Introduction

13

biosphere and compensates the losses that are induced by denitrification. N2 fixation

can occur in three ways:

I. Geochemical processes i.e. lightning [88]

II. Industrial processes i.e. The Haber−Bosch process [89]

III. Biological process i.e. via nitrogenase enzyme [90] found only in selected

groups of microorganisms [91]

However conversion of N2 to NH3 or NO3 is high energy demanding process as

triple bond between N2 make it one of the most inert element comparatively. Nitrogen

fixed through lightning process is very low and on the other hand, constant increase in

in oil prices and global attempts to alleviate greenhouse gas (GHG) emissions

associated with the agricultural use of mineral fertilizers containing N formed by an

energy intensive Haber-Bosch process is not appreciated.

However a group of prokaryotes contain nitrogenase; an enzyme complex,

involved in the reduction of atmospheric dinitrogen to ammonium, thus replenishing

the biological nitrogen pool (Figure 1-6). Scientists are focusing to substitute chemical

fertilizers with the increased use of biological nitrogen fixation (BNF) in leguminous

and non-legume cropping systems as it can be cheaper and environmental friendly

alternate.

1.5.2 Biological Nitrogen Fixation

Biological nitrogen fixation stands for the conversion of ammonia from atmospheric

nitrogen, carried out by free-living/associative and symbiotic bacteria which is

important for agriculture and environment. Enzymatic reduction of N2 was carried out

in BNF to produce NH3 and NH4 which act as initial molecules for N containing

biomolecules like amino acids [92]. Only domain Bacteria and Archaea are able to

perform BNF among all prokaryotes and eukaryotes [93]. Animals and plants are

dependent upon the BNF activity of N-fixing prokaryotes known as diazotrophs, for

their nitrogen supply.

1. Introduction

14

Figure 1-6 The nitrogen cycle [94]

1.6 The Diazotrophs

Microorganisms which have the ability to reduce atmospheric N2 to NH3 are known as

diazotrophs. Thus diazotrophs make a very important functional group of N cycle.

There are numerous well established associative relations between plants and

diazotrophs which allow them to directly harness the power of nitrogen reduction.

However there are two main mode of actions by which diazotrophs fix N2 for plants.

Symbiotic (Nodule forming diazotrophs)

Asymbiotic (associative diazotrophs)

Where symbiotic diazotrophs include Rhizobium, Bradyrhizobium, Mesorhizobium,

Sinorhizobium, Allorhizobium, Azorhizobium and associative diazotrophs include

Azospirillum, Azotobacter, Azoarcus, Enterobacter, Pseudomonas, Bacillus as well

known diazotrophs. We shall discuss in detail only the asymbiotic diazotrophs. There

are three different ways by which asymbiotic diazotrophs interact with plants.

1. Introduction

15

1.6.1 Associative and Free Living Diazotrophs

Free living diazotrophs share a significant portion of atmospheric nitrogen to the

biosphere but still there importance in agriculture is least. The absence of glucose

partner in soil explains this, as high energy is required to cover the vital necessities of

nitrogenase, which is more increased in shielding this enzyme from aerobic conditions.

Moreover, biologically fixed NH3 is also not transferred to plants directly by bacteria.

These diazotrophs are used as biofertilizer from mid of the 20th century on commercial

scale. Azotobacter and Azospirillum inoculants are designated to reduce N fertilizer

application [95, 96]. However, it is still debatable whether atmospheric nitrogen fixed

by these bacteria is considerably incorporated by the host [97]. Because the mechanisms

involved in the improvement of crop plants is not yet cleared, since these bacteria

possesses numerous mechanism involved in the mineralization and plant growth

promotion like P solubilization and phytohormone production. Examples of such free

living diazotrophs include Azospirillum, Azotobacter, Bacillus, Burkholderia,

Herbaspirillum and Paenibacillus [47, 98-101].

1.6.2 Endophytic Diazotrophs

Diazotrophs which have the ability to infect, multiply and spread inside the plant parts

without causing harm or risking their hosts are known as endophytic diazotrophs. The

best focused endophytic diazotrophs include individuals from Azoarcus, Burkholderia,

Gluconobacter, Herbaspirillum and Klebsiella [102] There are many studies about the

relation of endophytic diazotrophs with many crops e.g. sugarcane [103], rice [104],

maize [105], sunflower [106]. A few reports have shown their capacity to reduce N2 in

plants, by detecting the expression of nitrogenase genes in plant cells [107] and by

isotope investigation [108]. It is suggested that up to 70% of basic required N of plants

can be mediated by BNF through endophytic diazotrophs [102]. Despite the fact that

advancements has been made on the comprehension of this kind of beneficial plant

microbe association, the commitment of single species to the host nitrogen balance is

still indistinct. An ineffectively comprehended association between plant genotype, soil

and ecological conditions appear to decide the commitment of diazotrophic endophyte

nitrogen fixation to plant efficiency [109]. Thus more learning is required to completely

comprehend nitrogen fixation in these essential crop systems and to extend their

advantages further.

1. Introduction

16

1.6.3 Actinorhizal Diazotrophic Symbioses

Actinorhizal Frankia spp. and plant symbiosis is third way of diazotrophic interaction.

This type of association is observed in more than 8 dicotyledonous plant families and

25 genera [110]. The specificity of the association, contrary to Rhizobium-legume

symbiosis, is relatively low which increased interest for the potential expansion of the

nitrogen-fixing capacity to different frameworks. However little information on both

microbes and host plants hinders understanding of these frameworks.

1.6.4 General Biochemistry of Biological Nitrogen Fixation

A number of functional and regulatory genes are involve in BNF [111]. Nitrogenase

enzyme plays the most important role in the reduction of atmospheric N2. This enzyme

is a combination of two Fe-proteins, one is nitrogenase Fe-protein and second is

nitrogenase Mo-Fe-protein [92, 112, 113]. Nitrogen (N2) bounds to Mo-Fe-S

homocitrate part of Mo-Fe-protein as substrate (Figure 1-7). Same procedure is used

for other substrates like protons, acetylene etc. [114]. Whereas Fe-protein transports

electrons to Mo-Fe-protein at the cost of two Mg-ATP per electron [92]. BNF process

can be expressed completely as fellow, as described by Ferguson [83];

N2 + 10H++ 8e- + nMgATP → 2NH4++ H2 + nMgADP + nPi, n ≥ 16

Along these lines, this procedure is energy demanding and consumes 8 mol of

ATP to produce 1 mol of NH4+, even in normal conditions this proportion might be

increased [115]. The ability of a diazotroph to fix atmospheric nitrogen can be

quantified by acetylene reduction assay [116].

1. Introduction

17

Figure 1-7 Protein model of molybdenum nitrogenase.

(A) One catalytic half of the Fe protein: MoFe protein complex with the Fe protein

homodimer shown in tan, the MoFe protein α subunit in green, and the β subunit in

cyan. (B) Space filling and stick models for the 4Fe−4S cluster (F), P-cluster (P), and

FeMo-co (M) [92].

The Genetic of Biological Nitrogen Fixation (Nif Genes)

Nitrogen fixation is a complex process, carried out by combination of numerous gene

products (Figure 1-8). The number and ways of expression of genes is very diverse,

depending upon the type of species. The most abundant and important Mo-Fe protein

(α2β2) is encoded by nifDK whereas Fe-protein (α2) is expressed by nifH [117]. There

are numerous other accessory and regulatory genes which are responsible for synthesis

of nif regulon [118]. The nifD and nifK genes are part of same operon as well as nifH

1. Introduction

18

is additionally considered a portion of this operon often. Nitrogenase metal groups are

discovered together on the operon nifEN, integrated by nifE and nifN, considered as the

center operons which may have developed from the replication of two operons nifDK

and nifEN [119]. Moreover several different genes are present to add-on these operons

which are responsible to code proteins involved in electron transport i.e. nifF, nifJ

(Klebsiella pneumoniae), regulation i.e. nifA or Fe-Mo cofactor i.e. nifB, nifV

(Klebsiella pneumoniae) synthesis [120]. Few systems don't carry molybdenum,

however in one case vanadium (vnfDK and vnfH), while for other only iron is present

(anfDK and anfH). These two alternative systems are directed under Mo deficient

conditions [121].

Among all the nif genes, nifH gene is used as an important marker gene for the

studies of diazotrophs. Because nifH gene sequence is highly conserved and a huge

sequence data, generated studies in various conditions, is available for comparative

studies [32, 122, 123]. Correlation of 16S rRNA and nifH phylogenies gives no solid

proof for lateral exchange nifH gene [124] however, rare differences were observed [8,

32]. Use of recent molecular techniques like next generation sequencing and whole

genome sequencing, are producing reasonable information which will increase the

understanding about phylogenetic relations of diazotrophs in differing environmental

situations.

Figure 1-8 Cluster of nif genes involved in atmospheric nitrogen fixation.

1.6.5 Physiological and Phylogenetic Diversity of Diazotrophs

Diazotrophs are diverse in nature (Figure 1-9), found among phototrophic

microorganisms e.g. aerobic phototrophic Cyanobacteria [125], anaerobic purple-sulfur

1. Introduction

19

phototrophs (Chromatium) and green-sulfur phototrophs (Chlorobium) [126]. These

have been observed in organisms growing chemolithotrophically (Alcaligenes,

Thiobacillus, Methanosarcina, or Azospirillum lipoferum) [126, 127] and in a great

number of heterotrophic bacterial strains such as anaerobes (Clostridium), micro-

aerophiles (Herbaspirillum) and aerobes (Azotobacter) [128]. Despite the large number

of known diazotrophic genera, many isolates have never been assessed for BNF

capacity, so diazotrophs may be even more widespread than currently known [126].

Only a minority of diazotroph species is involved in symbioses, yet free-living

diazotrophs have received comparatively little attention from researchers. Although

free-living Cyanobacteria and their contribution to the N-budget of oceans and lakes

have been studied in some detail [129, 130], free living soil diazotrophs are still poorly

characterized which includes aerobic N-fixing bacterial genera found in soil e.g.

Azotobacter, Beijerinckia and Derxia, nevertheless the majority are microaerophilic

(e.g. Azospirillum, Herbaspirillum) or facultative and obligate anaerobes (e.g.

Klebsiella, Clostridium, Erwinia) [131].

Figure 1-9 Phylogenetic tree of diazotrophs based on 16S rRNA [132].

1. Introduction

20

1.7 Biological Nitrogen Fixation and Human Demand

The prospect of utilizing BNF for agricultural purposes has long been the driving force

behind N-fixation research. The total N-demand all over the world has been estimated

1.15×108 tonnes in 2015, which is supposed to increase up to 1.194×108 tonnes in 2018

where 4% of the total N demand is required by Pakistan (FAO, 2014). Moreover

excessive use of nitrogenous fertilizer is causing serious environmental issues (Figure

1-10) that can be grouped into below major groups [133];

Air quality – including shortening of human life through exposure to air

pollutants including particulate matter formed from NOx and NH3 emissions,

and from increased concentrations of nitrogen dioxide (NO2) and ground level

ozone (O3).

Greenhouse gas balance – including emissions of N2O plus interactions with

other reactive nitrogen (Nr) forms, particulate matter and atmospheric Nr

deposition, plus tropospheric O3. N2O is now also the main cause of

stratospheric ozone depletion, increasing the risk of skin cancer from UV-B

radiation.

Ecosystems and biodiversity – including the loss of species of high conservation

value naturally adapted to few nutrients. Eutrophication from atmospheric Nr

deposition is an insidious pressure that threatens the biodiversity of many

‘protected’ natural ecosystems.

Soil quality – over-fertilization and too much atmospheric Nr deposition acidify

natural and agricultural soils.

1. Introduction

21

Figure 1-10 The cascade of reactive nitrogen (Nr) forms and associated

environmental problems [133].

So modern emphasis on sustainable development and the known negative side

effects of mineral N-fertilizer have stimulated research in the field of BNF throughout

the past decades. This research has focused mainly on legumes and other symbiotic

systems, which represent a well-studied and widely used method of biological N-

fertilization [114, 134, 135]. However free-living diazotrophs got little attention, as they

mostly incorporate the N they fix into their own biomass and its availability to plant

will be indirect through subsequent mineralization of the biomass [136]. Although there

are many questions concerning their utility remain open. On a global scale annual

nitrogen fixed by associative diazotrophs share up to 30% of the total biologically fixed

N [96]. It can be a significant source in many terrestrial ecosystems [14], however, in

an agricultural context there is a room for more research to explore their potential to

increase crop productivity [96, 135, 137, 138].

As a result of all these beneficial attributes, PGPR should be and have been used

as bio-inoculant all over the world specifically in developing countries [135] as these

could be a better alternative to chemical fertilizer, pesticides (table 1.1) and will help

in organic farming with low production cost.

1. Introduction

22

Table 1.1 Important diazotrophic genera reported for their nitrogen fixation

ability with different crop plants.

Genera Crop

Acetobacter Sugarcane [139],

Achromobacter Sugarcane [140], wheat [141]

Azoarcus Kallar grass [142]

Azospirillum Cereals [143-146], potato [47], grasses [147]

Azotobacter Maize [144]

Bacillus Wheat [148], rice, maize [149]

Burkholderia Rice [150], switchgrass [151]

Enterobacteriaceae Sugarcane [152], rice [146], wheat [149], grasses [147]

Rhizobium Wheat, rice [146, 149, 153]

Herbaspirillum Rice [146, 150], wheat [109]

Pseudomonas Rice, wheat [149], corn [145]

Rhodococcus Rice [146]

1.8 Potato

Potato (Solanum tuberosum L.), belong to the family Solanaceae, is the third most

important food crop in the world after rice and wheat. More than a billion people

worldwide consume potato, and global crop production exceeds 350 million metric tons

(FAO, 2014). China and India produce 1/3 of the total potato produced all over the

world, where Pakistan stand 19th among the top 25 potato producing countries. At the

time of Independence in 1947, potato was cultivated on <3000 hectare with the

production of <30000 tonnes [154]. However an accelerated increase was observed

both in its production and area under cultivation with a production of 3.8×106 tonnes

with an area under cultivation of 1.74×105 ha (FAO, 2014) and it became the significant

source of rural income (worth some $300 million in 2005). Potato use per capita is

increasing with passing year in Pakistan as shown in the figure 1-11. Potato is grown

in all the irrigated areas of Pakistan, However Punjab shares 80% of the total production

of potatoes. Areas of Okara, Sahiwal, Jhang, Gojra, Sheikhupura and Gujranwala are

known for potato cultivation in Punjab, Pakistan [154].

1. Introduction

23

Figure 1-11 Potato consumption per capita per year in Pakistan.

Potato is a high fertilizer-demanding crop, which requires 250 kg ha-1 of

nitrogen and 150 kg ha-1 of phosphorus to get an optimum yield [155]. These

requirements not only increase the cost of production but also cause severe

environmental problems [156]. A more ecologically-friendly and economical approach

to this problem may lie in the exploitation of the rhizosphere microbiome. PGPR

constitute a largely unexplored biochemical wealth which have a profound role in

biogeochemical cycles and may directly or indirectly impact on the nutrient status of

soil [157].

1.8.1 PGPR Interaction with Potato

The rhizosphere engineering of potato using PGPR still leaves a lot to be desired.

Limited data are as yet available with respect to PGPR colonization, disease

suppression [158, 159] and growth promotion [160] in potato. The reported PGPR from

potato include mostly Azospirillum, Bacillus, Pseudomonas and Rhizobium spp. that

have been used for nitrogen fixation [47], improving phosphorus uptake [37],

production of indole acetic acid (IAA) [47] and biocontrol activity [161, 162] and

induced systemic resistance [163, 164]. However most of these studies are limited to

controlled conditions. Although for metagenomics, soils under potato cultivation are

well characterized both for structural and functional communities especially in The

1. Introduction

24

Netherlands [8-10, 32, 122, 165] but potato rhizosphere is yet to be explored and there

are no data available for Pakistan soils. It is important to explore the local soils for

metagenomics as many abiotic and biotic factors affect the diversity and abundance of

microbial communities in a specific vicinity. So we hypothesized that there will be

potential diazotrophic bacteria which will have the potential to fix atmospheric nitrogen

and can be used as substitute to chemical nitrogenous fertilizer. As these diazotrophs

will be from the local area so there will be more chances of their competency to perform

well. Keeping in mind these facts, the objectives of my study were;

▪ To study the plant root-microbial interaction in potato.

▪ To study the nitrogen fixation potential and biodiversity of the diazotrophs on

the basis of morphological, biochemical and molecular characterization.

▪ To study the colonization potential of selected diazotrophs.

▪ Evaluation of potential diazotrophs under controlled and field conditions.

2. Materials and Methods

2.1 Sample Collection

Soil samples were collected from the rhizosphere of potato plants in plastic bags (25 x

30 cm) from major potato growing areas of Punjab, Pakistan i.e. Gojra, Gujranwala,

Jhang, Okara, Sahiwal, Sheikhupura (Figure 2-1). Samples were kept on dry ice during

transportation to National Institute for Biotechnology and Genetic Engineering

(NIBGE). Each sample was collected in triplicate and finally pooled to get one sample.

The table 2.1 shows the geological information of sampling sites from each location.

Figure 2-1 Geological information of sampling sites.

The dots show the points from where the rhizosphere soil samples were collected.


26

Table 2.1 Geographical information of sampling sites from potato growing areas

Parameters Gojra Gujranwala Jhang Okara Sahiwal Sheikhupura

Geographical

position

31º

05.33

N,

72º

6.62 E

31º

31.94

N,

72º

6.62 E

32º

03.324

N,

74º 7.40

E

32º

03.509

N,

74º

7.259 E

31º

13.712

N,

72º

16.08 E

31º

13.715

N,

72º

16.097 E

30º

57.107

N,

73º

16.66 E

30º

55.472

N,

73º

19.537 E

30º

48.985

N,

73º

11.145 E

30º

37.707

N,

73º

01.203 E

31º

44.883

N,

74º

00.618 E

31º

46.43

N,

74º

01.61 E

Elevation 563 547 223 212 134 141 170 165 164 157 203 203

Agro-climate

condition

Semi-arid

Semi-arid

Humid subtropical

Semi-arid

Semi-arid

Subtropical

Soil temperature

(ºC) 9.3 9.1 11.3 11.7 10.9 10.4 8.1 8.6 9.0 9.4 9.9 10.4


27

2.2 Physiochemical Analysis of Soil Samples

Soil samples were analyzed for physical and chemical properties after drying at 40°C

for 24 h followed by sieving through a 2 mm mesh.

2.2.1 Soil Texture

In 500 mL beaker, 50 g of soil sample was soaked overnight in 40 mL of sodium

hexametaphosphate solution (1%) with 150 mL of distilled H2O. Soil sample was

stirred for 10 min and was shifted to 1 L cylinder. After 40 s of shaking, preliminary

reading was recorded whereas final reading was taken after 2 h with Bouyoucous

Hydrometer. International textural classification system [166] was used to recognize

the soil textural class.

2.2.2 pH of Soil (pH)

Soil paste was prepared by 250 g of soil and distilled H2O and allowed to stand for 1 h.

Then the pH of soil paste was measured by pH meter (JENCO Model-671 P). The pH

meter was calibrated using buffers of pH of 4.1 and 9.2 [167].

2.2.3 Electrical Conductivity (EC) of Soil

Electrical conductivity was recorded from the clear extract, extracted by using vacuum

pump from the soil paste with the help of EC meter (Jenway), following Rhoades, et al.

[168].

