MAKERERE UNIVERSITY

INCIDENCE AND MOLECULAR VARIABILITY OF CASSAVA BROWN STREAK DISEASE IN RWANDA

Jeanine UMFUYISONI BSc in Biology, Option: Biotechnology (University of Rwanda)

A THESIS SUBMITTED TO THE DIRECTORATE OF RESEARCH AND GRADUATE TRAINING IN PARTIAL FULFILMENT OF THE REQUIREMENTS FOR THE AWARD OF A MASTER OF SCIENCE DEGREE IN CROP SCIENCE (CROP PROTECTION) OF MAKERERE UNIVERSITY

May 2018 DECLARATION

I hereby declare that the work in this report is my own and has never been submitted to any other university

Signed: Date 24 October 2018. Jeanine Umfuyisoni MSc. Crop Science

This report has been submitted with the approval of the university supervisor

Signed Date 3rd October 2018.

Assoc. Prof. Jeninah Karungi Tumutegyereize Department of Agricultural Production College of Agricultural and Environmental Sciences School of Agricultural Production Makerere University

This report has been submitted with my approval as a supervisor

Signed Date 3rd October 2018 Dr. Peter Sseruwagi Mikocheni agriculture Research institute P.O Box 6226 Dar-es-Salaam,Tanzania

i

DEDICATION

To my wonderful husband Richard Manishimwe and my lovely son Rickson Béni Ishimwe, for their constant support, encouragement, love, patience and prayers during my study period. The pursuit for my goals would not be possible without you.

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ACKNOWLEDGEMENT

I would never have been able to finish my dissertation without the guidance of Almighty God. Thank you for keeping me strong and healthy throughout my academic endeavour. Thank you for giving me other people to whom I owe my deepest gratitude for their support.

I am grateful to the Rwanda Agriculture Board specifically Mrs Marie Claire Kanyange, the Cassava Disease Diagnostic Project country team leader and the whole Cassava Program team for selecting me as one of project beneficiaries (student).

My gratitude goes to Bill and Melinda Gates Foundation for funding the project “Cassava Disease Diagnostics” through Mikocheni Agricultural Research Institute (MARI) from which I got the full fund for my studies.

I would like to sincerely thank my supervisors Associate Professor Jeninah T. Karungi from Makerere University and Dr. Peter Sseruwagi from MARI for their guidance, patience and understanding during my studies. Despite their many other academic and professional commitments, they were able to provide their time for scientific advices, knowledge, many insightful discussions and suggestions.

I would like to thank Dr. Joseph Ndunguru, Dr. Fred Tairo, Dr. Laura Boykin and Dr. Mildred O. Ssemakula for useful guidance, constructive criticism, supervision and providing me an excellent atmosphere for doing my course work and research. Thank you for sourcing my molecular laboratory work at NaccRI and your support in Bioinformatic linkage analysis to Western Australia University.

Special thanks to my husband my son and my sister (Claire Uwineza) for your patience, practical and emotional support. I thank you for sticking by my side when I was in need. May almighty God bless you abundantly. Lastly I would like to express my sincerely appreciations to my friends and colleagues Andrew Mtonga, Veneranda Ngazi and Brenda Muga for inspiring my effort despite enormous work pressure we faced together.

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TABLE OF CONTENTS

DECLARATION ...... i ACKNOWLEDGEMENT ...... iii TABLE OF CONTENTS ...... iv LIST OF FIGURES ...... vii LIST OF TABLE ...... viii ABSTRACT ...... ix CHAPTER ONE ...... 1 1.0 INTRODUCTION ...... 1 1.1 Background ...... 1 1.2 Problem statement ...... 2 1.3 Justification ...... 3 1.4 Objectives ...... 4 1.4.1 Main objective ...... 4

1.4.2 Specific objectives ...... 4

1.4.3 Hypotheses ...... 4

CHAPTER TWO ...... 5 2.0 LITERATURE REVIEW ...... 5 2.1 Ecology and Agronomy of cassava ...... 5 2.2 Viruses and virus diseases of cassava in Africa ...... 5 2.3 CBSVs of cassava ...... 6 2.3.1 History of CBSVs in Africa ...... 6 2.3.2 Genetic diversity of CBSV and UCBSV ...... 7

2.3.3 Symptomology of cassava brown streak disease ...... 7

2.3.4 Cassava brown streak viruses transmission ...... 9

2.3.5 Host range of Cassava brown streak viruses ...... 10

2.3.6 Diagnosis and Detection ...... 10

2.3.7 Economical damage of Cassava brown streak viruses ...... 11

2.3.8 Management of Cassava brown streak disease ...... 11

CHAPTER THREE ...... 14

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3.0 INCIDENCE AND SEVERITY OF CASSAVA BROWN STREAK DISEASE IN MAJOR CASSAVA GROWING DISTRICTS OF RWANDA ...... 14 3.1 Introduction ...... 14 3.2 Study areas ...... 14 3.2.1 Sampling procedure ...... 15

3.2.2 Data Analysis ...... 16

3.3 Results ...... 17

3.3.2 Cassava brown streak disease Severity ...... 18

3.3.3 Whitefly abundance ...... 18

3.4 Discussion ...... 19 CHAPTER FOUR ...... 22 4.0 MOLECULAR VARIABILITY OF ISOLATES OF CBSVs INFECTING CASSAVA IN RWANDA...... 22 4.1 Introduction ...... 22 4.2 Detection of cassava brown streak viruses in collected samples ...... 22 4.2.1 Sample collection ...... 22

4.2.2 Nucleic acid extraction ...... 22

4.2.4 PCR Amplification ...... 23

4.2.5 Sequencing and phylogenetic analysis ...... 24

4.3 Results ...... 25 4.3.1 Detection of CBSV ...... 25 4.3.2 Geographical distribution of CBSVs ...... 26

4.3.3 Cassava brown streak virus molecular diversity...... 27

4.4 Discussion ...... 29 5.0 ALTERNATIVE HOSTS FOR CASSAVA BROWN STREAK VIRUSES (CBSVs) IN RWANDA...... 31 5.1 Introduction ...... 31 5.3 Detection of CBSV and UCBSV ...... 33 5.3.1 Nucleic Acid extraction, Complementary DNA synthesis and Polymerase chain reaction ...... 33

5.3.2 Sequencing and phylogenetic analyses ...... 33

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5.4 Results ...... 33 5.4.1 Plant characteristics and virus disease symptoms ...... 33 5.4.2 Ecology and distribution of non-cassava plant species ...... 35

5.4.3 PCR Results ...... 38

5.4.4 Sequencing ...... 38

5.5 Discussion ...... 42 6.0 GENERAL DISCUSSION, CONCLUSION AND RECOMMENDATIONS...... 45 6.1 General discussion ...... 45 6.2 Conclusions ...... 48 6.3 Recommendations ...... 49 REFERENCES ...... 50 APPENDICES ...... 59

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LIST OF FIGURES

Figure 1. Cassava leaves showing characteristic leaf symptoms (A&B) vein and inter-vein chlorosis and (C) chlorotic spots of cassava brown disease ...... 8

Figure2. Cassava stems showing characteristic stem symptoms (A) stem necrosis with streaks and (B) stem die-back of cassava brown disease ...... 8

Figure 3. Cassava roots showing characteristic root symptoms (A) radial constrictions and (B) yellow brown corky necrosis of cassava brown disease ...... 9

Figure 4. Map of Rwanda showing the surveyed areas……………………………………...15

Figure 5. CBSD incidence in the ten cassava growing districts of Rwanda, 2015 ...... 17

Figure 6. CBSD severity in the ten districts of Rwanda, 2015 ...... 18

Figure 7. 1.2% agarose gel of PCR amplified products at 344 bp and 440bp of fragments of CBSV and UCBSV coat protein genes respectively, using the specific primer pair: CBSDDF2 and CBSDDR and a 1 Kb ladder...... 25

Figure 8. Maps of Rwanda showing the distribution of CBSD in A, 2012 and B,2015 ...... 27

Figure 9. Virus-like symptoms on CBSV positively tested shrubs: A, Asystacia gangetica (Acanthaceae) B, Centella asiatica (Apiaceae) C, Physalis peruviana L. (Solanaceae ) D, Acanthus pubescens (Acanthaceae) E, Carica papaya L.( Caricaceae)F, Sida cordifolia L. (Malvaceae) G, Senna spectabilis (DC.) (Caesalpinioideae) ...... 35

Figure10: A) Gel picture of UCBSV amplifications from shrub leaf samples, B) Gel picture of CBSVs amplifications from cassava ...... 38

Figure 11: Phylogenetic tree (cladogram) inferred from the MrBayes method using Geneious 9.1.5 for nucleotide identities of partial coat protein of CBSVs. UCBSV and CBSV isolates determined in this study are all highlighted in yellow with those from non cassava plant sp. circled in red…………………………………………………… 41

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LIST OF TABLE

Table 1: Occurrence and distribution of cassava brown streak viruses in surveyed districts of Rwanda, 2015 ...... 26

Table 2: Distribution of CBSVs in plants with different symptom severity scores ...... 27

Table 3. Isolates detected in cassava ...... 28

Table 4: Plant species and ecology of PCR amplified samples for CBSVs in non-cassava plants ...... 36

Table 5: Isolates detected in cassava and alternative hosts………………………………... ..39

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ABSTRACT

In Rwanda, despite the importance of cassava as staple food, its production is greatly limited by biotic and abiotic constrains leading to poor yields. One of the most important constraints to cassava production is the cassava brown streak disease (CBSD). Cassava Brown Streak virus Ugandan valiant (UCBSV) was the first reported to occur in Rwanda in 2009. However, the Rwandan isolates of CBSVs have not been fully characterised which is a big gap in knowledge on epidemiology and management of CBSD in Rwanda. In addition no studies have been conducted in the country to understand the potential role of non-cassava plant species acting as virus reservoirs or alternative hosts for the viruses causing CBSD. This study therefore aimed at determining the incidence and severity of CBSD in the ten major cassava producing districts located in different agriculture zones of Rwanda. It also aimed at determining the diversity of cassava brown streak viruses; and to identify the possible alternative hosts for CBSD causal agents in Rwanda. In a survey conducted in 2015, 279 cassava leaf samples were collected from 93 cassava young field (3 to 6 MAP) and 101 leaf samples from shrubs and herbs with virus-like symptoms growing in or around cassava fields, as well as from areas with no nearby cassava fields. Sampling was undertaken in the ten districts of Rwanda: Bugesera, Nyagatare, Kayonza, and Kirehe (Eastern agriculture zone),

Rusizi, Nyamasheke, (Western agriculture zone), Gisagara, Nyanza, Ruhango and Kamonyi

(Southern agriculture zone) that grow large acreages of cassava.

The CBSD incidence was calculated as a percentage number of plants diseased relative to the total number of plants assessed, while a scale of 1 to 5 was used to determine the disease severity where 1 = no visible CBSD symptoms and 5 = severe foliar symptoms and/or defoliation and plant die. Total RNA was extracted using CTAB (cetyltrimethyl ammonium bromide) method and amplified using reverse transcriptase polymerase chain reaction (RT-

PCR). Positively amplified samples were partially sequenced using the Sanger method.

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Findings from this study showed that there were a significant differences (P<0.05) in the incidence of CBSD between surveyed districts. The disease was widely distributed in the surveyed areas with an average incidence of 27.39%, which is a marked increase from the

18.8% in 2012. There were no significant differences (p>0.05) in CBSD severity between the surveyed districts mean severity score was 2.4 indicating that all the districts had moderate symptoms. Based on the RT-PCR and sequencing results, this study revealed further spread of UCBSV, reports for the first time a new appearance CBSV and UCBSV- CBSV co- infections.

For alternative host, no CBSV was detected in the study materials. However, five species:

Asystacia gangetica (Acanthaceae), Physalis peruviana L. (Solanaceae), Carica papaya L.

(Caricaceae), Sida cordifolia L. (Malvaceae) and Senna spectabilis (DC.) (Caesalpinioideae) tested positive for UCBSV. This is the first report of non-cassava plant species as hosts for

UCBSV in Rwanda. The findings suggest that providing clean (virus-free) planting material alone might not offer an effective solution to management of CBSD in areas where such alternative host plants are rampant. It is therefore also advisable to create awareness on the importance of alternative host plants in the management of CBSD, with emphasis to reduce or remove the plants in cassava fields and the surrounding environments.

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CHAPTER ONE

1.0 INTRODUCTION

1.1 Background

Cassava (Manihot esculenta Crantz, family Euphorbaiceae), a native to South America

(Abraham, 1956; Karakacha, 2001), is believed to have been introduced into Sub-Saharan

Africa (SSA) by Portuguese navigators in the 16th the century from Brazil to the west coast of Africa (Jones, 1959) and later to East Africa through Madagascar and Zanzibar (Hillocks,

2002). Currently, it is an important staple food crop in many countries of the subcontinent

(FAOSTAT, 2012) and provides an affordable source of carbohydrate for over 800 million people around the world (Nassar et al., 2010).It is reported to occupy the second place worldwide after maize (Zea mays L.) for production of starch (Howeler et al., 2013).

