Diversity, symbiotic efficiency and effect of water hyacinth compost on population of rhizobia nodulating phaseolus vulgaris in Lake Victoria Basin

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HYACINTH COMPOST ON POPULATION OF RHIZOBIA

NODULATING Phaseolus vulgaris IN LAKE VICTORIA BASIN

MORRIS MUTHINI (B.Sc. Plant Biotech.)

I56/24336/2011

A THESIS SUBMITTED IN PARTIAL FULFILLMENT OF THE REQUIREMENTS

FOR THE AWARD OF THE DEGREE OF MASTER OF SCIENCE

(BIOTECHNOLOGY) IN THE SCHOOL OF PURE AND APPLIED SCIENCES OF KENYATTA UNIVERSITY.

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DEDICATION

I dedicate this work to my wife Juliet Gathigia, children Victor Mwendwa and Abigael

Mutheu, my father Muthini Maingi and mother Agnes Ngii who have been of great

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ACKNOWLEDGEMENTS

I would like to acknowledge my supervisors the late Dr. Allan Jalemba Mgutu, Dr.

Omwoyo Ombori and Dr. John M. Maingi for their research ideas, moral support,

motivation and guidance throughout the project. Thanks to the Chairman Department of

Plant Sciences, Kenyatta University, Dr. M. Gathaara for allowing me to use the

departmental laboratory and greenhouse facilities. My sincere gratitude goes to Mr.

Stephen Gichobi, Mr Lawrence Alaro, Mr. Patrick Mwangi, Mr. Daniel Ng’ang’a, Mr.

Roy Mulanda and Ms. Wambui Mwangi for their technical support. I appreciate

smallholder farmers from Korando B Sublocation in Kenya, for allowing me to use their

farms. Thanks also go to my fellow students in the IUCEA (VicRes) funded research

project; Newton Osoro, Towet Gideon and Fanuel Kawaka, for their ideas that helped

me achieve my project objectives. I appreciate my colleagues in the lab, Daniel Agyrifo,

Jane Kimani, Olive Sande and Sylvia Nawiri for their advice and encouragement. I

express my appreciation to Dr. John Muoma (Department of Biological Sciences,

Masinde Muliro University of Science and Technology), Dr. Alice Amoding

(Department of Soil Science, Makerere University) and Ms. Dative Mukaminega

(Faculty of Applied Sciences, Kigali Institute of Science and Technology)

(Collaborating researchers) for their input. This project work would not have been

possible without funding support from IUCEA/VICRES and I greatly appreciate their

support. I thank the National Commision for Science, Technology and Innovation for

awarding me a research grant to carry out my research work. I thank my wife Juliet

Ngathigia and children Victor Mwendwa and Abigael Mutheu, my parents, brothers and

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TABLE OF CONTENTS

TITLE……. ... i

DECLARATION ... ii

DEDICATION ... iii

ACKNOWLEDGEMENTS ... iv

TABLE OF CONTENTS ... v

LIST OF TABLES ... vii

LIST OF FIGURES ... ix

LIST OF PLATES ... x

ABBREVIATIONS AND ACRONYMS ... xi

ABSTRACT ... xiii

CHAPTER ONE ... 1

INTRODUCTION ... 1

1.1 Background of the study ... 1

1.2 Problem statement and justification of the study ... 4

1.3 Research questions ... 6

1.4 Hypotheses ... 6

1.5 Objectives ... 7

1.5.1 General objective ... 7

1.5.2 Specific objectives ... 7

1.6 Significance of the study ... 7

CHAPTER TWO ... 9

LITERATURE REVIEW ... 9

2.1 Common bean Phaseolus vulgaris L ... 9

2.2 Bean agronomy ... 9

2.3 Uses of P. vulgaris ... 10

2.4 Biological nitrogen fixation (BNF) ... 11

2.4.1 Rhizobia ... …… 11

2.4.2 Role of rhizobia in biological nitrogen fixation ... 13

2.4.3 Indigenous rhizobia as source of rhizobia inocula for common beans ... 14

2.5 Factors affecting indigenous rhizobia populations in the soil and Rhizobium–legume symbiosis ... 15

2.5.1 Effect of soil amelioration on soil microbial abundance ... 17

2.6 Estimation of rhizobia population in the soil ... 20

2.7 Morphological characterization of rhizobianodulating P. vulgaris ... 20

2.8 Genetic diversity of P. vulgaris nodulating rhizobia isolates from LVB ... 21

CHAPTER THREE ... 23

MATERIALS AND METHODS ... 23

3.1 The study area ... 23

3.1.1 Collection of soil samples ... 23

3.2 Soil analysis ... 24

3.3 Rhizobia trapping ... 25

3.3.1 Greenhouse rhizobia trapping experiments ... 25

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3.4 Isolation and identification of rhizobia using morphological characteristics ... 26

3.5 Gram staining of the rhizobia isolates ... 27

3.6 Bromothymol blue test ... 27

3.7 Authentication of the rhizobia isolates ... 27

3.8 Determination of symbiotic efficiency of representative rhizobia isolates ... 30

3.9 Effect of soil treatment on indigenous rhizobia populations in the soil ... 31

3.10 Genetic diversity of common bean indigenous rhizobia isolates from LVB ... 32

3.10.1 Preparation of rhizobia isolates and the reference strains for colony PCR... 32

3.10.2 PCR amplification of 16S rDNA (16S rRNA gene) ... 33

3.10.3 Agarose gel electrophoresis ... 34

3.10.4 Restriction fragment analysis of the 16S rDNA (16S rRNA gene) ... 34

3.11 Data analysis ... 34

CHAPTER FOUR ... 37

RESULTS.. ... 37

4.1 Soil analysis ... 37

4.2 Whole soil indigenous rhizobia trapping ... 39

4.3 Morphological characteristics of the rhizobia isolates... 41

4.4 Bromothymol Blue (BTB) and Congo red reaction of the rhizobia isolates ... 49

4.5 Gram reaction... 50

4.6 Authentication of rhizobia isolates ... 50

4.7 Morphological diversity of the rhizobia isolates ... 55

4.7.1 Morphological diversity of rhizobia isolates from whole soil trapping ... 55

4.7.2 Morphological diversity of farm trapping rhizobia isolates ... 59

4.8 Genetic diversity of the rhizobia isolates ... 63

4.8.1 Genetic diversity of whole soil trapping rhizobia isolates ... 63

4.8.2 Genetic diversity of farm trapping rhizobia isolates ... 70

4.9 Symbiotic efficiency of representative isolates ... 78

4.10 Effect of water hyacinth compost, DAP and commercial rhizobia inoculum (Rhizobia leguminosarum (strain 446)) treatment on rhizobia population in the soil 84 CHAPTER FIVE ... 92

DISCUSSION, CONCLUSIONS AND RECOMMENDATIONS ... 92

5.1 Discussion ... 92

5.1.1 Whole soil indigenous rhizobia trapping ... 92

5.1.1.1 Effect of soil on P. vulgaris variety rosecoco nodulation ... 92

5.1.2 Morphological and biochemical characteristics of the rhizobia isolates ... 94

5.1.3 Authentication of rhizobia isolates ... 95

5.1.4 Morphological diversity of the rhizobia isolates ... 97

5.1.5 Genetic diversity of the rhizobia isolates ... 100

5.1.6 Symbiotic efficiency of selected rhizobia isolates ... 103

5.1.7 Effect of water hyacinth compost, DAP and rhizobia inoculum on common bean indigenous rhizobia populations in the soil ... 105

5.2 Conclusions ... 110

5.3 Recommendations ... 112

REFERENCES ... 114

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LIST OF TABLES

Table 4.1: Soil characteristics of experimental sites compared with critical values for East African soils ... 39

Table 4.2: Effect of soil on common bean nodulation and biomass in the greenhouse ... 40

Table 4.3: Whole soil trapping rhizobia isolates from nodules of P. vulgaris

plants before the application of water hyacinthcompost ... 42

Table 4.4: Rhizobia isolates obtained from the nodules of P. vulgaris variety Rosecoco grown in farms amended with water hyacinth compost in Korando B sulocation in Kisumu, Kenya ... 45

Table 4.5: Abundance (%) of rhizobia isolates from nodules of P. vulgaris

plants grown in farms treated with water hyacinth compost in Korando B sub-location in Kisumu, Kenya ... 46

Table 4.6: Infectiveness and effectiveness of representative isolates obtained from whole soil trapping experiments in the greenhouse ... 51

Table 4.7: Infectiveness and effectiveness of representative rhizobia isolates obtained from farm trapping experiments ... 52

Table 4.8: Diversity indices of rhizobia isolates from whole soil trapping ... 56

Table 4.9: Pearson correlation coefficients of soil characteristics and morphological diversity indices of rhizobia isolates from whole soil trapping experiment from Korando B Sublocation in Kisumu- Kenya ... 58