2.2.4 Organic Matter

Soil solution was made by 1 g soil sample, 10 mL of 1N potassium dichromate solution,

20 mL of conc. H2SO4, 150 mL of distilled H2O and 25 mL of FeSO4 solutions (0.5 N).

This solution was then titrated with potassium permanganate solutions (0.1 N) to

develop pink color as an end point [169].

2.2.5 Total Nitrogen in Soil

To measure the total N, 10 g of soil was digested with 30 mL of conc. H2SO4 and 10 g

of digestion mixture contained K2SO4: FeSO4: CuSO4 = 10: 1: 0.5, in Kjeldahl’s

digestion tubes. Digested material was then cooled and total volume was made up to

250 mL. An aliquot of 10 mL was used for distillation of ammonia from this, with 4%

boric acid solution and indicator (boromocresol green and methyl red) in a receiver. To


28

increase the pH of the contents NaOH was added to the distillation flask. The material

was titrated against N/10 H2SO4 by Gunning and Hibbard’s method in the receiver

using micro Kjeldahl apparatus after distillation [170].

2.2.6 Extractable Phosphorus

Extractable phosphorous was measured by the method described by Olsen [171] using

spectrophotometer.

2.2.7 Extractable Potassium

To measure the extractable K, 5 g of soil solution was prepared with ammonium acetate

solution (1N) where the volume was made upto 100 ml. After continuous shaking, the

extractable K was determined on Flame Photometer [172] from extracts, obtained from

filtering suspension by Whatman filter paper No. 1.

2.3 Determination of Indigenous Bacterial Population

The bacterial population of rhizospheric soil was determined by serial dilution

technique [173]. One gram of rhizospheric soil was added to 9 mL of 0.89% (w/v) saline

solution following the serial dilution. For general bacterial population, CFU counting

on LB agar [1% (w/v) tryptone, 0.5% (w/v) yeast extract, 0.5% (w/v) NaCl, 1.5 – 2%

(w/v) Agar in 1 L with pH 7± 2] plate was carried out. 100 µL of dilution 3rd, 5th and

7th was spread on LB agar plate. The agar plates were incubated at 30°C for 2-6 days

and colony forming units (CFU/g) were counted.

For diazotrophic bacteria most probable number (MPN) was done on nitrogen

free malate (NFM) semi solid medium [Malic acid 5g, K2HPO4 0.5g, MgSO4.7H2O

0.2g, CaCl2 0.02g, NaCl 0.1g, NaMoO4.2H2O 0.002g, KOH 4.5g, Biotin 10µg, Agar

2g in 1L with pH 7± 2] [174]. 100 µL of suspension from 1st – 5th dilution was

inoculated to 900 µL NFM semi solid in eppendorf in five replicate for each dilution.

Inoculated eppendorfs were incubated for 14 days at 28±2°C and growth was observed

under microscope. The MPN was determined by the eppendorf tubes had culture growth

[175].

2.4 Isolation of Rhizospheric Bacteria

One gram of tightly adhering soil from root was added to 9 mL 0.89% (w/v) NaCl

solution for serial dilution [173]. 100 μL from dilutions, 10-4, 10-5 and 10-6 was spread


29

on LB-agar plates using sterilized glass spreader. The plates were incubated for 48 h at

28±2 ºC. Morphologically different bacterial isolates were selected from each plate and

re-streaked in order to get pure cultures.

2.4.1 Isolation of Nitrogen Fixing Bacteria from Rhizosphere

Soil with roots (0.1 g) was added to 1.5 mL eppendorf tubes containing NFM semi solid

medium [174] for the isolation of nitrogen fixing bacteria and incubated at 28±2 ºC.

Enrichment of cultures was carried out by adding 20 μL from each tube to fresh

eppendorf containing same medium after 48 h and the procedure was repeated 5-6

times. Culture from semi solid medium was then streaked on NFM agar plates and LB

agar plates. Single colonies with different morphological characters were obtained and

purified by re-streaking. Pure bacterial colonies were maintained on both LB and NFM

agar plates for further studies.

2.4.2 Preservation of Bacteria

The bacterial isolates were grown at 28±2 ºC for 24 h on respective media and preserved

in 20% glycerol at -80 ºC.

2.5 Morphological Characterization

Bacterial isolates were streaked on LB agar plates and incubated for 24 h at 28±2ºC for

colony morphology studies. Light microscope (Nikon LABOPHOTO-2, Japan) was

used to observe the shape and motility of bacterial isolates. Method described by

Vincent [176] was followed for Gram’s reaction.

2.6 Biochemical Characterization

2.6.1 Reference Strain

Azospirillum brasilense strain ER20 (Accession no. HE662867) [177] obtained from

the NBRC culture collection NIBGE, Faisalabad, Pakistan, was used as positive control

for in-vitro studies for nitrogen fixation and IAA production.

2.6.2 Nitrogen Fixation

Bacterial isolates were assessed for their nitrogen fixation ability using acetylene

reduction assay (ARA) as described by Hardy, et al. [178]. Each isolate was grown in

NFM semisolid medium for 72 h at 28±2°C and evaluated for nitrogenase activity by


30

gas chromatograph (Thermoquest, Trace GC, Model K, Rodon Milan, Italy) fitted with

Porapak N column and flame ionization detector (FID), following the standard protocol

described by Park, et al. [179]. The nitrogenase activity of the isolates was expressed

as nmoles of ethylene formed per hour per milligram of the protein. Method described

by Bradford [180] was adopted for estimation of protein concentration.

2.6.3 Analysis of Indole Acetic Acid (IAA) Production by Bacterial

Isolates

Indole acetic acid production ability was determined by colorimetric analysis and

HPLC. The test was performed both in the presence and absence of L-tryptophan as

precursor of IAA. One hundred mL LB broth in 250 ml Erlenmeyer flasks was

inoculated with 100 ml of overnight bacterial culture adjusted to optical density 0.6

(107–108 CFU ml-1) measured at 600 nm (Camspec M 350 double beam UV Visible,

UK). The bacteria were grown at 28±2ºC for 72 h with continuous shaking at 150 rpm.

Supernatant was collected by centrifuging at 4000×g for 15 min. Half of the supernatant

(≈50 ml) was filtered through 0.2 mm nylon filters (Millipore, USA). IAA was detected

by mixing 100 mL of Salkowski reagent (1 ml, 0.5 M FeCl3, 30 ml concentrated H2SO4

and 50 ml distilled H2O) with 100 mL of filtered supernatant and allowed to react at

room temperature for 20 min. IAA production was confirmed by pink color

development [181]. The remaining half of the supernatant (≈50 ml) was acidified to pH

2.8 with 1 N hydrochloric acid and extracted three times with equal volumes of ethyl

acetate [182]. The extract was evaporated to dryness, collected in 1 mL ethanol and

passed through 0.2 μm nylon filters (Millipore, USA). The extract was analyzed on

HPLC (Perkin Elmer, USA) at 260 nm, fitted with C-18 column and UV detector, using

methanol (30: 70 v/v) mobile phase at a flow rate of 0.5 mL min-1.

2.6.4 Phenotypic Microarrays

Metabolic potential of the bacterial isolates was evaluated by using BIOLOG GN2

microplates [183]. Strains were grown on LB agar plates for 48 h at 28±2 ºC. Culture

was then harvested from the surface of agar plates in 1.5 mL eppendorf containing 1

mL of DEPC water and starved for 3 h. The culture was then mixed with inoculation

fluid IF-0a and redox indicator as per the manufacturer’s instruction. 100 µL was added

to each of the 96 wells in the carbon utilization plate, PM2A (Biolog, Hayward, CA).

The plates were incubated at 28±2 °C for 24 h and observed on VERSA max micro-


31

plate reader (Molecular Devices, USA) with softmax pro-software for qualitatively as

well as quantitative analysis (Line et al., 2011).

2.7 Molecular Characterization

2.7.1 Extraction of Genomic DNA from Bacterial Isolates

Ultra-clean microbial DNA isolation kit (MoBio, CA) was used for the extraction of

DNA from pure cultures. Bacterial isolates were grown in LB broth at 28±2 °C for 48

h and centrifuged at 10,000 x g to get the bacterial cell pellet. Genomic DNA was then

extracted according to the manufacturer protocol. The DNA was quantified by

ultraspec™ 3100 (OD260, 260/280). Extracted DNA was stored at -20 °C.

2.7.2 Molecular Marker

1 kb ladder (Fermentas, Germany) was used as marker to compare and analyze the DNA

products (Figure 2-2).

Figure 2-2 1kb DNA ladder used to measure the size of the bands.

2.7.3 Analysis of Bacterial Fingerprints using BOX and ERIC PCR

The bacterial isolates with similar colony and cell morphology were differentiated by

using BOX and ERIC PCR. Colony PCR was adopted for both BOX and ERIC PCR.


32

Bacterial cultures were grown on LB agar plates at 28±2 °C for 24 h and single colony

was picked and dissolved in 100 µL of sterilized distilled water in 500 µL eppendorf

and homogenized by vortex to get cell suspension. The cell suspension was incubated

on 98 °C for 5 min for cell lysis. The resulting lysates were used as DNA template in

PCR reactions. BOX PCR was carried out following the protocol described by Gevers,

et al. [184] whereas for the ERIC PCR method used by Rasschaert, et al. [185] was

employed. Primers details, recipe of PCR reactions and thermal conditions for both

BOX and ERIC-PCR were given in table 2.2.

The PCR products were separated on 2 % agarose gel at 100 V for 3 h in 1X

Tris-EDTA (TAE) buffer (20mM Tris, 10 mM acetate, 0.5 mM EDTA, pH 8.0). The

20 µL of PCR product was mixed with 5 µL of 6X loading dye [0.05% of bromophenol

blue, 0.25% of xylene cyanol FF, 40% (w/v) of sucrose in water] onto the gel. 5 µL of

1 kb ladder (Fermentas, Germany) was used as marker to compare the banding pattern.

The gel was stained with ethidium bromide (5%) and observed under UV light and

photographed using gel documentation system (Vilbour Lourmat, France).

2.7.4 Identification of Bacterial Isolates using 16S rRNA Sequence

Analysis

The 16S rRNA of the bacterial isolates was amplified by PCR using the primers 968F

[186] and 1406R [187]. A total volume of 50 µL was used for amplification of 16S

rRNA gene following PCR conditions given in table 2.2. PCR product was analyzed by

1% (w/v) agarose Tris-acetate-EDTA (TAE) gel electrophoresis and visualized under

UV light after 5% Ethidium Bromide staining.

2.7.5 Amplification of NifH Gene

A nested PCR was conducted for the amplification of nifH gene form the pure cultures

[122]. A 25 µL reaction was prepared using primer set FGPH19 [188] and PolR [189].

The PCR product of the 1st PCR was used as template in the second PCR with primers

set of PolF and AQER [189]. PCR conditions used for both the PCRs are explained in

table 2.2. The product was analyzed on 1% (w/v) agarose TAE gel, stained with

ethidium bromide with 1 kb ladder (Fermentas, Germany).


33

2.7.6 DNA Sequencing and Sequence Analysis

The PCR products were purified using Wizard® SV Gel and PCR Clean-up System

(Promega) following the manufacturer instruction and sent for sequencing to LGC

Genomics, Berlin, Germany. The 16S rRNA gene and nifH gene sequences were

analyzed using Sequence scanner software package and phylogeny was determined by

BLASTn technique. The sequences were submitted to GenBank EMBL.

2.7.7 Phylogenetic Analysis

The software package MEGA6 was used for phylogenetic analysis. The sequences of

the isolates were compared and analyzed using alignment tool CLUSTAL W, by

downloading the closely related sequences from NCBI data base [190]. Phylogenetic

analysis were carried out using maximum likelihood (ML), however the bootstrap

values of 70% or greater were maintained to represent well supported nodes [191].


34

Table 2.2 PCR and thermal conditions for BOX-PCR and ERIC-PCR analysis,

16S rRNA and nifH genes.

Primers BOX-PCR (5’-3’) PCR mixture Thermal conditions

GTG5

(GTGGTGGTGGTGGTG)

1x buffer (Roche),

2.6 µL dNTPs (25mM),

5 µL DMSO (100%),

0.8 µL BSA (20mg/ml),

0.5μM each primer,

0.8U Taq polymerase (Roche).

95°C, 2 min

94°C 4 s, 92°C 30 s,

50°C 1 min 65°C 8 min,

35 cycles

final extension 65ºC 16

min

Primers ERIC-PCR (5’-3’) PCR mixture Thermal conditions

ERICR

(ATGTAAGCTCCTGGGGAT

TCAC)

ERIC2

(AAGTAAGTGACTGGGGTG

AGCG)

1x buffer (Roche),

0.5 µL dNTPs (10mM),

2.5 µL MgCl2 (50 mM),

0.5μM each primer,

0.16U Taq polymerase (Roche).

95°C, 1 min

95°C 1 min, 45°C 1 min,

72°C 1 min,

35 cycles


min

Primers 16S rRNA (5’-3’) PCR mixture Thermal conditions

968F

(GCACGGGGGGAACGCGA

AGAACCTTAC)

1406R

(ACGGGCGGTGTGTRC)

1X buffer (Bioline),

3.5 µL MgCl2 (50 mM),

0.4 µL dNTP’s (25 mM),

0.5 µL formamide, 0.05 µL T4,

0.5μM each primer,

0.1U Taq polymerase (Bioline).

95°C, 5 min

94°C 1 min, 57°C 1 min,

72°C 2 min

35 cycles


min

Primers nifH (5’-3’) PCR mixture Thermal conditions

FGPH19

(TACGGCAARGGTGGNATH

G)

PolR

(ATSGCCATCATYTCRCCG

GA)

PolF

(TGCGAYCCSAARGCBGAC

TC)

AQER

(GCCATCCATCTGTATGTC

CA)

0.20mM dNTPs, 1x buffer

(Roche),

0.01mg BSA (20mg/ml),

0.5μM each primer,

0.5U Taq polymerase (Roche)

0.25mM dNTPs, 1x buffer

(Roche),


0.5μM each primer,

0.8U Taq polymerase (Roche)

94°C 5 min,

94°C 60 s, 56°C 1 min,

72°C 2 min

30 cycles


min

94°C 5 min,

94°C 60 s, 48°C 1 min,

72°C 2 min

30 cycles


min


35

2.8 Plant Inoculation Studies

2.8.1 Preparation of Seeds

The medium sized potato tubers (2-3 cm), had minimum two growth buds, were surface

sterilized by 8% (v/v) sodium hypochlorite for 10 min followed by thorough washing

with sterilized distilled water.

2.8.2 Preparation of Bacterial Inoculum

The selected bacterial strains were grown in 250 Erlenmeyer flasks containing 100 mL

of LB broth with constant shaking at 180 rpm at 28±2°C upto 109 CFU mL-1. Bacterial

cultures were then centrifuged, at 8000 x g and washed by 0.89% (w/v) saline and again

centrifuged for pelleting. The bacterial cells pellet was re-suspended in equal amount

of 0.89% saline and sterilized potato tubers were immersed in bacterial inoculum for

30 min and sown.

2.8.3 Growth Room Experiments

Two pot experiments were conducted under control conditions (one in sterilized sand

and other in sterilized soil culture) to determine the potential effect of bacterial isolates

on the growth of potato plants. The Kuroda cultivar of potato was used as inoculation

in these studies. Experiments were laid in controlled conditions; 25°C temperature and

light and dark period of 16/8 with photon flux density of 400 mol m-2 S-1 in completely

randomized design (CRD) with 3 replicates of each treatment. Inoculated potato seeds

were sown into pots. The bacterial inoculum was again applied to potato roots after 7

days of sowing with sterilized syringe @ 5ml/plant. Growth parameters like shoot

length, root length, shoot fresh weight, root fresh weight, shoot dry weight, root dry

weight and N contents of plants was recorded after 60 days after sowing. Bacterial

population in potato rhizosphere at different time intervals was also recorded by serial

dilution plating [173].

2.8.4 Experiment 1: Potato Plant Inoculation in Sand Culture

Bacterial isolates were tested for their plant growth promoting effects on potato plants

in sterilized sand culture. There were 15 treatments each with 3 replicates, 2 un-

inoculated control treatments; positive control (recommended full dose of NF) and one

negative control (N0 = without nitrogen) and 13 inoculated treatments (Inoculated with

bacterial isolates + N0). Sand sterilized by soaking in 0.5 N nitric acid for 24 h then


36

washed to remove acid, air dried and autoclaved. Pots were filled and placed in growth

room (16/8 h light/dark and 20/8ºC day/night temperature). Hoagland’s solution [192]

without N, (1/2 strength) was applied at the rate of 2 mL per pot. Data was taken on

plant growth parameters and analyzed to select best performing isolates for further

studies after 60 days after sowing (DAS).

2.8.5 Experiment 2: Potato Plants Inoculated in Soil Culture

This experiment was conducted to estimate the nitrogen fixation potential of the

bacterial isolates and their effects on potato plant growth parameters. Three bacterial

strains which performed best in sand culture were inoculated to potato plants in

sterilized soil. (Sandy loam, EC 2.5 d S m-1, pH 8.1, OM 0.6% and total N 0.06%). Soil

was air dried, grinded and sieved (2 mm). Soil was the put at 60°C for 48 h and

autoclaved at 121°C for 30 min. Soil was mixed thoroughly and autoclaving process

was repeated 5 times. Each pot contained 8 kg of soil, had one medium size surface

sterilized potato with more than 2 growth buds. Pots were placed in growth room (16/8

h light/dark and 20/8ºC day/night temperature). Half dose of recommended 15N labelled

ammonium sulphate (5% abundance) was applied in solution 3–5 cm below the soil

surface by sterilized syringe and full dose of P was applied (mixed with the soil before

sowing). Sterilized distilled water was used for irrigation of plants. There were 4

replicates per each treatment. Plants were harvested 60 DAS and data was taken on

growth parameters and plant N contents.

2.9 Analysis of Plant Samples

2.9.1 Estimation of Total N in Plant Samples

The plant material were oven dried at 60°C for 48 h and cut into small pieces and ground

in a stainless steel grinding mill. The total N in plants was determined by Kjeldahl

method. The plants were first mixed with 2-2.5 g digesting mixture containing K2SO4,

CuSO4 and Se (100: 10: 1) and 5-7 ml of concentrated H2SO4 was added and digested

(Digestion system 40, Digester 1016, Tecator). The digested material was steam

distilled in the presence of concentrated NaOH solution and NH3 evolved was collected

into 2% boric acid solution. The ammonium-boric acid complex was titrated against

standard H2SO4 solution to calculate the total N of the solution. Following equation was

used;


37

N (mg g-1) = acid used (blank sample) × 14 × normality of acid used for titration

wt. of sample

2.9.2 Estimation of 15N Abundance in Potato Plants

Titrated solution, obtained after estimation of total N in plant samples, was acidified by

adding few drops of standard H2SO4. All the samples were concentrated to a volume of

5 mL by evaporation of water and analyzed for 15N by Rittenburg method on a mass

spectrometer fitted with double inlet system. Sodium hypobromite was used for

releasing of 15N [193]. Nitrogen fixation was calculated on the basis of % 15N a.e

(atomic excess) as described by Malik, et al. [194]. Below are the equations used,

Natural abundance: %Ndfa = (δ15NNFS - 15NFS / δ15NNFS) × 100

15N isotope dilution method: %Ndfa = (1 - % 15NFS / % 15NNFS) × 100

Where %Ndfa = % nitrogen fixation, δ15N = natural abundance of 15N, FS = fixing

system and NFS = non fixing system.