However, cassava production is affected by both abiotic (physical factors) and biotic (pests and diseases) constraints. Among biotic constraints, cassava mosaic geminiviruses (CMGs) and Cassava brown streak virus (CBSV), which cause cassava mosaic disease (CMD) and cassava brown streak disease (CBSD), respectively, are presently of social and economic importance in East Africa (Hillocks et al., 2003; Monger et al., 2001).

In Rwanda, cassava roots are a staple food for almost eleven million inhabitants and cassava leaves are a popular vegetable. It plays a major role in food security and is the third most important food crop after banana and maize (MINAGRI, 2014). Despite the great importance of the crop, many constraints hamper the production of cassava and threaten food security of the producing countries. In general, according to Legg et al. (2006), low yields of cassava in

Africa are attributed to both abiotic and biotic constraints. The major abiotic constraints are poor soils and unpredictable weather. The soils in most of the cassava producing countries are generally poor, sandy, and frequently mineral deficient (Hillocks, 2002); erratic rainfall

1 and prolonged dry spells with no reliable irrigation systems in place, compound the productivity problems. Biotic constraints primarily comprise a range of pests and diseases

(Legg et al., 2006). Arthropod pests including cassava green mite (Mononychellus tanajoa,

Bonder), cassava mealybug (Phenaccocus manihoti, Matile-Ferrero) and whitefly (Bemisia tabaci, Gennadius) seriously damage the crop and affect the final yield. Of the diseases,

Cassava mosaic disease (CMD), caused by cassava mosaic begomovirus (CMBs) (Family

Geminiviridae; genus Begomovirus) (Hong et al., 1993), and cassava brown streak disease

(CBSD) caused by Cassava brown streak virus (CBSV) (family Potyviridae; genus

Ipomovirus,) (Fauquet et al., 2005; Monger et al., 2001) , are currently the major threats to the crop’s health and productivity.

Cassava brown streak disease has been known in cassava production systems in Tanzania since early 1930s (Storey, 1936) , but became more damaging in areas away from the coast in the early 1990s (Hillocks et al., 2000). It is a devastating disease that causes loss of root production and quality and can render susceptible varieties unusable if cassava roots are left in the ground for over nine months (Ntawuruhunga et al., 2007). An important challenge in cassava brown streak disease management is that in Uganda, inland areas of Tanzania and

Western Kenya, the incidence is highest and severity greatest in CMD-resistant varieties that are being promoted for the management of the CMD pandemic (Mukasa, 2012).

1.2 Problem statement

Cassava brown streak disease is the second major constraint to cassava production in Rwanda after cassava mosaic disease. A report of a 2012 country survey (unpublished data) revealed a mean of 24.8% for CMD incidence and a severity mean of 2.9 while for CBSD; means were

18.8% and 2.1 for incidence and severity, respectively. Cassava Brown Streak virus Ugandan valiant (UCBSV) was the first reported to occur in Rwanda in 2009 (ISAR, 2009). However,

2 the Rwandan isolates of CBSVs have not been fully characterised which is a big gap in the knowledge of the epidemiology and management of CBSD in Rwanda. In addition, no studies have been conducted in the country to understand the potential role of non-cassava plant species acting as virus reservoir or alternative hosts for the viruses causing CBSD.

Available information on the host range of CMBs indicates that they are not restricted to cassava or to its wild relatives such as Manihot glaziovii Müll. Arg. only, since recently

African cassava mosaic virus (ACMV) was detected in Jatropha multifida L.

(Euphorbiaceae) (Ramkat et al., 2011) and Laportea (=Fluerya) aestuans (Urticaceae) in

Nigeria (Alabi et al., 2007). For CBSD, the existence of wild alternative host plants in the region was considered most likely. Documented evidence indicated the natural occurrence of cassava brown streak disease in tree cassava, Manihot glaziovii (Storey, 1936) and recently

(Mbanzibwa et al., 2011) have confirmed it.

1.3 Justification

In Rwanda, cassava is considered among the most important staple food in many regions

(MINAGRI, 2014). Unfortunately, in the areas of production, cassava brown streak disease is becoming a major threat to food security and is of great concern to farmers, researchers and cassava development agencies. Recent research efforts have only focused on CBSV detection and determination of incidence, severity and whitefly counts through conducting monitoring surveys. Possible wild plants harbouring CBSVs have not been identified and Rwandan

CBSV strains and isolates are not elucidated. Availability of sequences of isolates from wild species and cassava could give better explanation on the evolution of CBSVs and thus shed some light on the infection movement between cassava and other plant species. This is an important aspect especially in predicting the potential of CBSV isolates to evolve into new strains or even distinct viruses. Therefore, this study was conducted to confirm the identity

3 and genetic diversity of cassava brown streak viruses, and will generate information on their alternative hosts. In addition, sequencing of CBSV genomes will be a key to developing molecular diagnostic tools for early detection of CBSD associated virus isolates. Knowledge of alternative hosts will guide in weed management system that aims at preventing introduction and perpetuation of CBSVs across cassava fields. This study will give a clear understanding of the epidemiology of CBSD in Rwanda, which is an important key to designing proper management strategies of the cassava viral diseases.

1.4 Objectives

1.4.1 Main objective

This study aims at providing information for the management of Cassava Brown Streak

Disease by characterising CBSD causal agents from both cassava and non cassava plant species in Rwanda.

1.4.2 Specific objectives

1. To determine the incidence and severity of cassava brown steak disease in major cassava

growing districts of the different agrroecological zones of Rwanda.

2. To determine the genetic diversity of CBSVs associated strains infecting cassava in

Rwanda.

3. To identify the alternative hosts for CBSVs.

1.4.3 Hypotheses

1. There is no difference in distribution of CBSD associated viruses from one agro-

ecological zone to another.

2. Cassava is not the only host for CBSVs in Rwanda.

3. There is no relationship between CBSV sequences from wild species with corresponding

CBSV sequences in cassava.

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CHAPTER TWO

2.0 LITERATURE REVIEW

2.1 Ecology and Agronomy of cassava

Cassava is a shrub reaching 1-4 m height and is commonly known as tapioca, manioc, mandioca and yucca in different parts of the word. It belongs to the dicotyledonous family

Euphorbiaceae, and the Manihot genus (Hillocks, 2002). The Manihot genus is reported to have about 100 species, among which the only commercially cultivated one is Manihot esculenta Crantz. Cassava can be propagated from either stem cuttings or sexual seed, but the former is the commonest practice. Although cassava is a perennial crop, the storage roots can be harvested from 6 to 24 months after planting, depending on cultivar and the growing conditions (El-Sharkawy, 1993). In the humid lowland tropics the roots can be harvested after

6-7 months. In regions with prolonged periods of drought or cold, the farmers usually harvest after 18-24 months (Cock et al., 1979). In addition, the roots can be left in the ground for a long period of time, making it a very useful crop as a security against famine.

Cassava is found over a wide edaphic and climatic area between 30oN and 30oS latitude, growing in regions from sea level to 2300m a.s.l, mostly in areas considered marginal for other crops: lower fertility soils, annual rainfall from <600 mm in the sub-humid and humid tropics. Given the wide ecological diversity, cassava is subjected to highly varying temperatures, photoperiods, solar radiation and rainfall (Howeler et al., 2013)

2.2 Viruses and virus diseases of cassava in Africa

Cassava propagates vegetatively, which facilitates the dissemination of viral diseases by carrying viruses from one crop cycle to the next through the cuttings used as planting material. Without intervention, infection can therefore readily build up from one crop cycle to

5 the next, particularly where there is also a significant level of vector transmission (Legg et al., 2003). About twenty viruses have been identified in cassava fields in Africa and elsewhere in the world (Mbanzibwa, 2011). Of all the cassava viruses known in Africa, including Cassava mosaic geminiviruses (Geminiviridae: Begomovirus), Cassava brown streak virus (Potyviridae: Ipomovirus)l, Cassava Ivorian Bacilliform virus (unassigned),

Cassava Kumi viruses A and B, Cassava “Q” virus, Cassava common mosaic virus

(Potexvirus)(Calvert et al., 2002); only Cassava common mosaic virus (CsCMV) is known to occur in other continents, which suggests that cassava viruses are indigenous to Africa and moved to cassava from other plants of the region (Calvert et al., 2002). Moreover, only cassava mosaic begomoviruses (CMBs) and Cassava brown streak virus (CBSV), which cause cassava mosaic disease (CMD) and cassava brown streak disease (CBSD), respectively, are presently of social and economic importance in East Africa (Hillocks et al.,

2003; Monger et al., 2001) .

2.3 CBSVs of cassava

2.3.1 History of CBSVs in Africa

Cassava brown streak disease (CBSD) is caused by two viruses including: Cassava brown streak virus (CBSV) and Ugandan cassava brown streak virus (UCBSV) [picorna-like (+) ssRNA viruses; genus Ipomovirus; family Potyviridae] (Alicai et al., 2007; Mbanzibwa et al.,

2009)). It was first described by Storey (1936) in cassava from the foothills of the Usumbara mountains of Tanzania. (Nichols, 1950) reported the presence of the disease in all East

African coastal cassava growing areas; ranging from the North Eastern border of Kenya to

Mozambique and it was widespread at lower altitudes of Malawi. (Nichols, 1950) also recorded incidence of the disease in inland East Africa up to altitudes of 1000 m above sea level. Recently, CBSD has been reported at mid-altitude levels (1200-1500 m asl) in the

Democratic Republic Congo (Mahungu et al., 2003), Uganda, Mozambique, Zambia and

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Malawi (Alicai et al., 2007; Campo et al., 2011) and the Lake zone areas of Tanzania

(Jeremiah et al., 2008; Legg et al., 2011) , areas which were not reported to be at risk earlier.

2.3.2 Genetic diversity of CBSV and UCBSV

Genetic variability of CBSV and CBSUV for isolates infecting cassava has been studied widely (Mbanzibwa et al., 2009; Monger et al., 2010; Winter et al., 2010). There are 12 complete genomes in the GenBank and over 70 complete coat protein sequences and several partial sequences of different genes of CBSV and UCBSV. Recently, in Tanzania, Ndunguru et al (2015) give rise to the possibility that there may be as many as four distinct virus species associated to CBSD. Also, Sequence comparisons revealed stronger similarity to an isolate from nearby Tanzania (93.4% pairwise nucleotide identity) than to those previously reported from Malawi (Mbewe et at, 2017).

However, there are no sequences of the isolates from wild plants. A few partial sequences of

CBSV isolates from M. glaziovii sub-clustered in the main clade of CBSV suggesting a continuous evolution and therefore genetic distinctness (Mbanzibwa, 2011).

2.3.3 Symptomology of cassava brown streak disease

Symptoms of CBSD can be confused with those of CMD (Hillocks, 1997). However, in cassava brown streak virus affects the entire cassava plant; leaves, stems and the roots. CBSD symptoms include various patterns of chlorosis in the absence of distorted leaves, quite unlike the symptoms of CMD. There are different types of CBSD leaf symptoms described; symptoms may appear along the edges of secondary veins, later affecting tertiary veins and possibly developing into chlorotic blotches or may show also circular chlorosis between the main veins (Fig. 1). In the advanced stages, a large area of the lamina may be affected and the diseased leaves remain attached for weeks. Foliar symptoms vary according to specific variety, age of the plant, growing conditions (temperature, rainfall, and altitude) and the virus isolate that

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cause symptoms. Tolerant varieties show foliar symptoms with no or delayed root symptoms. At

high temperature new leaves do not show symptoms. In older plants, CBSD symptoms are hard

to recognize as leaf senesce is similar to the symptoms (Mohammed et al., 2012).

A C B

Figure1. Cassava leaves showing characteristic leaf symptoms (A&B) vein and inter-vein chlorosis and (C) chlorotic spots of cassava brown disease

Stem lesions have been observed on CBSD-affected plants (Fig.2). According to (Hillocks et

al., 2000) purple or brown lesions appear on the exterior surface of young green stems. This

is followed by formation of sepia necrotic lesions in the leaf scars (Nichols, 1950). The other

notable stem symptom is the death of node and internodes, which results into the so-called A dieback (Hillocks et al., 2000).

A B

Figure2. Cassava stems showing characteristic stem symptoms (A) stem necrosis with streaks and (B) stem die-back of cassava brown disease.

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Root symptoms may appear as radial constrictions and/or pits and fissures on the bark surface. Under these pits necrotic tissues can be seen in the cortex. Internal symptoms consist of yellow brown corky necrosis of the starch tissue (Hillocks, 1997). In some cases roots appear healthy, but reveal necrosis when opened (Hillocks, 1997) which is sometimes accompanied by root constriction (Alicai et al., 2007; Kanju et al., 2002) (Fig.3). Plants without foliar symptoms for CBSD may be found associated with root necrosis, which is otherwise common for plants with foliar symptoms (Hillocks et al., 1996). According to (Bock, 1994), leaf symptoms are not necessarily associated with root symptoms, and in general all symptoms may not appear on the affected plants.