Table 4.10: Diversity indices of rhizobia isolates from farm trapping ... 59

Table 4.11: Pearson correlation coefficients of soil characteristics and morphological diversity indices of rhizobia isolates from farm trapping experiments from Korando B Sublocation in Kisumu- Kenya ... 62

Table 4.12: Mean of different alleles (Na), number of effective alleles (Ne), Shannon's Information Index I (H), expected heterozygosity (He), unbiased expected heterozygosity (UHe) and percentage of polymorphic loci (% P ) of 5 wholesoil trapping rhizobia isolates populations from Kisumu , Kenya, Uganda and Rwanda based on ARDRA data ... 65

Table 4.13: Analysis of molecular variance (AMOVA) for 97 rhizobia isolates from whole soil trapping for five populations from Kenya, Uganda and Rwanda based on restriction digestion of 16S rDNA... 66

Table 4.14: Pairwise Population Matrix of Nei Unbiased Genetic Distance of five whole-soil trapping rhizobia populations from Kenya, Uganda and Rwanda ... 67

Table 4.15: Mean number of different alleles (Na), number of effective alleles (Ne), Shannon's Information Index I (H), expected Heterozygosity (He), Unbiased Expected Heterozygosity (UHe) and percentage of Polymorphic Loci (% P ) of farm trapping rhizobia populations from Kisumu , Kenya based on ARDRA data ... 73

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Table 4.17: Pairwise Population Matrix of Nei Unbiased Genetic Distance of four rhizobia populations from farm trapping from Korando B Sublocation in Kisumu, Kenya ... 75

Table 4.18: Symbiotic efficiency of indigenous rhizobia isolates from Lake Victoria Basin ... 80

Table 4.19: Pearson correlation coefficients among investigated parameters in common beans ... 82

Table 4.20: Effect of water hyacinth compost, inorganic fertilizer (DAP) and commercial rhizobia inoculum on common bean nodulating indigenous rhizobia populations in Korando B Sublocation in Kisumu-Kenya ... 86

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LIST OF FIGURES

Figure 4.1: Dendrogram showing morphological relationship of whole soil trapping rhizobia isolates from Korando B Sublocation Kisumu, Kenya, Rwanda and Uganda. ... 48

Figure 4.2: Dendrogram showing morphological relationship of farm trapping rhizobia isolates from four farms in Korando B Sublocation in Kisumu- Kenya. ... 49

Figure 4.3: Diversity profile of whole soil trapping rhizobia isolates from LVB. ... 56

Figure 4.4: Distance dendogram and principle componet diagram showing morphological diversity of whole soil trapping rhizobia isolates.. ... 57

Figure 4.5: Diversity profiles of rhizobia isolates from farm trapping. ... 60

Figure 4.6: Distance dendogram and principle component diagram showing morphological diversity of farm trapping rhizobia isolates.. ... 61

Figure 4.7: Principle coordinate analyses (PCA) of 97 rhizobia isolates from whole soil trapping.. ... 66

Figure 4.8: A neighbour joining dendrogram based on Nei’s 1978 unbiased genetic distance and Euclididian similarity index showing the genetic distance between whole soil trapping rhizobia populations from Kenya, Uganda and Rwanda.. ... 68

Figure 4.9: Evolutionary relationships among 97 whole soil trapping indigenous rhizobia isolates from LVB and 4 reference rhizobia strains (CIAT 899, WSM 1385, USDA 2667, R. leguminosarum

(strain 446) inferred using the Neighbor-Joining method. ... 69

Figure 4.10: Principal coordinate analyses (PCA) of farm trapping isolates.. ... 74

Figure 4.11: A neighbour joining dendrogram based on Nei’s 1978 unbiased genetic distance and Euclidian similarity index showing the genetic relationship between farm trapping rhizobia populations from Korando B Sublocation in Kisumu-Kenya. ... 75

Figure 4.12: Phylogenetic relationship of 125 farm trapping rhizobia isolates from LVB and 4 reference rhizobia strains (CIAT 899, WSM 1385, USDA 2667, R. leguminosarum (strain 446) inferred using the Neighbor-Joining method. ... 77

Figure 4.13 I: The relationship between soil characteristics and rhizobia populations in soils obtained from Korando B Sublocation in Kisumu, Kenya. ... 90

Figure 4.13 II: The relationship between soil characteristics and rhizobia populations in soils obtained from Korando B Sublocation in

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LIST OF PLATES

Plate 4.1: Effect of soils from farms in Korando B sub-location Kisumu, Kenya on growth and nodulation of P. vulgaris ... 40

Plate 4.2: Rhizobia isolates from P. vulgaris nodules in whole soil trapping experiment ... 43

Plate 4.3: Rhizobia isolates from P. vulgaris nodules from farm trapping after soil amendment with water hyacinth compost, DAP and Rhizobium

inoculum. ... 47

Plate 4.4: Effect of representative whole soil trapping isolates on P. vulgaris. ... 53

Plate 4.5: Effect of representative farm trapping rhizobia isolates on P. vulgaris

variety Rose coco. ... 55

Plate 4.6: Agarose gel electrophoresis of PCR amplified 16S rDNA of rhizobia isolates from whole soil trapping from farm A stained with SYBR green and ran on 1% agarose gel. ... 63

Plate 4.7: Gel electrophoresis of restriction digestion of 16S rDNA of rhizobia isolates from whole soil trapping stained with SYBR green and ran on 2.4 % agarose gel. ... 64

Plate 4.8: Gel electrophoresis of PCR amplified 16S rDNA of rhizobia isolates from farm trapping on 1 % agarose gel ... 70

Plate 4.9: Gel electrophoresis of the restriction digestion products of 16S rDNA from farm trapping rhizobia isolates in 2.4 % agarose gel. ... 72

Plate 4.10: Symbiotic effectiveness of selected rhizobia isolates. ... 83

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ABBREVIATIONS AND ACRONYMS

AFLP Amplified Fragment Length Polymorphism

AMOVA Analysis of Molecular Variance

ANOVA Analysis of Variance

ARDRA Amplified rDNA (Ribosomal DNA) Restriction Analysis

ASARECA Association for Strengthening Agricultural Research in Eastern

and Central Africa

BNF Biological Nitrogen Fixation

BTB Bromothymol blue (BTB)

CIAT International Center for Tropical Agriculture

DAP Diammonium Phosphate

DNA Deoxyribonucleic Acid

EM Effective Microorganisms

FAO Food and Agricultural Organization

ICRAF World Agroforestry Centre

KM Kilometers

LVB Lake Victoria Basin

LVBC Lake Victoria Basin Commission

LVEMP Lake Victoria Environmental Management Program

Masl Meters above sea level

MIRCEN Microbiological Resources Centre (MIRCEN), University of

Nairobi

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MPNES Most Probable Number Enumeration System

MUARIK Makerere University Agricultural Research Institute, Kabanyolo

PAST Paleontological Statistics Software Package

PCA Principal coordinate analysis

PCR Polymerase Chain Reaction

RAPD Randomly Amplified Polymorphic DNA

RFLP Restriction Fragment Length Polymorphism

SE Symbiotic Efficiency

SSP Single Superphosphate

SSR Single Sequence Repeats

TBE Tris-Borate-EDTA

TSP Triple Super Phoshate

UNEP United Nations Environmental Programme

UPGMA Unweighted Pair Group Method with Arithmetic Mean

W/V Weight/Volume

WMO World Climate Data and Monitoring Programme, United Nations

YMA Yeast Manitol Agar media

YMB Yeast Manitol Broth media

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ABSTRACT

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CHAPTER ONE INTRODUCTION 1.1Background of the study

Lake Victoria is the second largest fresh water lake in the world and is about 69000 km2

in size. Lake Victoria Basin (LVB) has an area of approximately 251,000 km2 (UNEP,

2006). Twenty two percent (22 %) of the catchment area is in Kenya, 11 % in Rwanda,

16 % in Uganda, 7 % in Burundi and 44 % in Tanzania (Matsuishi et al., 2006). The LVB

is characterized by high human population growth and by the year 2008 the population

was estimated to be more than 40 million (Albinus et al., 2008). Most of the inhabitants

in the LVB rely on subsistence agriculture (Ondigi et al., 2008).

Continued increase in human population, poor agricultural production methods, and

deforestation are major causes of land degradation and low food productivity in the LVB

(Aseto, 2003; ICRAF, 2004). Nitrogen requirements in the soil are usually higher as

compared to other major soil nutrients for sustainable food production (Otieno et al.,

2009). Studies have shown that despite availability of other nutrient sources to enhance

nitrogen in soil for improved crop yield, chemical fertilizers have been prioritized as a

solution to nutrient deficiencies in the soil (Gentili et al., 2006; Otieno et al., 2009). Use

of inorganic fertilizers in agricultural crop production leads to greenhouse gas emissions,

reduced water quality, reduced biodiversity and is a potential health and environmental

hazard (Khanal, 2009; Pearson et al., 2010). The cost of inorganic fertilizers has also

been in upward trend making it unaffordable by many small scale farmers (Argaw, 2012).

