The Molecular Basis of the Interactions of Rhabdoviruses with Their Insect and Plant Hosts

Home , Dicistroviridae, Drosophila X virus, Luteovirus, Rhabdoviridae, RNA virus, Viroplasm

THE MOLECULAR BASIS OF THE INTERACTIONS OF RHABDOVIRUSES WITH THEIR INSECT AND PLANT HOSTS

DISSERTATION

Presented in Partial Fulfillment of the Requirements for

the Degree Doctor of Philosophy in the Graduate

School of The Ohio State University

Chi-Wei Tsai, M.S.

* * * * *

The Ohio State University 2006

Dissertation Committee:

Professor Saskia A. Hogenhout, Adviser Approved by

Professor David M. Francis

Professor Margaret G. Redinbaugh ______

Professor Thomas G. Wilson Adviser Graduate Program in Entomology

ABSTRACT

Maize fine streak virus (MFSV) and Maize mosaic virus (MMV) are insect

transmitted plant rhabdoviruses. Many rhabdoviruses are economically important pathogens of human, livestock, and crops worldwide. Insects transmit many animal and all plant rhabdoviruses. These viruses invade and replicate in cells of various tissues in

insects, vertebrates, and plants. Therefore, insects are not only vectors but also replication

hosts for rhabdoviruses. Sigma rhabdovirus only infects Drosophila flies and is

transmitted to the progeny of flies through germinal cells. D. melanogaster is extensively

used as a model organism because of its traceable genetics and fully sequenced genome.

MFSV, MMV, and Sigma virus, and interactions with their hosts provide unique systems

for characterization of molecular factors determining rhabdovirus host ranges.

First, I completed the genome sequences of two maize-infecting rhabdoviruses,

MFSV and MMV. The MFSV genome encodes seven genes on the antigenomic strand,

whereas the MMV genome encodes six genes. More information about possible functions

of MFVS and MMV proteins was obtained through in planta cellular localization of

fluorescent-protein fusions. The results showed that the MFSV N, P4, M, and MMV P

proteins target the nuclei of plant cells, whereas the MFSV P, P3, and MMV N, P3, M

proteins do not. These findings are consistent with the presence of nuclear localization

signals (NLSs) only in nuclear targeting proteins. Co-introductions of rhabdovirus

proteins revealed that the rhabdovirus N and P proteins interact in which the P proteins follow the distribution of the N proteins, and that the N and P protein interaction is specific to cognate proteins of each virus.

MFVS and MMV replicate in the nucleus and assemble at the inner nuclear

membrane in insect and plant cells, and therefore nuclear import of viral proteins is

critical to complete virus morphogenesis. Using virus-induced gene silencing (VIGS) combined with in planta cellular localization experiments, I discovered that silencing of

Importin αs in plant cells inhibits the nuclear localization of the MFSV N protein and the

MFSV N-P complex in plant cells, suggesting that the MFSV N protein and the MFSV

N-P complex are dependent on Importin αs for nuclear import in plants. In addition, the

MFSV N protein and the MFSV N-P complex also targeted the nuclei of insect cells,

consistent with the hypothesis that the MFSV proteins interact with conserved nuclear

import pathways of plants and insects. Studies to determine whether the Importin αs are

involved in nuclear import of the MFSV N protein and the MFSV N-P complex into the

nuclei of drosophila cells are ongoing.

To investigate the drosophila response to Sigma virus infection, hybridization

experiments with RNAs from virus-infected and virus-free drosophila adults and cDNA

microarrays that contain whole drosophila genome were conducted. The Imd signaling

pathway was identified as the main component of the drosophila anti-Sigma virus

iii

response, whereas there were no indications of activation of the Toll pathway. This is in contrast with Drosophila C virus (DCV) and Drosophila X virus (DXV), which mainly activate the Jak-STAT and Toll pathways, respectively. This is the first comparative study showing that viruses can induce different immune pathways in drosophila, similarly to

Gram-negative bacteria and Gram-positive bacteria, which predominantly activate the

Imd and Toll pathways, respectively. These findings and future studies of Sigma virus-drosophila interactions with will help to elucidate innate immune response pathways against enveloped viruses in vertebrates. A comparison of how different insects, including drosophila, leafhoppers and planthoppers, respond to rhabdovirus infections should prove interesting.

