Biodiversity of Diazotrophs in Rhizosphere of Potato (Solanum...
Transcript of Biodiversity of Diazotrophs in Rhizosphere of Potato (Solanum...
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
Designation: Professor
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
E-mail Address: [email protected]
2. Prof. Dr. M. Kaleem Abbasi
Designation: Professor
University: Department of Soil & Environmental Sciences, University of Poonch.
Mailing Address: Faculty of Agriculture, Rawalakot Azad Jammu and Kashmir,
Pakistan
Telephone: +92 (5824) 960046
E-mail Address: [email protected]
3. Prof. Dr. Amer Jamil
Designation: Professor
University: University of Agriculture Faisalabad
Mailing Address: Molecular Biochemistry Lab, Department of Chemistry,
University of Agriculture Faisalabad
Telephone: +92 (41) 9201104
E-mail Address: [email protected]
Head of the Department (Name): Prof. Dr. Shahid Mansoor (S.I)
Signature with Date: _________________________________
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
Date: 16 December, 2016
Place: NIBGE, Faisalabad
Head, Department of Biotechnology: ___________________
Name: Dr. Shahid Mansoor
Date: 16 December, 2016
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
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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
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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
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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
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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
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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
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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
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Figure 3-9 Phylogenetic tree base on 16S rRNA sequences (1119 bp) of
Brevundimonas sp. isolated from potato rhizosphere .......................... 68
Figure 3-10 Phylogenetic tree base on 16S rRNA sequences (1159 bp) of
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
Figure 3-18 Electron micrographs of ultra-thin sections of potato root inoculated
with Rhizobium sp. TN04 .................................................................... 81
Figure 3-19 Electron micrographs of ultra-thin sections of potato root inoculated
with Azospirillum sp. TN09 ................................................................. 82
Figure 3-20 Electron micrographs of ultra-thin sections of potato root inoculated
with Azospirillum sp. TN10 ................................................................. 83
Figure 3-21 Electron micrographs of ultra-thin sections of potato root inoculated
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
Figure 3-24 Confocal image potato (variety Kuroda) root inoculated with YFP-
labelled Rhizobium sp. TN04 ............................................................... 88
Figure 3-25 Confocal image potato (variety Kuroda) root inoculated with YFP-
labelled Azospirillum sp. TN09............................................................ 89
Figure 3-26 Confocal image potato (variety Kuroda) root inoculated with YFP-
labelled Brevundimonas sp. TN37 ....................................................... 90
Figure 3-27 Principal component analysis showing the relationship between
different experimental locations and treatments. ............................... 101
Figure 3-28 Principal component analysis showing the relationship between
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
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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
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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
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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
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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.
2. Materials and Methods
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
2. Materials and Methods
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
2. Materials and Methods
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
2. Materials and Methods
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
2. Materials and Methods
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-
2. Materials and Methods
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.
2. Materials and Methods
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).
2. Materials and Methods
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].
2. Materials and Methods
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
final extension 72ºC 10
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
final extension 72ºC 10
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),
0.01mg BSA (20mg/ml),
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
final extension 72ºC 30
min
94°C 5 min,
94°C 60 s, 48°C 1 min,
72°C 2 min
30 cycles
final extension 72ºC 30
min
2. Materials and Methods
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
2. Materials and Methods
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;
2. Materials and Methods
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.
2. Materials and Methods
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
T5 = Inoculated with strain TN04 + N1/2
T6 = Inoculated with strain TN09 + 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
2. Materials and Methods
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
2. Materials and Methods
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
2. Materials and Methods
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.
2. Materials and Methods
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).
2. Materials and Methods
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
2. Materials and Methods
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
2. Materials and Methods
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.