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and it accounts for 75 % of the total nitrogen flow into the lake from the lake’s catchment

leading to increase in nutrient concentration, turbidity and reduction of dissolved oxygen

(LVEMP, 2002; Tamatamah, 2002; Kiwango and Wolanski, 2008; Kimwaga et al.,

2012). This in turn has led to algae blooms, infestation with waterweeds especially the

water hyacinth, death of fish and water-borne diseases (Kimwaga et al.,2012).

Water hyacinth (Eichhornia crassipes) is an aquatic macrophyte that is invasive and with

deleterious effects to the local environment (Kateregga and Sterner, 2007). Water

hyacinth causes de-oxygenation of the water body, interferes with fishing, water

transportation, rural and urban water supply (Kateregga and Sterner, 2009). It also serves

as a breeding site for mosquitoes and disease-causing organisms (Opande et al., 2004).

However, water hyacinth provides refuge for smaller fish which may promote

biodiversity (Opande et al., 2004; Kateregga and Sterner, 2009).

To enhance food crop production, there is need to adopt cheaper and environmentally

friendly means of improving soil fertility (Bundy and Andraski, 2005; Otieno et al.,

2009; Argaw, 2012). Other than the use of inorganic fertilizers in crop production,

biological nitrogen fixation (BNF) using rhizobia have been beneficial (Ogutcu et al.,

2008). Rhizobia have the ability to fix nitrogen (N2) through their symbiotic relationship

with leguminous plants (Gyorgy et al., 2010). Biological nitrogen fixation (BNF) is a climate change resilient farming system that boosts adequate management of soil, water

and biodiversity and is also cost effective compared to inorganic fertilizers (Ajouri et al.,

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soil nitrogen via symbiotic nitrogen fixation and potentially lead to increase in yields of

succeeding and associated non-nodulating plants (Brockwell et al., 1995). BNF is also important especially in regions with great farmland pressure and where fallow system is

not possible (Kiros and Singh, 2006).

The human population in the LVB relies on natural rainfall for crop production. They

mainly cultivate maize (Zea mays) and common beans (Phaseolus vulgaris) which are

their staple food (Ondigi et al., 2008). Nitrogen inputs through legume’s symbiotic

relationship with rhizobia improves common bean yield in Africa where it is a key source

of proteins for rural populations (Oliveira et al., 2010).

Symbiotic relationship between common beans and rhizobia is more effective when the

rhizobia strains are well adapted to existing environmental conditions (Alves, 2009).

Rhizobia inoculum should therefore be adapted to existing environmental conditions and

should be more competitive than the indigenous rhizobia strains in the soil (Torres et al.,

2009; Lindstrom et al., 2010). Isolation and characterization of indigenous rhizobia from

Lake Victoria basin can help identify more effective rhizobia strains that can be used as

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1.2 Problem statement and justification of the study

Lake Victoria Basin is experiencing serious land degradation due to conversion of

forests, wetlands and woodlands into farm land to increase and improve agricultural

production, to cater for increasing human and livestock population (LVBC, 2012).

Utilization of land for agricultural purposes in LVB has led to the discharge of

agrochemicals (Chemical fertilizers, nitrogen, phosphorus, pesticides and herbicides) into

Lake Victoria leading to eutrophication (Kimwaga et al., 2011; Kimwaga et al., 2012).

Eutrophication of Lake Victoria is the cause of the invasion and establishment of water

hyacinth in the lake (Williams et al., 2005). Due to the environmental challenges caused

by water hyacinth infestation in LVB there is need to control it (Villamagna and Murphy,

2010).

Proper management of agricultural practices in the LVB catchment can help decrease

agricultural runoff into the Lake (Mwanuzi et al., 2005; Williams et al., 2005; UNEP,

2013). One solution to the reduction of nutrient inflows into Lake Victoria is the use of

rhizobia. Rhizobia are economically and environmentally friendly microorganisms that

have been utilized as biofertilizers (Bhattarai et al., 2011).

Indigenous rhizobia are very diverse at species and strain levels in most soils and are

important sources of inoculants (Lindstrom et al., 2010). Identification and utilization of

effective indigenous rhizobia nodulating Phaseolus vulgaris from Lake Victoria Basin

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into Lake Victoria. According to Ndimele and Jim (2011), composted water hyacinth can

also be used to improve soil quality.

There is limited information on rhizobia nodulating common beans, their morphological

and genetic diversity, effect of water hyacinth compost on rhizobia populations in the soil

and presence or absence of more effective nitrogen fixing rhizobia strains from Lake

Victoria Basin.

In this present study, the morphological and genetic diversity of indigenous rhizobia

isolates nodulating common beans variety rosecoco from Lake Victoria Basin water

hyacinth compost testing farms was determined. The effect of water hyacinth compost

prepared using EM, water hyacinth compost prepared using manure, DAP and

commercial rhizobia inoculum (Rhimbium leguminosarum (strain 446) on indigenous

rhizobia populations was also determined. Representative rhizobia isolates from Korando

B Sublocation in Kisumu, Rwanda and Uganda farms were assessed for their nitrogen

fixing potential in association with common beans variety rosecoco. Isolates with high

symbiotic efficiency can be utilized together with water hyacinth compost to sustainably

improve soil fertility and common bean yield at affordable cost and minimize pollution in

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1.3Research questions

i. Are there morphological and genetic differences between P. vulgaris L variety

rosecoco nodulating rhizobia isolates from Lake Victoria Basin (LVB) water

hyacinth compost testing farms?

ii. Are there superior and effective indigenous nitrogen fixing rhizobia nodulating P.

vulgaris in LVB water hyacinth compost testing farms?

iii. Does water hyacinth compost, DAP, and commercial rhizobia inoculum Rhizobia

leguminosarum (strain 446) application influence rhizobia population in LVB water

hyacinth compost testing farms?

1.4 Hypotheses

i. Indigenous rhizobia that nodulate Phaseolus vulgaris L variety rosecoco from LVB

water hyacinth compost testing farms are not morphologically and genetically

diverse.

ii. Indigenous rhizobia isolates nodulating P. vulgaris variety rosecoco from LVB

water hyacinth compost testing farms have no differences in their symbiotic

efficiency.

iii. Water hyacinth compost, DAP and commercial rhizobia inoculum (Rhizobia

leguminosarum (strain 446) does not affect the population of indigenous rhizobia

that are effective in nodulating Phaseolus vulgaris variety rosecocoin Lake Victoria

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1.5 Objectives

1.5.1 General objective

To determine the genetic and morphological diversity and effectiveness of rhizobia

nodulating common bean variety rosecoco and the effect of selected soil treatments on

their population in water hyacinth compost testing farms in Lake Victoria Basin (LVB).

1.5.2 Specific objectives

i. To determine the morphological and genetic diversity of indigenous rhizobia

isolatesfrom LVB water hyacinth compost testing farms.

ii. To determine the symbiotic efficiency of representative indigenous rhizobiaisolates

from LVB water hyacinth compost testing farms.

iii. To assess the effect of water hyacinth compost, DAP and commercial rhizobia

inoculums (Rhizobia leguminosarum strain 446) application on indigenous rhizobia

populations in LVB water hyacinth compost testing farms.

1.6 Significance of the study

In this study effective indigenous rhizobia strains that can be packaged as common bean

inocula for use by smallholder farmers in Lake Victoria Basin were identified. The

enhancement of rhizobia populations in the soil by water hyacinth compost, demonstrated

that rhizobia and water hyacinth compost can be used together to improve common bean

crop yield. Use of water hyacinth compost together with effective rhizobia innocula will

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the lake. Reducing nitrogen fertilizer usage will also help mitigate greenhouse gas (GHG)

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CHAPTER TWO LITERATURE REVIEW

2.1 Common bean Phaseolus vulgaris L

Common bean (P. vulgaris L) is an annual leguminous plant belonging to the genus,

Phaseolus which originated from Central and South America and was domesticated in

6000 and 5000 BC in Peru and Mexico, respectively (Wortmann, 2006). It is an

important food legume grown in the tropics with the bulk of it being produced in

developing countries practicing low input agriculture (Miklas et al., 2006).