A B

Figure3. Cassava roots showing characteristic root symptoms (A) radial constrictions and (B) yellow brown corky necrosis of cassava brown disease

2.3.4 Cassava brown streak viruses transmission

The first observation by (Storey, 1936) Storey (1936) revealed that the disease was spreading in vegetative propagation and later that the invisible causal agent was both graft and sap transmissible to cassava plants (Bock, 1994; Rwegasira, 2009). Several attempts to transmit

CBSV through the suspected vectors including: aphids, Myzus persicae Sulz. (Lennon et al.,

1985); and whiteflies, B. afer (Bock, 1994) were not successful. Despite the failure,

9 speculation on whitefly involvement persisted and (Maruthi et al., 2005) reported for the first time that B. tabaci is a vector of CBSV, although at low frequencies. Contaminated farm implements such as knives used for cutting cassava stems may also spread the virus when used on infected plants and then on the healthy ones (Rwegasira, 2009).

According to (Legg et al., 2011), there are fewer data describing the distribution and spread of the CBSD pandemic than there are for CMD. However, in the Great Lakes region, published records for new occurrences of CBSD, do illustrate a strong association with the occurrence of high whitefly populations, and a general trend of spread similar to that for the

CMD pandemic (Alicai et al., 2007).

2.3.5 Host range of Cassava brown streak viruses

According to Storey (1939), the natural occurrence of cassava brown streak disease was believed to be in the cassava tree (Family: Euphorbiaceae, Genus Manihoti, specie Manihot esculenta Crantz). However, M. glaziovii was found infected with CBSV isolates in Tanzania

(Mbanzibwa et al., 2011) and in Kenya (Bock, 1994). In studies conducted in screen house by (Bua et al., 2009) , using different species of Nicotiana, they demonstrated that all species showed symptoms (general chlorosis, vein chlorosis and necrotic lesions), where N. benthamiana and N. tabacum recorded the highest level of CBSD and N. glutamia the lowest level. All of these speculations show that the natural hosts of the viruses causing CBSD are not known hence the need for research to generate information upon which formulation of effective CBSD management strategies would be based.

2.3.6 Diagnosis and Detection

Manifestation of CBSD symptoms on sensitive cultivars has often been used in diagnosing for CBSV infections. The disease symptoms are expressed on foliar, stem and storage roots.

However, Symptomatic leaves are usually sampled for the reverse transcriptase-polymarase

10 chain reaction (RT-PCR) based CBSV detection due to the fact that reliance on symptoms for diagnosis of CBSD is not reliable because some cultivars may phenotypically remain symptomless despite being infected (Abarshi et al., 2010). Thus, several RT-PCR diagnostic protocols for CBSV have been developed for diagnosis of the disease (Abarshi et al., 2010;

Mbanzibwa, 2011; Monger et al., 2001; Rwegasira, 2009) .

2.3.7 Economical damage of Cassava brown streak viruses

Cassava brown streak disease has a direct effect on the quality of tuberous roots and the disease can be more of a threat to food security than CMD (Alicai et al., 2007). It can render susceptible varieties unusable if cassava roots are left in the ground for over nine months

(Ntawuruhunga et al., 2007). The overall effect of CBSD is reduction of root yield by up to

74% (Muhana et al., 2004) and quality (Hillocks et al., 2001). When combined with cassava mosaic disease, 100% yield loss can result. This is generally caused by the fact that the symptoms are elusive, i.e. cannot be recognized easily by untrained or inexperienced people, and may sometimes not appear until the cassava plant has stayed longer than 9 months

(Hillocks, 1997). Storage roots showing CBSD symptoms are not fit for human and animal consumption.

2.3.8 Management of Cassava brown streak disease

Storey (1936) and Nichols (1950) recommend selection and use of disease free cuttings for planting material. However (Rwegasira et al., 2012), CBSD diagnosis is not always straightforward and apparently symptomless plants could be latently infected (Rwegasira et al.,

2012). Breeding for resistance (Jennings, 1957; Storey, 1936), phytosanitation (Nichols,

1947) and quarantine (Nichols, 1950) were the major undertakings in attempts to manage

CBSD. For the management of CMD, a broad range of approaches have been developed (Thresh et al., 2005; Vanderschuren et al., 2007). Such strategies include heat treatment to eliminate

ACMV and EACMV from growing shoots(Walter et al., 1982) , use of meristem-tip culture (Ng,

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1992) , roguing infected plants during early stages of plant growth (Thresh et al., 1994), and the use of cultivars with appreciable resistance to both the virus (Fargette et al., 1996; Jennings,

1960; Ogbe et al., 2003; Otim-Nape et al., 1996) and the vector (Otim-Nape et al., 1996).

Chemical control of the whitefly vector has seldom been practiced by farmers in Africa for economic reasons. In addition, pesticides are least effective in controlling arthropod-borne viruses if the main spread is from external sources and not within crops (Thresh et al., 2005). The negative impact of pesticides on the environment and risks to beneficial organisms including natural enemies and farmers’ health makes pesticides useless appealing (Thottappilly et al.,

2003).

Currently, the principal approach to CBSD management is the use of disease free planting material and resistant varieties. Research in developing or selecting resistant varieties is currently on-going in several countries (Vanderschuren et al., 2012), with some promising results in Tanzania, Uganda and Mozambique.

Apart from breeding, selection and roguing have been attempted with less success because different cultivars vary in symptom expression (Hillocks et al., 2000). The relationship between cultivar characteristics and seasonality in expression of CBSD symptoms is not well understood. Most farmers tend to select planting materials at harvesting when most of CBSD symptomatic leaves have been shed (Katinila et al., 2003). This makes it difficult to avoid using the infected materials.

Plant quarantine has shown itself to be the first line of defence, issued legislatively and intended to exclude pests from the host or geographic area (Ebbels, 2003). Legislation establishes the statutory authority for the government to engage in limiting further disposal of the pest or treating the localized infestation (Mohamed, 2003).

12

In Rwanda, the current management of the disease is based on visual inspection of CBSD symptoms in cassava fields and the distribution of varieties that are of moderate resistance to

CBSD infection. This study will provide a CBSV distribution map and information on incidence, severity and alternative hosts; the key tools in development of new policies and objectives that aim at sustainable management of the disease.

13

CHAPTER THREE

3.0 INCIDENCE AND SEVERITY OF CASSAVA BROWN STREAK DISEASE IN

MAJOR CASSAVA GROWING DISTRICTS OF RWANDA

3.1 Introduction

Cassava brown streak disease (CBSD) is the second most important constraint affecting cassava production in Eastern Africa after cassava mosaic disease (CMD) (Mbanzibwa et al.,

2009). It is an important virus disease that damages the starch-bearing tubers of cassava crops. It is associated with two viruses; cassava brown streak virus (CBSV) and Ugandan cassava brown streak virus (UCBSV) (Alicai et al., 2007). In Rwanda, Cassava Brown Streak

Disease was first reported in 2009 in only three districts (ISAR, 2009). A report of 2012 country survey (Gashaka et al., unpublished results) revealed that the country mean of CMD incidence was 24.8% with a severity mean of 2.9, while CBSD means were 18.8% and 2.1 for incidence and severity score, respectively. In 2013, in cassava young fields (3-6 months after planting - MAP), the CBSD means were at 26.8% and 2.2 for incidence and severity, respectively (unpublished data). It is therefore very important to monitor the disease incidence and severity changes regularly, in time and space, for better management of CBSD.

3.2 Study areas

In 2015, a survey was carried out in farmer’s fields in the 10 major cassava-producing districts in three different agricultural zones, in order to establish the occurrence and distribution of cassava brown streak viruses in the ten major cassava growing districts of

Rwanda; including: Bugesera, Nyagatare, Kayonza, and Kirehe (Eastern Province), Rusizi,

Nyamasheke, (Western Province), Gisagara, Nyanza, Ruhango and Kamonyi in Southern

Province. These districts were selected according to the importance of cassava as a food

14 crop, and where the disease under study has caused serious problems (Fig. 4 shows the sampled areas on a map)

Figure 4. Map of Rwanda showing the surveyed areas, 2015

3.2.1 Sampling procedure

The survey was conducted on 3 to 6 months young cassava fields. The approach was to stop at intervals along the itinerary traversing each sampling area (Sseruwagi et al., 2004); the minimum proximity was approximately 5 km. In each Province, three districts were surveyed and in each district 10 fields were randomly selected. In each region, a particular representative route that captures the area of interest was discussed and agreed upon by the survey team and adopted. Amongst issues considered included the sample area and availability of suitable cassava fields. During sampling, cassava farms or fields with either pure stand or intercrops with other crops were selected and randomly visited and 30 representative plants were selected along X-shaped transects in each field. For fields having more than one variety, 30 plants from the predominant variety were assessed for CBSD along

15 the two diagonals. Fields that were too narrow were assessed along a diagonal and fields that had fewer than 30 plants all plants were assessed (Sseruwagi et al., 2004). The following parameters were measured during observations: a) Disease incidence: number of plants with disease relative to the total number of plants

assessed, {i.e. incidence % = (number of plants with symptoms/total number of plants

assessed) x 100}. b) Disease severity: area or volume of plant tissue that is diseased relative to the total area

or volume. Normally, it is expressed using a scale that indicates the extent of symptoms

damage. The scale of 1 to 5 was used, where 1 = no visible CBSD symptoms, 2 = mild

foliar symptoms, on some leaves, 3 = pronounced foliar symptoms, but no die-back, 4 =

extensive foliar symptoms which might include slight dieback of terminal branches, and

5 = severe foliar symptoms and/or defoliation and plant die-back (Hillocks et al., 1996). c) Adult whitefly count: Whitefly populations were assessed by counting the number of

adults whitefly on the top five fully-expanded apical leaves for the tallest shoot of each

of the 30 plants sampled.

Also, geo-coordinates (altitude, latitude and longitude) of the sampled fields were recorded using Geographical Positioning System (GPS) loggers.

3.2.2 Data Analysis

Data were analysed using GenStat 12th Edition (VSN Int., 2011). Incidence, severity and whitefly abundance data were subjected to Analysis of variance (ANOVA) to compare means between districts and the means separated using LSD at 5% level of significance and correlation analysis was done to see the association between different variables. Geo- coordinates (altitude, latitude and longitude) data of CBSV/CBSD were transformed into geographical information maps to show the occurrence degree and distribution.

16

3.3 Results

A total of 93 young fields (3-6 MAP) from 10 most cassava producing districts (Kayonza,

Nyagatare, Kirehe, Kamonyi, Bugesera, Ruhango, Nyanza, Gisagara, Rusizi and

Nyamasheke) were visited during the 2015 survey.

3.3.1 Cassava brown streak disease incidence

Cassava brown streak disease was observed in all the major cassava production areas in

Rwanda. The disease is widely distributed in the surveyed districts with an average incidence of 27.39%. There was a significant difference (P<0.05) in the incidence of CBSD between districts, with Bugesera district having the highest average CBSD incidence (70.33%), followed by Nyanza, Kamonyi, Gisagara and Ruhango at 60.66%, 39.33%, 35.33% and

34.07%, respectively. Kirehe district had the lowest CBSD incidence of 1.33% (Fig.5).

80

70

60 50 40 30 20 10

Incidence mean(%) Incidence 0

Districts

Figure 5. CBSD incidence in the ten cassava growing districts of Rwanda, 2015

17

3.3.2 Cassava brown streak disease Severity

The results on CBSD severity are presented in Figure 6 which showed that cassava brown streak disease severity scores moderate symptoms (score 2.1-2.7). The lowest mean severity was in Nyagatare district (Eastern Province) at 2.1 while the highest was recorded in the district of Kamonyi (Southern Province) at 2.7. However, there was no significant difference

(p>0.05) in CBSD severity between the surveyed districts and a mean severity of 2.4 was observed for all the surveyed areas. Though there were no plants that had severity score 5

(very severe disease symptoms), 7.8% of the sampled plants had a score of 4.

5

4

3

2

1

Severity mean Severity 0

Districts

Figure 6. CBSD severity in the ten districts of Rwanda, 2015

3.3.3 Whitefly abundance

The study results showed that there were significant differences (P<0.05) in B.tabaci abundance between the surveyed districts. Adult whitefly populations were generally high and averaged 22.16 adult whiteflies per plant shoot in the study. The highest mean B. tabaci number was recorded in Nyanza district at 50.56 adults and the lowest was in Rusizi at 0.6 adults per plant shoot.

18

3.4 Discussion

To the best of our knowledge, this survey of cassava brown streak disease infecting cassava in Rwanda was the most comprehensive so far covering the ten cassava most producing districts of Kayonza, Nyagatare, Kirehe, Bugesera in the Eastern province, Kamonyi,

Nyanza, Ruhango, Nyanza in Southern province and Nyamasheke and Rusizi in the Western province. The plants affected by cassava brown streak disease were easily identified due to the symptoms they exhibited. Typical symptoms of CBSD observed were venal chlorosis, intervenal chlorosis, mottling and the diebacks associated with more severe symptoms of the disease as previousely described by Hillocks et al. (2000).