ACKNOWLEDGMENTS

I am grateful to my advisor, Dr. Saskia A. Hogenhout, for her intellectual guidance,

encouragement, and constant support for this project, and for her assistance in editing this

dissertation.

I would like to thank my advisory committee members, Drs. David M. Francis,

Margaret G. Redinbaugh, and Thomas G. Wilson, for their advice and support of my graduate study.

I am also grateful for the assistance of Drs. El-Desouky Ammar, Xiaodong Bai, Ralf

G. Dietzgen, Roy E. Gingery, Michael M. Goodin, Sophien Kamoun, John A. Lindbo,

Elizabeth A. McGraw, Tea Meulia, Scott L. O’Neill, Markus Riegler, Anna E. Whitfield,

and Joe Win.

I also thank John J. Abt, Valdir R. Correa, Tatiana Fazzolari, Dave E. Fulton, Mark

W. Jones, Roger Mitchell, Sharon E. Reed, Angela D. Strock, William E. Styer, Jane C.

Todd, Tania Y. Toruno, and Kristen J. Willie for their help and friendship.

This research was supported by the OARDC Graduate Research Enhancement Grant

Program, the National Research Initiative of the USDA Cooperative State Research,

Education and Extension Service Grant 2002-35302-12653, and the Australian Research

Council Linkages International Grant LX0452397.

VITA

1995 ...... B.S. Entomology, National Taiwan University

1997 ...... M.S. Entomology, National Taiwan University

1997-1999 ...... Second lieutenant, Army Reserve Officers' Training Corps (ROTC), Taiwan

1999-2001 ...... Research assistant, Department of Entomology, National Taiwan University

2001-present ...... Graduate Research Associate, The Ohio State University

PUBLICATIONS

1. Tsai CW, Redinbaugh MG, Willie KJ, Reed S, Goodin M, and Hogenhout SA. 2005. Complete genome sequence and in planta subcellular localization of Maize fine streak virus proteins. J. Virol. 79:5304-5314.

2. Reed SE, Tsai CW, Willie K, Redinbaugh MG, and Hogenhout SA. 2005. Shotgun sequencing of the negative-sense RNA genome of the rhabdovirus Maize mosaic virus. J. Virol. Methods 129:91-96.

3. Tsai CW and Lee HJ. 2001. Analysis of specific adaptation to a domicile habitat: a comparative study of two closely related cockroach species. J. Medical Entomol. 38:245-252.

4. Tsai CW and Lee HJ. 2000. Circadian locomotor rhythm masked by the female reproduction cycle in cockroaches. Physiol. Entomol. 25:63-73.

5. Tsai CW and Lee HJ. 1997. Volatile pheromone detection and calling behavior exhibition: secondary mate-finding strategy of the German cockroach, Blattella germanica (L.). Zoological Studies 36:325-332.

FIELDS OF STUDY

Major Field: Entomology

vii

TABLE OF CONTENTS Page Abstract ………………………………………..……….…………….…………... ii Acknowledgements …………………………..……….………………….………. v Vita …………………………………………..……….…………….…………….. vi List of Tables ………………………………………………………………….….. x List of Figures ………………...……………………………………………...…… xi

Chapters:

1. Literature Review …………………………………………………...…….. 1

1.1 Abstract ...………………………………………………………...…… 2 1.2 Introduction ……………………………………………………..……. 3 1.3 MFSV and MMV genome organization ……………...……...... ……. 6 1.4 Gene expression and genome replication ……………...…...…….…... 7 1.5 Protein function ………………………………………...... …………... 9 1.6 Virus-insect interactions ………………………………...….…...……. 15 1.7 Virus-plant interactions ………………………………...……....…….. 22 1.8 Research objectives ……………………………….………...... ……... 23 1.9 References …………………………………………………...... ……... 25

2. Complete genome sequence and in planta subcellular localization of Maize fine streak virus proteins …………..…….………………………… 36