2. Materials and Methods
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
final extension 72ºC 30
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),
0.01mg BSA (20mg/ml),
0.5μM each primer,
0.5U Taq polymerase
(Roche),
0.25mM dNTPs,
1x buffer (Roche),
0.01mg BSA (20mg/ml),
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
final extension 72ºC 30
min
94°C 5 min,
94°C 60 s, 48°C 1 min,
72°C 2 min
30 cycles
final extension 72ºC 30
min
Primers qPCR-nifH (5’-3’) PCR mixture Thermal conditions
FGPH19
(TACGGCAARGGTGGNATHG)
PolR
(ATSGCCATCATYTCRCCGGA)
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
(ATSGCCATCATYTCRCCGGA)
RoeschF
(ACCCGCCTGATCCTGCACGCCA
AGG)
RoeschR
(ACGATGTAGATTTCCTGGGCCT
TGTT)
0.20mM dNTPs,
1x buffer (Roche),
0.03mg BSA (20mg/ml),
0.5μM each primer,
0.25U FastStart High
Fidelity PCR System,
50ng template DNA,
0.20mM dNTPs,
1x buffer (Roche),
0.03mg BSA (20mg/ml),
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
final extension 72ºC 10
min
95°C 5 min,
94°C 45 s, 50°C 45 s,
72°C 45 s
20 cycles
final extension 72ºC 10
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
TN40 Sahiwal Medium, round, white Short rods -ve
TN41 Sahiwal Small, round, white Short rods -ve
TN42 Sahiwal Medium, 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
TN3 Gujranwala 138.21±15.21 TN25 Okara 85.21±9.66
TN4 Gujranwala 151.70±16.11 TN26 Okara 86.01±7.28
TN5 Gujranwala 69.16±6.15 TN27 Okara 0
TN6 Gujranwala 0 TN28 Sheikhupura 85.97±8.03
TN7 Gujranwala 0 TN29 Sheikhupura 99.81±8.76
TN8 Gujranwala 127.80±14.45 TN30 Sheikhupura 98.79±11.04
TN9 Jhang 143.07±11.98 TN31 Sheikhupura 124.22±7.43
TN10 Jhang 134.95±13.99 TN32 Sheikhupura 116.28±9.9
TN11 Jhang 132.59±12.72 TN33 Sheikhupura 112.99±12.94
TN12 Jhang 102.30±7.08 TN34 Sheikhupura 0
TN13 Jhang 105.48±8.61 TN35 Sheikhupura 88.98±9.71
TN14 Jhang 0 TN36 Sheikhupura 0
TN15 Gojra 91.92±8.29 TN37 Sahiwal 135.07±8.97
TN16 Gojra 82.60±8.38 TN38 Sahiwal 96.60±7.61
TN17 Gojra 0 TN39 Sahiwal 127.56±11.63
TN18 Gojra 100.78±7.7 TN40 Sahiwal 0
TN19 Gojra 96.78±8.41 TN41 Sahiwal 0
TN20 Gojra 0 TN42 Sahiwal 92.60±8.42
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)
TN1 Gujranwala 24.40±2.06 TN23 Okara 0
TN2 Gujranwala 13.51±0.7 TN24 Okara 0
TN3 Gujranwala 30.43±3.16 TN25 Okara 4.82±0.46
TN4 Gujranwala 3.50±0.41 TN26 Okara 3.45±0.33
TN5 Gujranwala 5.46±0.65 TN27 Okara 0
TN6 Gujranwala 0 TN28 Sheikhupura 16.35±1.11
TN7 Gujranwala 0 TN29 Sheikhupura 14.56±0.72
TN8 Gujranwala 12.49±0.6 TN30 Sheikhupura 11.32±0.51
TN9 Jhang 19.77±1.45 TN31 Sheikhupura 15.36±0.8
TN10 Jhang 25.25±2.1 TN32 Sheikhupura 6.30±0.43
TN11 Jhang 1.35±0.32 TN33 Sheikhupura 0
TN12 Jhang 15.51±0.62 TN34 Sheikhupura 3.43±0.29
TN13 Jhang 9.72±0.35 TN35 Sheikhupura 1.43±0.3
TN14 Jhang 0 TN36 Sheikhupura 1.51±0.26
TN15 Gojra 14.45±1.71 TN37 Sahiwal 0
TN16 Gojra 2.14±0.76 TN38 Sahiwal 2.21±0.32
TN17 Gojra 1.42±0.42 TN39 Sahiwal 0
TN18 Gojra 4.23±0.25 TN40 Sahiwal 0
TN19 Gojra 0 TN41 Sahiwal 2.85±0.43
TN20 Gojra 0 TN42 Sahiwal 3.57±0.5
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 + - - - - + + - -
3. Results
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).
3. Results
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 =
TN29, 28 = TN30, 29 = TN31, 30 = TN32, 31 = TN33, 32 = TN20, 33 = TN34, 34 = TN35, 35 = TN37, 36 = TN39, 37 = TN40, 38 = TN36, 39 = TN38, 40 =
TN41, 41 = TN43, 42 = TN44, 43 = TN42, 44 = TN42, B = Blank
3. Results
58
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 =
TN15, 15 = TN14, 16 = TN16, 17 = TN17, 18 = TN18, 19 = TN19, 20 = TN10, 21 = TN21, 22 = TN22, 23 = TN23, 24 = TN25, 25 = TN26, 26 = TN27, 27 =
TN28, 28 = TN29, 29 = TN30, 30 = TN31, 31 = TN32, 32 = TN33, 33 = TN20, 34 = TN34, 35 = TN35, 36 = TN37, 37 = TN39, 38 = TN40, 39 = TN36, 40 =
TN38, 41 = TN41, 42 = TN43, 43 = TN44, 44 = TN42, B = Blank, +ve = Positive control
3. Results
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.
3. Results
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.
3. Results
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)
TN02 Gujranwala Azospirillum sp. LN833442
Azospirillum
brasilense strain Az72
(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)
TN08 Gujranwala Azospirillum sp. LN833447
Azospirillum
brasilense strain Gr22
(99)
TN09 Jhang Azospirillum sp. LN833448
Azospirillum
brasilense strain Gr22
(99)
TN10 Jhang Azospirillum sp. LN614537
Azospirillum
brasilense strain Gr42
(99)
TN11 Jhang Sphingobacterium sp. LN833449
Sphingobacterium
mizutaii strain Ht8-22
(99)
TN12 Jhang Azospirillum sp. LN833450
Azospirillum
brasilense strain Az72
(99)
TN13 Jhang Azospirillum sp. LN833451
Azospirillum
brasilense strain Az72
(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
brasilense strain Gr42
(99)
TN30 Sheikhupura Azospirillum sp. LN833464
Azospirillum
brasilense strain Gr42
(99)
TN31 Sheikhupura Azospirillum sp. LN833465
Azospirillum
brasilense strain Gr42
(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
Figure 3-6 Phylogenetic tree base on 16S rRNA sequences (1014 bp) of
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.