2.10 Field Experiments

Selected bacterial strains were further tested for their effectiveness and growth

promotion under field conditions. Experiments were carried out in potato growing

season 2012-2013 where seed of potato cultivar “Kuroda” was used.

2.10.1 Soil Analysis and Bacterial Population

The composite soil samples were taken (0-15 cm depth) before sowing from field area.

The data was taken on soil physicochemical properties and bacterial populations as

describe above in section 2.2 and 2.3 respectively.

2.10.2 Locations of Field Experiments

Location 1: National Institute for Biotechnology and Genetic Engineering (NIBGE),

Faisalabad.

Location 2: Potato Research Institute (PRI), Sahiwal.


38

2.10.3 Layout Plan of Experiments

Randomized complete block design (RCBD) was used with six treatments and four

replicates for each treatment, in field experiments. The treatments included three

control (un-inoculated) and three inoculated treatments as follow;

T1 = Un-inoculated + N0 (-ve control treatment)

T2 = Un-inoculated + N1/2 (+ve control treatment)

T3 = Un-inoculated + NF (+ve control treatment)

T4 = Inoculated with strain TN03 + N1/2



2.10.4 Agronomy of Crop

Field was irrigated well two weeks before sowing in order to ensure the provision of

ample moisture, required for the germination of seed tubers. Soil was cultivated 3-4

times by ploughing followed by planking to prepare field. Potato tubers were dipped in

bacterial inoculum for 30 min before sowing for inoculation purpose. Manual sowing

was carried out on ridges. Each plot size was 18.9 m2 (3.6 m × 5.25 m) where R×R

distance was 75 cm and P×P distance was 30 cm. Nitrogen (as urea) was applied (full

dose = 250 Kg ha-1, Half dose = 175 Kg ha-1) in two splits, half at the time of sowing

and half at first irrigation, by broadcasting. Phosphorus (as single super phosphate) and

Potassium minerals (as muriate of potash) were applied @ 150 kg ha-1 each at the time

of sowing. All the other agronomic practices, plant protection measures (weed, insect,

pest and disease control) was kept similar for all the treatments.

2.10.5 Parameters Studied

Data was taken on following growth parameters;

1. Plant height (cm)

2. Number of branches plant-1

3. Number of compound leaves plant-1


39

4. Number of tubers plant-1

5. Plant fresh weight (g)

6. Plant dry weight (g)

7. Tuber fresh weight (g)

8. Tuber dry weight (g)

9. Number of tubers per plant

10. Tuber yield (Kg ha-1)

11. Plant N contents (mg g-1)

2.11 Root Colonization Studies

2.11.1 Ultra-structure Studies using Transmission Electron

Microscopy (TEM)

The root colonization ability of isolates was studied by inoculating the bacterial isolates

to the potato plants and observations under TEM. The surface sterilized potato tubers

variety Kuroda, were inoculated by selected bacterial inoculum. The inoculated

potatoes were grown in sterilized sand culture as described in section 2.8. Plants were

harvested after 30 days of sowing. Roots were washed with sterilized water and cut

with sterilized cutter into pieces of approximately 1-3 cm. the cut roots were embedded

in 1.5% (w/v) water agar in form of cubes of approximately 2-3 mm3. The cubes were

put in the 2 mL eppendorf containing fixative; 5% gluteraldehyde dissolved in 0.2 M

PIPES buffer [0.58 g NaCl, 3 g PIPES, 1 M NaOH, 0.2 g MgCl2.6H2O, pH 6.8] with

pH 8.0. The fixative was replaced with 0.2 M buffer after 16-18 h. The samples were

kept/dipped in PIPES buffer for 16-18 h [195], then washing was done in fresh buffer

for 1-2 h. Samples were treated with 0.2% osmium tetraoxide, dissolved in PIPES

buffer for 16–18 h and again washed twice for 30 min with sterile distilled water. After

being treated with 5% aqueous uranyl acetate for 16–18 h, the samples were washed

twice with sterile distilled water for 30 min. The samples were then immersed twice in

absolute ethanol for 30 min, followed by immersion in propylene oxide (100%) for 30

min for dehydration. Infiltration of samples was carried out with propylene oxide at a

ratio of 1:1 for 24–48 h and then with spur resin for a further 24–72 h, using benzyl


40

dimethyl amine as the accelerator in all infiltration steps. The samples were

polymerized for 72 h at 70ºC on flat embedded molds, followed by incubation at room

temperature for at least 24 h before cutting. Sections (150–200 nm) were cut using an

ultra-microtome (RMC-7000; Boeckeler Instruments, USA) and carefully placed on

copper grids. Sections were then double-stained with uranyl acetate (30 min) and lead

citrate (10min), washed with deionized water and observed under transmission electron

microscope (TEM; JEOL JEM1010, USA).

2.11.2 Root Colonization Studies using Confocal Laser Scanning

Microscope (CLSM)

Preparation of Electro-competent Cells

Electro-competent cells of selected bacterial strains were prepared, following

methodology of Wu, et al. [196]. Selected bacterial strains were grown in LB broth with

continuous shaking at 180 rpm at 30±2°C for 24 h to obtain the desired CFU of bacteria

i.e. 1×108. Chilled falcon tubes (50 mL) containing 40 mL of each bacterial culture

was pelleted by centrifugation at 6000 × g for 15 min at 4°C. The cells were re-

suspended very gently in 20 mL of chilled 10% (v/v) glycerol. The pelleting and re-

suspending process was repeated and the volume of 10% (v/v) glycerol was reduced

gradually (15 mL, 10 mL, 5 mL for 2nd, 3rd and 4th washing respectively). The cells

were finally re-suspended in 250 µL of 10% glycerol. Aliquots of 50 µL of cell

suspension were flashed frozen and stored at -80°C.

Transformation of Bacterial Strains with Yellow Fluorescent Protein (YFP)

Plasmid DNA, containing pBBRIMCS-4 vector, was isolated from E. coli strain DH5a.

It was a 4.95 kb vector with broad range ampicillin resistant along with the cassette of

yellow fluorescence protein (YFP) [197], shown in figure 2-3. Plasmids were isolated

from 1-5 mL of overnight grown culture using QIAGEN QIA Miniprep kit, following

the standard protocol provided by the manufacturer. The concentration of plasmid was

checked by UltraspecTM 3100 at OD 260, 260/280. Electro-poration was carried out on

Gene Probe (200 Ω resistor, 12.5 KV cm-1, 25 µF capacitor) with isolated plasmid DNA

and electro-competent cells [196]. Selection of the transformed colonies was done by

spreading 100 µL on LB ampicillin agar plates containing the 50 µg mL-1 of ampicillin.

The grown colonies were further confirmed by observing the cells of bacteria under

CLSM on glass slide yfp filter. The transformed cells of bacterial isolates were


41

preserved in 20 % (v/v) glycerol at -80 °C. The transformed bacterial strains were

grown in 100 ml of LB ampicillin (50 µg/mL) broth for 24 h at 28±2 °C upto 1×108

CFU, and harvested by centrifugation at 8000 x g and re-suspended in 0.89 % (w/v)

NaCl. Medium size potato tubers were surface sterilized (as described in section 2.8.1)

and inoculated with yfp-transformed strains. Inoculated potato tubers were grown in

sand (soaked in 0.5 N nitric acid for 24 h then washed, dried and autoclaved) culture

for 30 days. Roots were cut, washed with a sterilized blade in to ½ cm pieces and

observed under CLSM (Olympus fluoview Ver. 1.3), on glass slide with sterilized water

covered with cover slip, to detect the colonization of yfp-labelled bacterial strains [198].

Figure 2-3 Restriction map of vector pBBR1MCS-4.

2.12 Metagenomics Studies

Metagenomic approach was adopted to study the both culturable and non-culturable

bacterial community in the rhizosphere of potato. Bacterial diversity was evaluated

using denaturing gradient gel electrophoresis (DGGE) and bacterial abundance was

quantified by qPCR of 16S rRNA and nifH genes.


42

2.12.1 Rhizospheric Soil DNA Extraction

Soil DNA was extracted by PowerSoil DNA extraction kit (Mo Bio Laboratories Inc.,

NY). Soil sample (0.5 g) was collected from the rhizosphere of potato plants and soil

DNA was extracted following the manufacturer’s protocol, with little modifications as

glass beads of 0.1 mm diameter @ 0.25 g were added to the soil. The cells were broken

three time for 1 min using (BioSpec Products, USA). The quantity and purity of

extracted DNA was checked out by running it on agarose gel (1.5% w/v) at 80 V for 90

min in 1X TAE buffer (20 mM Tris, 10 mM acetate, 0.5 mM EDTA; pH 8.0) against 5

μL of 1-kb DNA molecular size marker (Promega, Leiden, Netherlands). DNA quality

was determined after dyeing with ethidium bromide, on the basis of degree of DNA

shearing (average molecular size) and the amounts of co-extracted compounds.

Quantification of extracted DNA was done using Nanodrop 2000 (Thermo Fisher

Scientific).

2.12.2 DGGE Analysis for Microbial Diversity

PCR Amplification of 16S rRNA Gene for DGGE Analysis

16S rRNA gene was amplified using the forward primer F968 [199] with a GC-clamp

attached to 5’ and the universal reverse primer R1401.1b [200]. Primer sequences and

PCR conditions are given in table 2.3. The concentration and size of the PCR products

was determined by running samples in agarose TAE gel (1.5% w/v). Ethidium bromide

was used for staining and PCR products were compared with a molecular weight marker

(Smart ladder; Eurogentec) to confirm the size. A single band of 450bp was amplified

by all DNA samples.

Amplification of NifH Gene for DGGE Analysis

The amplification of nifH genes was carried out by nested PCR as described above in

section 2.7.5 except forward primer PolF, used in second PCR contained a GC clamp

(table 2.3) as used by e Silva, et al. [122]. The concentration of the PCR products was

confirmed by 1.5% (w/v) agarose TAE gel followed by staining with ethidium bromide.

Amplicons were compared with a molecular weight marker (Smart ladder; Eurogentec).

All DNA samples yielded PCR amplification, resulting in one single band of the

expected size (360bp).


43

DGGE System Conditions

DGGE profiles were generated with the Ingeny Phor-U system (Ingeny International,

Goes, The Netherlands). For DGGE analysis of 16S rRNA, about 200 ng of amplicons

were loaded on 45-65% denaturant gradient (100% denaturant corresponded to 7 M

urea and 40% (v/v) deionized formamide).

For nifH PCR products (250-300 ng/ lane) were loaded onto 6% (w/v)

polyacrylamide gels for nifH gene, 1 mm thick, in 0.5X TAE buffer with a 40-65%

denaturant gradient to separate the generated amplicons.

Electrophoresis and Gel Documentation

Electrophoresis was carried out at 60ºC for 16 h at 100 V. 0.5 μg L-1 of SYBR Gold

(Invitrogen, Breda, The Netherlands) in 0.5x TAE buffer was used for staining the gels

and images were taken using Imagemaster VDS (Amersham Biosciences,

Buckinghamshire, UK). GelCompar 6 software package (Applied Mathematics, Sint-

Martens Latem, Belgium) was used for the analysis of resulted fingerprints.

Computer-assisted Analysis of DGGE Fingerprinting

DGGE patterns were compared by Pearson’s correlation coefficient by clustering the

different lanes using the GelCompar 6 software, by the unweighted-pair group method

with arithmetic mean, rolling-disk background subtraction, and no optimization [201].

Range-weighted richness (Rr) values [202] were calculated using the following

equation;

Rr = N2 x Dg.

Where N = number of bands and Dg = the denaturing gradient. Matrix derived

were transformed to square roots and nonmetric multi-dimensional scaling was applied

using PAST software package [203].

2.12.3 Quantitative PCR

Quantification of the Total Bacterial Community

For the abundance of total bacterial community, 16S rRNA gene was quantified using

primers 16SFP and 16SRP [204]. The qPCR assays were done on an ABI Prism 7300

sequence detection system (Applied Biosystems) in 96-well plates, following the PCR

conditions given in table 2.3. Specificity and expected size of amplicons (263 bp) were


44

confirmed by melting curve analysis and products were run in 1.5% agarose gel and

ethidium bromide was used for staining.

Quantification of the Diazotrophic Community

To quantify the diazotrophic community, nifH gene was quantified using the primers

FGPH19 [188] and PolR [189]. Entire quantification of the nifH gene was done on the

ABI Prism 7300 Cycler (Applied Biosystems, Germany) in three replicates, following

PCR mixture and thermal cycling conditions described in table 2.3 [205]. Specificity

and expected size of amplicons (263 bp) were confirmed by melting curve analysis and

products were run in 1.5% agarose gel and ethidium bromide was used for staining.

Standard curves were formed by serial dilutions (107 to 102 gene copy numbers

μL-1) of nifH gene from Bradyrhizobium liaoningense, cloned in plasmid JM 109

(Promega, Madison, WI, EUA) derived from Escherichia coli. Below formula was used

to calculate the efficiency;

Eff = [10(-1/slope) -1]

*Two independent qPCRs were performed and the results were similar for all samples.

Extracted soil DNA was diluted and mixed with known amount of standard DNA before

qPCR to test inhibition in the PCR reactions. No change in Ct values in the presence

diluted soil DNA indicated the absence of severe inhibition.

2.12.4 Pyrosequencing of NifH Gene

Barcoded pyrosequencing approach was used to study the diversity of nitrogen-fixing

bacteria. A nested approach was used to amplify from total community DNA with nifH

gene-specific primers PolF/PolR [189] and RoeschF/ RoeschR [206] by the FastStart

High Fidelity PCR system and PCR Nucleotide Mix (Roche Diagnostics GmbH,

Mannheim, Germany). PCR amplifications were carried out in triplicates on each soil

DNA, following the PCR conditions given in table 2.3 and pooled together. PCR

amplicons were run on agarose gel, required bands were excised and purified by

Qiaquick PCR purification Kit (Qiagen) to remove primer dimers. Sample-specific tags

and adapter sequences were added using normal primers in an additional PCR

amplification of 20 cycles following the same PCR conditions for 454 pyrosequencing.

PCR products were additional refined using AMpure beads (Beckman Coulter) and as

specified by Roche, pooled in an equimolar ratio. Sequencing was performed from only


45

5’ (forward) ends of amplicons. A second-generation pyrosequencer (454 GS FLX

Titanium; Roche) was used for emulsion PCR. Emulsion breaking of DNA-enriched

beads, and sequencing runs of the pooled amplicons were carried out using titanium

reagents and titanium procedures following the protocol suggested by manufacturer.

Data generated from the 454-pyrosequencing have been submitted to the National

Center for Biotechnology Information (NCBI).

2.12.5 NifH Gene Metagenomics Data Analysis

The low quality and failed reads were removed from raw data by automatic amplicon

pipeline of the GS Run Processor (Roche) for quality filtering of the pyrosequencing

reads. The FunGene Pipeline of RDP server

(http://fungene.cme.msu/edu/FunGenePipeline) with default settings was used to

compare the nifH gene libraries. Primer sequences were cut from the resulted sequences

and data was further cleaned from low quality and shorter (<350 bp) reads. Filtered

nucleotide sequences were translated into amino acid and all further analyses were

processed with amino acid sequences. Frame-shifts errors affected by deletion or

insertion of bases can be recognized by targeting a protein-coding gene [207]. Amino

acid sequences containing in-frame stop codon(s) were removed by visually inspection

and sequences were then aligned using MUSCLE 3.8 [208].

DOTUR [209] was used to classify the operational taxonomic units (OTUs) and

to construct the rarefaction curves, using 90% sequence similarity cutoff of amino acids

[210, 211]. Diversity indices and richness estimates were measured for both the total

number of sequences and also for the subsets normalized to the same number of

sequences by the Perl script daisychopper.pl (available at

http://www.genomics.ceh.ac.uk/GeneSwytch/Tools.html; [212]. The representative

sequences for nifH gene were blasted using BLASTP database against a non-redundant

protein sequence.


46

Table 2.3 PCR and thermal conditions for PCR-DGGE analysis and qPCR for

16S rRNA and nifH genes and pyrosequencing for nifH.

Primers DGGE-16S rRNA (5’-3’) PCR mixture Thermal conditions

F968-GC *

(AACGCGAAGAACCTTAC)

R1401.1b

(CGGTGTGTACAAGACCCG

GGAACG)

1X buffer (Bioline),

1.5 µL MgCl2 (50 mM),

1 µL dNTP’s (10 mM),

1 µL DMSO (100%),

0.5 µL BSA,

0.5μM each primer,

0.3U Taq polymerase

(Bioline).

94°C 5 min,

94°C 1 min, 57°C 1 min,

72°C 2 min,

35 cycles


min

Primer qPCR-16S rRNA (5’-3’) PCR mixture Thermal conditions

16SFP

(GGTAGTCYAYGCMSTAAACG)

16SRP

(GACARCCATGCASCACCTG)

12.5μl Power Sybr Green

PCR Master mix,

0.5ul BSA (20mg/ml),

0.8μM each primer,

2ul DNA template

95°C 10 min, 1 cycle

95°C for 27 s, 62°C for 1

min,

72°C for 30 s, 39 cycle

Primers DGGE-nifH (5’-3’) PCR mixture Thermal conditions

FGPH19

(TACGGCAARGGTGGNATHG)

PolR

(ATSGCCATCATYTCRCCGGA)

PolF-GC *

(TGCGAYCCSAARGCBGACTC)

AQER

(GCCATCCATCTGTATGTCCA)

0.20mM dNTPs,

1x buffer (Roche),


0.5μM each primer,

0.5U Taq polymerase

(Roche),

0.25mM dNTPs,

1x buffer (Roche),


0.5μM each primer,

0.8U Taq polymerase

(Roche)

94°C 5 min,

94°C 60 s, 56°C 1 min,

72°C 2min

30 cycles


min

94°C 5 min,

94°C 60 s, 48°C 1 min,

72°C 2 min

30 cycles


min

Primers qPCR-nifH (5’-3’) PCR mixture Thermal conditions

FGPH19

(TACGGCAARGGTGGNATHG)

PolR


12.5μl Power Sybr Green

PCR

Master mix,

0.5ul BSA (20mg/ml),

0.25μM each primer,

2ul DNA template

95°C 10 min, 1 cycle

94°C for 60 s, 55°C for 27

s,

72°C for 60 s, 40 cycle

Primers pyrosequencing-nifH (5’-3’) PCR mixture Thermal conditions

PolF

(TGCGAYCCSAARGCBGACTC)

PolR


RoeschF

(ACCCGCCTGATCCTGCACGCCA

AGG)

RoeschR

(ACGATGTAGATTTCCTGGGCCT

TGTT)

0.20mM dNTPs,

1x buffer (Roche),


0.5μM each primer,

0.25U FastStart High

Fidelity PCR System,

50ng template DNA,

0.20mM dNTPs,

1x buffer (Roche),


0.5μM each primer,

0.25U FastStart High Fidelity

PCR System,

50 ng template DNA

95°C 5 min,

94°C 45 s, 50°C 45 s,

72°C 45 s,

20 cycles


min

95°C 5 min,

94°C 45 s, 50°C 45 s,

72°C 45 s

20 cycles


min

*GC clamp according to Muyzer [213].