Phaseolus vulgaris has over 40,000 varieties that differ in growth habits seed

characteristics, maturity and adaptation (Schwartz et al., 1996; Jones, 1999). The plant

has pinnately compound and trifoliate leaves. It is self-pollinated; however

cross-pollination can occur via insect cross-pollination when the stigma is extended. The seeds are

non-endospermic and diverge in size and color ranging from the small black wild type to

the large white, brown, red, black or mottled seeds (Wortmann, 2006). Growth habits

vary from determinate bush to indeterminate, extreme climbing types. The determinate

bush varieties are commonly grown in Africa (Buruchara, 2007).

2.2 Bean agronomy

Common bean grow well in warm seasons and is not favored by long exposure to very

low temperatures or frost in all stages of growth (Katungi, et al., 2009). In places where

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temperatures at night constrain pollination. Common bean needs adequate amounts of

rainfall (300 – 600 mm) especially during and just after the flowering stage. Most

common bean varieties mature between 65 to 110 days from emergence to physiological

maturity making it a short season crop (Buruchara, 2007). Some varieties especially the

climbers mature in 200 days after planting and are planted mainly in cool upland

elevations (Katungi et al., 2009). Common bean requires fertile well-drained soils that

do not interfere with germination and emergence (Katungi et al., 2009). In most African

countries the crop does well in soils with pH 5.5, altitudes of 1200–2200 Meters above

sea level (Masl), mean temperature of 15–23 °C and more than 400 mm of rain

(Wortmann, 2006; Katungi et al., 2009).

2.3 Uses of P. vulgaris

Common bean (Phaseolus vulgaris L.) is a major source of protein, complex

carbohydrates, folic acid and dietary fiber (Oliveira et al., 2010; Buruchara et al., 2011).

Common bean also contains essential minerals including, zinc, iron, phosphorus and

calcium (Costa et al., 2006). They are a cheaper alternative source of protein as compared

to animal or fish products and greatly consumed in the world to control malnutrition

(FAO, 2011). Half of the world’s common bean production occurs in developing

countries where it contributes to food security (Amin et al., 2014). According to Katungi

et al. (2010), common beans have high commercial potential and can be utilized to

improve soil fertility in Sub-Saharan Africa. The value of the common bean crop

surpasses that of other legumes, including lentil, chickpea, cowpea and pea, emphasizing

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2014). Beans are cultivated for subsistence use in the great lakes region that has the

highest recorded per capita consumption in Africa (Blair et al., 2010). They are also a

source of steady income for scores of rural households (FAO, 2011).

2.4 Biological nitrogen fixation (BNF)

Biological nitrogen fixation is the process through which atmospheric molecular nitrogen

(N2) is reduced to ammonia (NH3) by a specialized group of prokaryotes mainly bacteria

using the enzyme nitrogenase to catalyze the conversion process (Franche, 2009, Wagner,

2011). Prokaryotes responsible for biological nitrogen fixation (BNF), include free living

soil bacteria in the genera Azotobacter, Bacillus, Beijerinckia, Chromatium, Clostridium,

Klebsiella, Rhodospirillum and free living aquatic cyanobacteria (Garg, 2007; Wagner,

2011). Biological nitrogen fixers of great agricultural importance include those that form

symbiotic relationships with legumes and other plants. These include α-proteobacteria,

the order Rhizobiales, the family Rhizobiaceae, including species of Azorhizobium,

Bradyrhizobium, Mesorhizobium, Rhizobium, and Sinorhizobium. β-proteobacteria have

also been reported to form symbiotic relationships with plants and fix nitrogen (Franche,

2009).

2.4.1 Rhizobia

Soil microorganisms represent the world’s major pool of biological diversity and are vital

for functioning of terrestrial ecosystems with plant diversity enhancing the rates of

microbial processes that mediate carbon (C) and nitrogen (N) cycling (Mukerji et al.,

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Rhizobia also referred to as ‘root nodule bacteria’ were first identified by Hellriegel and

Wilfarth (1888) as the source of fixed nitrogen in root nodules of legumes (Hirsch et al.,

2001; Howieson and Brockwell, 2005). They are medium sized Gram-negative,

rod-shaped bacteria, which do not form endospores. Their lengths varry from 1.2 - 3.0 μm

and their width is about 0.5 - 0.9 μm. Gram stained cells have a uniform gram negative

reaction when young but parts of older cells may not stain. Rhizobia are motile with a

polar or subpolar flagellum or 2-6 peritrichous flagella (Somasegaran and Hoben, 1994).

Rhizobia that infect the host plant but do not fix nitrogen are considered parasitic to the

host plant which supply them with nutrients and protect them within nodule structures

(Denison and Kiers, 2004).

Rhizobia are a taxonomically diverse and phylogenetically heterogeneous group

belonging to the alpha and beta subclasses of Proteobacteria (Gyaneshwar et al., 2011;

Moulin et al., 2013; Bakhoum et al., 2014). Alpha rhizobia form symbiotic relationship

with most legumes while the beta rhizobia have symbiotic interaction with the Mimosa

genus (Gyaneshwar et al., 2011). The woody plant Parasponia sp. is the only nonlegume

nodulated by rhizobia bacteria (Franche et al., 2009). According to Gyaneshwar et al.

(2011), rhizobia are in the class Alphaproteobacteria and family Rhizobiacea in a single

order Rhizobiales. Rhizobial species are in the genera; Allorhizobium previously genus

Methylobacterium, Azorhizobium, Rhizobium, Mesorhizobium, Sinorhizobium,

Bradyrhizobium, Devosia, Burkholderia, Cupriavidus and Shinella (Pereira de Lyra et

al., 2013). P. vulgaris is promiscuous in its symbiotic interactions because it has the

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et al.,2007). Rhizobial species nodulating P. vulgaris include Rhizobium leguminosarum

bv. phaseoli, R. gallicum (bv. phaseoli and bv. gallicum), R. tropici, R. giardinii (bv.

phaseoli and bv. giardinii) and R. etli bv. Phaseoli (Grange et al., 2007; Elbanna et al.,

2009).

2.4.2 Role of rhizobia in biological nitrogen fixation

Available nitrogen is low in most environments and molecular nitrogen from the

atmosphere is the main reserve of nitrogen in the biosphere (Franche et al., 2009). Plants

cannot assimilate molecular nitrogen directly but it is made available through the nitrogen

fixation process that is well developed in some prokaryotic cells (Franche et al., 2009).

Approximately 200 million tons of nitrogen is converted to ammonia by nitrogen fixing

micro-organisms, especially bacteria of the genus Rhizobium as compared to 100 million

metric tons of inorganic nitrogen fertilizers produced annually (Glazer and Nikaidog,

2007).

Nitrogen fixation rates vary with plant species, and the environment and potential rates of

0.41 to 0.82 kg/acre per day have been reported, which are suitable for all of the plant’s

nitrogen needs (Graham et al., 2004). According to Lindstrom et al. (2010) biological

nitrogen fixation by rhizobia is an important pathway for sustainable nitrogen input into

agro-ecosystem. Biologically fixed nitrogen is used directly by the plant, and therefore

not lost through leaching, denitrification and volatilization (Garg, 2007). The use of BNF

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of inorganic fertilizer use on the global nitrogen cycle, ground and surface-water

contamination and global warming (Garg and Geetanjali, 2007).

Common bean yields are low in East Africa, mainly due to low soil fertility with most of

the soils having low available nitrogen and phosphorus (Lunze et al., 2007). Like other

leguminous plants, common bean is able to establish nitrogen fixing symbiotic

relationship with rhizobia, which can improve yields (Torres et al., 2009; Oliveira et al.,

2010). In addition to nitrogen fixation, rhizobia enhance the plants defense against

pathogens and pests and adaptation to environmental stress (Franche et al., 2009). Due to

their agricultural and environmental significance, the diversity of indigenous rhizobia in

their natural populations has been greatly studied in different parts of the world (Agrawal

et al., 2012).

2.4.3 Indigenous rhizobia as source of rhizobia inocula for common beans

Common bean (Phaseolus vulgaris L.) is an important food crop and many studies have

been carried out to identify efficient and competitive strains of rhizobia for nitrogen

biofertilizer production to cope with the nitrogen requirements of this crop (Gonzalez et

al., 2008). Indigenous rhizobia are found naturally in soils of a particular locality and

have a great diversity and enhanced abundance in areas where compatible legumes are

grown and the soils are fertile (Shamseldin et al., 2005; Zengeni et al., 2006; Lindstrom

et al., 2010). A soil may have different species and different strains within a species and

analogous isolates may be found in various localities (Bala et al., 2001, Abaidoo et al.,

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Most soils have high numbers of indigenous rhizobial strains that may be ineffective in

symbiosis but highly competitive due to their adaptation to the environmental conditions

(Shamseldin et al., 2005). Isolation and identification of new rhizobia isolates from a

wide diversity of indigenous rhizobia populations ensures a sustainable source of

effective replacement rhizobia strains with few geographical limitations to the areas of

use and can be developed as commercial inoculums (Musiyiwa et al., 2005; Lindstrom et

al., 2010).