2.1 Abstract ………………………..……………………………………… 37 2.2 Introduction …………………………………………………………… 38 2.3 Materials and methods ………………………………………………... 40 2.4 Results ………………………………………………………………… 48 2.5 Discussion …………………………………………………………….. 54 2.6 Acknowledgments …………………………………………………….. 58 2.7 References……………………………………………………………... 60

viii

3. Shotgun sequencing of the negative-sense RNA genome of the rhabdovirus Maize mosaic virus …………..……………………………… 74

3.1 Abstract ………………………..……………………………………… 75 3.2 Introduction …………………………………………………………… 76 3.3 Materials and methods ………………………………………………... 77 3.4 Results and discussion ……….……………………..………………… 80 3.5 Acknowledgments …………………………………………………….. 87 3.6 References……………………………………………………………... 88

4. Subcellular localization and nuclear import of Maize fine streak virus and Maize mosaic virus proteins …………..……………………….…………. 96

4.1 Abstract ………………………..……………………………………… 97 4.2 Introduction …………………………………………………………… 98 4.3 Materials and methods ………………………………………………... 100 4.4 Results ………………………………………………………………… 105 4.5 Discussion …………………………………………………………….. 110 4.6 Acknowledgments …………………………………………………….. 113 4.7 References……………………………………………………………... 114

5. Sigma rhabdovirus activates the innate immune response of drosophila …. 123

5.1 Abstract ………………………..……………………………………… 124 5.2 Introduction …………………………………………………………… 125 5.3 Materials and methods ………………………………………………... 128 5.4 Results ………………………………………………………………… 132 5.5 Discussion …………………………………………………………….. 138 5.6 Acknowledgments …………………………………………………….. 143 5.7 References……………………………………………………………... 144

Bibliography …………………………………………………………….………… 159

LIST OF TABLES

Table Page 1.1. Six genera of the Family Rhabdoviridae …………….……...... …. 31

1.2. Predicted nuclear localization signal of plant rhabdovirus proteins with PSORT ………………………………………………………………….….... 32

2.1. Features of the encoded proteins of the MFSV genome …………...... 63

2.2. Genome organization of plant rhabdoviruses ……………...... ……...…… 64

3.1. Results of shotgun library construction, sequencing, and assembly ...... 90

3.2. Comparison of the genome organizations of the livestock pathogen VSIV, and the plant infecting nucleorhabdoviruses and cytorhabdoviruses ...... 91

3.3. Time comparison of the shotgun and primer walking sequencing rocedures ... 92

3.4. Cost comparison of the shotgun and primer walking sequencing procedures .. 93

4.1. Predicted nuclear localization signal and cellular localization of Maize fine streak virus (MFSV) and Maize mosaic virus (MMV) proteins with PSORT..…….…………………...... ……… 117

5.1. Primers used in this study for amplifying Northern blot probes ………...…… 148

5.2. Overview of differentially regulated genes in response to Sigma virus infection in drosophila ………..……………………………………...... 149

5.3. Comparison of upregulated immunity genes of virus-infected drosophila ...... 150

LIST OF FIGURES

Figure Page 1.1. A. Transmission electron micrograph of Maize mosaic virus (MMV) infected maize leave. B. Transmission electron micrograph of Maize fine streak virus (MFSV) infected maize leave ...... 33

1.2. A. Schematic diagram of the MFSV and MMV genomes. B. Representations of the minimal rhabdovirus genome and virion composition ...... 34

1.3. Persistent propagative transmission of plant rhabdovirus by a leafhopper ...... 35

2.1. Schematic diagram of the MFSV genome organization ...... 65

2.2. Sequences of the 3’ and 5’ termini of plant rhabdovirus genomes ...... 66

2.3. Comparison of rhabdovirus gene junctions ...... 67

2.4. RLM-RACE of the MFSV G and L mRNAs ...... 68

2.5. Detection of MFSV gene transcripts in infected maize by Northern blot analysis ...... 69

2.6. Epifluorescence microgaphs of subcellular localizations of fluorescent- protein fusions of the MFSV N and ORF2 proteins ...... 70

2.7. Epifluorescence micrographs of subcellular localizations of fluorescent-protein fusions of the MFSV ORF3, ORF4, and ORF5 proteins in the cellular and nuclear views ...... 71