This study revealed an increase of CBSD incidence and severity from 18.8% and 2.1, respectively in 2012 to 27.39% and 2.4 in 2015, respectively. It was observed that mean

CBSD incidence was highest in the southern districts (Kamonyi, Ruhango, Nyanza and

Gisagara) and lowest in western surveyed districts (Nyamasheke and Rusizi). Possible reasons for the high level of disease in South and East provinces include the relatively high density of cassava cultivation, which makes virus spread between crops easier, and the mid- altitude and associated higher average temperatures favouring whitefly vector populations

(Hillocks et al. 1999).

This study also showed an inverse relationship between altitude and whitefly abundance and also between altitude and CBSD severity. These associations provide a strong indirect indication that CBSD is most prevalent in agro-ecologies favourable for whitefly population increase. The virtual low incidence and severity of CBSD from higher altitude parts of the surveyed areas confirms that in Rwanda the disease is still behaving in the manner established during the early years of CBSD research, in which it was noted that CBSD was not a problem at higher elevations (Nichols 1950). At that time, it was hypothesized that this

19 was due to the lack of transmission of the causal virus(es) at higher altitudes. Studies in

Tanzania also linked decreasing abundance of the whitefly vector with increasing altitude

(Legg and Raya 1998; Hillocks et al. 1999). Findings from this study also match with the studies by Alicai et al. (2007) who explained that the re-emergence of CBSD in central

Uganda coincided with high B. tabaci population in the area and studies by Maruthi et al.

(2005) who noted that the period of spreads of CBSD in coastal Tanzania closely coincided with abundance in whitefly population at the time.

Results from the survey indicate moderate CBSD severity at an average of 2.4. However, it should be noted that there was an increase from 2.1 in 2012 to the present 2.4. The moderate severity may suggest a non aggressive CBSD virus species/ strain- in the surveyed districts.

This was the case with the CMB recombinant variant (EACMV-UG) of the EACMV, which has been associated with severe CMD that spread from Uganda in 1988 to neighbouring countries as described by Gibson et al. (1996) and Legg (1999). The results may also herald a gradual increase in severity with time and the need to institute containment procedures. Plant age could also have influenced the severity of CBSD symptoms. Mature plants (up to six months) generally had severe leaf symptoms compared to young ones. Increasing shoot symptoms severity with plant age is a notable feature of CBSD (Hillocks et al., 2001;

Nichols, 1950).

Five districts (Bugesera in East, Kamonyi, Ruhango, Nyanza and Gisagara in South) that are known to be the highest cassava producers in the country exhibit high CBSD incidence, severity and whitefly abundance, a combination of main factors for further CBSD spread.

This is a great challenge in management of the disease since these districts are ecologically

20 favourable for cassava production and multiplication; i.e. if measures are not taken the whole country may become a CBSD hotspot, resulting into a threat to food security in Rwanda.

In conclusion, results from this study indicate that cassava CBSD infections are on the rise and the rise is being promoted by high populations of whiteflies. As such, management options should include options that target the vector.

21

CHAPTER FOUR

4.0 MOLECULAR VARIABILITY OF ISOLATES OF CBSVs INFECTING

CASSAVA IN RWANDA

4.1 Introduction

In Rwanda, recent research efforts have only focused on CBSV detection and determination of incidence, severity and whitefly counts by conducting diagnostic surveys. However, the

Rwandan isolates of CBSVs are not yet known, which presents a big gap in knowledge of epidemiology and management of CBSD in Rwanda. It is therefore very important to know the different isolates of CBSD causal agents, and then develop appropriate diagnostic tools for proper management of the disease in Rwanda.

4.2 Detection of cassava brown streak viruses in collected samples

4.2.1 Sample collection

During the survey as described in Study I. section 3.1.3, in each selected field, at least 3 leaf samples were collected randomly on CBSD diseased and symptomless plants and a total of

279 leaf samples were collected from 93 fields. The leaf samples were pressed using herbarium sheets and boards, and left to dry which allowed for longer storage time prior to detection of

CBSVs.

4.2.2 Nucleic acid extraction

RNA was extracted from approximately 100mg of cassava leaf samples using CTAB

(cetyltrimethyl ammonium bromide) (Doyle et al., 1987). The leaves were placed in 2ml eppendorf tubes with metal beads to facilitate grinding for 40 seconds in an automated grinder machine. 1 ml of extraction buffer with 0.2% β-mercaptoethanol was added and vortexed to disperse the tissue in the buffer. Then the mix was incubated at 65°C for 15 min,

22 while shaking vigorously several times. The extract was then mixed with 800 μl of chloroform: isoamyl alcohol (24:1); vortexed briefly and centrifuged at 12,000 rpm for 10 min at 4°C. The top aqueous solution (400 μl) was removed and transferred into new micro- centrifuge tubes to which an equal volume of cold isopropanol was added to precipitate the

RNA. The content was then incubated at -20 for 20 to 30 min followed by centrifugation at

13,000 rpm for 10 min at 4°C to precipitate the pellet and the supernatant was discarded. The

RNA pellet was then washed in 500 μl of 70% ethanol and the tubes vortexed briefly before being incubated at -20°C for at least 10 min. The tubes were then centrifuged for 5 min at

13,000 rpm. The ethanol was removed and the pellet air-dried. Finally the dried pellet were re-suspended in 50 μl of sterile nucleic-free water and stored at -20. DNA residue was digested using a DNase enzyme in order to remain with RNA only.

4.2.3 Complementary DNA synthesis

The first strand of a complementary DNA was synthesised according to Thermo Scientific

RevertAid First strand Synthesis Kit by mixing in each tube 1μl of RNA template, Oligo dt 1

μl and 10 μl of nuclease free water into a sterile nuclease free tubes and the mix was incubated at 65°C for 5min. After that, vials were returned on ice and the following components were added; 5x Reaction buffer 4 μl, RiboLock RNase Inhibitor (20U/μl) 1μl,

10mM dNTP Mix 2μl, and 1 μl RevertAid M-MuLV RT (200U/ μl). The total volume of 20

μl in each tube was incubated at 40°C for 60 min. the reaction was terminated by heating the mix at 70°C for 5min.

4.2.4 PCR Amplification

The amplification of cDNA was performed using one pair of specific primers CBSDDF2 (5-

GCTMGAAATGCYGGRTAYACAA-3) and CBSDDR (5-

GGATATGGAGAAAGRKCTCC-3) primers to amplify partial 344 bp fragments of the coat protein (CP) of CBSV and a 430-440 bp fragment for UCBSV (Mbanzibwa et al.,2011). The

23

PCR reaction mix consisted of 24 μl made up of 16.0 μl sterile distilled water, 2.5 μl of 10 X

PCR buffer, 1.5 μl of MgCl2 (25 mM), 0.5 μl of 10mM of deoxynucleotide triphosphate

(dNTPs), 0.2 μl of Tween 20 (5%), 1.0 μl of each of the primers CBSDDF2/CBSDDR (10 mM), 0.3 μl of Taq DNA polymeramase, and 1.0 μl of cDNA template.

PCR cycling programme was performed using the thermocycler ICycler Biorad® version

4.006 as follows: 94 oC for 2 min for initial denaturation followed by 35 cycles of 94 oC (30 sec), 51 oC (30 sec) and 72oC (30 sec) for denaturation, annealing and extension, respectively.

PCR products were analyzed by electrophoresis in a 1×TAE buffer on a 2% agarose gel, stained with ethidium bromide (0.1 mg/ml). The amplified DNA fragments were electrophoresed in a 1.2 % agarose gel stained with Ethidium bromide (0.01 μl/ml) at a voltage of 80 V and visualisation was done under ultraviolet (UV) light and recorded using an image analyser.

4.2.5 Sequencing and phylogenetic analysis

The PCR products of the positive amplified samples were sent for Sanger’s sequencing method (Sanger et al., 1977) at MACROGEN S.A (Amsterdam, the Netherlands). Sequences were imported to Geneious 9.1.5(Kearse et al., 2012) for further analyses. Contigs were generated using De Novo Assemble and consensus sequences were built in Geneious.

Sequence alignment was conducted using the MAFFT alignment option (default parameter) and the aligned sequences were used to generate phylogenetic trees. Phylogenetic trees were generated using the Bayesian method and this approach provides many trees by sampling the best ones using MrBayes (Ronquist et al., 2012). MrBayes 3.2.1 was run in parallel on the

Magnus supercomputer (located at Pawsey Supercomputer Centre, Perth, Western Australia) utilising the BEAGLE library. MrBayes 3.2.1 was run with a GTR + I + G model of molecular evolution, utilizing four chains for 30 million generations and trees were sampled every 1000 generations. All runs reached a plateau in likelihood score, which was indicated

24 by the standard deviation of split frequencies (0.0015), and the potential scale reduction factor (PSRF) was close to one, indicating that the Markov chain Monte Carlo (MCMC) chains converged.

4.3 Results

4.3.1 Detection of CBSV

Using the species specific primer pair CBSDF2 and CBSDR to screen for viruses, CBSVs were confirmed to be present in symptomatic and non symptomatic samples. In a total of 279 cassava leaf samples collected, only 158 were having clear symptoms and the remaining 121 were asymtomatic. Moreover, among symptomatc samples, 36 tested CBSV negative and 31 of asymtomatic tested CBSV positive. A total 154 out of 279 samples had the virus with 102

UCBSV, 40 CBSV and 12 dual infection.

On agalose gel, amplifed CBSVs were electrophoresed and band appeared at the expected sizes 344 bp for CBSV and 440 for UCBSV (Fig.7).

L C+ 1 2 3 4 5 6 7 8 9 10 11

440bp

344bp

Figure 7. Agarose gel results of PCR products of CBSV(344bp) and UCBSVs (440bp) coat protein. L: ladder, C+: positive control, 1-11: sample one to ten.

25

4.3.2 Geographical distribution of CBSVs

Based on the GPS coordinates of the sampled points and RT-PCR test results of 279 samples, a map indicating the presence of CBSVs in the surveyed areas was constructed (Fig.8b).

Ugandan Cassava Brown Streak Virus was more widespread than Cassava Brown Streak

Virus in the surveyed districts (Table 1). Single infections of UCBSV occurred in all the 10 cassava producing districts (Kayonza, Kirehe, Nyagatare, Kamonyi, Bugesera, Ruhango,

Nyanza, Gisagara, Nyamasheke and Rusizi). Also, single infections of CBSV were detected in 7 of the district including Kayonza, Kirehe, Kamonyi, Bugesera, Ruhango, Nyanza,

Gisagara. Dual infections (CBSV-UCBSV) were detected in only 4 districts: Kayonza,

Bugesera, Ruhango, and Nyanza. Results also show the distribution of CBSVs in plants with different symptom severity scores (Table 2), where the predominant score is 3 for CBSV and

UCBSV, and 2 for dual infection (UCBSV+CBSV). The disease was also detected in the symptomless sample at 19.5% of the total samples.

Table 1: Occurrence and distribution of cassava brown streak viruses in surveyed districts of Rwanda, 2015 (Figures in parentheses are percentages; +, positive)

Districts + UCBSV + CBSV Dual infection Total Kayonza 8(5.2) 6(3.9) 3(1.9) 17(11.0) Kirehe 8(5.2) 4(2.6) 0(0.0) 12(7.8) Nyagatare 6(3.9) 0(0.0) 0(0.0) 6(3.9) Kamonyi 17(11.0) 7(4.5) 0(0.0) 24(15.6) Bugesera 10(6.5) 7(4.5) 4(2.6) 21(13.6) Ruhango 11(7.1) 5(3.2) 4(2.6) 20(13.0) Nyanza 12(7.8) 6(3.9) 1(0.6) 19(12.3) Gisagara 15(9.7) 5(3.2) 0(0.0) 20(13.0) Nyamasheke 11(7.1) 0(0.0) 0(0.0) 11(7.1) Rusizi 4(2.6) 0(0.0) 0(0.0) 4(2.6) TOTAL 102(66.2) 40(26) 12(7.8) 154(100)

26

B A

Figure8. Maps of Rwanda showing the distribution of the viruses causing CBSD in 2012 (A) and 2015 (B).

Table 2: Distribution of CBSVs in plants with different symptom severity scores

Virus detected Plants with the severity score (scale 1-5) Score 1 2 3 4 5 Total UCBSV 24(15.6) 36(23.4) 36(23.4) 6(3.9) 0(0.0) 102(66.2) CBSV 6(3.9) 15(9.7) 17(11.0) 2(1.3) 0(0.0) 40(26) CBSV + UCBSV 0(0.0) 5(3.2) 3(1.9) 4(2.6) 0(0.0) 12(7.8) Total 30(19.5) 56(36.4) 56(36.4) 12(7.8) 0(0.0) 154(100) Figures in parentheses are percentages

4.3.3 Cassava brown streak virus molecular diversity

4.3.3.1 Sequencing

The PCR products of 25 randomly selected cassava samples that positively amplified by RT-

PCR were sequenced by Sanger’s method and blasted in NCBI. Only seventeen were

27 confirmed to be very closely identical to CBSD associated viruses that are already in the

Genbank (Table 3).