2.5 Factors affecting indigenous rhizobia populations in the soil and Rhizobium

legume symbiosis

Agronomic and ecological impacts of rhizobia inoculated in the soil depends on their

symbiotic properties (nodulation, efficiency, specificity and host range) and their

adaptation to environmental constraints affecting nitrogen fixation like salinity, acidity,

high nitrogen content, phosphorus deficiency, drought and soil temperature

(Workalemahu, 2009; Riah et al., 2014). Regular soil fertility management and legume

cropping enhances rhizobia population and diversity (Lindstrom et al., 2010).

About 4 % of the world soils have salinity problems and most of the saline areas are in

the Tropics and the Mediterranean region (Jadhav et al., 2010). High salinity in the soil

may have a harmful effect on soil microbial populations due to direct toxicity or through

osmotic stress (Ogutcu et al., 2010). Soils with pH higher than 8.0 have high sodium

chloride, borate and bicarbonate associated with high salinity and this reduces nitrogen

fixation (Jadhav et al., 2010). Saline conditions in the soil limit symbiosis by; affecting

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infection process, affecting root nodule function, reducing plant growth, photosynthesis

and demand for nitrogen (Ogutcu et al., 2010). Soil acidity constrains symbiotic nitrogen

fixation in both tropical and temperate soils limiting Rhizobium survival and persistence

in soils and reducing nodulation (Graham et al., 1994).

Symbiotic legume nitrogen fixation requires essential mineral nutrients for normal

establishment and functioning of the symbiosis. Chemical elements such as C, H, O, N,

P, S, K, Ca, Mg, Fe, Mn, Cu, Zn, Mo, B, C1, Ni and Co are essential for the

legume-Rhizobium symbiosis (Weisany et al., 2013). The nutrients have specific physiological

and biochemical roles and there are minimum nutrient concentrations required within

both legumes and rhizobia to sustain metabolic function at rates which do not limit

growth (Weisany et al., 2013). Availability of mineral N in the soil inhibits nodule

formation and nitrogenase activity (Weisany et al., 2013). Studies have shown that,

nitrogen fixation reduces with age of the plant mainly due to nitrogen accumulation in the

soil (Mohammadi et al.,2012). In soils with high phosphorus concentration inhibition of

nodulation by high nitrogen concentration is reduced (Gentili and Huss-Danell, 2003).

Phosphorus is important for molecular and biochemical plant processes, especially in

energy acquisition, storage and utilization (Weisany et al., 2013). Nodules are strong

sinks for phosphorus and nodulation and nitrogen fixation are greatly influenced by P

availability (Weisany et al., 2013). Reports show that phosphorus P in the soil enhances

nodulation, nitrogen fixation and plant growth (Gentili and Huss-Danell, 2003).

Deficiency of phosphorous supply and availability is a major limitation on nitrogen

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Water content in the soil directly influences the growth of microorganisms in the

rhizosphere, including rhizobia. Decreasing water activity below critical tolerance limits

and alters root architecture, plant growth and exudations (Mohammadi et al., 2012).

Drought results in osmotic stress, which causes changes in rhizobia morphology

(Elboutahiri et al., 2010). Drought also affects rhizobia root-hair colonization and

infection persistence and survival in soil, as well as dehydration of cells (Monica et al.,

2013). Rhizobium usually cannot tolerate or function under high levels of osmotic stress

caused by drought (Monica et al., 2013). Drought conditions that influence nitrogen

fixation and poor nodulation of legumes has been reported in arid soils possibly due to

decreases in rhizobia population during the dry season (Mohammadi et al., 2012).

Rhizobia survival in the soil is influenced by high temperatures compared to low

temperatures. High temperatures can be deleterious and affecting both the legume and the

rhizobia and all steps in the development of an efficient nitrogen fixation (Abdullah and

Al-Falih, 2002; Monica et al., 2013). According to Boboye et al. (2011) temperature

affects root hair infection, bacteroid differentiation, nodule structure and the functioning

of the legume root nodule. Environmental temperatures of 32 - 33 °C are harmful to most rhizobia species; however some rhizobia strains have shown adaptation to high

temperatures of up to 42 °C (Boboye et al., 2011; Monica et al., 2013).

2.5.1 Effect of soil amelioration on soil microbial abundance

The root system of higher plants is composed of inanimate organic substances and a vast

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2010). High bacterial population in the rhizosphere has been reported to improve the

efficiency in nutrient cycling (Iqbal et al.,2012). Rhizosphere microflora promote plant

growth by producing plant growth stimulating substances, releasing elements in organic

forms via mineralization of organic complexes and formation of stable soil structure (Das

et al., 2010).

To maximize on agricultural production and high economic returns farmers use various

agricultural inputs. These inputs include mineral fertilizers like sulfates, urea, ammonium

nitrate, and phosphates; organic fertilizers such as farm yard manures, composts and

biosolids; other organic products like humic acids and microbial inoculants (Bunemann et

al., 2006). Agricultural inputs can affect soil organisms through changes in nutrient

availability or toxicity in the first season after the application or longer term if repeated

additions are required to reach a threshold above which effects occur (Bunemann et al.,

2006).

Inorganic fertilizers are important sources of plant nutrients for increased crop production

(Atere and Olayinka, 2012). They can have both negative and positive effects on plant

nodulation, nitrogen fixation and growth of leguminous plants (Huda et al., 2007;

Ruiz-Valdiviezo et al., 2009). There are reports on a decrease in population or activity of soil

organisms after mineral fertilization that has been attributed to the toxicity of metal

contaminants present in phosphate fertilizers (Bunemann et al., 2006). Nitrogen, sulphur

and potassium fertilizers have low levels of contaminants but phosphate fertilizer such as

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considerable amounts of lead, mercury, cadinium and other contaminants from the

phosphate rocks utilized in their manufacture (McLaughlin et al., 2000). Tests on

ammonium fertilisers have also shown decrease in soil pH and cation exchange capacity

(CEC) which results in degradation of hydraulic properties of the soil and reduction in

microbial activity(Bunemann et al., 2006).

Use of organic soil amendments can lead to increase in soil microbial activity, soil

bacterial densities and microbial diversity (Das and Dkhar, 2012; Zaccardelli et al.,

2013). The application of manure has been reported to increase rhizobial population in

the soil (Zengeni et al., 2006). Studies have shown that soil amendment with raw and

composted organic materials results in increased microbial abundance, microbial activity

and microbial diversity in the soil although interval of observed increases in soil

organisms depends on the amount and proportions of readily decomposable carbon

substrates added and the availability of nutrients, mainly nitrogen (Bunemann et al.,

2006; Das and Dkhar, 2012).

Compost increases the carbon/nitrogen ratio which influences bacteria and eukaryotic

community structure (Watts et al., 2010). The biomass of water hyacinth can be

composted and utilized as organic fertilizer for crop production, especially grain legumes

(Atere and Olayinka, 2012). Water hyacinth has a low C: N ratio (17:1) which make it

compost quickly and therefore promote high bacterial growth (Basu et al., 2011).

Application of water hyacinth compost improves physical and chemical properties of the

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biological processes like N fixation and P solubilization in the soil (Cousin et al., 2002;

Das et al., 2010; Basu et al., 2011; Reid et al., 2011).

2.6 Estimation of rhizobia population in the soil

Rhizobia have a saprophytic phase in the soil between symbiotic phases and their

populations depend on the physical and chemical properties of the soil environment as

well as the frequency of planting of legumes in a particular area (Martyniuk and Oron,

2008). A serial soil dilution-plant infection technique is used to estimate, the most

probable number (MPN) of indigenous or introduced rhizobia in the soil for there is no

selective medium available for making plate counts of these bacteria in the soil

(Sadowsky and Graham, 1998, Martyniuk and Oron, 2008). In this technique, seedlings

of the legume of interest are grown in enclosed Leonard jar assembly or enclosed glass

tube containing nutrient free vermiculite, agar or sand to support roots and nitrogen free

plant growth medium as a source of nutrients. The seedlings are inoculated with serial

dilutions of the soil sample and assessed for nodulation after 4-6 weeks of growth.

Nodule formation on the host roots shows the presence of rhizobia in a soil dilution and

the total number of setups with nodulated plants is utilized in the calculation of the MPN

of rhizobia in the soil sample (Bala et al., 2010).

2.7 Morphological characterization of rhizobianodulating P. vulgaris

The diversity of rhizobia has been uncertain mainly due to the enormous leguminous

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(Wolde-Meskel et al., 2004). Rhizobia diversity can be investigated using morphological,

physiological, biochemical assay and genotyping techniques (Alexandre et al., 2006;

Marinkovic et al., 2013). Phenotypic methods utilize cultural and morphological

characteristics including growth rate and colony characteristics on yeast extract-mannitol

mineral salts medium (Zahran et al., 2012). Molecular techniques are more preferred

because they are quicker methods in characterization and can effectively differentiate

genera, species and strains (Giongo et al., 2008).