3. Results

Part 3.1 Bacterial Isolation and Characterization

3.1.1 Physico-chemical Characteristics and Bacterial Population

Analysis of Soil Samples

Rhizospheric soil samples of potato plants were collected from the major potato

growing areas of Punjab, Pakistan, including Gojra, Gujranwala, Jhang, Sahiwal and

Sheikhupura (table 3.1). EC value ranged from 0.6 to 3.7 d S m-1 in Sahiwal and

Sheikhupura soils respectively. Overall pH of soil was alkaline with a pH range of 7.8

to 8.4 in different areas. Maximum organic matter content 1.84% was observed in

Sheikhupura soils while Okara soils had minimum 0.77% organic contents. Total

nitrogen was found in the range of 0.05–0.13%. Total bacterial population (CFU) and

population of diazotrophs (MPN) per gram of soil is represented in Figure 3-1.

3.1.2 Bacterial Isolation and Morphological Characterization

Forty four bacteria were isolated from six different sites. Details of morphological

characteristics of all the isolates is given below in table no. 3.2. Eight isolates (TN01-

TN08) were obtained from Gujranwala, six isolates (TN09-TN14 and TN15-TN20)

were purified each from Jhang and Gojra respectively, seven (TN21-TN27) from Okara

region, nine from Sheikhupura (TN28-TN36) and eight (TN37-TN44) from Sahiwal

soils. These isolates exhibited different colony morphologies (table 3.2). Most of the

bacteria were rods and short rods while some were vibroid.

3. Results

48

Table 3.1 Physico-chemical properties of soil samples obtained from potato rhizosphere, from different potato growing areas of Punjab.

Parameters Gojra Gujranwala Jhang Okara Sahiwal Sheikhupura

Gr1 Gr2 Gj1 Gj2 Jg1 Jg2 Ok1 Ok2 Sw1 Sw2 Sk1 Sk2

EC (d S m-1) 1.01 0.92 2.3 1.8 1.2 1.2 3.3 0.9 1 0.6 3.2 3.7

Soil pH 8.4 8.2 8.2 7.8 8.3 8.4 8.2 8.3 8.3 8.4 8 8.1

Soil texture Sandy

Loam

Sandy

Loam Loam Loam

Sandy

Loam

Sandy

Loam

Sandy

Loam

Sandy

Loam Loam Loam Loam Loam

Organic matter

(%) 1.39 1.14 1.15 1.04 0.87 1.32 1.08 0.77 1.04 1.53 1.84 1.11

Total N (%) 0.13 0.1 0.09 0.08 0.08 0.08 0.05 0.07 0.07 0.06 0.1 0.08

Total mineral

N (mg kg-1) 10.2 9.31 8.11 6.02 5.61 6.13 5.91 6.75 4.13 5.31 8.29 7.27

Total P

(mg Kg -1) 59.8 63.3 96.3 98.9 62.5 75.6 51.6 27.6 50.6 29.3 97.7 115.7

Total K

(mg Kg -1) 360 260 140 260 140 560 240 200 220 140 340 320

Saturation (%) 30 32 34 36 30 26 30 20 36 30 32 31

3. Results

49

Figure 3-1 Total bacterial population (CFU) of bacteria and diazotrophs (MPN)

in the rhizospheric soil samples from potato.

CFU recovered on LB while MPN in NFM vails. Value are average of 3 replicates each.

Bar represents standard deviation of replicates.

Table 3.2 Morphological characteristics of bacterial isolates purified from

rhizosphere of potato.

Isolate

code Isolation site Colony morphology

Cell

morphology

Gram’s

reaction

TN01 Gujranwala Small, wrinkled, pinkish Rods -ve

TN02 Gujranwala Small, round, white creamy Rods -ve

TN03 Gujranwala Small, round, white Vibroid -ve

TN04 Gujranwala Small, round, white Short rods -ve

TN05 Gujranwala Medium, round, yellow Rods -ve

TN06 Gujranwala Medium, round, creamy Short rods -ve

TN07 Gujranwala Medium, irregular, brownish rods +ve

TN08 Gujranwala Small, round, orange Vibroid -ve

TN09 Jhang Small, round, white Vibroid -ve

TN10 Jhang Medium, round, pinkish Rods -ve

TN11 Jhang Medium, round, yellow Rods +ve

TN12 Jhang Small, irregular, pinkish Vibroid -ve

TN13 Jhang Small, round, creamy Round -ve

TN14 Jhang Medium, round, white creamy Short rods -ve

TN15 Gojra Small, round, transparent Rods -ve

TN16 Gojra Medium, round, orange Rods -ve

TN17 Gojra Small, round, pale yellow Short rods -ve

TN18 Gojra Small, round, brownish Short rods -ve

TN19 Gojra Medium, round, creamy Rods +ve

TN20 Gojra Small, round, white Vibroid +ve

TN21 Okara Small, irregular, yellowish Rods -ve

TN22 Okara Small, irregular, pinkish Vibroid -ve

TN23 Okara Medium, irregular, yellowish Rods -ve

TN24 Okara Small, irregular, grayish Short rods -ve

TN25 Okara Medium, round, brownish Short rod -ve

TN26 Okara Medium, round, yellowish Short rods -ve

3. Results

50

TN27 Okara Small, round, yellowish Rods -ve

TN28 Sheikhupura Small, round, pinkish Vibroid -ve

TN29 Sheikhupura Small, irregular, whitish Rods -ve

TN30 Sheikhupura Medium, round, whitish Vibroid -ve

TN31 Sheikhupura Small, irregular, orange Vibroid -ve

TN32 Sheikhupura Medium, smooth, brownish creamy Rods +ve

TN33 Sheikhupura Medium, smooth, greenish yellow Rods +ve

TN34 Sheikhupura Large wavy, dark, creamy Rods +ve

TN35 Sheikhupura Medium, round, creamy Short rods -ve

TN36 Sheikhupura Small, smooth, creamy Short rods -ve

TN37 Sahiwal Medium, round, pale yellow Short rods -ve

TN38 Sahiwal Medium, round, white Short rods -ve

TN39 Sahiwal Medium, round, creamy Short rods -ve


TN41 Sahiwal Small, round, white Short rods -ve


TN43 Sahiwal Small, smooth, creamy Short rods -ve

TN44 Sahiwal Medium, irregular, grayish Short rods -ve Bacterial isolates were obtained on NFM medium and observed on LB medium.

3.1.3 Analysis of Plant Growth Promoting (PGP) Traits In vitro

Nitrogenase Activity

Nitrogenase activity of bacterial isolates was estimated by acetylene reduction assay.

Six bacterial isolates from Gujranwala, five from Jhang, four from Gojra, four from

Okara, Seven from Sheikhupura and six from Sahiwal showed nitrogenase activity

(table 3.3). Nitrogen fixation among the bacteria ranged from 69.0-151.7 nmoles mg-1

protein h-1. Isolate TN04 from Gujranwala showed maximum nitrogen fixation (151.70

nmoles mg-1 protein h-1). Twelve isolates showed nitrogenase activity below 50 nmoles

which were regarded as non-nitrogen fixer (table 3.3).

3. Results

51

Table 3.3 Nitrogenase activity of bacterial isolates purified from potato growing

areas of Punjab.

Isolate

name

Isolation

site

ARA (nmoles

mg-1 protein h-1)

Isolate

name

Isolation

site

ARA (nmoles

mg-1 protein h-1)

TN1 Gujranwala 130.47±16.35 TN23 Okara 84.94±11.56

TN2 Gujranwala 135.42±13.3 TN24 Okara 0




TN6 Gujranwala 0 TN28 Sheikhupura 85.97±8.03


TN8 Gujranwala 127.80±14.45 TN30 Sheikhupura 98.79±11.04

TN9 Jhang 143.07±11.98 TN31 Sheikhupura 124.22±7.43



TN12 Jhang 102.30±7.08 TN34 Sheikhupura 0


TN14 Jhang 0 TN36 Sheikhupura 0

TN15 Gojra 91.92±8.29 TN37 Sahiwal 135.07±8.97


TN17 Gojra 0 TN39 Sahiwal 127.56±11.63

TN18 Gojra 100.78±7.7 TN40 Sahiwal 0



TN21 Okara 0 TN43 Sahiwal 95.99±12.66

TN22 Okara 127.21±12.07 TN44 Sahiwal 93.36±6.38 NA = No activity/ <50 nmoles

The nitrogenase activity was detected by acetylene reduction assay. Bacterial isolates

were inoculated to NFM medium. The data is mean of 3 replicates. ± shows the standard

deviation of the replicates.

Production of Indole-3-acetic Acid (IAA)

Twenty two isolates showed the production of pink color by colorimetric method

indicating IAA production. Quantification of IAA was done on HPLC (table 3.4). The

in vitro synthesis of IAA in isolates ranges from 30.43 - 1.35 µg mL-1. Among the IAA

producing isolates, maximum IAA production was observed by isolated TN03 which

belonged to Gujranwala while minimum was by isolate TN11 which belonged to Jhang.

Fifteen isolates did not show IAA production.

3. Results

52

Table 3.4 Quantification of IAA produced by bacterial isolates in LB medium

supplemented with L-Tryptophan.

Isolate

name Isolation site

IAA

production

(µg mL-1)

Isolate

name Isolation site

IAA

production

(µg mL-1)








TN8 Gujranwala 12.49±0.6 TN30 Sheikhupura 11.32±0.51



TN11 Jhang 1.35±0.32 TN33 Sheikhupura 0



TN14 Jhang 0 TN36 Sheikhupura 1.51±0.26







TN21 Okara 0 TN43 Sahiwal 0

TN22 Okara 8.67±0.7 TN44 Sahiwal 0

The IAA quantification was done by HPLC. Bacterial isolates were inoculated to LB

medium supplemented with tryptophan. Data was taken after 72 h of inoculation. The

data is mean of 3 replicates. ± shows the standard deviation of the replicates.

3. Results

53

3.1.4 Phenotypic Microarray Analysis

Phenotypic microarray of nine morphologically different isolates was carried out

through BioLog PM2A microplate system represented that isolate TN19 from Gojra

metabolize maximum, 80 out of 95 different carbon sources which were evident from

the conversion of colorless tetrazolium dye to violet formazan. Different bacterial

isolates showed differential ability to utilize a number of substrates of different nature.

Isolate TN03, TN37 and TN42 utilized only 9 type of substrates while isolate TN9

utilize 10 substrates, however nature of the carbon sources varied from isolate to isolate.

Isolate TN1, TN4, TN14 and TN18 consumed 52, 28, 55 and 54 carbon sources

respectively (table 3.5).

Table 3.5 Differential metabolic profiling of bacterial isolates associated with

potato roots (Biolog PM2A Microplate analysis)

Carbon Source TN

01

TN

03

TN

04

TN

09

TN

14

TN

18

TN

19

TN

37

TN

42

Chondroitin

Sulfate C - - + - - - + - -

α-Cyclodextrin + - - - - - + - -

β-Cyclodextrin + - + - - - + - -

γ-Cyclodextrin - - - - - - + - -

Dextrin + + - - + - + - -

Gelatin + - - - - - + - +

Glycogen - + - - + - + + -

Inulin - - - - + - + - -

Laminarin + - + - - - + - -

Mannan - + - + + - + + -

Pectin - - - - - + + + -

N-Acetyl-D-

Galactosamine - - - - - + - - -

N-Acetyl-

Neuraminic Acid - - - - - - + - -

β-D-Allose - - + - - + + - -

Amygdalin - - - - + - + - -

D-Arabinose - - + - + + + + +

D-Arabitol + - - - + + - - -

L-Arabitol - - - - + + + - -

Arbutin + + - + + + - - -

2-Deoxy-D-

Ribose + - - - - - + - +

i-Erythritol + - - - - - + - -

D-Fucose + + - + + + + + +

3-0-β-D-Galacto-

pyranosyl-D-

Arabinose

- - - - - + + - -

Gentiobiose + - - - + + - - -

L-Glucose + - - - - - + - -

Lactitol + - - - + + + - -

3. Results

54

D-Melezitose - - + - + + + - -

Maltitol - - - - + + + - -

α-Methyl-D-

Glucoside - - + - + + + - -

β-Methyl-D-

Galactoside + - + - + + + - -

3-Methyl

Glucose + - + - - - + - -

β-Methyl-D-

Glucuronic Acid + - - - + + - - -

α-Methyl-D-

Mannoside + - + - + - + - -

β-Methyl-D-

Xyloside + - - - + + + - -

Palatinose + - - - + + + - -

D-Raffinose + - + + + + + - -

Salicin - + - - + + - - -

Sedoheptulosan - - - - - - + - -

L-Sorbose + + - + + + + - -

Stachyose + - - - + + + - -

D-Tagatose - - - - + + + - +

Turanose - - - - + + - - -

Xylitol + - + - + + + - -

N-Acetyl-D-

Glucosaminitol + - - - + - + - -

γ-Amino Butyric

Acid - + - - + + + - -

δ-Amino Valeric

Acid - - - - - + + - -

Butyric Acid - - - + + + + - -

Capric Acid + - - - - - + - -

Caproic Acid - - - - - - + - -

Citraconic Acid + - + - + - + - -

Citramalic Acid + - - - - - + - -

D-Glucosamine - + - - + + + + +

2-Hydroxy

Benzoic Acid + - + - - - + - -

4-Hydroxy

Benzoic Acid + - + - - + + - -

β-Hydroxy

Butyric Acid + - + - - + + - -

γ-Hydroxy

Butyric Acid + - + - - - + - -

α-Keto-Valeric

Acid + - - - - - + - -

Itaconic Acid + - - - - - + - -

5-Keto-D-

Gluconic Acid - - + + + + + + +

D-Lactic Acid

Methyl Ester - - + - - - + - -

Malonic Acid + - + - + - + - -

Melibionic Acid + - - + - + + - -

Oxalic Acid - - + - - - + - -

3. Results

55

No bacterial isolate showed utilization of acetamide.

Oxalomalic Acid - - + - + + + + +

Quinic Acid - - + - + + + - -

D-Ribono-1,4-

Lactone + - - - + + + - -

Sebacic Acid + - + - - - + - -

Sorbic Acid + - - - + + + - -

Succinamic Acid + - - - + + + - -

D-Tartaric Acid + - - - + + + - -

L-Tartaric Acid + - + - + + + - -

L-Alaninamide - - + + - - + - -

N-Acetyl-L-

Glutamic Acid - - - - + + - + +

L-Arginine + - - - + + + - -

Glycine + - - - - - + - -

L-Histidine + - - - + + + - -

L-Homoserine + - - - - + + - -

Hydroxy-L-

Proline + - + - + + + - -

L-Isoleucine - - - - - + + - -

L-Leucine - - + - + + + - -

L-Lysine - - - - + + - - -

L-Methionine - - - - - - + - -

L-Ornithine - - - - + + + - -

L-Phenylalanine + - - - - - - - -

L-Pyroglutamic

Acid + - - - + - - - -

L-Valine - - - - - - - - -

D,L-Carnitine - - - - - - + - -

Sec-Butylamine + - - - + - + - -

D.L-Octopamine - - - - + - + - -

Putrescine + - - + + + - - -

Dihydroxy

Acetone - - - - + + + - -

2,3-Butanediol - - - - + + - - -

2,3-Butanone + - - - + + + - -

3-Hydroxy 2-

Butanone + - - - - + + - -

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56

3.1.5 Molecular Characterization

DNA Extraction

The extraction of the genomic DNA for screened isolates was carried out using Ultra

clean microbial DNA isolation kit (MoBio, CA), following the manufacture protocol

and run on 1% agarose gel for confirmation.

ERIC and BOX-PCR

Purified isolates from different sites were compared by ERIC and BOX-PCR

fingerprinting. Banding patterns generated by different isolates after amplification of

genomic DNA with BOX and ERIC primers is shown in figure 3-2 and 3-3 respectively.

The 1 kb ladder was run with the amplified product in order to compare the patterns

among the isolates. The amplified product ranged 250-4000 bp with BOX and <250-

2000 bp with ERIC primers. The finger printing pattern generated by isolates were

compared by using Gel Compare 6 by Jaccard correlation coefficient. PCA was applied

on the data derived on the basis Jaccard correlation (a band-based analysis).

Five clusters were formed by the isolates in dendrogram, formed on the basis of

data generated from BOX-PCR showed that bacterial isolates may be from 5 major

groups. Different groups of bacteria were isolated from each site and mixed clustering

was observed (Figure 3-4). Isolate TN06 and TN07 showed clustering at 100%

similarity, same pattern was found for isolate TN09, TN31 from Jhang and TN28, TN30

from Sheikhupura. All the other isolates showed <90% similarity among them.

Dendogram formed on the basis of ERIC-PCR analysis showed only 3 main

clusters where Isolates TN28, TN29, TN30 (Sheikhupura) and TN31 (Jhang) showed

100% similarity with each other. Isolate TN09 from Jhang clustered at 100% with

TN24. Isolate TN03 (Gujranwala) with TN22 (Okara), isolate TN26 and TN27 from

Okara resulted 100% similar to one another. Whereas isolate TN42 and TN44 from

Sahiwal and Isolate TN36 (Sheikhupura) and Isolates TN40 (Sahiwal) showed 100%

similarity with each other. All the remaining isolates were clustered on <90% with each

other (Figure 3-5).

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57

Figure 3-2 BOX-PCR fingerprinting patterns of different isolates from potato rhizosphere. M = 1kb marker, 1 = TN1, 2 = TN2, 3 = TN8, 4 = TN3, 5 = TN4, 6 = TN5, 7 = TN6, 8 = TN7, 9 = TN9, 10 = TN24, 11 = TN12, 12 = TN13, 13 = TN15, 14 =

TN14, 15 = TN16, 16 = TN17, 17 = TN18, 18 = TN19, 19 = TN10, 20 = TN21, 21 = TN22, 22 = TN23, 23 = TN25, 24 = TN26, 25 = TN27, 26 = TN28, 27 =


TN41, 41 = TN43, 42 = TN44, 43 = TN42, 44 = TN42, B = Blank

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Figure 3-3 ERIC-PCR fingerprinting patterns of different isolates from potato rhizosphere. M = 1kb marker, 1 = TN2, 2 = TN8, 3 = TN3, 4 = TN4, 5 = TN5, 6 = TN6, 7 = TN7, 8 = TN8, 9 = TN9, 10 = TN24, 11 = TN11, 12 = TN12, 13 = TN13, 14 =



TN38, 41 = TN41, 42 = TN43, 43 = TN44, 44 = TN42, B = Blank, +ve = Positive control

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59

Figure 3-4 Composite dendrogram generated from the data for BOX-PCR

fingerprints of bacterial isolates from potato rhizosphere.

Different colors indicate the sites of isolation.

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60

Figure 3-5 Composite dendrogram generated from the data for ERIC-PCR

fingerprints of bacterial isolates from potato rhizosphere.

Different colors indicate the sites of isolation.

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61

Sequence and Phylogenetic Analysis of 16S rRNA Gene

The 16S rRNA of all the isolates was amplified and sequenced commercially from

Eurofins, Germany. BLASTn analysis of 16S rRNA gene revealed that isolates belongs

to the already reported bacterial genera i.e. Achromobacter, Advenella, Agrobacterium,

Azospirillum, Bacillus, Brevundimonas, Enterobacter, Pseudomonas, Rhizobium,

Stenotrophomonas etc. with upto 99% sequence similarity. Chimeras were detected and

removed from the sequences and cleaned sequences were submitted to EMBL database

and accession numbers were obtained (table 3.6).