2.8. Subcellular localization of fluorescent-protein fusions of the MFSV N and ORF2 proteins and the SYNV N and P proteins ...... 72

2.9. Epifluorescence microgaphs of infiltrated DsRed-SYNV P alone, coinfiltrated CFP-MFSV N and DsRed-SYNV P, and coinfiltrated GFP-SYNV N and YFP-MFSV 2 ...... 73 xi

3.1. Schematic representation of the genome organization of Maize mosaic virus 94

3.2. Comparative sequence analysis of gene junctions of rhabdovirus genomes .... 95

4.1. A. Cellular localization of fluorescent protein fusions of the MMV N, P, P3 and M proteins in N. benthamiana leaves. B. Cellular localization of fluorescent protein fusions of the MFSV N and P proteins in N. benthamiana leaves ...... 118

4.2. RT-PCR analysis confirming the silencing of two NbImp-α homologs of N. benthamiana ...... 119

4.3. A. Cellular localization of the MFSV N and P proteins in the NbImp-α1 and α2 silenced N. benthamiana leaves. B. Cellular localization of the MFSV N-P complex in the NbImp-α1 and α2 silenced N. benthamiana leaves ...... 120

4.4. Cellular localization of the MMV N and P proteins in the NbImp-α1 and α2 silenced N. benthamiana leaves ...... 121

4.5. Cellular localization of fluorescent protein fusions of the SV40 NLS and the MFSV N and P proteins in drosophila S2 cells ...... 122

5.1. Total up- and down-regulated genes in response to Sigma virus infection in drosophila ...... 151

5.2. Northern blot analysis of immunity genes expressing in Sigma virus-infected and virus-free drosophila ...... 152

5.3. Upregulated defense-related genes in response to Sigma virus infection in drosophila ...... 153

5.4. Downregulated defense-related genes in response to Sigma virus infection in drosophila ...... 154

5.5. Up- and downregulated cell fate-related genes of Sigma virus-infected drosophila ...... 155

xii

5.6. Up- and downregulated metabolism genes of Sigma virus-infected drosophila 156

5.7. Up- and downregulated structural genes of Sigma virus-infected drosophila .. 157

5.8. Up- and downregulated sensing genes of Sigma virus-infected drosophila ..... 158

xiii

CHAPTER 1

LITERATURE REVIEW

MAIZE FINE STREAK VIRUS AND MAIZE MOSAIC VIRUS:

A GOOD SYSTEM TO STUDY THE INTERACTIONS AMONG

RHABDOVIRUS, PLANT, AND INSECT VECTOR

1.1. Abstract

Rhabdoviruses are among the most serious pathogens of human, livestock, and crops

worldwide. Insects transmit many animal and all plant rhabdoviruses to their hosts, and insects are not only vectors but also replication hosts for rhabdoviruses. Many rhabdoviruses are also passed on to the progeny of their insect hosts. Insect transmission of plant rhabdoviruses is highly specific. For example, Maize fine streak virus (MFSV) is specifically transmitted by the black-faced leafhopper, Graminella nigrifrons, and Maize mosaic virus (MMV) by the corn planthopper, Peregrinus maidis. These two maize-infecting rhabdoviruses that differ in vector specificity provide a unique system to characterize the molecular factors that determine vector specificity and host range of rhabdoviruses among insect hosts. Insects transmit rhabdoviruses from plant to plant in a persistent propagative manner. Viruses first infect the midgut and other organs, and subsequently the salivary glands from where viruses are introduced into plants along with insect saliva. The midgut cells are the first virus entry sites from where the viruses spread to other tissues presumably via the nervous system, trachea, and hemolymph. For transmission to occur, viruses must cross several membrane barriers, including midgut and salivary gland-infection and escape barriers. In addition to these membrane barriers, the success of rhabdovirus infection in insects is also determined by other factors, such as efficiency of virus replication and effectiveness or onset of various insect immune response pathways. The MFSV and MMV genomes have been sequenced to completion.

The MFSV genome encodes seven genes whereas MMV genome encodes six genes.