2.8 Genetic diversity of P. vulgaris nodulating rhizobia isolates from LVB

Advances in molecular methods have enabled the revision of taxonomic classification of

many bacterial groups (Rincon et al., 2008). Several molecular approaches have been

used to study the biodiversity of rhizobia which include; Amplified Ribosomal DNA

Restriction analysis (ARDRA) of both 16S and 23S rDNA, DNA finger printing using

REP and BOX primers (Shamseldin et al., 2008); RAPD (Randomly Amplified

Polymorphic DNA), AFLP (Amplified Fragment Length Polymorphism), SSR (Single

Sequence Repeats) and 16S rRNA gene sequencing (Williams et al., 1990; Young et al.,

1991; Mehmood et al., 2008).

The ribosomal DNA and proteins are highly conserved, showing their essential role in

translation processes (Shamseldin et al., 2005). Genes encoding the rDNA in bacteria are

organized into an operon with three main parts; genes coding for the 16S rDNA, the 23S

rDNA and the region between them, named internally transcribed spacer (ITS) (Woese et

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identification and characterization of new isolates at the genus or species level, a process

called ribotyping (Shamseldin et al., 2005).

According to Laguerre et al. (1994) variations in 16S rRNA genes can be estimated by

polymerase chain reaction (PCR) amplified ribosomal DNA restriction analysis

(ARDRA) of 16S rDNA. Amplified ribosomal DNA restriction analysis (ARDRA) of

ribosomal genes is suitable for phylogenetic studies as it is highly discriminatory,

reproducible and agrees with partial or complete gene sequencing (Germano et al., 2006;

Gyorgy et al., 2010). RFLP analysis of 16S rDNA has been used to study rhizobial

isolates from P. vulgaris, Sesbania sp. and soybean (Herrera-Cervera et al., 1999;

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CHAPTER THREE MATERIALS AND METHODS

3.1 The study area

Field experiments were carried out at Korando B sub-location in Kisumu (Kenya).

Laboratory and greenhouse experiments were carried out at Kenyatta University.

3.1.1 Collection of soil samples

Soil samples were collected from farmers’ fields in the LVB. In Kenya, farm A (S 00°

05.404; E 034° 41.862’), Farm B (S 00° 05.120’; E 034° 41.613’), farm C (S 00°

05.325’; 034° 41.796’) and farm D (S 00° 05.167; E 034° 42.084’), all in Korando B

sub-location in Kisumu County were used. Soils were also collected in four farms in

Kabanyolo (Uganda) (N 00° 27' 00" E 32° 37' 00") and two farms from Nyabarongo

(Rwanda) (S 02o 29'29'' E 030o 46' 95''). The farms were approximately 120 m2 in size.

The soil samples were collected before soil amendment for all the farms, and at the end

of the cropping season after soil amendment for farm A, B, C and D in Kenya.

Before soil amendment, soil samples were collected from 20 points diagonally and across

each farm keeping a radius of at least 6 m from each point. Five hundred grams of the

soil subsamples were taken at a depth of 0-30 cm after clearance of the soil debris from

the soil surface (Abaidoo et al., 2002). A kilogram of homogenous soil sample from each

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avoid cross contamination. The soil was then transported to Kenyatta University lab for

greenhouse experiments.

At the end of the cropping season, after soil amendment soil sampling was carried out

from the four farms in Kenya diagonally and across 20 subplots in each farm representing

the various treatments on the soil (Soil treated with conventional fertilizer (DAP), water

hyacinth compost prepared using EM (effective microorganisms), water hyacinth

compost prepared using cow manure, control soil with no amendment and commercial

rhizobia). Six point samples were collected from each subplot. Before collecting the soil,

organic matter on the soil surface was cleared. Soil was dug to a depth of 30 cm from the

soil surface. The soils were collected with different equipment to avoid cross

contamination. A kilogram of composite soil sample for each treatment from every farm

was packed independently in khaki bags-size 4 and transported to Kenyatta University for

greenhouse experiments. Soil samples that were not used immediately were stored at a

temperature 4 oC.

3.2 Soil analysis

Soil texture was determined by the hydrometer method. Soil pH was determined using a

pH meter in a 1:2.5 soil: water suspension. The soil organic matter was determined

through the Walkley and Black oxidation method. Total soil nitrogen was determined by

the macro-Kjedahl digestion, distillation and titration method. Available P was

determined using the Olsen extraction method. Exchangeable cations were determined

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leachate analyzed using atomic absortion spectroscopy (AAS) while K+ and Na+ were

determined using flame emission spectroscopy (FES) according to procedures by

Okalebo et al. (2002).

3.3 Rhizobia trapping

3.3.1 Greenhouse rhizobia trapping experiments

In the greenhouse soil collected from each site in Kenya, Uganda and Rwanda was mixed

to form a homogenous composite sample for each site. The soil samples were potted in

six different sterilized pots accommodating one kilogram of soil. This was replicated for

each farm. Rose coco bean that is mainly planted in the study region was used as the

trapping host. The bean seeds were surface sterilized using 3 % (v/v) sodium

hypochlorite and pre-germinated on a nutrient free agar media before planting. Two

seedlings were planted per pot after three days pre-germination of the seeds. The pots

were arranged in a randomized complete block design. Watering was carried out at one

day interval because of the high water holding capacity of the soils. Nodulation

assessment of the plants was carried out 35 days after planting. The roots were carefully

washed in running tap water and the nodules detached and wrapped with absorbent tissue

paper to dry at room temperature.

3.3.2 Rhizobia trapping in the farms

Nodules were obtained from P. vulgaris variety Rosecoco plants from the four farms

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season starting from March 2012. Three bean plants were sampled for nodule analysis

from plots which had been treated with the following: water hyacinth compost made

using cattle manure to provide the starter culture, Effective Microorganisms (EM), DAP

(Commercial fertilizer) applied at a rate of 40 kg/ha. In the control experiment, the soil

was not amended. Harvesting of the plants was carried out at the onset of flowering. The

roots were carefully washed, nodules detached and wrapped with absorbent tissue paper

to dry at room temperature. After drying the nodules were stored in a fridge at 4 °C.

3.4 Isolation and identification of rhizobia using morphological characteristics

Nodules representing each host plant from the field soil treatments and whole soil

trapping experiment in the greenhouse were selected from the preserved nodules. They

were placed in sterile distilled water and let to imbibe water for one hour. They were

rinsed with sterile distilled water and dipped for 5 seconds in 95 % (v/v) ethanol to

reduce the surface tension and remove air bubbles from the tissues. The nodules were

then sterilized by dipping them in 3 % (v/v) sodium hypochlorite solution for 4 minutes.

They were then rinsed in five changes of sterile distilled water and crushed with a sterile

glass rod in a drop of sterile distilled water. A loop full of the nodule suspension was

streaked onto Yeast-Mannitol agar (YMA) plates containing 25 µg/ml Congo red and

incubated at room temperature in the dark. Observations on colony emergence were made

after three days. After five days of incubation, well isolated colonies were streaked on

YMA plates containing Congo Red. The isolates were grouped using the procedure

described by Odee et al. (1997). The morphologies of the different colonies were

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consistency, color, texture, size of the first independent colonies and shape of the

margins.

3.5 Gram staining of the rhizobia isolates

Gram staining of the pure 24 hour old rhizobia isolates was carried out using the method

described by Beck et al. (1993).

3.6 Bromothymol blue test

The isolates were evaluated for their ability to alkalize or acidify the media by growing

them on YMA media containing Bromothymol blue (BTB) (25 mg/L). After three days

growth at room temperature the isolates were grouped as either acid producing, neutral or

alkali producing based on the color changes observed on the media. Fast growing

rhizobia are acid producing and turn YMA media with BTB color from green to yellow.

Slow growing rhizobia are alkali producing and turn the media color from green to blue

(Laurette et al., 2015).

3.7Authentication of the rhizobia isolates

Representative isolates from whole soil trapping and on farm trapping experiment were

tested to confirm their nodule forming ability on the host legume under bacteriololgically

controlled conditions. Leornard jar assemblies were prepared according to the procedure

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The assembly was composed of a plastic cup of 8 cm brim diameter tapering to a bottom

diameter of 4 cm with a 1.5 cm 2 rectangular hole at the bottom. The cups were swabbed

with 70 % (v/v) ethanol and fit with a 20 cm long sponge in the hole at the bottom of the

cup. Prior to fitting, the sponge was sterilized in 3 % (v/v) NaoCl for 4 minutes and

rinsed in five changes of sterilized distilled water. The cup assembly was then suspended

in the brim of a larger plastic vessel whose inner surface had been decontaminated by

swabbing using 70 % (v/v) ethanol.