Isolate TN01, TN02, TN03, TN08, TN09, TN10, TN12, TN13, TN15, TN22,

TN29, TN30 and TN31 revealed upto 99% similarity with different strains of

Azospirillum brasilense. These isolates also clustered with Azospirillum sp. and A.

brasilense when phylogenetically analyzed with other species of genera Azospirillum

where A. halopraeferens (NR_044859) was taken as root (Figure 3-6). Isolates TN04,

TN18, TN27, TN42 showed similarity with Rhizobium sp. with 95% similarity in case

of isolate TN18 with already reported Rhizobium sp. NIASMXIII whereas isolates

TN06, TN14, TN26 and TN41 were highly similar (99%) to Agrobacterium sp. except

isolate TN26 and TN41 which had 94% and 98% sequence similarity with

Agrobacterium sp. QW10 and A. tumefaciens strain 12b3 respectively. In phylogenetic

tree of Agrobacterium and Rhizobium sp., isolates TN04, TN06, TN41 and TN42 were

clustered with Rhizobium sp. (KR653316) and A. tumefaciens (KP410818) while

isolates TN18 made separate cluster with Rhizobium sp. (JX514405). A separate cluster

was observed by isolate TN26 and TN27 with each other where A. brasilense

(AY324110) was placed as root (Figure 3-7). Isolate TN19, TN32 and TN33 were

identified as Bacillus sp. with 99% sequence identity except isolate TN33 (95%).

Isolate TN19 made cluster with Bacillus sp. (KR998239), B. pumilus (NR_043242) and

B. pumilus (EU302128), strain TN32 with B. altitudinis (NR_042337) while strain

TN33 showed clustering with many strains of already reported Bacillus sp. with

bootstrap value of 97 (Figure 3-8). Four isolates TN37, TN39, TN40 and TN44 were

identified as Brevundimonas sp. (99% sequence similarity) out of which isolate TN37

clustered with three different Brevundimonas sp. i.e. B. naejangsanensis (KF453785),

B. diminuta (AB680378) and B. naejangsanensis (NR_116722) while isolate TN40 and

TN44 were branched with B. vesicularis (KM873029), B. diminuta (KF624717) and B.

naejangsanensis (KC634247) while B. vesicularis (NR_037104) was kept as root for

3. Results

62

the phylogenetic tree (Figure 3-9). Isolate TN17, TN21 and TN23 revealed 99%

sequence identity with Stenotrophomonas sp. Phylogenetic analysis of those isolates

showed clustering of isolate TN17 with S. maltophilia (KF254512), strain TN21 with

S. maltophilia (KM248329) and Stenotrophomonas sp. (AB508865) whereas isolate

TN23 was clustered with S. maltophilia (LC040948) and Stenotrophomonas sp.

(KF737373). P. stutzeri (NR_118798) was placed as root (Figure 3-10).

Table 3.6 Identification of bacterial isolates from rhizosphere of potato based on

16S rRNA gene sequence analysis.

Strain

code Strain origin

16S rRNA based

identification

GenBank

accession

number

Closest GenBank

match

(% identity)

TN01 Gujranwala Azospirillum sp. LN833441

Azospirillum

brasilense strain Az23

(98)


Azospirillum


(99)

TN03 Gujranwala Azospirillum sp. LN833443 Azospirillum sp. YM

249 (99)

TN04 Gujranwala Rhizobium sp. LN833444 Rhizobium sp.

IRBG74 (99)

TN05 Gujranwala Advenella sp. LN833445 Advenella sp.

NTC-1NF (99)

TN06 Gujranwala Agrobacterium sp. LN833446

Agrobacterium

tumefaciens strain

12b3 (99)


Azospirillum

brasilense strain Gr22

(99)

TN09 Jhang Azospirillum sp. LN833448

Azospirillum


(99)


Azospirillum


(99)

TN11 Jhang Sphingobacterium sp. LN833449

Sphingobacterium

mizutaii strain Ht8-22

(99)


Azospirillum


(99)


Azospirillum


(99)

TN14 Jhang Agrobacterium sp. LN614534

Agrobacterium

tumefaciens strain

AF114 (99)

TN15 Gojra Azospirillum sp. LN833452 Azospirillum

brasilense (99)

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63

TN16 Gojra Achromobacter sp. LN833453 Achromobacter sp.

CCBAU 45284 (99)

TN17 Gojra Stenotrophomonas sp. LN833454

Stenotrophomonas

maltophilia strain

MTH20 (99)

TN18 Gojra Rhizobium sp. LN833455 Rhizobium sp.

NIASMXIII (95)

TN19 Gojra Bacillus sp. LN833456 Bacillus safensis

strain AF44 (99)

TN21 Okara Stenotrophomonas sp. LN833457

Stenotrophomonas

maltophilia strain

Akg1 (99)

TN22 Okara Azospirillum sp. LN833458 Azospirillum sp. YM

132 (99)

TN23 Okara Stenotrophomonas sp. LN833459 Stenotrophomonas sp.

QW30 (99)

TN25 Okara Shinella sp. LN833460 Shinella sp. TV7Nov

(99)

TN26 Okara Agrobacterium sp. LN833461 Agrobacterium sp.

QW10 (94)

TN27 Okara Rhizobium sp. LN833462 Rhizobium sp. T45

(99)

TN29 Sheikhupura Azospirillum sp. LN833463

Azospirillum


(99)


Azospirillum


(99)


Azospirillum


(99)

TN32 Sheikhupura Bacillus sp. LN833466 Bacillus sp.

UYFA152 (99)

TN33 Sheikhupura Bacillus sp. LN833467 Bacillus altitudinis

strain OSR50 (95)

TN34 Sheikhupura Bacillus sp. LN833468 Pseudomonas sp. S12

(99)

TN35 Sheikhupura Achromobacter sp. LN833469 Achromobacter sp.

F32 (99)

TN36 Sheikhupura Pseudomonas sp. LN614533 Pseudomonas sp.

VET-5 (99)

TN37 Sahiwal Brevundimonas sp. LN833470

Brevundimonas

naejangsanensis

strain HWG-A15 (99)

TN38 Sahiwal Enterobacter sp. LN614535 Enterobacter cloacae

strain RU07 (99)

TN39 Sahiwal Brevundimonas sp. LN833471 Brevundimonas terrae

strain KSL-145 (99)

TN40 Sahiwal Brevundimonas sp. LN833472 Brevundimonas sp.

X60 (99)

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64

TN41 Sahiwal Agrobacterium sp. LN833473

Agrobacterium

tumefaciens strain

12b3 (98)

TN42 Sahiwal Rhizobium sp. LN614536 Rhizobium sp.

IRBG74 (99)

TN43 Sahiwal Enterobacter sp. LN833474 Enterobacter cloacae

strain WL19 (99)

TN44 Sahiwal Brevundimonas sp. LN833475 Brevundimonas sp.

MM68May (99)

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65


Azospirillum sp. isolated from potato rhizosphere (●) and published sequences.

Neighbor joining method was adopted. Bootstrap values greater than 50 were given and

were based on 1000 replicates. Azospirillum halopraeferens (NR_044859) was used as

outgroup.

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66

Figure 3-7 Phylogenetic tree base on 16S rRNA sequences (603 bp) of Rhizobium

sp. and Agrobacterium sp. isolated from potato rhizosphere (♦) and published

sequences.


were based on 1000 replicates. Azospirillum brasilense (AY324110) was used as

outgroup.

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67

Figure 3-8 Phylogenetic tree base on 16S rRNA sequences (979 bp) of Bacillus sp.

isolated from potato rhizosphere (■) and published sequences.


were based on 1000 replicates. Paenibacillus larvaeT (AY530294) was used as

outgroup.

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68


Brevundimonas sp. isolated from potato rhizosphere (■) and published

sequences.


were based on 1000 replicates. B. vesicularis (NR_037104) was used as root.

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69


Stenotrophomonas sp. isolated from potato rhizosphere (♦) and published

sequences.


were based on 1000 replicates. Pseudomonas stutzeri (NR_118798) was used as

outgroup.

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70

Sequence and Phylogenetic Analysis of NifH Gene

Total 16 isolates out of 44 isolates showed positive amplification for nifH gene (Figure

3-11). The nifH sequences of Azospirillum sp. strains TN01, TN02, TN03, TN10,

TN12, TN13, TN15, TN22, TN30 and TN31 revealed 98 - 100 sequence similarity with

already reported nifH gene of A. brasilense. Rhizobium sp. strain TN04 showed 100%

nifH gene sequence similarity with already reported Rhizobium sp. S1SS148 while

Stenotrophomonas sp. strain TN21 showed 99% sequence similarity of nifH with

uncultured bacterium clone Spruce2009_F1. NifH gene sequenced amplified from

Brevundimonas sp. strain TN37 was highly similar to nifH of A. brasilense Gr42 while

Brevundimonas sp. strain TN40 and TN44 had similarity to the nifH gene sequence of

uncultured bacterium clone OTU-31 (table 3.7).

Phylogenetic analysis of nifH gene sequences showed three major groups where

Azospirillum made Group I. NifH of isolates TN10, TN15, TN30, TN31 and TN37 were

clustered with A. brasilense (FR669137) and isolates TN01, TN02, TN03, TN12, TN13

and TN22 were branched with A. formosense (HM193519), A. brasilense (X51500) and

A. lipoferum (AF216882) with high bootstrap value. Although Stenotrophomonas sp.

strain TN21 belongs to γ-proteobacteria and Rhizobium sp. strain TN04 belongs to α-

proteobacteria were grouped together in Group II, however Stenotrophomonas sp.

TN21 formed sub cluster with uncultured nitrogen-fixing bacterium clone (KF656868),

B. thiooxidansT (DQ431163), S. maltophilia (DQ431162). Even the BLASTn results of

nifH gene sequence showed homology with the nitrogenase gene from Rhizobium sp.,

Stenotrophomonas sp. and uncultured bacteria. On the other hand Rhizobium sp. TN04

clustered with Rhizobium sp. (KU527070) and uncultured Rhizobium sp. (FN666266).

Group III included isolate TN40 and TN44 which were branched with uncultured

bacterium clone (KF541088) with bootstrap value 75 whereas nifH sequence of M.

lacustera (AF296355) was placed as root (Figure 3-12).

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71

Figure 3-11 Agarose gel photograph showing amplified nifH gene from bacterial

strains from potato rhizosphere.

M = 1 kb ladder, 1 = Azospirillum sp. TN01, 2 = Azospirillum sp. TN02, 3 =

Azospirillum sp. TN03, 4 = Rhizobium sp. TN04, 5 = Azospirillum sp. TN10, 6 =

Azospirillum sp. TN012, 7 = Azospirillum sp. TN13, 8 = Azospirillum sp. TN15, B =

Blank

Table 3.7 NifH gene sequence similarity of nitrogen fixing bacteria from

rhizosphere of potato.

Strain Closest GenBank match based on nifH

sequence (% identity)

Accession

No.

Azospirillum sp. TN01 Azospirillum brasilense strain Gr52 (99) LT596584



Rhizobium sp. TN04 Rhizobium sp. S1SS148 (100) LT596587


Azospirillum sp. TN10 Azospirillum brasilense strain Gr42 (100) LN681358




Stenotrophomonas sp. TN21 Uncultured bacterium clone Spruce2009_F1

(99) LT596592




Brevundimonas sp. TN37 Azospirillum brasilense Gr42 (100) LT596596

Brevundimonas sp. TN40 Uncultured bacterium clone OTU-31 (83) LT596597

Brevundimonas sp. TN44 Uncultured bacterium clone OTU-31 (83) LT596598

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Figure 3-12 Phylogenetic tree based on nifH sequences (211 bp) of the bacterial

strains isolated from potato rhizosphere (♦) compared with related published

sequences.

Neighbor joining method was adopted. Bootstrap values greater than 50 are given and

were based on 1000 replicates. An archeal species Methanosarcina lacustera

(AF296355) was used as outgroup.

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73

Part 3.2 Plant Inoculation and Colonization Studies

3.2.1 Pot Experiment (Sand Culture)

Thirteen potential rhizobacteria were inoculated to potato (variety Kuroda) to evaluate

the plant growth promoting potential in sand. Two control treatments were also

included, one with recommended dose of N and second with zero N. Generally

inoculation resulted in improved growth (table 3.8 and 3.9).

Inoculation with Brevundimonas sp. TN37 resulted in maximum increase in

shoot length (20.48 cm). Pseudomonas sp. TN36 inoculated plants showed maximum

increase in root length (24.75 cm) as shown in figure 3-13.

In case of shoot fresh weight, Rhizobium sp. TN04 exhibited maximum shoot

fresh weight (51.93 g). Maximum root fresh weight (43.33 g) was produced by

Azospirillum sp. TN09. Highest shoot dry weight (5.1 g) was exhibited by Rhizobium

sp. TN04 and Azospirillum sp. TN09 had maximum root dry weight (3.95 g) which

were statistically similar to the dry weights of shoot and root, produced by positive

control (Figure 3-14).

Plant nitrogen contents were also increased by inoculation. Rhizobium sp. TN04

had maximum increase in N contents of shoot (4.16%) positive control, whereas

Rhizobium sp. TN04 and Azospirillum sp. TN09 resulted in maximum root N contents

(2.39 g) that were similar to the positive control (Figure 3-15).

Different bacterial strains showed differential colonization potential at different

time intervals. Overall bacterial population tended to decrease as the time after sowing

increased. Azospirillum sp. TN09 showed the best colonization potential upto 60 days

among all the isolates (Figure 3-16 a) which showed relatively stable association in

terms of CFU g-1 of rhizospheric sand however Agrobacterium sp. TN14, Bacillus sp.

TN32 and Pseudomonas sp. TN36 resulted gradual decrease of association with potato

plants, led to poor colonization (3.16 b, c).

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74

Table 3.8 Effect of inoculation of different nitrogen fixing strains on shoot

growth parameters of potato plant grown in sterilized sand.

Treatments Shoot

length (cm)

Shoot fresh

weight (g)

Shoot dry

weight (g)

Shoot N

contents (%)

T1 22.28 a 54.35 a 5.35 a 4.20 a

T2 10.50 i 24.75 h 3.00 g 3.08 e

T3 13.33 efg 39.35 def 3.93 cdef 3.52 d

T4 12.85 efgh 38.8 def 3.52 efg 3.53 d

T5 16.38 c 46.03 bc 4.38 bcd 4.04 abc

T6 19.20 b 51.93 a 5.10 ab 4.16 a

T7 12.35 fgh 50.53 ab 4.65 abc 4.10 ab

T8 13.65 ef 38.9 def 3.83 def 3.98 abc

T9 17.25 c 37.33 efg 3.55 efg 3.50 d

T10 11.90 ghi 33.60 g 3.25 fg 3.43 de

T11 14.18 de 36.70 efg 3.48 fg 3.45 de

T12 19.68 b 35.15 fg 3.25 fg 3.42 de

T13 20.48 b 43.60 cd 4.33 bcde 3.74 bcd

T14 11.40 hi 41.68 cde 3.98 cdef 3.74 bcd

T15 15.60 cd 38.45 efg 3.35 fg 3.65 cd

LSD 5% 1.699 5.112 0.807 0.400

SEC 0.844 2.538 0.401 0.198

Data on plant parameters were taken after 60 day of sowing. Values are the mean of 4

replicates. Values in same column sharing same letter do not differ significantly (P≥

0.05) according to Fisher’s LSD.T1 = Un-inoculated + full dose of N (Positive control)

T2 = Un-inoculated + zero N (Negative control)

T3 = Azospirillum sp. TN01 + zero N



T6 = Rhizobium sp. TN04 + zero N



T9 = Sphingobacterium sp. TN11 + zero N

T10 = Agrobacterium sp. TN14 + zero N

T11 = Bacillus sp. TN32 + zero N

T12 = Pseudomonas sp. TN36 + zero N

T13 = Brevundimonas sp. TN37 + zero N

T14 = Enterobacter sp. TN38 + zero N


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75

Table 3.9 Effect of inoculation of different nitrogen fixing strains on root growth

parameters of potato plant grown in sterilized sand.

Treatments Root length

(cm)

Root fresh

weight (g)

Root dry

weight (g)

Root N contents

(%)

T1 25.75 a 45.68 a 4.65 a 2.40 a

T2 10.43 g 19.56 j 1.77 e 1.26 g

T3 14.28 e 27.80 f 3.20 bc 2.13 bc

T4 19.2 d 27.38 fg 2.18 de 2.09 bc

T5 14.53 e 40.98 bc 3.85 ab 2.35 a

T6 21.98 bc 40.00 bc 3.88 ab 2.39 a

T7 12.70 ef 43.33 ab 3.95 ab 2.39 a

T8 14.40 e 37.35 cd 2.65 cd 2.1 bc

T9 23.85 ab 26.68 fgh 2.43 cde 2.01 cd

T10 20.13 cd 23.53 ghi 1.98 de 1.62 ef

T11 11.70 fg 24.33 fghi 2.28 de 2.11 bc

T12 24.75 a 23.38 hij 1.85 de 1.57 f

T13 19.1 d 34.28 de 3.78 b 2.28 ab

T14 24.2 a 31.88 e 3.5 b 1.97 cd

T15 13.93 e 22.43 ij 1.82 de 1.82 de

LSD 5% 2.033 3.852 0.841 0.213

SEC 1.009 1.912 0.418 0.105

Data on plant parameters were taken after 60 day of sowing. Values are the mean of 4

replicates. Values in same column sharing same letter do not differ significantly (P≥

0.05) according to Fisher’s LSD.

T1 = Un-inoculated + full dose of N (Positive control)

T2 = Un-inoculated + zero N (Negative control)







T9 = Sphingobacterium sp. TN11 + zero N

T10 = Agrobacterium sp. TN14 + zero N

T11 = Bacillus sp. TN32 + zero N

T12 = Pseudomonas sp. TN36 + zero N

T13 = Brevundimonas sp. TN37 + zero N

T14 = Enterobacter sp. TN38 + zero N


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76

Figure 3-13 Effect of bacterial inoculation on potato plants. Plants were

harvested 60 days after sowing.

A = Un-inoculated + full dose of N, B = Un-inoculated + zero N

C = Azospirillum sp. TN01 + zero N, D = Azospirillum sp. TN02 + zero N

E = Azospirillum sp. TN03 + zero N, F = Rhizobium sp. TN04 + zero N

G = Azospirillum sp. TN09 + zero N, H = Azospirillum sp. TN10 + zero N

I = Sphingobacterium sp. TN11 + zero N, J = Agrobacterium sp. TN14 + zero N

K = Bacillus sp. TN32 + zero N, L = Pseudomonas sp. TN36 + zero N

M = Brevundimonas sp. TN37 + zero N, N = Enterobacter sp. TN38 + zero N

O = Rhizobium sp. TN42 + zero N

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Figure 3-14 Effect of different bacterial strains on potato shoot and root dry

weight.

Value are average of 4 replicates each. Bar represents standard deviation of the data.

Means containing the same letter are not different from each other at 5% LSD.

Figure 3-15 Effect of different bacterial strains on potato shoot and root N

contents.

Value are average of 4 replicates each. Bar represents standard deviation of the data.

Means containing the same letter are not different from each other at 5% LSD.

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78

Figure 3-16 Population dynamics of inoculated bacterial strains in rhizosphere of

potato at different time intervals after sowing.

Value are average of 4 replicates each. Bar represents standard deviation of the

replicates.

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79

3.2.2 Root Colonization Studies through Transmission Electron

Microscopy (TEM)

Five bacterial strains showing good plant growth potential were further evaluated for

plant colonization studies at ultrastructural level. Transmission electron microscopic

observations showed that Azospirillum sp. strain TN03, TN09, TN10, Rhizobium sp.

TN04 and Brevundimonas sp. TN37, showed good colonization with potato roots. The

Azospirillum sp. strain TN03 resided in the niches formed by root hairs of potato plant

(Figure 3-17).