4.3.3.2 Phylogenetic analysis

The phylogenetic analysis of the 17 sequences of CBSVs from this study together with selected reference sequences and an outgroup (Cucumber Vein Yellowing Virus), using

MrBayes revealed a clear partitioning into two main groups; UCBSV and CBSV groups. The isolates are grouped into three clusters (Fig. 12 of section 5.4.4.1). This included two clusters for isolates of UCBSV and one for CBSV. The UCBSV isolates from this study are sub- clustered into 3 sub-clusters whereas CBSV isolates are in just one and same clade.

Four isolates (UCBSV_RW_C14_Rus, UCBSV_RW_C17_Bug, UCBSV_RW_C10_Nyz,

UCBSV_RW_C15_Rus) are grouped together and clusters with the old published isolates

(Mbanzibwa et al., 2011) from the Genbank, other four (UCBSV_RW_C19_Kay,

UCBSV_RW_C6_Bug, UCBSV_RW_C16_Kir, UCBSV_RW_C18_Nyt) cluster much more with the CP sequences from newly published isolates by Ndunguru et al (2015). The last

UCBSV isolates under this study (UCBSV_RW_C2_Kam, UCBSV_RW_C5_Bug), closely clustered with the CP sequences from newly published work by Kathurima et al. (2016).

Table 3. Isolates detected in cassava Isolate ID Species Identity (%) to the Accession of the Host reference reference RW-C2- Kam UCBSV 99.3 FJ039520 Cassava RW-C3- Kam CBSV 99.3 HM453034 Cassava RW-C4-Bug CBSV 99.7 HM453034 Cassava RW-C5- Bug UCBSV 95.3 FJ039520 Cassava RW-C6- Bug UCBSV 98.1 LT560277 Cassava

28

RW-C7-Ruh CBSV 79.8 KR911743 Cassava RW-C9-Ruh CBSV 80.2 KR911743 Cassava RW-C10-Nyz UCBSV 85.7 KJ606226 Cassava RW-C11-Nyz CBSV 72.7 HM453034 Cassava RW-C12-Gis CBSV 81.6 HM453034 Cassava RW-C14-Rus UCBSV 99.7 KJ606226 Cassava RW-C15-Rus UCBSV 99.4 KJ606226 Cassava RW-C16-Kir UCBSV 98.1 LT560227 Cassava RW-C17-Bug UCBSV 99.4 KJ606226 Cassava RW-C18-Nyt UCBSV 87.1 LT560277 Cassava RW-C19-Kay UCBSV 98.3 LT560277 Cassava RW-C20-Kay CBSV 78.4 KR911743 Cassava

4.4 Discussion

Ugandan Cassava Brown Streak Virus (UCBSV) was the first reported to occur in Rwanda in

2009 (ISAR, 2009). In this study, we report for the first time the occurrence of cassava brown streak virus and dual infections of UCBSV+CBSV. The occurrence of the new virus (CBSV) in Rwanda may suggest new spread or recent introduction of the virus. However, the detection of only the two known CBSD-associated virus species is in line with Winter et al.

(2010) who showed that there are only two virus species (CBSV and UCBSV) associated with CBSD in the East African region.

It was also observed that in mixed infections, both or at least one of the viruses may not accumulate to a largely increased level, but may also broaden virus distribution in the host thereby increasing virus availability to the feeding vectors as indicated by Ateka et al (2017) and Mascia et al.(2010). The high prevalence of UCBSV in the study areas suggests that

UCBSV could be limiting the multiplication and distribution of CBSV in the host plant when in mixed infection thereby increasing the availability of UCBSV, but limiting that of CBSV to the vector resulting into the low prevalence of CBSV. Sequence analysis indicated that there is greater genetic variability amongst UCBSV than CBSV isolates. This fact, coupled

29 with the much wider distribution of UCBSV than CBSV suggests that UCBSV is likely to be the endemic species that has been associated with cassava in Rwanda since the earliest records of CBSD in the country (Gashaka et al., 2012 unpublished results). Substantial within country movements of cassava planting materials, which occur informally from farmer to farmer, lead to further dispersal of cassava infecting virus species over relatively short periods of time (Finn et al 2017). CBSV would therefore seem to be a relatively recent introduction from a neighbouring country, most likely Tanzania where CBSV is the predominant species (Mbanzibwa et al., 2009; Rwegasira et al., 2011). Comparisons of full genome sequences of Rwandan CBSVs with those of CBSVs obtained from CBSD-affected areas of neighbouring countries (Tanzania and Uganda) would likely clarify questions about the origins of CBSVs currently occurring in Rwanda. The results from this study highlight the need for the development and implementation of effective management strategies.

30

CHAPTER FIVE

5.0 ALTERNATIVE HOSTS FOR CASSAVA BROWN STREAK VIRUSES (CBSVs)

IN RWANDA

5.1 Introduction

Cassava production is affected by both abiotic (physical factors) and biotic (pests and diseases) constraints. Among biotic constraints, only cassava mosaic begomoviruses (CMBs) and Cassava brown streak viruses (CBSVs), which cause cassava mosaic disease (CMD) and cassava brown streak disease (CBSD), respectively, are presently of social and economic importance in East Africa (Monger et al. 2001a; Hillocks & Jennings, 2003).

About twenty viruses have been identified to infect cassava crops in Africa and elsewhere in the world (Mbanzibwa, 2011a). Of all the cassava viruses known in Africa, including

Cassava mosaic begomoviruses (Geminiviridae: Begomovirus), Cassava brown streak viruses (Potyviridae: Ipomovirus), Cassava Ivorian Bacilliform virus (unassigned), Cassava

Kumi viruses A and B, Cassava “Q” virus, Cassava common mosaic virus (Potexvirus)

(Calvert and Thresh, 2002); only Cassava common mosaic virus (CsCMV) is known to occur in other continents, which suggests that most cassava viruses are indigenous to Africa and moved to cassava from other plants (Calvert and Thresh, 2002). Thus, it is plausible that native plant species could act as alternative and/or reservoir hosts for cassava viruses and may contribute to cassava virus evolution and disease epidemics.

Available information on the natural host range of CMBs indicates that they are not restricted to cassava and its wild relatives such as Manihot glaziovii only. Recently, African cassava mosaic virus (ACMV) was detected in Jatropha multifida L. (Euphorbiaceae) (Ramkat et al.,

2011) and Laportea (=Fluerya) aestuans (Urticaceae) in Nigeria (Rossel et al., 1987). In a recent study both ACMV and east African cassava mosaic virus (EACMV) were documented in

M. glaziovii (a wild relative of cassava) (Ogbe et al., 2006), a weed Combretum confertum

31

(Benth.)(Combretaceae), a leguminous plant, Senna occidentalis (L.) Link (Fabaceae), and only

ACMV in Ricinus communis L. (Euphorbiaceae) in the humid forest and derived/coastal savannah agroecological zones of Nigeria. Moreover, according to Storey (1939), the natural occurrence of CBSD is in the cassava tree (Manihot esculenta Crantz). However, M. glaziovii was found to be infected with CBSV isolates in Tanzania (Mbanzibwa et al., 2011) and in

Kenya (Bock, 1994). To date, there have been no reports on the potential roles of non-cassava plant species acting as virus reservoir or alternative hosts in the perpetuation of CBSVs in

Rwanda and at region level. Increased availability of sequences of isolates from wild species may provide more information on the evolution of CBSV and UCBSV and thus improve our awareness of the adaptation of these viruses to cassava as a new host. Therefore, this study aimed to identify alternative hosts for CBSV and UCBSV in Rwanda.

5.2 Sampling procedure

During the survey as described in chapter three section 3.2, leaf samples with virus-like symptoms were collected from shrubs growing in cassava fields or in bushes surrounding cassava fields ( within 50m from the field). Assessment was done on potential alternate hosts using typical virus-like foliar symptoms; i.e., mottling, curling, vein clearing, inter-veinal chlorosis among others. A total of 101 leaf samples with clear virus-like symptoms, from forty-two (42) different plant species (appendice 2) were collected from 48 villages of the ten surveyed districts. The leaf samples were kept in 70% ethanol in eppendorf tubes to keep them fresh for long-time storage.

32

5.3 Detection of CBSV and UCBSV

5.3.1 Nucleic Acid extraction, Complementary DNA synthesis and Polymerase chain reaction

Total nucleic acid was extracted according the Cety-trimethyl ammonium bromide (CTAB) method (Doyle and Doyle, 1987) as described in chapter four section 4.2.2. The Pellet was not clear, so it was subjected to an advanced cleaning to remove the plants chlorophyll and other impurities by adding in the dried pellet 200 ul of Low salt TAE buffer and left at room temperature for 30 min. After mixing gently by tapping, 200 ulvolume of chloroform-isoamyl alcohol was added, mixed well and spinned at 12,000 rpm. About 180ul of supernatant was transferred to a fresh tube where 200ul of 3M sodium acetate and a double of that volume

[(180ul +200ul)x2] of cold absolute ethanol was added. The mix was left over night at -20°C without shaking or mixing, and then was spin at 13,000 rpm for 10 min, the supernatant was carefully discarded and the pellet kept. The pellet was washed with 500ul of 70% ethanol and dried by air. The complementary DNA synthesis (cDNA) and Polymerase chain reaction

(PCR) were done as described in chapter four sections 4.2.3 and 4.2.4.

5.3.2 Sequencing and phylogenetic analyses

The 12 positive amplified samples were sent to MACROGEN S. A (Amsterdam, the

Netherland) for CP partial sequencing using Sanger’s sequencing method and the sequences obtained were analysed as described in chapter four section 4.2.5.

5.4 Results

5.4.1 Plant characteristics and virus disease symptoms

The non-cassava plant species had a range of virus-like disease symptoms (Fig.10 & table 4).

The foliar symptoms included: feathery chlorosis on either sides of the small veins, yellowing on the older leaves on the apex with intermitted mosaic (yellow and green patches), leaf

33 mottling, leaf curling and spotted yellow patches commonly near the apex and stunting. Most of the symptoms appeared on the mature leaves and the young expanding leaves were often symptomless. A brief description of the characteristics of the non-cassava plant species reported to be CBSD infected in the study is presented in Table 4. The plants were either annual and/or perennial deciduous or herbaceous trees or shrubs. Chinese violet (Asystacia gangetica T. Anders)(Acanthaceae) is a rapidly growing perennial shrubby herb, with oval shaped leaves, which grows to 10m height (Figure 9A). Cassia (Senna spectabilis)(Caesalpinioideae) is a deciduous perennial tree 7-15 m tall, with a spreading crown, and alternate compound leaves (Figure 9G). For most of the year, its leaves are green, but shades old leaves in the dry season. Pawpaw (Carica papaya) (Caricaceae) is an annual or perennial herbaceous plant, that is widely cultivated as a fruit tree by both smallholder and commercial farmers. Pawpaw has compound leaves attached to hollow petioles. The leaves are mostly green until senescence when they change colour to yellow and drop off the stem as the plant matures (Figure 9E). Acanthus pubescens (Acanthaceae), a perennial herbaceous shrub with spiny leaves. It grows to 4 m tall. Its vegetation stays green for most of its growth life (Figure 9D). Malva-branca (Sida cordifolia) (Malvaceae) a short perennial shrub (Figure

9F).

Some of the other plants that had typical virus-like disease symptoms, but tested negative for CBSVs included: Cape gooseberry (Physalis peruviana L.), an annual or perennial herbaceous 0.4-3 m tall plant like wild tomato, but usually with a stiffer, more upright stem. It may be cultivated for its fruits but grows mainly as a wild plant. It has compound broad leaves (Figure 9C). Centella or Asiatic pennywort (Centella asiatica L.), is a herbaceous, frost-tender perennial aquatic plant with slender, creeping stems, rounded leaves and rhizomes, growing vertically down (Figure 9B).

34

A

B

C D

E F

G

Figure 9. Virus-like symptoms on CBSV positively tested shrubs: A, Asystacia gangetica (Acanthaceae) B, Centella asiatica (L) (Apiaceae) C, Physalis peruviana L. (Solanaceae ) D, Acanthus pubescens (Acanthaceae) E, Carica papaya L.( Caricaceae)F, Sida cordifolia L. (Malvaceae) G, Senna spectabilis (DC.) (Caesalpinioideae)

5.4.2 Ecology and distribution of non-cassava plant species

The ecology within which the non-cassava plant species were sampled is described in brief in

Table 4. Cassia (Senna spectabilis DC.) grows mainly as a garden ornamental or shade tree in compounds and along roads. It is a highly invasive species that aggressively invades natural forests in East Africa, threatening the environment. Physalis peruviana (Cape gooseberry),

35

Acanthus pubescens, Carica papaya (Pawpaw), Asystacia gangetica T. Anders (Chinese

violet) and Sida cordifolia (Malva-branca.) were found growing in open flat grasslands and

along hill sides with short shrubs. In contrast, Centella or Asiatic pennywort (Centella

asiatica L.) occurred in wetlands in the valleys, where cassava was produced together with

sweet potato, arrow roots, beans and maize inter crops.