Seven hundred mls of sterilized nitrogen-free nutrient solution (Broughton and Dilworth

1971) (Appendix 1) was added into the lower container of the Leonard jar assembly. The

pH of the nutrient solution had been adjusted to 6.8 with 1.0 M NaOH or 1.0 M HCL

before autoclaving for 15 minutes at 121 °C.

Vermiculite was used as a rooting medium. The vermiculite was soaked in tap water for

two days followed by washing in running tap water to remove dissolved nutrients. The

vermiculite was then rinsed with sterile distilled water and the pH was adjusted to 6.8.

The vermiculite was then sterilized by autoclaving and transferred to the cups of the

Leonard jar assemblies maintaining the sponge wick upright in the middle of the cup. The

top cups were covered with aluminum foil swabbed with 70 % (v/v) ethanol. The

complete Leonard jar assemblies were then put into khaki paper bags for insulation. The

assemblies were steamed in an autoclave for ten minutes to get rid of any possible

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Rose coco bean seeds were selected for uniformity in size shape and colour and then

surface-sterilized in 3.0 % (v/v) sodium hypochlorite after soaking for 6 min, followed by

rinsing in five changes of sterile water. The sterilized seeds were pregerminated in kilner

jars containing damp sterile vermiculite at a temperature of 28 °C in growth room. The

seedlings were then transplanted into the Leonard jar assemblies (Beck et al., 1993;

Somasegaran and Hoben, 1994). Three seedlings were planted into each Leonard jar and

then later thinned to two. Four replicates were used for each treatment. The seedlings

were maintained for eight days in Leonard jar assemblies before inoculation with the

representative rhizobia isolates. The rhizobia isolates were cultured in YMB media for

three days before inoculation. One milliliter (1ml) of each culture was inoculated per

plant. The treatments were laid out in a randomized block design (RBD) in a greenhouse.

Jars of uninoculated seedlings were used as negative controls. Jars inoculated with

commercial rhizobia R. leguminosarum (strain 446) were used as positive control. The

nitrogen free media in the Leonard jars was replenished after every seven days.

The plants were harvested after 45 days of growth in the Leonard jar assemblies. The

plant roots were washed with tap water to remove vermiculite and then the attached wick

was removed taking care not to destroy the roots and nodules. The plants were scored for

the presence or absence of nodules and the number of nodules recorded per plant.

Presence of a single nodule in a Leonard jar for any plant was considered as a

confirmation that the isolate is rhizobia (Zahran et al., 2012). The nodules were then

counted wrapped in tissue paper and stored at room temperature. Shoots were separated

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constant dry weight and their respective biomass determined according to the procedure

described by Bala et al. (2010).

3.8 Determination of symbiotic efficiency of representative rhizobia isolates

Determination of symbiotic efficiency of representative rhizobia isolates was carried out

as described by Somasegaran and Hoben (1994). Common beans variety Rosecoco seeds

were sterilized and pregerminated as described in section 3.7. The seedlings were then

transplanted into the Leonard jar assemblies as described in section 3.7 (Beck et al.,

1993; Somasegaran and Hoben, 1994). Three seedlings were planted into each Leonard

jar and then later thinned to two.

The seedlings were maintained for eight days in Leonard jar assemblies before

inoculation. Rhizobia isolates obtained in section 3.4 and a reference strain R.

leguminosarum (strain 446) were cultured in YMB media for three days before being

used for inoculation. One milliliter (1ml) of each culture was inoculated on to the roots of

each seedling. Plants inoculated with the reference strain R. leguminosarum (strain 446)

and those grown in Leonard jars with nitrogen supplemented media (Nitrogen-free media

with 140 ppm nitrogen as KNO3) were used as positive control. Non-inoculated plants

grown in Leonard jars with nitrogen free media were used as negative control. Each

treatment had four replicates and the jars were arranged in a randomized complete block

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Harvesting and processing of the samples was carried out 45 days after planting according

to method described by Beck et al. (1993). Performance of the plants under different

treatments was assessed based on the plant shoot, root, nodule dry matter and number of

nodules. Symbiotic effectiveness (SE %) was calculated by comparing the total dry

weight of inoculated plants with the total dry weight of N-applied uninoculated control

(Total dry weight of inoculated plant/Total dry weight of N-applied plant) x 100 (Karaca

and Uyanoz, 2012). The best isolates were prepared and packaged using the method

described by Beck et al. (1993) for field trials in LVB.

3.9 Effect of soil treatment on indigenous rhizobia populations in the soil

Rhizobia populations in soils that had been treated with DAP ((NH4)2 HPO4) containing

18-21 % N, 6-54 % P2O5 and 2 % S, Rhizobium leguminosarum inocula (Rhizobium

leguminosarum (strain 446)), water hyacinth compost prepared using EM (WH+EM),

water hyacinth compost prepared using cattle manure (WH+M) and untreated control soil

were determined using the plant infection technique ( Maingi et al., 2001). The Leonard

jar assemblies were prepared as described in section 3.7.

Common bean variety rosecoco seeds were selected, sterilized and pregerminated as

described in section 3.7. The seedlings were then transplanted into the Leonard jar

assemblies (Beck et al., 1993; Somasegaran and Hoben, 1994). Two seedlings were

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The seedlings were grown in the Leonard jar assemblies for 5 days before inoculation

with serially diluted soil inocula. Soil inocula were prepared by suspending 10 g of

thoroughly mixed composite soil sample for the various soil treatments in 90 ml of sterile

distilled water in a 500 ml flask and shaken for 30 min at room temperature (25 ºC). One

ml of each suspension was aseptically pipetted into 9 ml sterile water diluent in

McCartney bottle and shaken for 2 min. The resulting suspension was serially-diluted

tenfold from 10-1 to 10-6 with four replications at each dilution level. Aliquots of one ml

were used to inoculate 5 day old P. vulgaris seedlings.

The plants were harvested after 35 days of growth in the greenhouse. The plant roots

were carefully washed with tap water to remove vermiculite and then the attached wick

was carefully removed taking care not to destroy the roots and nodules. The plants were

scored for the presence or absence of nodules. The number of nodulated plants at each

dilution was recorded and used as an ordered code (from low to high dilution) and used to

estimate the most-probable-number (MPN). Most Probable Number (MPNES) computer

program by Bennet et al. (1990) was used to calculate the populations.

3.10 Genetic diversity of common bean indigenous rhizobia isolates from LVB

3.10.1 Preparation of rhizobia isolates and the reference strains for colony PCR

The rhizobia isolates obtained in section 3.4 and reference rhizobia strains were grown on

yeast mannitol agar (YMA) media for 24 hours. The reference rhizobia strains were R.