More information about possible functions of MFVS and MMV proteins is obtained

through in planta cellular localization of fluorescent-protein fusions. The results showed

that the MFSV N, P4, M, and MMV P proteins only target the nuclei of plant cells,

whereas the MFSV P, P3, and MMV N, P3, M proteins do not. These findings are consistent with the presence of nuclear localization signals only in nuclear targeting proteins. Co-introductions of rhabdovirus proteins revealed that the rhabdovirus N and P

proteins interact in which the P proteins follow the distribution of the N proteins, and that

the N and P protein interaction is specific to cognate proteins of each virus. The nuclear

localization of MFSV and MMV proteins is consistent with the replication site of MFSV

and MMV in plant cells. MFSV and MMV predominantly replicate in the nucleus and

bud through the inner nuclear membrane of various cell types of plants. The viral proteins

are probably multifunctional and also inhibit host gene expression, suppress RNA

silencing, and interact with host proteins causing cytopathogenesis in host cells.

1.2. Introduction

Many rhabdoviruses are economically important pathogens of humans, livestock,

and crops, of which Rabies virus (RABV) is probably the most well known species (Rose

and Whitt, 2001). The rhabdoviruses that cause rabies and fish diseases appear to be

confined to vertebrate hosts, whereas vesiculo-, ephemero-, cyto-, and

nucleo-rhabdoviruses are transmitted to their vertebrate or plant hosts by insects

(Hogenhout et al., 2003; Jackson et al., 2005; Rose and Whitt, 2001). Rhabdoviruses also

infect and replicate in their vector insects. Thus, most rhabdoviruses have two natural

hosts: either insects and plants, or insects and vertebrates. In most cases, rhabdoviruses

are transmitted to the progeny of the insect hosts via the female and male germinal cells

(Brun, 1991; Nault, 1997; Tesh et al., 1972).

Insect transmission of plant rhabdoviruses is highly specific; a given rhabdovirus is

transmitted only by one or a few closely related insect species. Maize fine streak virus

(MFSV) is specifically transmitted by the black-faced leafhopper, Graminella nigrifrons,

and Maize mosaic virus (MMV) is specifically transmitted by the corn planthopper,

Peregrinus maidis (Falk and Tsai, 1985; Redinbaugh et al., 2002). These two maize-infecting rhabdoviruses that differ in vector specificity provide a unique system to characterize the molecular interaction of viruses and their vector insects. Insects play two roles in this system: they are vectors as well as replication hosts for these viruses.

Therefore, the system allows for the characterization of factors that determine vector competence and the virus host range among insects. Unlike mammalian rhabdoviruses, the risk to human health is not an obstacle when studying plant rhabdoviruses. In addition, the research does not involve experiments with live vertebrate animals.

The family Rhabdoviridae consists of six genera that are all associated with disease

of vertebrates or plants (Table 1.1) (Walker et al., 2000). The four genera of

vertebrate-infecting rhabdoviruses (Vesiculovirus, Ephemerovirus, Lyssavirus, and

Novirhabdovirus) include the livestock pathogens Vesicular stomatitis Indiana virus

(VSIV) and Bovine ephemeral fever virus (BEFV), the human pathogen RABV, and the fish pathogen Infectious hematopoietic necrosis virus (IHNV). MFSV and MMV belong to plant nucleorhabdoviruses (Fig. 1.1) (Jackson et al., 2005; Redinbaugh et al., 2002).

The plant-infecting rhabdoviruses are separated into the two genera, Nucleorhabdovirus

and Cytorhabdovirus, depending on their sites of replication and morphogenesis in plant

cells (Jackson et al., 2005; Walker et al., 2000). Nucleorhabdoviruses replicate in the

nucleus, bud through the inner nuclear membrane, and accumulate in the perinuclear

space of plant cell. In contrast, cytorhabdoviruses replicate in the cytoplasm and undergo

morphogenesis from the endoplasmic reticulum, and accumulate in the cytoplasm of

plant cells.

MFSV and MMV have a typical morphology common among rhabdoviruses (Fig.

1.1) (Bradfute and Tsai, 1983; Falk and Tsai, 1983; Redinbaugh et al., 2002).