The distribution of the non-cassava plant species was studied and seems to indicate that the

plants occurred either within cassava fields or in the surrounding bushes, up to 50 m from the

nearest cassava field. Some plant species occurred more frequently than others, while some

were a rare find. The frequency of occurrence was categorized as: ‘very high’ (more than

70% incidence), ‘moderately high’ (20-50% incidence) and ‘low’ (less than or equal to 10%

incidence) (Table 4). Rough estimates show that the incidence of species Senna spectabilis

Asystacia gangetica, and Sida cordifolia to be very high (>70%), while that of species

Centella asiatica and Acanthus pubescens was high (>50%<70%), and that of species Carica

papaya and Physalis peruviana was low (≤10%), and the plants occurred rarely

Table 4: Plant species and ecology of PCR amplified samples for CBSVs in non-cassava plants Local name in Plant species Plant family Key features Virus Occurre Frequenc Kinyarwanda and common and ecology of disease nce in y of name* the plant species symptom relation occurrenc description to e in the cassava collection plants site**

Kasiya Senna Fabaceae/ Deciduous tree, feathery Same Very High spectabilis Caesalpinioi 7-15 m tall, with chlorosis field (DC.) (Cassia) deae a spreading crown, and on either alternate sides of compound leaves. the small Perennial plant veins grown as a shade (Sev. 3) tree in compounds and wild in light forests. Highly invasive. 36

Gaperi Physalis Solanaceae Herbaceous plant, Green In Low peruviana 0.4-3 m tall, like mosaic vicinity (Cape tomato, but (Sev. 3) gooseberry) usually with a stiffer, more upright stem. Are annual or perennial and may be cultivated or grow wild even on poor soils.

Gutwikumwe Centella Apiaceae Herbaceous, Vein In High asiatica Urban frost-tender banding vicinity (Centella/ perennial aquatic (Sev. 3) Asiatic plant with pennywort) slender, creeping stems, rounded leaves and rhizomes, growing vertically down. Common in wetlands.

Igitovu Acanthus Acanthaceae Herbaceous Stunting In High pubescens perennial shrub (Sev. 3) vicinity with spiny leaves growing to 4 m tall with origin in East Africa (Uganda).

Ipapayi Carica papaya Caricaceae Fruit tree widely Venal In Low (pawpaw) cultivated and chlorosis vicinity may be annual or (Sev. 4) perennial.

Ijojwe Asystacia Acanthaceae Rapidly growing Veinal and Same Very high gangetica T. perennial shrubby inter-veinal field and Anders herb which grows chlorosis in (Chinese to 10 m height.. (Sev. 4) vicinity violet)

Umucundura Sida cordifolia Malvaceae Short perennial Leaf Same Very high (Malva- shrub yellowing field and branca) and curling in (Sev. 4) vicinity

** The frequency of occurrence was categorized as: ‘very high’ (more than 70% incidence), ‘moderately high’ (20-50% incidence) and ‘low’ (less than or equal to 10% incidence)

37

5.4.3 PCR Results

Leaf samples were collected from a total of 43 different plant species that exhibited virus-like disease symptoms. Only two plants were from the same family with cassava. Using the species-specific primer pair CBSDF2 and CBSDR (Mbanzibwa et al., 2011) to screen for viruses, CBSVs were confirmed to be present in 55% of cassava and 7% of the non-cassava samples. Out of a total of 101 leaf samples collected from non-cassava plants, only 12 samples from seven different plant species amplified positively (Table 4). On agalose gel, the bands were not consistent and some of the non-cassava samples showed bands located below the expected size of CBSVs. However, in general the bands were located at around

440bp, confirming the presence of UCBSVin these samples. No fragment of CBSV was amplified from the non-cassava leaf samples. On the other hand, out of the 279 cassava leaf samples analysed, 154 were positive, with 102, 40 and 12 testing positively for UCBSV,

CBSV and dual infection (UCBSV+CBSV), respectively.

Figure10: A) Gel picture of UCBSV amplifications from shrub leaf samples, B) Gel picture of CBSVs amplifications from cassava

5.4.4 Sequencing A

The PCR products of the 12 RT-PCR amplified samples from non-cassava plants and 27 randomly selected RT-PCR products from cassava were sequenced. A BLAST program was used to retrieve similar sequences in the NCBI GenBank, only 7 samples from 5 different non-cassava plant species and 20 out of 27 from cassava were confirmed to be closely

38 matched to CBSD-associated viruses (Table 5) that are already in the GenBank. All the isolates from shrubs were UCBSV, whereas those from cassava were both UCBSV and

CBSV.

Table 5: Isolates detected in cassava and alternative hosts

Isolate ID Species Similarity (%) Accession numbers Host to the reference of the reference RW-CP-Car UCBSV 99.5 KJ606226 Pawpaw (Carica papaya) RW-AG-Ac_1 UCBSV 99.2 KJ606226 Chinese violet (Asystacia gangetica) RW-SS-Cae_1 UCBSV 99.3 LT560272 Cassia (Senna spectabili) RW-SC-Mal UCBSV 97.5 LT560272 Malva-branca (Sida cordifolia) RW-AP-Ac UCBSV 98.6 KJ606226 Acanthus pubescens RW-SS-Cae_2 UCBSV 99.2 KJ606226 Cassia (Senna spectabilis) RW-AG-Ac_2 UCBSV 99.3 KJ606226 Chineseviolet(Asystaesia gangetica) RW-C1-Kam CBSV 99.4 KR911743 Cassava (Manihot esculenta) RW-C2- Kam UCBSV 99.3 FJ039520 Cassava (Manihot esculenta) RW-C3- Kam CBSV 99.3 HM453034 Cassava (Manihot esculenta) RW-C4-Bug CBSV 99.7 HM453034 Cassava (Manihot esculenta) RW-C5- Bug UCBSV 95.3 FJ039520 Cassava (Manihot esculenta) RW-C6- Bug UCBSV 98.1 LT560277 Cassava (Manihot esculenta) RW-C7-Ruh CBSV 79.8 KR911743 Cassava (Manihot esculenta) RW-C8-Ruh UCBSV 72.9 LT560277 Cassava (Manihot esculenta) RW-C9-Ruh CBSV 80.2 KR911743 Cassava (Manihot esculenta) RW-C10-Nyz UCBSV 85.7 KJ606226 Cassava (Manihot esculenta) RW-C11-Nyz CBSV 72.7 HM453034 Cassava (Manihot esculenta) RW-C12-Gis CBSV 81.6 HM453034 Cassava (Manihot esculenta) RW-C13-Gis UCBSV 84.1 KJ606226 Cassava (Manihot esculenta) RW-C14-Rus UCBSV 99.7 KJ606226 Cassava (Manihot esculenta) RW-C15-Rus UCBSV 99.4 KJ606226 Cassava (Manihot esculenta) RW-C16-Kir UCBSV 98.1 LT560227 Cassava (Manihot esculenta) RW-C17-Bug UCBSV 99.4 KJ606226 Cassava (Manihot esculenta) RW-C18-Nyt UCBSV 87.1 LT560277 Cassava (Manihot esculenta) RW-C19-Kay UCBSV 98.3 LT560277 Cassava (Manihot esculenta) RW-C20-Kay CBSV 78.4 KR911743 Cassava (Manihot esculenta)

5.4.4.1 Phylogenetic analyses

Phylogenetic analyses revealed that isolates obtained from cassava plants and those from non-cassava plant species clustered with the respective CBSV and UCBSV isolates available in GenBank (Table 5) and thus are clustered into two main groups. The UCBSV isolates from

39 non-cassava hosts were closely related to the corresponding virus isolates infecting cassava

(Fig.11: Table 5). These findings showed that isolates fromed non-cassava plants in this study, expressed high similarities between themselves (97.5 to 99.5%) and form their own sub-clade (Fig. 12). However, they were closer to some Rwandan isolates reported in the current study including UCBSV_RW_C14_Rus, UCBSV_RW_C15_Rus,

UCBSV_RW_C10_Nyz and UCBSV_RW_C17_Bug, and an old Rwandan isolate

(UCBSV_RW_JX570676) and one from the Democratic Republic of Congo

(UCBSV_DRC_JX291144) that are reported in the Genbank.

40

Figure 11: Phylogenetic tree (cladogram) inferred from the MrBayes method using Geneious 9.1.5 for nucleotide identities of partial coat protein of CBSVs. UCBSV and CBSV isolates determined in this study are all highlighted in yellow with those from non cassava plant sp. circled in red.

41

5.5 Discussion

I report here for the first time the occurrence of five non-cassava plant species including

Asystacia gangetica (Acanthaceae), Senna spectabilis (Caesalpinioideae), Carica papaya

(Caricaceae), Acanthus pubescens (Acanthaceae) and Sida cordifolia (Malvaceae) that tested positive with UCBSV, confirming that these plants are alternative host plants for the virus in

Rwanda.

The five non-cassava plants that have tested positive for UCBSV are not from

Euphorbiaceae, the plant family that contains cassava. These 5 non-cassava plants are from very divergent plant families. They represent 5 different plant orders: 1) Asystacia gangetica

(Acanthaceae), Lamiales 2) Physalis peruviana (Solanaceae), Solanales 3) Carica papaya

(Caricaceae), Brassicales 4) Sida cordifolia (Malvaceae), Malvales, 5) Senna spectabilis

(Caesalpinioideae), Fabales and 2 major higher level clades the Asterids and Rosids (

Stevens, 2001). Why does this matter? Most have thought the CBSD alternative hosts would be closely related to cassava (Mbanzibwa, 2011 and Bock ,1994). In fact, reservoirs for

UCBSV are found throughout the angiosperm phylogeny. Practically, this means that future research need to focus on all plants as possible sources for the viruses – focusing on cassava relatives could overlook the main sources for CBSD.

Knowledge of the host range of crop-infecting pathogens and insect pests is key in the development of sustainable integrated pest and disease management (IPDM) strategies. For years, it was believed that cassava-infecting viruses in sub Saharan Africa, including CMBs and CBSVs, have their original hosts in native African plant species. Following the introduction of cassava from the Americas into Western and Eastern Africa in the 13th and

16th centuries, respectively by Portuguese sailors, the crop was massively adopted and produced as a major subsistence food staple (Jones, 1969 (Hillocks, 2002); these results

42 confirm the fact that existing non-cassava plants with the cassava-infecting viruses served as sources of inoculum spreading the viruses into the cassava crops.

The virus-like disease symptoms observed on the non-cassava plant species in our study were diverse and included all the shades characteristic of CBSD (Nichols, 1950; Storey, 1936).

The symptoms included: feathery chlorosis on either side of the small veins, yellowing on the older leaves on the apex with intermitted mosaic (yellow and green patches), leaf mottling, leaf curling and spotted yellow patches commonly near the apex and stunting. Most of the symptoms appeared on the mature leaves and the young expanding leaves were often symptomless. The affected leaves always retained some green patches amidst chlorosis or necrosis. Knowledge of virus disease symptoms is key, and this should be extended to smallholder farmers and extension personnel to ensure adequate and timely monitoring and reporting of suspected non-crop disease hosts, and hence reduce the emergence and growth of crop disease threats.

The non-cassava host plant species were diversely distributed ranging from cultivated areas near or within cassava fields. However, many farmers in Rwanda use Senna spectabilis as a hedge crop around different fields because of its value as an agroforestry tree. Due to its perennial nature, this plant species could offer a constant source of inoculum to spread

CBSD-associated viruses to new plantings of cassava in the vicinity. Similarly, Carica papaya L. which is a fruit-tree found in and around fields would provide a reservoir for

CBSVs year round. Asystacia gangetica, Sida cordifolia L. and Acanthus pubescens are also perennial weeds and are most likely to occur in many regions of Rwanda which may keep the

CBSVs in the country if management measures are not taken seriously.

43

In summary, this study confirms for the first time the occurrence of CBSD causal agents in other plant species than cassava and underscores the importance of documenting the natural host range of different CBSVs to better understand the role of indigenous non-cassava plant species in the epidemiology of CBSD.

44

CHAPTER SIX

6.0 GENERAL DISCUSSION, CONCLUSION AND RECOMMENDATIONS

6.1 General discussion

This study aimed at determining the incidence and severity of CBSD in the ten districts of

Rwanda. It also aimed at determining the diversity of cassava brown streak viruses; and to determine the possible alternative hosts for CBSD causal agents in Rwanda. Results from this study have shown that CBSVs and the disease that they cause occur throughout the major cassava growing districts of Rwanda. Fifty-five percent of the samples collected were diagnosed positive for the virus in the surveyed districts. It suggests that CBSD is a problem throughout the cassava production systems in the country. The results in this study indicated that CBSD incidence in young cassava fields (3 to 6 MAP) was high in southern surveyed districts, while lowest in western surveyed districts of Rwanda. CBSV species occurs in

Rwanda not only in single infection but also in co-infections (UCBSV+CBSV), though prior studies had reported only UCBSV species to be associated with CBSD in Rwanda (Gashaka et al., 2012 unpublished results).

In this study, the lack of a clear association between CBSD symptom severity and the virus types shows the absence of any virulent CBSD-associated virus on cassava in the study areas.