The Rhizobium sp. strain TN04 showed excellent colonization and was present

among the root hairs as shown in figure 3-18. Azospirillum sp. strain TN09 also showed

strong association with potato roots and observed in the rhizosphere, close to roots cell

wall (Figure 3-19).

Similarly Azospirillum sp. strain TN10 was also localized in the rhizosphere of

potato plant near root surface (Figure 3-20). A strong colonization potential of

Brevundimonas sp. strain TN37 was also observed (Figure 3-21) where bacterial cells

were colonized with the potato roots. No bacterial cells were observed in the roots of

un-inoculated control plants (Figure 3-22).

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80


with Azospirillum sp. TN03, 30 days post inoculation.

RS = Rhizosphere, RC = Root cell and B = Bacterium

Azospririllum sp. TN03 can be seen as micro-colonies out side the root cells and within

the grooves of root hairs (A). Clusters of bacterial cell are present on to the epidermal

cells of roots (B, C and D) forming several cell layers on it. This layering help bacteria

to strongly colonize with the roots.

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81


with Rhizobium sp. TN04, 30 days post inoculation. Boxes in the Figure 3-15 (A and B) represent the area with higher resolution in figure 3-15 B, C and D, respectively. A significant portion of Rhizobium sp.TN04 colonized in the patches along niches formed at the border region of adjacent epidermis cells (A, B, C, D, and F). Growth within this protected area might lead to more friendly contact to the root surface and better utilization of nutrients excreted by the roots. Bacterial cell aggregates were also formed over the cell wall of root cells (E). RS = Rhizosphere, RC = Root cell and B = Bacterium

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82




Azospirillum sp. TN09 cells were established in the slots made among the root cells

where multiple bacterial cell masses made micro-colonies (A, B). Multicellular

aggregates, embedded in extracellular matrix were also found on the root surface which

showed strong adhesion of bacterial cells with the potato roots (C, D).

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83



Box in the Figure 3-17 (A) represents the area with higher resolution (figure 3-17 B)


Azospirillum sp. TN10 were closely attached to the root surface, forming layer of

bacterial cells all over the root area (A, B). Bacterial cells stick to the cell wall of the

root cells forming strong relationship (C, D) which benefit both bacteria and plant.

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84


with Brevundimonas sp. TN37, 30 days post inoculation.


Brevundimonas sp. TN37 resident over the root surface of potato. Bacterial cells form

sheets in the grooves, formed by the root cells (A, B) and also closely attached to the

cell wall of plant cells (C, D).

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85

Figure 3-22 Electron micrographs of un-inoculated potato root control, 30 days

post inoculation.

RS = Rhizosphere and RC = Root cell

No bacteria were found on the root surface, intracellular spaces (B) and onto surface of

epidermal cells of root (A, B, D). The rhizosphere and endosphere on non-inoculated

control plants are totally devoid of bacteria.

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86

3.2.3 Root Colonization Studies through Confocal Laser Scanning

Microscopy (CLSM)

Colonization potential of bacterial strains was also studied, using CLSM. Four bacterial

strains, Azospirillum sp. TN03, Rhizobium sp. TN04, Azospirillum sp. TN09 and

Brevundimonas sp. TN37 were transformed with YFP gene and YFP-tagged bacteria

were inoculated to potato plants grown in sterilized sand. Roots of inoculated plants

were observed under CLSM after 25 day of inoculation. Primary and secondary roots

of Azospirillum sp. TN03 inoculated plants showed the presence of bacterial isolate on

the potato roots giving green fluorescence. Colonization density decreased gradually

from the upper part of the root towards the root tip (Figure 3-23 A). Azospirillum sp.

TN03 was mostly colonize in the intercellular spaces over the surface of root cells

(Figure 3-23 B). Bacterial masses were detected over the cells surface of potato roots,

forming macro-colonies (Figure 3-23 C) that showed the close association of potato

with strain TN03.

Strong colonization of Rhizobium sp. TN04 was observed on the junctions

between the primary and secondary roots, which may be because this area provide a

better niche for the bacterial colonization (Figure 3-24 A). Bacterial aggregates onto

the epidermal cells of root were detected in the form of micro-colonies (Figure 3-24 C),

assured the association of Rhizobium sp. TN04 with potato.

Azospirillum sp. TN09 was spotted in the form of large bacterial masses on the

lateral roots and root hairs of potato (Figure 3-25). Bacterial clusters were present all

over the surface of roots (Figure 3-25 A, B). Dense colonization over the root hairs of

potato witnessed the strong attachment of Azospirillum sp. TN09 with potato roots.

Primary roots, the tip area of emerging lateral roots and root hairs were the preferred

targets of colonization by the Brevundimonas sp. TN37 (Figure 3-26). Bacterial

aggregates were detected on the root surface and root hairs of potato (Figure 3-26 A,

B) which was the result of establishment of associative relationship between

Brevundimonas sp. TN37 and potato plant.

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87


labelled Azospirillum sp. TN03, in sand culture 25 days post inoculation.

RC = Root cell, B = Bacteria

Fluorescence observed under CLSM on longitudinal sections of primary and lateral

roots of potato, showing the bacterial presence (A, B). Bacterial aggregates were clearly

seen on the root epidermis on higher magnification (C) which confirms its association

with potato roots.

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88


labelled Rhizobium sp. TN04, in sand culture 25 days post inoculation.

RC = Root cell, B = Bacteria

More fluorescence was detected around the junction between primary and secondary

roots which showed that bacteria preferred to colonize that area (A) over the primary

roots surface (B). Micro-colonies of bacteria on the roots assured the colonization of

Rhizobium sp. TN04 with potato (C).

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89


labelled Azospirillum sp. TN09, in sand culture 25 days post inoculation.

RC = Root cell, B = Bacteria, RH = Root hair

Bacterial cell were distributed all over the surface of potato root whereas Macro-

colonies in the form of thick bacterial aggregates were observed (A, B). Root hairs were

infected with Azospirillum sp. TN09 (C), which covered root hair tips and lateral area,

showed the strong establishment of the bacteria in the root zone of potato.

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90


labelled Brevundimonas sp. TN37, in sand culture 25 days post inoculation.

RC = Root cell, B = Bacteria, RT = Root tip, RH = Root hair

Primary root tips and root hairs of potato were preferred targets of colonization by

Brevundimonas sp. TN37 (A, C). Macro-colonies, forming bacterial aggregates were

observed on the epidermal root cells (B) that endorse the association of strain TN37

with potato.

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91

3.2.4 In vivo N2 Fixation Analysis using 15N Analysis (Soil Culture)

Three pot experiments were conducted in soil to quantify the N fixation ability of

bacterial strain and to check their effect on plant biomass production and N contents.

Plants were inoculated with three bacterial strains, Two Azospirillum strains TN03,

TN09 and one Rhizobium strain TN04 which showed excellent colonization potential

in previous experiments. In experiment 1, zero dose of N (N0) was applied for

quantification % nitrogen fixation (%Ndfa) using 15N natural abundance technique and

half and full doses of recommended N (N1/2 and NF respectively) was applied in

experiment 2 and 3 respectively for the use of 15N dilution techniques to estimate

%Ndfa. 15N was applied in the form of ammonium sulphate with 5% a.e (atomic

access). All three experiments were conducted simultaneously in similar conditions.

Plants growth parameters were positively affected by the inoculation of

bacterial strains (table 3.10). Maximum shoot length (22.6 cm) was recorded in case of

Azospirillum sp. TN09 + NF among the inoculated treatments which was statistically at

par to un-inoculated control with full dose of N. Root length showed significant

increase to inoculation with all strains. Maximum root length (27.3 cm) was also

observed in Azospirillum sp. TN09 + NF while minimum root length (12.9 cm) was

recorded in case of Azospirillum sp. TN03 + N0, which was statistically similar to un-

inoculated control without N.

Plant biomass production was increased with inoculation of all bacterial strains

as compared to un-inoculated control. Both Azospirillum sp. strains TN03 and TN09

showed maximum increase in shoot dry weight (7.3 g and 7.7g, respectively).

Maximum root dry weight (5.4g and 5.5g) was produced by Azospirillum sp. strains

TN03 and TN09 respectively.

Plant N contents were also significantly increased by inoculation of

diazotrophic strains both in exp. 1 and exp. 2 (in case of N0 and N1/2) but no significant

increase was observed in exp. 3 (NF). Among the inoculated treatments, Azospirillum

sp. TN09 + NF resulted in the maximum plant N contents (4.3%) which was statistically

similar to un-inoculated control with full dose of N where Azospirillum sp. TN03 and

Rhizobium sp. TN04 without N gave the minimum N contents (2.5%) in plant. 15N%

abundance of plants was affected by the N fertilization rate because the average for N0,

N1/2 and NF was 0.43, 0.96 and 1.05, respectively. 15N% abundance. of all inoculated

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92

plants were found to be lesser as compared to their respective un-inoculated controls

which indicates the dilution of 15N due to the uptake of biologically fixed nitrogen

(%Ndfa). Maximum dilution of 15N was recorded by Azospirillum sp. TN09 in all three

experiments.

The %Ndfa value varies from 44.8% to 13.5% amongst the bacterial strains.

Generally %Ndfa was decreased with external fertilization of N except for Azospirillum

sp. TN03 which did not show significant effect on %Ndfa on the addition of N fertilizer.

Maximum %Ndfa (44.8%) was observed by Azospirillum sp. TN09 + N0 however

minimum %Ndfa (13.5%) was recorded by Azospirillum sp. TN03 + N0 (table 3.10).

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93

Table 3.10 Effect of inoculation of nitrogen fixing strains on growth parameters,

N contents and 15N abundance of potato plant grown in sterilized soil and

estimates of biologically fixed nitrogen.

Shoot

length

(cm)

Root

length

(cm)

Shoot

dry

weight

(g)

Root

dry

weight

(g)

Plant N

contents

(%)

%15N

abundance %Ndfa

Exp. 1 δ15N method

Un-inoculated +

N0 16.47 g

11.48

f 3.63 e 2.53 f 1.68 h 0.43±0.002 -

Azospirillum sp.

TN03+ N0

17.6

efg

12.87

ef 4.17 e 2.87 ef 2.49 f 0.42±0.003 16.61 cd

Rhizobium sp.

TN04 + N0

17.07

fg

14.15

def 4.07 e 2.67 ef 2.48 f 0.41±0.003 28.48 b

Azospirillum sp.

TN09 + N0

17.53

efg

15.22

de 3.97 e 3.27 de 2.15 g 0.40±0.003 44.79 a

Exp. 2 15N isotope dilution method

Un-inoculated +

N1/2

17.6

efg

16.10

d 4.90 d

3.03

def 3.12 e 0.96±0.006 -

Azospirillum sp.

TN03 + N1/2

18.47

def

15.69

de 5.77 c 4.23 c 3.73 d 0.83±0.034 13.52 d

Rhizobium sp.

TN04 + N1/2

18.97

de

21.19

c 5.67 c 3.53 d 3.52 d 0.77±0.019 19.61 c

Azospirillum sp.

TN09 + N1/2

19.47

cd

21.05

c 5.97 c 4.43 bc 3.66 d 0.66±0.025 31.14 b

Exp. 3 15N isotope dilution method

Un-inoculated +

NF 23.83 a

23.98

bc 7.80 a 5.33 a 4.40 a 1.05±0.051 -

Azospirillum sp.

TN03 + NF

21.3

bc

26.25

ab 7.27 ab 5.37 a 4.01 c 0.90±0.057 13.77 d

Rhizobium sp.

TN04 + NF 21.83 b

26.77

ab 7.03 b 4.97 ab 4.09 bc 0.86±0.027 18.14 cd

Azospirillum sp.

TN09 + NF

22.63

ab

27.28

a 7.73 ab 5.50 a 4.32 ab 0.83±0.01 21.24 c

LSD 5% 1.860 3.064 0.706 0.624 0.254 - 5.539

SEC 0.901 1.485 0.342 0.303 0.123 - 2.636

Data on plant parameters were taken after 60 day of sowing. The data is mean of 3

replicates. Values in same column sharing same letter, do not differ significantly (P≥

0.05) according to Fisher’s LSD. ± shows the standard deviation of the data.

3.2.5 Field Experiments

All three isolates, TN03, TN04 and TN09 were further evaluated for their PGP potential

with potato in field conditions. Two field experiments were conducted on two different

locations i.e. National Institute for Biotechnology and Genetic Engineering (NIBGE),

Faisalabad and Potato Research Institute (PRI), Sahiwal with six treatments, three un-

inoculated (T1, T2 and T3) and three un-inoculated (T4, T5 and T6) treatments. All the

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94

inoculated treatments resulted in significant effect on all growth parameters of potato.

Data on soil analysis and CFU of the experiment sites were given in table 3.11.

Table 3.11 Physico-chemical analysis and bacterial population of soil from

experimental sites.

Parameter NIBGE, Faisalabad PRI, Sahiwal

Soil pH 7.3 7.2

Soil texture Sandy Loam Loam

Organic matter (%) 0.58 0.82

Total N (%) 0.06 0.08

Total P (ppm) 53.8 64.3

Total K (ppm) 260 170

Bacterial population (g-1 soil) 7 × 106 7 × 107

Plant Height (cm)

Plant height showed the significant effect of inoculation for all the bacterial strains.

Inoculation of Rhizobium sp.TN04 with half dose of recommended N resulted in the

maximum plant height (42.8 cm) among the inoculated treatments with half dose of N.

It showed 18% increase in plant height over the un-inoculated control with half dose of

N fertilizer. There was no significant difference of locations on plant height (table 3.12).

Similar trends were observed in the interaction of locations with treatments. Rhizobium

sp. TN04 with half dose of N at Sahiwal (L2×T5) gave the maximum plant height (43.5

cm) which was significantly higher than un-inoculated control with half dose of N on

both locations (L1×T2 and L2×T2), as presented in table 3.14.

Number of Branches per Plant

All the isolates supplemented with half dose of N, showed significant increase in

number of branches per plant which were statistically at par to un-inoculated control

treatment with recommended dose of N (T3). Azospirillum sp. TN09 gave the

maximum increase (18.5%) in number of branches per plant, within the inoculated

treatments over un-inoculated control with half dose of recommended N (table 3.12).

Locations showed non-significant effect on number of branches per plant. L2×T6

revealed the maximum number of branches (3.5) in interactive effect of inoculated

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95

treatments with locations. There was no significant difference in number of branches

per plant among the inoculated treatments coupled with half dose of N and un-

inoculated control with recommended dose of N on both the locations (table 3.14).

Number of Compound Leaves per Plant

Bacterial inoculation with half dose of recommended N applied a significant effect on

number of compound leaves of potato (table 3.12). Maximum number of compound

leaves (32.96) were observed by Azospirillum sp. TN09 with half dose of recommended

N. It (T6) cause upto 14% increase over un-inoculated control with half dose of N

fertilizer (T2). Relatively high number of compound leaves per plant were observed at

Sahiwal (L2) but it was statistically at par to Faisalabad (L1). When the interactive

effect of inoculation with different bacterial strains and locations (T×L) was considered,

statistically high number of compound leaves (33.5) were recorded by Rhizobium sp.

TN04 provided with half dose of N at Sahiwal (L2×T5), as shown in table 3.14.

Plant Fresh Weight per Plant (g)

A significant effect of different bacterial inoculants was recorded on plant fresh weight

with half dose of N as compared to plants without inoculations with half dose of

nitrogen fertilizer (table 3.12). Amongst the inoculated treatments, statistically higher

plant fresh weight (301.7 g) was produced by Rhizobium sp. TN04 + half dose of N

(T5) which increased the fresh weight upto 21% over the un-inoculated control with

half dose of N (T2). Keeping in view the locations, relatively high plant fresh weight

per plant (273.83 g) was produced on L2 than L1 but no significant difference was

observed. When interactions between treatments and locations (T×L) were studied,

maximum fresh weight per plant (307.1 g) was produced by Rhizobium sp. TN04

coupled with half dose of N at Sahiwal (L2×T5). Zero N with un-inoculated plants

resulted in least plant fresh weight on both the locations (table 3.14).

Plant Dry Weight (g)

Significantly higher plant dry weight was produced in all the inoculated treatments with

half dose of N over un-inoculated control plants with zero and half dose of nitrogen

fertilizer (table 3.12). Rhizobium sp. TN04 with half dose of N resulted as maximum

dry weight per plant (23.2 g) among the inoculated treatments. Plant dry weight was

significantly increased (18.5%) by Rhizobium sp. TN04 + half dose of N (T5) as

compared to un-inoculated control with half dose of recommended N. No significant

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96

effect of locations was observed on plant dry weights. So far interaction between

treatments and locations (T × L) was concerned, plants inoculated with isolate TN04

with half dose of N at Sahiwal field (L2×T5) produced highest plant dry weight (23.43

g) amid the inoculated treatments (table 3.14).

Tuber Fresh Weight (g)

Inoculation with different bacterial isolates along with half dose of N fertilizer

significantly increased tuber fresh weight (table 3.13). In comparison with un-

inoculated plants with half dose of N fertilizer (T2), significantly higher tuber fresh

weight (139.8 g) was recorded in plants inoculated with Rhizobium sp. TN04 with half

dose of recommended N (T5), as shown in table 3.9b. Isolate TN04 (Rhizobium sp.)

increased the tuber fresh weight upto 9% over un-inoculated treatment coupled with

half dose of N. There was not significant effect of locations on tuber fresh weight,

however as far as interaction of location and treatments (L×T) is concerned, maximum

tuber fresh weight (139.9 g) was produced by Rhizobium sp. TN04 at Faisalabad

(L1×T5) among the inoculated treatments (table 3.15).

Tuber Dry Weight (g)

Tuber dry weight was significantly affected both by inoculated treatments and un-

inoculated treatments with half and full dose of recommended N over un-inoculated

plants without N fertilizer (table 3.13). Amongst the inoculated treatments, maximum

dry weight (12.6 g) of potato tuber was observed in plants inoculated with Rhizobium

sp.TN04 along with half dose of nitrogen, caused 14% increase in tuber dry weight.

Un-inoculated plants with no nitrogen produced least tuber dry weight (7.94 g). No

significant effect of locations was observed on tuber dry weight. However treatment

inoculated with Rhizobium sp. TN04 along with half dose of N produced highest tuber

dry weight (12.7 g) at Sahiwal (L2), when the interactions of inoculated bacterial

treatments and locations (T × L) were studied (table 3.15).

Number of Tubers per Plant

Inoculation of potato plants with different bacterial strains along with half dose of

nitrogen at various locations significantly increased the number of tubers per plant over

un-inoculated treatment with zero and half dose of N fertilizer (table 3.13). Maximum

number of tubers per plant (3.9) was recorded in plants inoculated with Azospirillum

sp. TN09 supplemented with half dose of nitrogen which was statistically at par with

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97

un-inoculated treatments along with recommended dose of nitrogen (4.4). Number of

tubers per plant were increased upto 12% by inoculation with Azospirillum sp. TN09

(T6) over un-inoculated control with half dose of N (T2). In terms of locations,

significantly higher number of tubers per plant (3.96) were produced on Sahiwal field

(L2). Statistically higher number of tubers per plant was found Azospirillum sp. TN09

coupled with half dose of N (4.4) on Sahiwal location (L2×T6) which was similar to

L2×T3 (Un-inoculated + Recommended dose of N at Sahiwal) in the interaction

between treatments and locations (T × L) shown in table 3.15.