Rhabdovirus virions are bullet shaped or bacilliform with a knobby surface and helical

nucleocapsid core. A typical rhabdovirus particle is composed of a lipid envelope derived from host nuclear or cellular membrane, and a ribonucleocapsid core consisting of a negative-strand RNA genome bound to a complex of nucleocapsid protein (N), polymerase-associated phosphoprotein (P), and multifunctional polymerase (L) (Rose

and Whitt, 2001). Further, the transmembrane glycoprotein (G) protrudes from the

exterior of the lipid envelope, and the matrix protein (M) connects the G protein and

envelope to the ribonucleocapsid core. Virion sizes vary depending on the virus species

and fixation methods with lengths of 130 to 350 nm and widths of 45 to 100 nm (Jackson

et al., 2005). Similar to other plant rhabdoviruses, the MFSV and MMV virions are

bacilliform particles measuring 231 nm x 71 nm and 224 nm x 68 nm, respectively (Falk

and Tsai, 1983; Redinbaugh et al., 2002).

1.3. MFSV and MMV genome organization

The complete MFSV genome (GenBank accession number NC 005974) consists of

13,782 nt, including a 184 nt 3’ leader and 145 nt 5’ trailer (Tsai et al., 2005/ Chapter 2 of this dissertation). The MMV genome (GenBank NC 005975) consists of 12,133 nt, but the sequences of the 3’and 5’ ends have not been determined (Reed et al., 2005/ Chapter

3 of this dissertation). The MFSV and MMV genomes encode homologs of the five core genes found in the prototypical VSIV genome (Fig. 1.2) (Reed et al., 2005; Tsai et al.,

2005). The coding regions are flanked by leader and trailer sequences whose 3’ and 5’ termini contain sequences that have some degree of complementarity and therefore may give rise to a putative panhandle structure (Tsai et al., 2005). The five structural genes appear in the same order as the genes of VSIV, 3’-N-P-M-G-L-5’, and are thought to serve analogous roles in plant rhabdoviruses (Jackson et al., 2005). In addition to these five core genes, MFSV and MMV genomes carry one or two additional open reading frames (ORFs) residing between the P and M genes. The MFSV genome contains seven

ORFs in the order 3’ leader-N-P-3-4-M-G-L-5’ trailer, and the MMV genome contains six

ORFs in the order 3’ leader-N-P-3-M-G-L-5’trailer (Fig. 1.2) (Reed et al., 2005; Tsai et al., 2005). Other plant rhabdoviruses for which genome sequences are available also contain one to four ORFs between the P and M genes, and none or one ORF between the

G and L genes (Heaton et al., 1989; Huang et al., 2003; Reed et al., 2005; Revill et al.,

2005; Tanno et al., 2000; Tsai et al., 2005; Wetzel et al., 1994). Vertebrate-infecting rhabdoviruses do not have genes between the P and M genes. Some vertebrate

rhabdoviruses have one or more ORFs between the G and L genes, for example, rg gene

of RABV, NV gene of IHNV, and Gns, α1, α2, α3, β, γ of BEFV (Kurath et al., 1997;

McWilliam et al., 1997; Tordo et al., 1986).

Rhabdovirus genomes contain conserved “gene junction” regions between each gene.

The MFSV gene junction regions have the consensus sequence 3’-UUUAUUUUGUAG

UUG-5’ (Tsai et al., 2005), and the consensus sequence of MMV is 3’-AUUCUUUUU

GGGUUG-5’ (Reed et al., 2005). The intergenic gene junctions of rhabdovirus are

comprised of a poly U tract on the genomic strand (element I), thought to be the

intracellular polyadenylation signal or stutter sequence, followed by a short

nontranscribed region (element II), and a conserved transcription start site (element III).

The gene junction regions of rhabdoviruses are highly conserved within the viral genome

and among rhabdoviruses (Jackson et al., 2005; Redinbaugh and Hogenhout, 2005).

Further, similar gene junction sequences are found in Paramyxoviridae, Filoviridae,

Bornaviridae (Jackson et al., 2005). Indeed, these regions have been shown to play important roles in regulating mRNA transcription and genome replication.