Figure

Table 4.1: Soil characteristics of experimental sites compared with critical values for East African soils
Table 4 1 Soil characteristics of experimental sites compared with critical values for East African soils . View in document p.52
Table 4.3: Whole soil trapping rhizobia isolates from nodules of P. vulgaris plants before the application of water hyacinth compost
Table 4 3 Whole soil trapping rhizobia isolates from nodules of P vulgaris plants before the application of water hyacinth compost . View in document p.55
Table 4.4: Rhizobia isolates obtained from the nodules of P. vulgaris variety Rosecoco grown in farms amended with water hyacinth compost in Korando B sulocation in Kisumu, Kenya   Isolate Rhizobia isolate groups
Table 4 4 Rhizobia isolates obtained from the nodules of P vulgaris variety Rosecoco grown in farms amended with water hyacinth compost in Korando B sulocation in Kisumu Kenya Isolate Rhizobia isolate groups . View in document p.58
Table 4.5: Abundance (%) of rhizobia isolates from nodules of  P. vulgaris plants grown in farms treated with water hyacinth compost in Korando B sub-location in Kisumu, Kenya  Isolate morphological group  % Percentage of isolates per farm
Table 4 5 Abundance of rhizobia isolates from nodules of P vulgaris plants grown in farms treated with water hyacinth compost in Korando B sub location in Kisumu Kenya Isolate morphological group Percentage of isolates per farm . View in document p.59
Figure 4.1: Dendrogram showing morphological relationship of whole soil trapping rhizobia isolates from Korando B Sublocation Kisumu, Kenya, Rwanda and Uganda
Figure 4 1 Dendrogram showing morphological relationship of whole soil trapping rhizobia isolates from Korando B Sublocation Kisumu Kenya Rwanda and Uganda. View in document p.61
Figure 4.2: Dendrogram showing morphological relationship of farm trapping rhizobia isolates from four farms in Korando B Sublocation in Kisumu- Kenya
Figure 4 2 Dendrogram showing morphological relationship of farm trapping rhizobia isolates from four farms in Korando B Sublocation in Kisumu Kenya. View in document p.62
Table 4.6: Infectiveness and effectiveness of representative isolates obtained from whole soil trapping experiments in the greenhouse  1234
Table 4 6 Infectiveness and effectiveness of representative isolates obtained from whole soil trapping experiments in the greenhouse 1234. View in document p.64
Table 4.7: Infectiveness and effectiveness of representative rhizobia isolates obtained from farm trapping experiments ISOLATE Group Nod no.1 NDW2(g) SDW3(g) RDW4(g)
Table 4 7 Infectiveness and effectiveness of representative rhizobia isolates obtained from farm trapping experiments ISOLATE Group Nod no 1 NDW2 g SDW3 g RDW4 g . View in document p.65
Table 4.8: Diversity indices of rhizobia isolates from whole soil trapping Diversity indices Farm A Farm B Farm C Rwanda Uganda
Table 4 8 Diversity indices of rhizobia isolates from whole soil trapping Diversity indices Farm A Farm B Farm C Rwanda Uganda . View in document p.69
Figure 4.4: Distance dendogram and principle componet diagram showing morphological diversity of whole soil trapping rhizobia isolates
Figure 4 4 Distance dendogram and principle componet diagram showing morphological diversity of whole soil trapping rhizobia isolates. View in document p.70
Table 4.9: Pearson correlation coefficients of soil characteristics and morphological diversity indices of rhizobia isolates from whole soil trapping experiment from Korando B Sublocation in Kisumu- Kenya  Shannon DominancepH SOM Na N Av.P K
Table 4 9 Pearson correlation coefficients of soil characteristics and morphological diversity indices of rhizobia isolates from whole soil trapping experiment from Korando B Sublocation in Kisumu Kenya Shannon DominancepH SOM Na N Av P K . View in document p.71
Table 4.10: Diversity indices of rhizobia isolates from farm trapping  Diversity indices Farm A  Farm B Farm C Farm D
Table 4 10 Diversity indices of rhizobia isolates from farm trapping Diversity indices Farm A Farm B Farm C Farm D . View in document p.72
Figure 4.5: Diversity profiles of rhizobia isolates from farm trapping.
Figure 4 5 Diversity profiles of rhizobia isolates from farm trapping . View in document p.73
Figure 4.6: Distance dendogram and principle component diagram showing morphological diversity of farm trapping rhizobia isolates
Figure 4 6 Distance dendogram and principle component diagram showing morphological diversity of farm trapping rhizobia isolates. View in document p.74
Table 4.11: Pearson correlation coefficients of soil characteristics and morphological diversity indices of rhizobia isolates from farm trapping experiments from Korando B Sublocation in Kisumu- Kenya  pH SOM  N Av.P K Ca Na H
Table 4 11 Pearson correlation coefficients of soil characteristics and morphological diversity indices of rhizobia isolates from farm trapping experiments from Korando B Sublocation in Kisumu Kenya pH SOM N Av P K Ca Na H . View in document p.75
Figure 4.7: Principle coordinate analyses (PCA) of 97 rhizobia isolates from whole soil trapping
Figure 4 7 Principle coordinate analyses PCA of 97 rhizobia isolates from whole soil trapping. View in document p.79
Table 4.14: Pairwise Population Matrix of Nei Unbiased Genetic Distance of five whole-soil trapping rhizobia populations from Kenya, Uganda and Rwanda Uganda Rwanda   Farm B Farm C   Farm A
Table 4 14 Pairwise Population Matrix of Nei Unbiased Genetic Distance of five whole soil trapping rhizobia populations from Kenya Uganda and Rwanda Uganda Rwanda Farm B Farm C Farm A . View in document p.80
Figure 4.8: A neighbour joining dendrogram based on Nei’s 1978 unbiased genetic distance and Euclididian similarity index showing the genetic distance between whole soil trapping rhizobia populations from Kenya, Uganda and Rwanda
Figure 4 8 A neighbour joining dendrogram based on Nei s 1978 unbiased genetic distance and Euclididian similarity index showing the genetic distance between whole soil trapping rhizobia populations from Kenya Uganda and Rwanda. View in document p.81
Figure 4.9: Evolutionary relationships among 97 whole soil trapping indigenous rhizobia isolates from LVB and 4 reference rhizobia strains (CIAT 899, WSM 1385, USDA 2667, R
Figure 4 9 Evolutionary relationships among 97 whole soil trapping indigenous rhizobia isolates from LVB and 4 reference rhizobia strains CIAT 899 WSM 1385 USDA 2667 R. View in document p.82
Table 4.15: Mean number of different alleles (Na), number of effective alleles (Ne), Shannon's Information Index I (H), expected Heterozygosity (He), Unbiased Expected Heterozygosity (UHe) and percentage of Polymorphic Loci (% P) of farm trapping rhizobia populations from Kisumu , Kenya based on ARDRA data
Table 4 15 Mean number of different alleles Na number of effective alleles Ne Shannon s Information Index I H expected Heterozygosity He Unbiased Expected Heterozygosity UHe and percentage of Polymorphic Loci P of farm trapping rhizobia populations from Kisumu Kenya based on ARDRA data . View in document p.86
Figure 4.10: Principal coordinate analyses (PCA) of farm trapping isolates. Percentage of variation explained by the first two coordinates  (1) 24.66 %, (2)18.39 %
Figure 4 10 Principal coordinate analyses PCA of farm trapping isolates Percentage of variation explained by the first two coordinates 1 24 66 2 18 39 . View in document p.87
Table 4.16: Analysis of Molecular Variance (AMOVA) for 125 farm trapping rhizobia isolates from four populations from Kenya based on restriction digestion of 16S rDNA  Source       Df    SS    MS Est
Table 4 16 Analysis of Molecular Variance AMOVA for 125 farm trapping rhizobia isolates from four populations from Kenya based on restriction digestion of 16S rDNA Source Df SS MS Est. View in document p.87
Table 4.17: Pairwise Population Matrix of Nei Unbiased Genetic Distance of four rhizobia populations from farm trapping from Korando B Sublocation in Kisumu, Kenya
Table 4 17 Pairwise Population Matrix of Nei Unbiased Genetic Distance of four rhizobia populations from farm trapping from Korando B Sublocation in Kisumu Kenya . View in document p.88
Figure 4.12: Phylogenetic relationship of 125 farm trapping rhizobia isolates from LVB and 4 reference rhizobia strains (CIAT 899, WSM 1385, USDA 2667, R
Figure 4 12 Phylogenetic relationship of 125 farm trapping rhizobia isolates from LVB and 4 reference rhizobia strains CIAT 899 WSM 1385 USDA 2667 R. View in document p.90
Table 4.18: Symbiotic efficiency of indigenous rhizobia isolates from Lake Victoria Basin Isolate Nodule number Nodule dry weight Shoot dry weight Root dry weight
Table 4 18 Symbiotic efficiency of indigenous rhizobia isolates from Lake Victoria Basin Isolate Nodule number Nodule dry weight Shoot dry weight Root dry weight . View in document p.93
Table 4.19: Pearson correlation coefficients among investigated parameters in common beans
Table 4 19 Pearson correlation coefficients among investigated parameters in common beans . View in document p.95
Table 4.20: Effect of water hyacinth compost, inorganic fertilizer (DAP) and commercial rhizobia inoculum on common bean nodulating indigenous rhizobia populations in Korando B Sublocation in Kisumu-Kenya Farm Treatment MPN rhizobia Range at ( P 0.095)  Confidence
Table 4 20 Effect of water hyacinth compost inorganic fertilizer DAP and commercial rhizobia inoculum on common bean nodulating indigenous rhizobia populations in Korando B Sublocation in Kisumu Kenya Farm Treatment MPN rhizobia Range at P 0 095 Confidence . View in document p.99
Table 4.21: Comparison of the effect of soil treatment on rhizobia populations using Kruskal-Wallis H test Treatment n Mean rhizobia population rank
Table 4 21 Comparison of the effect of soil treatment on rhizobia populations using Kruskal Wallis H test Treatment n Mean rhizobia population rank . View in document p.102
Figure 4.13 I: The relationship between soil characteristics and rhizobia populations in soils obtained from Korando B Sublocation in Kisumu, Kenya
Figure 4 13 I The relationship between soil characteristics and rhizobia populations in soils obtained from Korando B Sublocation in Kisumu Kenya. View in document p.103
Figure 4.13 II: The relationship between soil characteristics and rhizobia populations in soils obtained from Korando B Sublocation in Kisumu, Kenya
Figure 4 13 II The relationship between soil characteristics and rhizobia populations in soils obtained from Korando B Sublocation in Kisumu Kenya. View in document p.104
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