This finding is in agreement with the study of Winter et al., 2010 and Mbanzibwa et al.,

2010, who suggested that the two virus species (CBSV and UCBSV) in single or mixed infection seem to induce and exhibit similar symptoms on cassava in the field suggesting lack of an aggressive species of CBSVs on cassava in the studied areas. Based on the phylogenetic analysis, it was clear that CBSV isolates in Rwanda are heterogeneous at the isolate level, and were clustered in two subgroups, this serves to demonstrate that genetic diversity of cassava brown streak viruses is high, and, the fact that CP sequences of isolates from Rwanda cluster well

45 with the newly published isolate from Tanzania by Ndunguru et al. (2015) and Kathurima et al.(2016), predict that wider screening of full length DNA sequences would be likely to reveal a higher degree of genetic diversities among the CBSVs and their associated isolates in Rwanda.

Results from this study confirmed that five non-cassava species including Asystaesia gangetica

(Acanthaceae), Physalis peruviana (Solanaceae), Carica papaya (Caricaceae), Sida cordifolia (Malvaceae), Senna spectabilis (Caesalpinioideae) are potential alternative host for

CBSVs. These findings have big implications for the epidemiology and management of the disease in Rwanda but also at the regional level:

The occurrence of several host plant species for cassava-infecting brown streak viruses presents a serious challenge in managing cassava virus diseases in smallholder farming systems in Rwanda and the region in general. Perennials plants have a long presence in the crop growing environments and may sustain the presence of virus inoculum if infected with crop-infecting viruses.

Cassava is usually grown together with and/or near bushes with several plant species including such potential host plants for viruses and associated insect vectors. In many areas, the fields are left un-weeded for long periods, especially after the crop attains the age of six months, to reduce on the cost of maintenance. The micro-ecology that builds in such environments encourages the build-up and flourishing of multiple flora and fauna, including crop-infecting pathogens and insect vectors, even in the periods when crops are not in production.

For example, in areas with two cropping seasons, where cassava is grown continuously, such as Rwanda, the presence of virus-infected cassava and alternative host plant species, maintains a high inoculum leading to high disease spread rates with devastating effects to cassava production. The situation is unpredictable and may equally be serious in areas with a

46 single cropping season, where cassava is grown and harvested uniformly, and replaced with other crops. In such environments, such as is the case for many countries in southern Africa including: Malawi, Mozambique and Zambia, the presence of alternative host plants serve as avenues for overwintering of both disease-causing pathogens and insects including plant viruses and associated insect vectors.

Unfortunately, the propagation of the belief that ‘cassava is a hardy plant’ that can do well in marginal environments, requiring minimal attention, only serves to worsen the situation.

Thus, many farmers ignore weeding cassava fields to keep them free of weeds and shrubs, counting on its ability to do well. But in so doing, this maintains potential sources of inoculum for plant-infecting viruses and insect vectors. On a broader level, the findings of our study have significant implications for the epidemiology of CBSD in the Great Lakes region, where the countries share a similar ecology. Not only is access to disease-free planting materials a major challenge to many smallholder farmers, the cost of management of cassava fields to rid them of potential alternative host plant species for cassava viruses and insect vectors can be high.

All five plants species are widely distributed in the four countries (Uganda, Tanzania,

Burundi and the Democratic Republic of Congo) neighbouring Rwanda. It will be important to assess the occurrence and incidence of the UCBSV-infected plant species in Rwanda

(initially) and in the neighbouring countries (where possible) to map the magnitude of the problem.

Since CBSD appears in the list of six serious biological threats to food security in the world

(Pennisi, 2010), the information generated will also benefit and guide CBSD resistance breeding work, all with a view of improving cassava production in the country and hence

47 improving food security and poverty among the rural poor in Rwanda and the region as a whole.

6.2 Conclusions

This study determined the incidence and severity of CBSD in the study areas of Rwanda and has revealed an increase of CBSD incidence and severity from 18.8% and 2.1, respectively in

2012 to 27.39% and 2.4 in this study, respectively. It was observed that CBSD incidence was highest in the southern districts (Kamonyi, Ruhango Nyanza and Gisagara) and lowest in western surveyed districts (Nyamasheke and Rusizi). The disease occurs at a moderate severity score of 2.4, which an increase of 0.3 in magnitude in three years. The study also established the diversity of CBSVs affecting cassava crops and the potential alternative hosts for UCBSV in the studied areas of Rwanda. This study therefore contributes to the understanding of the genetic diversity of CBSVs in Rwanda and has clearly demonstrated the occurrence of two distinct viruses CBSV and UCBSV that are associated with CBSD, i.e.

CBSV as a recently introduced virus and confirmed the perpetuation of UCBSV and also the occurrence of CBSV+UCBSV co-infections.

The study identified five divergent plants, i.e. Asystacia gangetica (Acanthaceae), Senna spectabilis (Caesalpinioideae), Carica papaya (Caricaceae), Acanthus pubescens

(Acanthaceae) and Sida cordifolia (Malvaceae) as non cassava plant species that host CBSD causal viruses in Rwanda; these results confirm the fact that existing non-cassava plants with the cassava-infecting viruses serve as sources of inoculum spreading the viruses into the cassava crops.

48

6.3 Recommendations

Recommended strategies for the protection of the cassava crop against CBSD outbreaks should include:

1. Studies on the ability of B. tabaci and other possible vectors to transmit CBSVs to non-

cassava hosts and vice versa would complement findings from this study to provide a

foundation for a detailed understanding of CBSD epidemiology and help to develop

sustainable strategies for the management of the disease in Rwanda and in the wider East

African Region.

2. Studies on the incidence and geographical distribution of the CBSD-affected non-cassava

plant species in Rwanda should be established, starting in the areas where the UCBSV-

infected plants reported in the current study were collected,

3. An IPM strategy that emphasizes the creation of awareness of virus disease symptoms

and the role of alternative plant species in the spread of plant virus diseases is paramount.

The IPM strategy should encourage the farmers to remove all identified alternative host

plant species in and around cassava fields. Even greater emphasis should be placed on

keeping crop fields free of weeds and shrubs that may have potential for serving as

alternative hosts for the crop-infecting viruses and associated insect vectors.

49

REFERENCES Abarshi, M., Mohammed, I., Wasswa, P., Hillocks, R., Holt, J., Legg, J., . . . Maruthi, M.

(2010). Optimization of diagnostic RT-PCR protocols and sampling procedures for

the reliable and cost-effective detection of Cassava brown streak virus. Journal of

virological methods, 163(2), 353-359.

Abraham, A. (1956). Tapioca cultivation in India: Indian Council Of Agricultural Research;

New Delhi.

Alabi, O., Ogbe, F., Bandyopadhyay, R., Dixon, A., Hughes, J., & Naidu, R. (2007). The

occurrence of African cassava mosaic virus and East African cassava mosaic

Cameroon virus in natural hosts other than cassava in Nigeria. Paper presented at the

Phytopathology.

Alicai, T., Omongo, C., Maruthi, M., Hillocks, R., Baguma, Y., Kawuki, R., . . . Colvin, J.

(2007). Re-emergence of cassava brown streak disease in Uganda. Plant Disease,

91(1), 24-29.

Stevens, P. F. (2001 onwards). Angiosperm Phylogeny Website. Version 12, July 2012 [and

more or less continuously updated since]." will do.

http://www.mobot.org/MOBOT/research/APweb/.

Bock, K. (1994). Studies on cassava brown streak virus disease in Kenya. Tropical Science,

34(1), 134-145.

Bua, B., & Namara, J. (2009). Reaction of Nicotiana species to Cassava Brown Streak Virus

from Uganda. Paper presented at the 9th African Crop Science, Conference

Proceedings, Cape Town, South Africa, 28 September-2 October 2009.

Calvert, L., & Thresh, J. M. (2002). The viruses and virus diseases of cassava. Cassava:

biology, production and utilization, 237-260.

50

Campo, B. V. H., Hyman, G., & Bellotti, A. (2011). Threats to cassava production: known

and potential geographic distribution of four key biotic constraints. Food Security,

3(3), 329.

Cock, J., Franklin, D., Sandoval, G., & Juri, P. (1979). The ideal cassava plant for maximum

yield. Crop Science, 19(2), 271-279.

Doyle, J., & Doyle, J. (1987). DNA extraction by using DTAB-CTAB procedures.

Phytochemical Bulletin, 19, 11-17.

Ebbels, D. L. (2003). Principles of plant health and quarantine: CABI.

El-Sharkawy, M. A. (1993). Drought-tolerant cassava for Africa, Asia, and Latin America.

BioScience, 43(7), 441-451.

Fargette, D., Colon, L., Bouveau, R., & Fauquet, C. (1996). Components of resistance of

cassava to African cassava mosaic virus. European Journal of Plant Pathology,

102(7), 645-654.

Fauquet, C., Mayo, M., Maniloff, J., Desselberger, U., & Ball, L. Virus Taxonomy, VIIIth

Report of the ICTV. 2005: Elsevier, Academic Press.

Fauquet, C., Mayo, M., Maniloff, J., Desselberger, U., & Ball, L. (2005). ICTV (2005).

VIRUS TAXONOMY: VIIIth Report of the International Committee on Taxonomy of

Viruses: Elsevier Academic Press, Hong Kong.

Gibson, R. W., LEGG, J. P., & OTIM‐NAPE, G. W. (1996). Unusually severe symptoms are

a characteristic of the current epidemic of mosaic virus disease of cassava in Uganda.

Annals of Applied Biology, 128(3), 479-490.

Hillocks, R. (1997). Cassava virus diseases and their control with special reference to

southern Tanzania. Integrated Pest Management Reviews, 2(3), 125-138.

51

Hillocks, R., & Jennings, D. (2003). Cassava brown streak disease: a review of present

knowledge and research needs. International Journal of Pest Management, 49(3),

225-234.

Hillocks, R., Raya, M., Mtunda, K., & Kiozia, H. (2001). Effects of brown streak virus

disease on yield and quality of cassava in Tanzania. Journal of Phytopathology,

149(7‐8), 389-394.

Hillocks, R., Raya, M., & Thresh, J. (1996). The association between root necrosis and

above‐ground symptoms of brown streak virus infection of cassava in southern

Tanzania. International Journal of Pest Management, 42(4), 285-289.

Hillocks, R., & Thresh, J. (2000). Cassava mosaic and cassava brown streak virus diseases in

Africa: a comparative guide to symptoms and aetiologies. Roots, 7(1), 1-8.

Hillocks, R. J. (2002). Cassava in Africa. Cassava: biology, production and utilization, 41-

54.

Hong, Y., Robinson, D., & Harrison, B. (1993). Nucleotide sequence evidence for the

occurrence of three distinct whitefly-transmitted geminiviruses in cassava. Journal of

General Virology, 74(11), 2437-2443.

Howeler, R., Lutaladio, N., & Thomas, G. (2013). Save and grow: cassava. A guide to

sustainable production intensification, 1, 87-97.

Jennings, D. (1957). Further studies in breeding cassava for virus resistance. The East African

Agricultural Journal, 22(4), 213-219.

Jennings, D. (1960). Observations on virus diseases of Cassava in resistant and susceptible

varieties. I. Mosaic disease. Empire Journal of Experimental Agriculture, 28(109).

Jeremiah, S., & Legg, J. (2008). Cassava brown streak virus. Available on:

http://www.youtube. com/watch.

Jones, W. O. (1959). Manioc in Africa: Stanford University Press.

52

Kanju, E., Mtunda, K., Muhanna, M., Raya, M., & Mahungu, N. (2002). Management of

Cassava brown streak virus disease in Tanzania. Paper presented at the Cassava

Brown Streak Virus Disease: Past, Present and Future. Proceedings of an international

conference, Mombasa, Kenya.

Karakacha, W. H. (2001). Serological and molecular characterization of begomoviruses

infecting cassava (Manihot esculenta Crantz) in Africa.

Katinila, N., Hamza, H., & Raya, M. (2003). Social Impact of Cassava Brown Streak Virus

Disease on Rural Livelihoods in Southern Tanzania. Cassava Brown Streak Virus

Disease: Past, Present, and Future.

Kearse, M., Moir, R., Wilson, A., Stones-Havas, S., Cheung, M., Sturrock, S., . . . Duran, C.

(2012). Geneious Basic: an integrated and extendable desktop software platform for

the organization and analysis of sequence data. Bioinformatics, 28(12), 1647-1649.

Legg, J. (1999). Emergence, spread and strategies for controlling the pandemic of cassava

mosaic virus disease in east and central Africa. Crop Protection, 18(10), 627-637.

Legg, J., Jeremiah, S., Obiero, H., Maruthi, M., Ndyetabula, I., Okao-Okuja, G., . . . Gashaka,

G. (2011). Comparing the regional epidemiology of the cassava mosaic and cassava

brown streak virus pandemics in Africa. Virus Research, 159(2), 161-170.

Legg, J., Owor, B., Sseruwagi, P., & Ndunguru, J. (2006). Cassava mosaic virus disease in

East and Central Africa: epidemiology and management of a regional pandemic.

Advances in virus research, 67, 355-418.

Legg, J., & Thresh, J. (2003). Cassava virus diseases in Africa. Paper presented at the

Proceedings of the First International Conference on Plant Virology in Sub-Saharan

Africa (4–8 June 2001, Ibadan, Nigeria), IITA, Ibadan, Nigeria.