Neighbor joining method was adopted. Bootstrap values greater than 50 were given and
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.
Neighbor joining method was adopted. Bootstrap values greater than 50 were given and
were based on 1000 replicates. Paenibacillus larvaeT (AY530294) was used as
outgroup.
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68
Figure 3-9 Phylogenetic tree base on 16S rRNA sequences (1119 bp) of
Brevundimonas 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. B. vesicularis (NR_037104) was used as root.
3. Results
69
Figure 3-10 Phylogenetic tree base on 16S rRNA sequences (1159 bp) of
Stenotrophomonas 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. 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).
3. Results
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
Azospirillum sp. TN02 Azospirillum brasilense strain Gr37 (100) LT596585
Azospirillum sp. TN03 Azospirillum brasilense strain Gr37 (99) LT596586
Rhizobium sp. TN04 Rhizobium sp. S1SS148 (100) LT596587
Azospirillum sp. TN09 Azospirillum brasilense strain Gr42 (100) LT596588
Azospirillum sp. TN10 Azospirillum brasilense strain Gr42 (100) LN681358
Azospirillum sp. TN12 Azospirillum brasilense strain Gr59 (100) LT596589
Azospirillum sp. TN13 Azospirillum brasilense strain Gr59 (100) LT596590
Azospirillum sp. TN15 Azospirillum brasilense strain Gr42 (99) LT596591
Stenotrophomonas sp. TN21 Uncultured bacterium clone Spruce2009_F1
(99) LT596592
Azospirillum sp. TN22 Azospirillum brasilense strain Gr59 (98) LT596593
Azospirillum sp. TN30 Azospirillum brasilense strain Gr42 (99) LT596594
Azospirillum sp. TN31 Azospirillum brasilense strain Gr42 (99) LT596595
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
3. Results
72
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.
3. Results
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).
3. Results
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
T4 = Azospirillum sp. TN02 + zero N
T5 = Azospirillum sp. TN03 + zero N
T6 = Rhizobium sp. TN04 + zero N
T7 = Azospirillum sp. TN09 + zero N
T8 = Azospirillum sp. TN10 + 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
T15 = Rhizobium sp. TN42 + zero N
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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)
T3 = Azospirillum sp. TN01 + zero N
T4 = Azospirillum sp. TN02 + zero N
T5 = Azospirillum sp. TN03 + zero N
T6 = Rhizobium sp. TN04 + zero N
T7 = Azospirillum sp. TN09 + zero N
T8 = Azospirillum sp. TN10 + 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
T15 = Rhizobium sp. TN42 + zero N
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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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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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Figure 3-17 Electron micrographs of ultra-thin sections of potato root inoculated
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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Figure 3-18 Electron micrographs of ultra-thin sections of potato root inoculated
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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Figure 3-19 Electron micrographs of ultra-thin sections of potato root inoculated
with Azospirillum sp. TN09, 30 days post inoculation.
RS = Rhizosphere, RC = Root cell and B = Bacterium
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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Figure 3-20 Electron micrographs of ultra-thin sections of potato root inoculated
with Azospirillum sp. TN10, 30 days post inoculation.
Box in the Figure 3-17 (A) represents the area with higher resolution (figure 3-17 B)
RS = Rhizosphere, RC = Root cell and B = Bacterium
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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Figure 3-21 Electron micrographs of ultra-thin sections of potato root inoculated
with Brevundimonas sp. TN37, 30 days post inoculation.
RS = Rhizosphere, RC = Root cell and B = Bacterium
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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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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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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Figure 3-23 Confocal image potato (variety Kuroda) root inoculated with YFP-
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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Figure 3-24 Confocal image potato (variety Kuroda) root inoculated with YFP-
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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Figure 3-25 Confocal image potato (variety Kuroda) root inoculated with YFP-
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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Figure 3-26 Confocal image potato (variety Kuroda) root inoculated with YFP-
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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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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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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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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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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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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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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100
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
3. Results
101
Figure 3-27 Principal component analysis showing the relationship between
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
3. Results
102
Figure 3-28 Principal component analysis showing the relationship between
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
3. Results
103
Figure 3-29 Field view of inoculated potato (variety Kuroda) plants grown at
PRI, Sahiwal.
3. Results
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).
3. Results
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.
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.
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.
Gj = Gujranwala, Gr = Gojra, Jg = Jhang, Ok = Okara, Sk = Sheikhupura, Sw = Sahiwal
3. Results
111
Figure 3-38 Relative abundance of diazotrophs in rhizosphere of potato from
different regions.
Gj = Gujranwala, Gr = Gojra, Jg = Jhang, Ok = Okara, Sk = Sheikhupura, Sw = Sahiwal
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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