Tuber Yield (tonnes ha-1)

A significant increase was observed in tuber yield by the inoculation of bacterial

isolates along with half dose of recommended N (table 3.13). Rhizobium sp. TN04 with

half dose of N significantly increased (19%) yield of potato in comparison to un-

inoculated control with half dose of N. It produced 15.3 tonnes ha-1 of potato which was

the maximum yield among inoculated treatments. Location also significantly affected

the tuber yield. Statistically higher tuber yield (14.53 tonnes ha-1) was recorded at

Sahiwal (L2) field area (table 3.15). Significant effect was also observed in the

interaction among treatments and locations (T × L). Maximum tuber yield (15.58 tonnes

ha-1) was produced by treatment inoculated with Rhizobium sp. TN10 + half dose of N

at Sahiwal (L2×T5) which was not significantly different from the yield (18.3 and 18.4

tonnes ha-1) produced by un-inoculated control with recommended dose of N at

Faisalabad and Sahiwal (L1×T3 and L2×T3, receptively).

N Contents of Tuber (%)

Plant N contents were significantly increased by the inoculation of nitrogen fixing

strains supplemented with half dose of N over un-inoculated treatments, had zero and

half dose of N. Maximum plant N contents (17.9 mg g-1) were observed by Rhizobium

sp. TN04 with half dose of N which was statistically similar to un-inoculated control

with recommended dose N. Rhizobium sp. TN04 caused 10% increase in plant contents

over the un-inoculated control treatment with half dose of N (table 3.13). There was no

significant effect of locations on the N contents of plants, however Rhizobium sp. TN04

provided with half dose of N at Sahiwal (L2×T5) produced maximum N contents in

plants (17.9 mg g-1) which was at par to N contents produced by the un-inoculated

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98

control with full recommended dose of N both on Faisalabad and Sahiwal (L1×T3 and

L1×T3, respectively) as given in table 3.15.

Principal component analysis (PCA) was also carried out to summarize the

relationship among agronomic parameters, yield and experimental locations. The effect

of different treatments remained different on the agronomic and yield parameters of

potato at two different experimental locations in the categorical PCA analysis

(CATPCA). The treatments were scattered differently at the PC1 and PC2, which

showed their effect according to the locations. All the treatments except T1 (un-

inoculated control with zero N), were loaded positively to the PC1 and PC2 showed

positive correlation within the parameters (Figure 3-27 and 3-28). Overall all the

inoculated treatments along with half dose of recommended fertilizer performed better

on Sahiwal field area (L2). Variance of (CATPCA) was found 81.41% with respect to

PC1 and PC2.

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Table 3.12 Comparison of treatment means and location means of potato

inoculated with bacterial isolates under field conditions.

Table 3.13 Comparison of treatment means and location means of potato

inoculated with bacterial isolates under field conditions.

T1 = Un-inoculated control + zero N T2 = Un-inoculated control + half dose of recommended N T3 = Un-inoculated control + full dose of recommended N T4 = Inoculated with Azospirillum sp. TN03 + half dose of recommended N T5 = Inoculated with Rhizobium sp. TN04 + half dose of recommended N T6 = Inoculated with Azospirillum sp. TN09 + half dose of recommended N L1 = Faisalabad L2 = Sahiwal

Plant height

(cm)

No. of main

branches

per plant

No. of

compound

leaves per

plant

Plant fresh

weight (g)

Plant dry

weight (g)

Effect of treatments

T1 27.46 d 2.04 c 18.1 d 148.31 e 12.83 d

T2 36.06 c 2.84 b 28.95 c 249.16 d 19.49 c

T3 49.95 a 3.69 a 40.61 a 342.98 a 28.34 a

T4 41.68 b 3.18 ab 31.08 bc 272.5 cd 21.08 bc

T5 42.83 b 3.28 ab 32.93 b 301.69 b 23.19 b

T6 42.23 b 3.38 ab 32.96 b 294.69 bc 22.8 b

LSD 5% 2.820 0.671 2.470 27.39 2.373

Effect of locations

L1 39.34 a 2.99 a 30.2 a 262.61 a 20.73 a

L2 40.72 a 3.14 a 31.34 a 273.83 a 21.84 a

LSD 5% 1.628 0.387 1.426 15.81 1.370

Tuber fresh

weight (g)

Tuber dry

weight (g)

No. of

tubers per

plant

Yield (tonnes

ha-1)

Tuber N

contents

(mg g-1)

Effect of treatments

T1 85.74 c 7.94 d 2.64 c 10.95 d 14.19 d

T2 127.6 b 11.01 c 3.5 b 12.76 c 16.21 c

T3 159.04 a 14.15 a 4.36 a 16.59 a 18.36 a

T4 133.79 b 11.76 bc 3.65 ab 14.44 b 17.52 abc

T5 139.84 b 12.64 b 3.89 ab 15.28 b 17.92 ab

T6 137.72 b 12.06 bc 3.94 ab 15 b 16.87 bc

LSD 5% 14.85 1.099 0.836 0.992 1.327

Effect of locations

L1 129.61 a 11.45 a 3.36 b 13.81 b 16.80 a

L2 131.63 a 11.74 a 3.96 a 14.53 a 16.88 a

LSD 5% 8.570 0.634 0.483 0.573 0.766

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Table 3.14 Potato growth parameters as affected by interaction between

bacterial treatments and locations under field conditions.

Plant height

(cm)

No. of

main

branches

per plant

No. of

compound

leaves per

plant

Plant fresh

weight (g)

Plant dry

weight (g)

L1×T1 26.68 d 1.95 c 17.250 e 143.45 g 12.18 e

L1×T2 35.55 c 2.8 bc 28.60 d 242.40 f 18.98 d

L1×T3 49.18 a 3.58 ab 39.65 a 337.40 ab 27.88 a

L1×T4 40.93 b 3.15 ab 30.53 bcd 267.70 def 20.15 bcd

L1×T5 42.20 b 3.18 ab 32.33 bc 296.25 cd 22.95 bc

L1×T6 41.53 b 3.3 ab 32.85 b 288.45 cde 22.25 bcd

L2× T1 28.25 d 2.13 c 18.95 e 153.18 g 13.48 e

L2× T2 36.58 c 2.88 abc 29.30 cd 255.93 ef 20.00 cd

L2× T3 50.73 a 3.8 a 41.58 a 348.55 a 28.80 a

L2×T4 42.43 b 3.2 ab 31.63 bcd 277.30 cdef 22.00 bcd

L2×T5 43.45 b 3.38 ab 33.53 b 307.13 bc 23.43 b

L2×T6 42.93 b 3.45 ab 33.08 b 300.93 bcd 23.35 bc

LSD 5% 3.989 0.949 3.494 38.728 3.356

Table 3.15 Potato growth parameters as affected by interaction between

bacterial treatments and locations under field conditions.

Tuber fresh

weight (g)

Tuber dry

weight (g)

No. of

tubers per

plant

Yield

(tonnes

ha-1)

Tuber N

contents

(mg g-1)

L1×T1 83.30 d 7.80 e 2.43 d 10.58 g 14.11 c

L1×T2 124.30 c 10.90 d 3.10 bcd 12.70 ef 16.18 b

L1×T3 157.83 ab 13.73 ab 4.28 ab 15.75 b 18.31 a

L1×T4 134.95 c 11.60 cd 3.40 abcd 14.23 cd 17.46 ab

L1×T5 139.88 abc 12.60 bc 3.45 abcd 14.98 bc 17.89 ab

L1×T6 137.40 bc 12.08 cd 3.53 abcd 14.65 bc 16.92 ab

L2× T1 88.17 d 8.08 e 2.85 cd 11.33 fg 14.27 c

L2× T2 130.90 c 11.13 cd 3.90 abc 12.83 de 16.25 b

L2× T3 160.25 a 14.58 a 4.45 a 17.43 a 18.42 a

L2×T4 132.63 c 11.93 cd 3.90 abc 14.65 bc 17.59 ab

L2×T5 139.80 abc 12.68 bc 4.33 a 15.58 bc 17.94 ab

L2×T6 138.05 bc 12.05 cd 4.35 a 15.35 bc 16.83 ab

LSD 5% 20.99 1.554 1.183 1.403 1.877 T1 = Un-inoculated control + zero N T2 = Un-inoculated control + half dose of recommended N T3 = Un-inoculated control + full dose of recommended N T4 = Inoculated with Azospirillum sp. TN03 + half dose of recommended N T5 = Inoculated with Rhizobium sp. TN04 + half dose of recommended N T6 = Inoculated with Azospirillum sp. TN09 + half dose of recommended N L1 = Faisalabad L2 = Sahiwal

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101


different experimental locations and treatments. T1 = Un-inoculated control + N0 T2 = Un-inoculated control + N1/2 T3 = Un-inoculated control + NF T4 = Inoculated with Azospirillum sp. TN03 + N1/2 T5 = Inoculated with Rhizobium sp. TN04 + N1/2 T6 = Inoculated with Azospirillum sp. TN09 + N1/2 L1 = Faisalabad L2 = Sahiwal

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different experimental locations and treatments. T1 = Un-inoculated control + N0 T2 = Un-inoculated control + N1/2 T3 = Un-inoculated control + NF T4 = Inoculated with Azospirillum sp. TN03 + N1/2

T5 = Inoculated with Rhizobium sp. TN04 + N1/2 T6 = Inoculated with Azospirillum sp. TN09 + N1/2 L1 = Faisalabad L2 = Sahiwal

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103

Figure 3-29 Field view of inoculated potato (variety Kuroda) plants grown at

PRI, Sahiwal.

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104

Part 3.3 Metagenomics Studies of Potato Rhizosphere

3.3.1 Extraction of Soil DNA

For metagenomics, soil DNA was extracted from the rhizosphere of potato in triplicate

from each sample and each DNA was treated as separate sample for further analysis.

3.3.2 Quantification of the Total Bacterial Community

Overall total bacterial community estimated by the copy number of 16S rRNA gene,

ranged from log value 11.5±0.26 to 11.9±0.07 gene copy numbers g-1 of rhizospheric

soil in regions under study (Figure 3-30). Significantly higher number of 16S rRNA

gene was observed in Gujranwala region whereas lowest values were recorded in

Sahiwal area.

3.3.3 Quantification of the N-fixing Community

Abundance of diazotrophs in the rhizosphere of potato varied significantly from region

to region (Figure 3-31). Sheikhupura region showed maximum copy number of nifH

gene g-1 which showed this region have highest diazotrophs. However Jhang region

bared least abundance of diazotrophs among the regions under study.

Total bacterial richness showed no major change overall when compared to the

abundances of diazotrophs in the rhizosphere of potato, which changes from region to

region in areas under study (Figure 3-32).

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105

Figure 3-30 Abundance of total bacterial community based on 16S rRNA in

potato rhizosphere.

Gj = Gujranwala, Gr = Gojra, Jg = Jhang, Ok = Okara, Sk = Sheikhupura, Sw = Sahiwal

Values are mean of 6 replicates. Column sharing same letter, do not differ significantly

(P≥ 0.01) according to Fisher’s LSD.

Figure 3-31 Abundance of diazotrophs based on nifH gene in potato rhizosphere.


Values are mean of 6 replicates. Column sharing same letter, do not differ significantly

(P≥ 0.01) according to Fisher’s LSD.

3. Results

106

Figure 3-32 Comparison of total bacterial population to the diazotrophic

community in rhizosphere of potato across different regions.

3.3.4 Changes in the Structural Community based on 16S rRNA

The structural community in the rhizosphere of potato from different sites were

characterized using PCR-DGGE analysis based on 16S rRNA gene. Analysis of DGGE

gel revealed that number of bands per sample were in range of 17±3, 21±3, 18±2, 21±2,

24±3 and 19±2, for soils of Gujranwala, Jhang, Okara, Sheikhupura Sahiwal and Gojra

respectively (Figure 3-33). Minimum number of bands were found in Gujranwala soils

which also showed divergence when data was analyzed using non-metric multi-

dimensional scaling from all the other regions which are grouped together at 95%

similarity (Figure 3-34). It shows that structural community associated with potato is

similar in all regions under study except Gujranwala region.

3.3.5 Changes in the Diazotrophic Community based on NifH Gene

In order to characterize the diversity of N fixing community within different regions,

PCR-DGGE analysis was performed based on nifH gene. Number of bands for nifH

gene varies from 12±2, 12±3, 9±2, 12±2, 13±3 and 9±2 in Gujranwala, Jhang, Okara,

Sheikhupura, Sahiwal and Gojra respectively (Figure 3-35). Minimum number of bands

were found in Gr soils. All the regions were clustered together at 90% when analyzed

in non-metric multi-dimensional scaling (Figure 3-36) which showed that potato

rhizosphere contains similar types of diazotrophs in all the regions under study.

3. Results

107

Figure 3-33 Denaturing gradient gel of amplified 16S rRNA from potato

rhizosphere.

M = Marker, 1, 2, 3 = Gj1, 4, 5, 6 = Gj2, 7, 8, 9 = Jg1, 10, 11, 12 = Jg2, 13, 14, 15 =

Ok1, 16, 17, 18 = Ok2, 19, 20, 21 = Sk1, 22, 23, 24 = Sk2, 25, 26, 27 = Sw1, 28, 29,

30 = Sw2, 31, 32, 33 = Gr1, 34, 35, 36 = Gr2

Figure 3-34 Non-metric multidimensional scaling analysis based on fingerprints

generated by PCR-DGGE of 16S rRNA from potato rhizosphere.

3. Results

108

Figure 3-35 Denaturing gradient gel of amplified nifH gene from potato

rhizosphere.

M = Marker, 1, 2, 3 = Gj1, 4, 5, 6 = Gj2, 7, 8, 9 = Jg1, 10, 11, 12 = Jg2, 13, 14, 15 =

Ok1, 16, 17, 18 = Ok2, 19, 20, 21 = Sk1, 22, 23, 24 = Sk2, 25, 26, 27 = Sw1, 28, 29,

30 = Sw2, 31, 32, 33 = Gr1, 34, 35, 36 = Gr2

Figure 3-36 Analysis based on fingerprints generated by PCR-DGGE of nifH

gene from potato rhizosphere.

3. Results

109

3.3.6 Pyrosequencing (NifH Gene)

Pyrosequencing was done to deeply sequence the nifH gene, in order to understand the

changes in the diazotrophic community across the regions under study. In total 218830

reads were obtained for samples from 6 sites in which maximum reads were for Gj1

(25171) and minimum were for Jg2 (14305). For the diversity and richness comparisons

among samples, data were resampled randomly to same sequencing level (12000

sequences for each treatment) because in any community assessments of richness and

diversity change with ways of sampling. It resulted total number of observed species

ranges from 291 to 446 (Figure 3-37) among which maximum diversity was obtained

for Sheikhupura, however Gujranwala soils were least divers for nifH gene as shown

by the phylogenetic diversity whole tree analysis (Figure 3-37). Whereas for the

Shannon index, nifH gene diversity was higher in Sahiwal soil but lowest was observed

again for Gujranwala region (Figure 3-37).

Structural diversity was estimated by applying similarity cut off at 90%. NifH

gene diversity was minimum for Gujranwala soils which was dominated mostly by

Azorhizobium, Azospirillum, Azotobacter and Bradyrhizobium species (Figure 3-38).

The highest diversity was observed in soils from Sheikhupura region; it contained

Azospirillum, Azotobacter and Bradyrhizobium species mainly but also many other

genera like Azorhizobium and Zoogloea species (Figure 3-38). The genera having

number of sequences below 100 were added to the rare genera for every sample. In total

the retrieved nifH sequences changes considerably among soils under study based on

amino acid sequences. These sequences were distributed among α-, β- and γ-

proteobacteria (Figure 3-38). The most abundant class was α-proteobacteria with

dominant genera Azorhizobium, Azospirillum, Bradyrhizobium and Rhizobium

followed by β-proteobacteria specifically Azoarcus, Burkholderia, Pseudacidovorax

and Zoogloea genera. The nifH genes associated with γ-proteobacteria were dominated

Azotobacter, Methylomonas, Methylobacter and Celerinatantimonas spp. and

Geobacteria belonged to delta-proteobacteria were also observed in soils under study.

3. Results

110

Figure 3-37 Refraction indexes of Shannon, chao1, abundance and diversity of

diazotrophs in rhizosphere of potato from different regions.


3. Results

111

Figure 3-38 Relative abundance of diazotrophs in rhizosphere of potato from

different regions.


4. Discussion

4.1 Bacterial Isolation and Characterization

The rhizosphere is a preferential niche for various types of microorganisms including

PGPR which are directly or indirectly beneficial for plant growth due to their PGP

properties. The ultimate benefit of the use of PGPR is not only their plant growth

promoting attributes, but also their environment friendliness and their cost-effective

nature [214]. Potato is growing rapidly, as an important food commodity, needs special

attention and requires extensive fertilization, mainly through chemical fertilizers. The

application of bio-inoculants could minimize environmental problems as they possess

many advantages over chemical fertilizers. So efforts are required to explore soil

microbial diversity, their distribution and behavior in soil habitats in order to get potent

bacterial isolates which can promote plant growth in given set of environmental

conditions.

Punjab shares 80% of the total production of potato in Pakistan. Gojra,

Gujranwala, Jhang, Okara, Sahiwal and Sheikhupura regions are located in central

Punjab, which is the prime area for potato production in Pakistan. Main crop rotations

include potato-maize-maize and potato-maize-fallow, with main potato varieties

including Simply Red, Kuroda, Cardinal, Rodio, Desiree, Mutta and Sante. In Punjab

area, about 250 Kg nitrogen and 150 Kg each of phosphorus and potash is used per

hectare for potato production. The objectives of present study were to explore potato

rhizosphere in order to get understanding of diversity and interaction of diazotrophs as

N is the major input in potato crop. The analysis were based on both culturable and

non-culturable fraction of community. We aimed to get potential diazotrophs from

major potato growing areas of Pakistan and assumed that bacteria associated with

potato roots would have maximum probability to exert PGP effects on potato crop being

native to the conditions.

Forty four bacteria were isolated from the potato rhizosphere using NFM

medium. Most of the isolates formed pellicle or ring on the surface of media. Bacterial

isolates showed different growth patterns, colony and cell morphology from the same

4. Discussion

113

crop as plants harbor diverse bacteria in their rhizosphere. Many isolates formed vibroid

rods with the cell motility of typical azospirilla in NFM medium [215] where Gram’s

negative dominated (84%) the total bacterial population obtained on NFM [216].

Phenotypic identification is not reliable and decisive because bacteria change

their morphology with change in the abiotic environment like pH, temperature, carbon

source etc. Moreover many bacterial species belonging to the same genus show similar

cell and colony morphology. The molecular methods provide more accurate tools for

bacterial identification, so isolates were further differentiated on the basis of rep-PCR

[217] using GTG5 and ERIC sets of primers. The reproducibility of isolates in one PCR

run, the discriminatory power and the genetic diversity of the fingerprint are very

similar for both ERIC and GTG5 [185]. A fairly high discriminatory power was found

for all the 44 isolates that could be differentiated from each other on the basis of at least

one band difference in their respective rep-PCR fingerprints [218, 219]. Dendogram

based on GTG5 revealed the clustering of isolate TN26 and TN27, clustering of isolates

TN09, TN31, TN28 and TN29 and grouping of TN24 with TN10 at 100% whereas for

ERIC-PCR TN24 was grouped with TN09, TN03 with TN22, TN42 with TN44 and

TN36 clustered at 100% with TN40.