Lennon, A., Aiton, M., & Harrison, B. (1985). Cassava viruses from Africa. Annual Report of

the Scottish Crop Research Institute. SCRI, Dundee, UK, 168.

53

Mahungu, N., Bidiaka, M., Tata, H., Lukombo, S., & N’luta, S. (2003). Cassava brown streak

disease-like symptoms in Democratic Republic of Congo. Roots, 8(2), 8-10.

Maruthi, M., Hillocks, R., Mtunda, K., Raya, M., Muhanna, M., Kiozia, H., . . . Thresh, J.

(2005). Transmission of Cassava brown streak virus by Bemisia tabaci (Gennadius).

Journal of Phytopathology, 153(5), 307-312.

Mascia, T., Cillo, F., Fanelli, V., Finetti-Sialer, M. M., De Stradis, A., Palukaitis, P., &

Gallitelli, D. (2010). Characterization of the interactions between Cucumber mosaic

virus and Potato virus Y in mixed infections in tomato. Molecular plant-microbe

interactions, 23(11), 1514-1524.

Mbanzibwa, D., Tian, Y., Tugume, A., Mukasa, S., Tairo, F., Kyamanywa, S., . . . Valkonen, J. (2011). Simultaneous virus-specific detection of the two cassava brown streak- associated viruses by RT-PCR reveals wide distribution in East Africa, mixed infections, and infections in Manihot glaziovii. Journal of virological methods, 171(2), 394-400. Mbanzibwa, D., Tian, Y., Tugume, A., Mukasa, S., Tairo, F., Kyamanywa, S., . . . Valkonen,

J. P. (2009). Genetically distinct strains of Cassava brown streak virus in the Lake

Victoria basin and the Indian Ocean coastal area of East Africa. Archives of virology,

154(2), 353-359.

Mbanzibwa, D. R. (2011). Molecular Characterization of Viruses Causing the Cassava Brown Streak Disease Epidemic in Eastern Africa.

Mbewe W, Winter S, Mukasa S, Tairo F, Sseruwagi P, Ndunguru J, Duffy S. 2017. Deep sequencing reveals a divergent Ugandan cassava brown streak virus isolate from Malawi. Genome Announc 5:e00818-17.

McQuaid CF, van den Bosch F, Szyniszewska A, Alicai T, Pariyo A, Chikoti PC, et al. (2017) Spatial dynamics and control of a crop pathogen with mixed-mode transmission. PLoS Comput Biol 13(7): e1005654.

54

Mohamed, R. (2003). Role of open quarantine in regional germplasm exchange. Cassava

Brown Streak Virus Disease: Past, Present and Future, 63.

Mohammed, I., Abarshi, M., Muli, B., Hillocks, R., & Maruthi, M. (2012). The symptom and

genetic diversity of cassava brown streak viruses infecting cassava in East Africa.

Advances in virology, 2012.

Monger, W., Seal, S., Isaac, A., & Foster, G. (2001). Molecular characterization of the

Cassava brown streak virus coat protein. Plant pathology, 50(4), 527-534.

Monger, W. A., Alicai, T., Ndunguru, J., Kinyua, Z., Potts, M., Reeder, R., . . . Glover, R.

(2010). The complete genome sequence of the Tanzanian strain of Cassava brown

streak virus and comparison with the Ugandan strain sequence. Archives of virology,

155(3), 429-433.

Muhana, M., Raya, M., Hillocks, R., & Kiozya, H. (2004). On farm performance and

farmers’ evaluation of cassava brown streak tolerant variety “Kiroba” in the eastern

zone of Tanzania’. Paper presented at the 9th Triennial Symposium of the

International Society for Tropical Root Crops–Africa Branch (ISTRC-AB).

Nassar, N., & Ortiz, R. (2010). Breeding cassava to feed the poor. Scientific American, 302(5), 78-84.

Ndunguru J, Sseruwagi P, Tairo F, Stomeo F, Maina S, Djinkeng A, et al. (2015) Analyses of Twelve New Whole Genome Sequences of Cassava Brown Streak Viruses and Ugandan Cassava Brown Streak Viruses from East Africa: Diversity, Supercomputing and Evidence for Further Speciation. PLoS ONE 10(10): e0139321.

doi:10.1371/journal. pone.0139321.

Ng, S. (1992). Tissue culture of root and tuber crops at IITA. Biotechnology: Enhancing

research on tropical crops of Africa CTA/IITA, 135-141.

55

Nichols, R. (1947). Breeding cassava for virus resistance. The East African Agricultural

Journal, 12(3), 184-194.

Nichols, R. (1950). The brown streak disease of cassava: distribution, climatic effects and

diagnostic symptoms. The East African Agricultural Journal, 15(3), 154-160.

Ntawuruhunga, P., & Legg, J. (2007). New spread of cassava brown streak virus disease and

its implications for the movement of cassava germplasm in the East and Central

African region. USAID, Crop Crisis Control Project C3P.

Ogbe, F. O., Atiri, G., Dixon, A. G., & Thottappilly, G. (2003). Symptom severity of cassava

mosaic disease in relation to concentration of African cassava mosaic virus in

different cassava genotypes. Plant pathology, 52(1), 84-91.

Otim-Nape, G., Thresh, J., & Fargette, D. (1996). Bemisia tabaci and cassava mosaic virus

disease in Africa. Bemisia: 1995, taxonomy, biology, damage, control and

management.

Ramkat, R., Calari, A., Maghuly, F., & Laimer, M. (2011). Occurrence of African cassava

mosaic virus (ACMV) and East African cassava mosaic virus–Uganda (EACMV-UG)

in Jatropha curcas. Paper presented at the BMC proceedings.

Ronquist, F., Teslenko, M., Van Der Mark, P., Ayres, D. L., Darling, A., Höhna, S., . . .

Huelsenbeck, J. P. (2012). MrBayes 3.2: efficient Bayesian phylogenetic inference

and model choice across a large model space. Systematic biology, 61(3), 539-542.

Rwegasira, G., Momanyi, G., Rey, M., Kahwa, G., & Legg, J. (2011). Widespread

occurrence and diversity of Cassava brown streak virus (Potyviridae: Ipomovirus) in

Tanzania. Phytopathology, 101(10), 1159-1167.

Rwegasira, G. M. (2009). Aspects of the epidemiology of cassava brown streak virus disease

in Tanzania. University of the Witwatersrand.

56

Rwegasira, G. M., & Rey, C. M. (2012). Relationship between symptoms expression and

virus detection in cassava brown virus streak-infected plants. Journal of Agricultural

Science, 4(7), 246.

Sanger, F., Nicklen, S., & Coulson, A. R. (1977). DNA sequencing with chain-terminating

inhibitors. Proceedings of the national academy of sciences, 74(12), 5463-5467.

Sseruwagi, P., Sserubombwe, W., Legg, J., Ndunguru, J., & Thresh, J. (2004). Methods of

surveying the incidence and severity of cassava mosaic disease and whitefly vector

populations on cassava in Africa: a review. Virus Research, 100(1), 129-142.

Storey, H. (1936). Virus Diseases of East African Plants: V.—Streak Disease of Maize. The

East African Agricultural Journal, 1(6), 471-475.

Thottappilly, G., Thresh, J., Calvert, L., & Winter, S. (2003). Cassava Virus and Virus-like

Diseases of Major Crops in Developing Countries (pp. 107-165): Springer.

Thresh, J., & Cooter, R. (2005). Strategies for controlling cassava mosaic virus disease in

Africa. Plant pathology, 54(5), 587-614.

Thresh, J., & Otim-Nape, G. (1994). Strategies for controlling African cassava mosaic

geminivirus Advances in Disease Vector Research (pp. 215-236): Springer.

Vanderschuren, H., Akbergenov, R., Pooggin, M. M., Hohn, T., Gruissem, W., & Zhang, P.

(2007). Transgenic cassava resistance to African cassava mosaic virus is enhanced by

viral DNA-A bidirectional promoter-derived siRNAs. Plant molecular biology, 64(5),

549-557.

Vanderschuren, H., Moreno, I., Anjanappa, R. B., Zainuddin, I. M., & Gruissem, W. (2012).

Exploiting the combination of natural and genetically engineered resistance to cassava

mosaic and cassava brown streak viruses impacting cassava production in Africa.

PloS one, 7(9), e45277.

57

WALTER, J. K., Head, P. Q. S., & RAYMOND, L. (1982). Heat therapy of cassava infected

with African cassava mosaic disease. Plant Disease, 66(6), 475.

Winter, S., Koerbler, M., Stein, B., Pietruszka, A., Paape, M., & Butgereitt, A. (2010).

Analysis of cassava brown streak viruses reveals the presence of distinct virus species

causing cassava brown streak disease in East Africa. Journal of General Virology,

91(5), 1365-1372.

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APPENDICES

1. ANOVA TABLES Severity and Districts

Change d.f. s.s. m.s. v.r. F pr.

+ District 9 360.9153 40.1017 93.93 <.001

Residual 2780 1186.816 0.4269

Total 2789 1547.731 0.5549

Incidence and Districts

Change d.f. s.s. m.s. v.r. F pr.

+ District 9 1564081 173786.8 250.77 <.001

Residual 2780 1926537 693

Total 2789 3490618 1251.6

Whitefly and Districts

Change d.f. s.s. m.s. v.r. F pr.

+ District 9 922224 102469 27.15 <.001

Residual 2780 10492011 3774

Total 2789 11414235 4093

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2. Non-cassava plants sampled for CBSD alternative hosts identification

No Sampled Plants Plant species Families 1 Senna spectabilis (DC.) Irwin & Fabaceae / Kasiya Barneby Caesalpinioideae 2 avoca Persea americana Mill. Lauraceae 3 Budaforoma Mirabilis jalapa L. Nyctaginaceae 4 Gaperi Physalis peruviana L. Solanaceae 5 Gutwikumwe Centella asiatica (L) Urban Apiaceae 6 Icyumya Aspilia pluriseta SCHWEINF. Asteraceae Melanthera scandens Asteraceae (SCHUM.et TH.) ROBERTY Salvia nilotica Lamiaceae 7 Gutenbergia cordifolia BENTH. Asteraceae Idoma ex OLIVER var. cordifolia 8 Igicucu Manihot graviozii Euphorbiaceae 9 Igicumucumu Leonotis nepetaefolia R. BR. Lamiaceae 10 Igihongore Terminalia mollis M.A.Lawson Combretaceae 11 Acanthus pubescens (T. Acanthaceae Igitovu THOMS. ex OLIV.) ENGL. 12 Ikawa Coffea arabica L. Rubiaceae 13 Ikibonobono Ricinus communis L Euphorbiaceae 14 Ikijumba Ipomoea batatas (L.) Lam. Convolvulaceae 15 Ikinini Cinchona officinalis L. Rubiaceae 16 Clerodendrum rotundifolium Lamiaceae Ikiziranyenzi Oliv. 17 Iminenere 18 Intobo Solanum aculeastrum DUNAL. Solanaceae Igitoborwa Solanum macrocarpon Raddi Solanaceae 19 Inturusu Eucalyptus ssp. Myrtaceae 20 Inyabarasanya Bidens pilosa L. Asteraceae 21 Ipapayi Carica papaya L. Caricaceae 22 Isogo Solanum nigrum L. Solanaceae 23 Itabi Nicotiana tabacum L. Solanaceae 24 Tithonia diversifolia (Hemsl.) Asteraceae Kimbazi A. Gray 25 Kinuka Tagetes minuta L. Asteraceae 26 Majojwe/Ijojwe Asystacia gangetica T. Anders Acanthaceae

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27 Spermacoce princeae Rubiaceae

(K.Schum.) Verdc. var. Ngingojana princeae

28 Thunbergia alata BOJ. ex Acanthaceae Nkurimwonga SIMS.

29 Ruhugura/umunaba Triumfetta rhomboidea Jacq. Malvaceae 30 Trichodesma zeylanicum Boraginaceae Sinkangwagasambu (Burm. f.) R.Br. 31 Oxygonum sinuatum Polygonaceae (HOCHST.et STEUD.ex Ubuhandanzovu MEISN.) DAMMER 32 Umubirizi Vernonia amygdalina Delile Asteraceae 33 Vigna vexillata (L.) A. RICH. Fabaceae Umucasuka var. vexillata Vigna frutescens A. RICH. 34 Umucundura M Sida cordifolia L. Malvaceae 35 Menispermaceae Stephania abyssinica (Quart.- Dill. & A. Rich.) Walp. var. Umuhanda abyssinica 36 Fabaceae Umukubayoka Senna occidentalis L. (Caesalpinioideae) 37 Umukuyu Ficus sycomorus L. Moraceae 38 Umumenamabuye Coffea eugenoides S. Moore Rubiaceae

39 Markhamia lutea (Benth.) Bignoniaceae Umusave K.Schum. 40 Solanecio manii (HOOK.f) Asteraceae Umutagara JEFFREY 41 Ocimum lamiifolium Hochst. Lamiaceae Umwenya ex Benth. 42 Uruteja Commelina africana L. Commelinaceae

61