ERIC and Rep-PCR fingerprinting can discriminate among the taxonomically

different bacterial strains but cannot distinguish among the closely related ones. 16S

small sub-unit ribosomal nucleic acid molecules is considered to be the authentic

taxonomic marker for the identification of bacteria at genus and species level. Its

greatest contribution is a provision of sequence based taxonomy. Identification based

on 16S rRNA gene sequence analyses showed that bacterial strains belong to known

PGPR genera Azospirillum, Achromobacter, Agrobacterium, Advenella, Bacillus,

Brevundimonas, Pseudomonas, Enterobacter, Rhizobium, Shinella, Sphingobacterium

and Stenotrophomonas. The overall population was dominated by proteobacteria

belonged to α, β and γ-proteobacteria among which Azospirillum shared 29% of the

total bacterial strains. This may be due to the biasness of the medium used for isolation

[174] as NFM medium is considered specific for isolation of azospirilla. Several

members of these genera are well documented for their plant growth promotion

activities at different plants like wheat [220, 221], grapevine [222], beech [223], maize

[145, 219], sunflower [36], sugarcane [152], potato [37, 47] etc.

4. Discussion

114

Bacterial strains were then evaluated for their N fixation potential by acetylene

reduction assay (ARA) which is an indirect method to quantify the nitrogen fixation

[178]. Since all the bacteria strains may not have the ability to fix nitrogen [224]

because enrichment in NFM medium increase the probability of isolation of diazotrophs

but the pellicle formed also includes bacteria which are unable to fix nitrogen. It is fact

that the authenticity of enrichment methods mainly depend on the easiness with which

the microbe of interest is able to develop from small inoculum [225]. Thirty two

bacterial strains showed the ability to reduce acetylene to ethylene in ARA among

which bacterial strains belonging to Azospirillum [47], Rhizobium [47, 226] and

Brevundimonas [227] genera showed higher nitrogenase activity. These groups are well

documented for their nitrogen fixation ability with many crops [219, 227, 228]. While

the remaining may be efficient scavengers of traces of reduced nitrogen or they may be

oligotrophs which grow on nitrogen fixed and released by other diazotrophs present in

the mixed culture during the enrichment process [229].

Bacterial isolates which showed ARA ability were further confirmed by

amplification of nifH gene. Nitrogenase enzyme plays key role in nitrogen fixation

whose various subunits are encoded by nif gene i.e. nifD, nifH and nifK [230]. However

nifH gene which encodes the nitrogenase reductase subunit, has become the marker

gene for study as it is the most sequenced gene among three nif genes. NifH gene is also

evolutionary conserved among the diazotrophs and could be used for their identification

[231]. NifH gene was amplified in 16 isolates and sequence analysis resulted in four

major clusters. NifH gene for all the isolated bacterial strains showed clustering with

their respective genera except Brevundimonas sp. TN37 which was clustered with

Azospirillum group that shows that it may have acquired its nif gene from Azospirillum

genus.

The effects of PGPR are often related to a manipulation of the network of plant

hormones that are involved in growth and stimulation of root formation because auxins

are involved in root growth and proliferation [96]. Here, only the biosynthesis of IAA

was observed as phenylacetic acid (PAA) is known for its weak auxin activity [232,

233]. There are many pathways resulting in IAA production in rhizobacteria; some have

tryptophan as a main precursor while others have tryptophan-independent pathways

[234]. Two main types of detection techniques are widely used for IAA quantification:

I. a colorimetric method using Salkowski reagents, II. A high performance liquid

4. Discussion

115

chromatography (HPLC) method [181]. Addition of tryptophan can affect the amount

of IAA produced in those bacteria which have tryptophan-dependent pathway.

However, the amount of IAA produced varies across strains and also with the

concentration of tryptophan added [235]. Twenty nine strains showed the production of

IAA on HPLC, which ranges from 1.35 to 30.43 µg mL-1. Maximum IAA was produced

in presence of tryptophan by Azospirillum sp. which is well explored for this aspect

[236]. Detection of IAA on spectrophotometer give false positive results and high range

of IAA because of the reason that spectrophotometer cannot differentiate among IAA

and its other indole derivatives [47, 237]. The indole derivatives have no role in root

elongation and proliferation hence falsify the real results. It is useful, though, for

screening large bacterial populations, as it is fast, thus reducing sample size, time and

cost of eventual subsequent HPLC analyses.

The plant host and root exudates actually shape and drive the soil bacterial

community structure. Bacteria evolve metabolic adaptations to occupy special niches

as particular carbon sources are generated by individual plant [238, 239]. A great

diversity of utilizing the carbon source pattern exist (metabolic potential) in aquatic,

soil and rhizosphere bacteria, however metabolically versatile strains are more

successful competitor in plant microbe interaction [240, 241]. To determine the ability

to utilize different carbon sources, strains from Azospirillum, Agrobacterium, Bacillus,

Brevundimonas and Rhizobium genera were tested on PM2A microplate [242]. Bacillus

sp. TN19 utilized 80% of the provided carbon sources which show its competency in

host plant rhizosphere over other associative bacteria. It is not clear how the

consumption of plant produced carbon sources by bacteria indirectly impacts the host,

though carbon source has been shown to dictate expression of Enterobacter genes

involved in motility, colonization, adhesion and biofilm formation, thus serving as

signal for interaction with host plant [243].

4.2 Plant Inoculation and Colonization

A positive and consistent benefit of any PGPR to plant is only materialized when

bacteria enters in a tight and close association with plant roots. It is very important to

evaluate the colonization potential of bacterial isolates both in vitro and in vivo.

Sophisticated and highly sensitive microscopic techniques like transmission electron

microscopy (TEM) and confocal laser scanning microscope (CLSM) are known for the

4. Discussion

116

ultrastructure and root colonization studies of PGPR strains [244, 245]. Azospirillum

spp. are well reported for their association with cereals as well as with non-cereal plant

roots [246, 247]. In present study Azospirillum sp. strains TN03, TN09 and TN10 were

localized in root surface of potato. These isolates formed micro-colonies in the grooves

formed in the root hairs. Multiple cell aggregates were observed in extracellular matrix

on the root surface. Potato roots inoculated with Rhizobium sp. TN04 showed strong

adhesion of bacterial cell with potato roots. Bacterial clusters were observed in the

niches formed by roots. Brevundimonas sp. TN37 was able to colonize potato roots and

cells were attached to the root surface in the extracellular matrix in the form of layers

with is the initial step to form biofilm. Adequate numbers of bacterial cells were thus

present in the rhizosphere, and this may be related to the root area being rich in nutrients

and having adequate micro-aerobic conditions that allow bacteria to carry out plant

growth promoting activities like nitrogen fixation, IAA production [228, 248-250].

CLSM is also widely used in microbiology studies. Yellow fluorescent protein

(yfp) was used as a reporter which allowed easy detection of marked strains as this

process don’t require application of destructive extraneous substances [48]. Plant roots

inoculated with yfp-labelled Azospirillum sp. strains TN03 and TN09, Rhizobium sp.

TN04 and Brevundimonas sp. TN37 when examined by CLSM, revealed strong

bacterium-root associations. Adequate numbers of bacterial cells were present in the

rhizosphere, where root hairs were preferred at early colonization stage. Root hairs are

known to be sites of increased rhizodeposition and have been suspected to be involved

in eliciting chemotaxis and specific attachment [251-253]. Micro and macro colonies

were also seen in the junctions of primary and secondary roots which may be because

it provides a better niche. Cells of Brevundimonas sp. TN37 were distributed all over

the root zone which is probably due to variation in the root exudates at different places

within root system [254].

In field, due to the presence of other bacteria the competition is tough and hence

bacterial survival was quantified by CFU in a time course study upto 60 days of

inoculation. Azospirillum spp., Enterobacter sp. TN38 and Rhizobium sp. TN04

showed high population densities and survival in rhizosphere. In spite of the fact that

other strains showed a decline, apparently their population sizes up to 60 days were

sufficient for the effects seen [47]. Similar patterns of bacterial population declines

(4.67 cfu g-1 of soil) were described by Fischer, et al. [255] in wheat. There may be

4. Discussion

117

several reasons for the fluctuations in bacterial population densities, with plant growth

stage being a strong factor that affects the indigenous plant-associated communities in

field-grown potato plants [165]. All the tested strains from Azospirillum spp. and

Rhizobium sp. TN04 showed relatively stable population size, which indicates it is a

good root colonizer and PGPR in potato, as has been also reported for other crops like

maize, strawberry, grasses and vegetables [47, 219, 248, 256, 257].

Plant inoculation assays showed that all bacterial strains improved plant growth

as compared to the un-inoculated control however bacterial strains from genera

Azospirillum spp. TN03, TN09, Rhizobium sp. TN04 and Brevundimonas sp. TN37

which may be attributed to the ability of strains to fix nitrogen as well as produce high

amounts of IAA. IAA can positively influence root elongation and lateral root

development, which helps plants to acquire maximum water and essential nutrients.

This may ultimately result in a well-established, vigorous and healthy plant. Bacterial

strains producing phytohormones are known to influence the balance of plant

phytohormones, eventually inducing different growth stages [258] as well as, in an

overall fashion, promoting plant growth [159]. Greater plant dry weight may also be

ascribed to the enhanced leaf areas of the inoculated plants, especially which were

inoculated with Azospirillum sp. TN09 and Rhizobium sp. TN04 whereas greater shoot

lengths reported may be positively correlated with greater photosynthetic rates and

intensified transpiration efficiency because of availability of sufficient nitrogen, as

compared to other plants [47, 219, 259]. These results are consistent with earlier studies

in which root and shoot lengths, fresh weights and dry weights were increased by the

application of Enterobacter on sunflower [36], bio-priming of Agrobacterium,

Pseudomonas enhance the germination of radish [260] and inoculation of Rhizobium

increased plant growth parameters in wheat [220], rice [149] and potato [47]. PGPR

exhibiting multiple traits, ultimately can provide more benefits to plants in terms of

stimulating growth. The variable innate PGPR potential of the strains may cause

differential growth responses in plants [159, 219]. In this study, bacterial strains were

tested only with respect to two traits, i.e. IAA production and nitrogenase activity.

However, they might have potential for other PGPR traits which still need to be

explored e.g. P solubilization and biocontrol.

There are contradictory reports about the ability of free-living Rhizobia to fix

nitrogen. However many reports are available regarding the nitrogen fixation activity

4. Discussion

118

of free living Rhizobium both in pure culture [261] as well as in in vivo conditions [149].

Those strains were isolated from maize [226], oat [149], potato [47] and rice [149].

Similarly in our study, Rhizobium sp. TN04 showed the highest acetylene reducing

activity (151.70±16.11nmoles mg-1 protein h-1) in the in vitro assay. Scientists

suggested the expression of nif gene under low oxygen conditions which are attained

by endophytic Rhizobia by residing in intercellular spaces and/or in non-growing dead

cells of plant [262, 263] which is further supported by Fischer, et al. [264] who observed

the expression of nifH genes which belonged to Rhizobium sp. in sugarcane.

Actual share of inoculated strains in provision of nitrogen used by the potato

plant in plant inoculated studies was quantified by using 15N dilution techniques, which

is a useful tool for selection and screening of diazotrophs associated with different crops

[194, 265-270]. To assess the availability of nitrogen from different sources, a non-

nitrogen fixing reference plant is required in 15N isotope methods. Bacterial strains were

inoculated in fumigated sterilized soil whereas un-inoculated plants were taken as non-

fixing control as it was difficult to find a plant which has similar root system like potato

[194]. The results for natural abundance experiment showed higher values of Ndfa than

both the 15N dilution experiments which may be due to the effective nitrogen fixation

by bacterial strains in the absence of supplemented N, since Ndfa was decreased in Exp.

3 where full dose of N was applied. The 15N in inoculated plants tend to dilute as

expected as nitrogen is fixed biologically [194]. Plant growth parameters and total N in

plant positively correlated to N inputs which showed that potato plant got benefit both

from supplemented and biologically fixed nitrogen, similar phenomenon was observed

in maize by Montañez, et al. [266]. The 15N dilution methodology has mostly been used

for the quantification of BNF in legumes and non-legumes, however these methods

poses certain problems which are often discussed by Chalk [136, 271, 272]. So we

observed relative response of potato to the selected bacterial strains and also the

comparative estimations of nitrogen fixation ability of inoculated strains side by side.

Considerable studies has been carried out to see the effect of PGPR on potato

[37, 47, 163, 164, 273, 274] however these studies are either from the past or restricted

to the controlled conditions. Whereas in natural soil and environmental conditions

sometimes rhizobacteria may not perform well as expected. To verify the positive

effects of selected strains on growth and yield of potato, field experiments were

conducted. The diazotrophs get less responsive with respect to nitrogen fixation in the

4. Discussion

119

presence of chemical N [266], thus 3 control treatments (NF, N1/2 and N0) were laid out

to get the clear understanding of the effect of bacterial isolates. All the growth

parameters were positively affected by the inoculation of both Azospirillum spp. and

Rhizobium sp. (table 3.12 and 3.13). The Rhizobium sp. TN04 gave the higher plants

height, plant and tuber fresh and dry weight, yield and nitrogen contents among

inoculated bacterial strains however the overall increase was not significantly different.

This may be due to the well persistence and competence of the Rhizobium sp. in natural

environment over the Azospirillum sp. Overall increase in potato growth cannot be

attributed to sole nitrogen fixation and IAA production of bacterial strains because there

are many other factors which affect the plant growth in presence of PGPR [37, 158,

162-164]. Inoculation with PGPR strains cannot substitute the exogenously applied

forms of nitrogen however gave significant increase when applied with half dose of

recommended nitrogen which in other way is beneficial if cost benefit ratio for the

experiment is observed. These associative rhizobacteria give better germination and

growth ultimately good start and better stand to respective plants when applied in field

conditions as many researcher reported this phenomenon [159, 160, 270, 275].

The study of culturable diazotrophs in potato rhizosphere showed that potato

harbor diverse kind of bacteria which have diverse metabolic potential, nitrogenase

activity and phytohormone production. Bacteria belong to 12 genera were isolated

(Achromobacter, Advenella, Agrobacterium, Azospirillum, Bacillus, Brevundimonas,

Enterobacter, Pseudomonas, Rhizobium, Shinella, Sphingobacterium and

Stenotrophomonas) of which Azospirillum and Rhizobium [47], were isolated first time

from potato rhizosphere. These both are good colonizer of potato and increase plant

growth and yield both in controlled and field conditions so can be use as bio-inoculant

for potato plant growth promotion.

4.3 Metagenomics Analysis of Potato Rhizosphere

Metagenomics studies were carried out to understand the structural and functional

community changes across the potato growing regions in terms of richness and diversity

of total as well as diazotrophic community. There are many abiotic and environmental

factors involved in shaping the bacterial community structure in soil of specific region

[33, 276]. Abundance of 16S rRNA gene was quite high than the normal range in all the

rhizospheric soils which showed that rhizosphere has higher bacterial populations as

4. Discussion

120

compared to normal soil. Since root exudates secreted by plant, provides good sources

of carbon. Bacterial abundance changes even with cultivar type and plant growth stages,

due to the change in composition of root exudates [9, 10]. Maximum bacterial

population was observed for Gujranwala and Sheikhupura soils which may be due to

relatively higher soil temperature at the time of sampling, since temperature is the most

important parameter affecting the bacterial population in soil [32, 277]. Soil nutrient

status (mineral N and P) is another important factor which affects the bacterial

population. Excessive availability of minerals support the bacterial growth and increase

in population was observed with increase in N and P availability as reported earlier by

Jorquera, et al. [278]. This might be the reason for higher abundance of bacteria in

Gujranwala and Sheikhupura soils. Effect of pH of soil and other abiotic factors studied,

on the bacterial abundance was not observed since all soil samples were alkaline with

similar range of pH except Gujranwala region. Although numerous reports are

available about pH as an important element of bacterial community abundance and

composition [8, 279] but in this particular study, we did not find any correlation of pH

and rhizospheric bacterial community abundance.

The relative abundance of total bacteria remained fairly constant when

compared with abundance of diazotrophic community. Significantly higher abundance

of nifH gene in Gujranwala and Sheikhupura might be the combined effect of soil

temperature, N availability and soil texture as higher temperature and clay contents

support diazotrophic population in specific soils [32, 277]. Nevertheless, the effect of

N is still controversial on the abundance of diazotrophs since reports are available both

for positive and negative effects of higher nitrate on abundance of diazotrophic

community [32, 280, 281]. Agricultural practices adopted in these two regions as it

come under Wheat-Cotton zone of Punjab, Pakistan, might also be the reason for higher

abundance of both structural and functional communities. Since it is well established

that management practices affect soil quality and nutrient dynamics which has ultimate

effect on abundance, diversity and composition of soil bacteria [33, 282].

For the structural studies of bacterial community PCR-DGGE was carried out

which showed similar diversity for the rhizospheric soils of all the regions except for

Gujranwala. However for functional community no significant difference was

observed, which suggests that factors which affect structural community do not

necessarily affect the diazotrophs. As specific plant species harbor specific groups of

4. Discussion

121

bacteria and on the other hand PCR-DGGE can detect most abundant taxa (more than

1% of community), so these might be possible explanations for less diversity obtained.

One organism may produce multiple bands on DGGE gel which may give the false

picture for number and intensity of amplicons and ultimately the parameters calculated

from these fingerprints. Due to these shortcomings, results from DGGE fingerprints

should be deduced as clues and not as complete measurements of structure and diversity

of microbial communities.

In order to get the deep understanding of the microbial structure,

pyrosequencing of nifH gene was carried out as diazotrophic communities were the

main focus of this study. Pyrosequencing is a high data comparable DNA sequencing

technique [283] that has revolutionized microbial detection due to its greater sample

coverage and potential to identify rare microbes. Analysis of pyrosequencing resulted

in strong diversity among the soils from different regions. Sheikhupura region showed

the most diverse diazotrophs while Gujranwala resulted least diversity among all the

sites. Few members of class α-proteobacteria i.e. Azorhizobium, Azospirillum and

Bradyrhizobium and γ-proteobacteria i.e. Azotobacter were found dominant across all

the soils which was not surprising as these all are well reported from potato rhizosphere

[10, 32, 122]. Abundance of Bradyrhizobium in potato rhizosphere is interesting, as

these all areas come under cotton-wheat, mix cropping and wheat-rice rotation zones

of Punjab and there was not significant cropping of nodulating crops. So here it may

serve as soil saprophyte as Bradyrhizobium is known only as symbiotic N-fixer [284].

On the other hand Azospirillum and Azotobacter are the genera of associative nitrogen

fixer found almost everywhere on Earth and also reported from potato rhizosphere [47,

285-287]. Some genera were abundant in soils of specific regions like Azohydromonas

were detected in Gujranwala, Geobacter and Methylocella in Jhang, Methylogaea in

Okara, Zoogloea in Sheikhupura and Pseudacidovorax for both Okara and Sahiwal.

Which show these genera were more sensitive to abiotic and environmental factors.

Bacterial communities structure specially diazotrophs in any region are shaped by many

factors which include soil structure, pH, nutrient composition of soil [32, 278], crop

rotation and agronomic practices adopted [33, 282] and even the cultivar of same plant

species [10]. However in our study higher electrical conductivity and potassium ion

concentration may be the reason of higher diversity in Sheikhupura and Okara soils as

these both factors are positively correlated with structural composition of bacteria in

4. Discussion

122

soil since salt concentration and electrical conductivity are known as strong gradients

involved in shaping microbial community structure at global scale [288].

The metagenomics analysis revealed that population of total bacteria remain

almost similar in the rhizosphere of potato but their relative abundance changes due to

many abiotic and biotic factors. Potato harbor diazotrophs from different genera, well

known for their nitrogen fixation ability which can be isolated and used for the

production of bio-inoculants for potato cultivation.

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