1.4. Gene expression and genome replication

The mechanism of gene expression and genome replication of plant rhabdoviruses

has not been analyzed in detail so far, mainly because infectious full-length clones for

reverse genetics experiments are not yet available for plant rhabdoviruses. However, it is

likely that gene expression and genome replication mechanism are conserved among the

rhabdoviruses. Some recent reviews about transcription and replication of rhabdoviruses

have been published, mostly based on studies with VSIV for which an infectious clone is

available (Barr et al., 2002; Rose and Whitt, 2001; Whelan et al., 2004; Jackson et al.,

2005). The RNA genomes of rhabdoviruses are templates for two distinct RNA synthetic

processes: transcription to generate mRNAs and genome replication via production of

positive-strand genome complements that act as templates to generate progeny

negative-strand viral genomes. For transcriptional control, the virus-encoded

RNA-dependent RNA polymerase enters at the 3’ proximal site and transcribes the viral

genes sequentially in the 3’ to 5’order. There is a gradient in the molar abundance of

transcripts for viral genes in the 3’ to 5’ direction, because ~ 30 % of the polymerase fails

to transcribe the downstream gene at each gene junction. Thus, levels of gene expression

are primarily regulated by the position of each gene relative to the 3’ leader and also by

cis-acting sequences located at the beginning and end of each gene and at the gene

junctions.

Following the primary transcription and translation, RNA replication occurs in the cytoplasm (most rhabdoviruses) or nucleus (nucleorhabdoviruses) and depends on the prior translation products. The N protein encapsulates nascent antigenomic and genomic

RNA. The P and L proteins form a RNA polymerase complex. Abundance of the 3’ leader

mRNA, combined with the accumulation of the N protein, and probably the P protein, are

thought to regulate the switch between mRNA synthesis and genomic RNA replication.

The N protein binds the 3’ leader mRNA, however when the N protein increases in larger amounts than the 3’ leader mRNA and in sufficient abundance for encapsidation of nascent antigenomic and genomic RNA, RNA replication is initiated. The depletion of

the N and P proteins in the replication phase triggers new rounds of mRNA transcription

to reload the N and P proteins. Thus, the availability of the N and P proteins provides an

ingenious feedback to regulate mRNA transcription and genomic RNA replication.

1.5. Protein function

Cellular localization studies of plant rhabdovirus proteins are available for MFSV

(Tsai et al., 2005), MMV (Chapter 4 of this dissertation) and Sonchus yellow net virus

(SYNV) (Goodin et al., 2001; 2002; Martins et al., 1998), and biochemical analyses are available for some proteins of SYNV, Lettuce necrotic yellows virus (LNYV), and Rice yellow stunt virus (RYSV) (Jackson et al., 2005). I will explain what is currently known about the function of plant rhabdovirus proteins in the following sections.

1.5.1. The N protein

The SYNV N protein is a major component of the viral nucleocapsid, viroplasm,

and polymerase complex (Wagner et al., 1996), and as mentioned above is involved in

regulation of the gene transcription and genome replication switch. The MFSV and

SYNV N proteins contain nucleoplasmin-like bipartite nuclear localization signals

(NLSs) close to the carboxyl terminus, and these proteins localize to the nuclei of

Nicotiana benthamiana leaf cells (Goodin et al., 2001; Tsai et al., 2005). Putative NLSs

are also identified in the N proteins of nucleorhabdoviruses MMV, RYSV, and Taro vein chlorosis virus (TaVCV) (Table 1.2). Although the MMV N protein contains two

SV40-like NLSs, the protein does not localize to the nuclei of N. benthamiana leaf cells

(Chapter 4 of this dissertation). In addition, it was found that Importin αs are involved in the nuclear import of the MFSV and SYNV N proteins (Goodin et al., 2001; Chapter 4 of this dissertation). Importin αs are karyophilic proteins that recognize the NLS. For nuclear import to occur, Importin α binds NLS-containing proteins and Importin β, and then Importin β docks to the nuclear pore complex translocating the NLS-containing proteins into the nucleus (reviewed in Goldfarb et al., 2004). Nucleorhabdoviruses replicate in the nucleus and assemble at inner nuclear membrane, and therefore nuclear import of viral proteins is critical to complete morphogenesis. Neither cytorhabdovirus nor vertebrate-infecting rhabdovirus N proteins harbor putative NLSs consistent with the cytoplasmic replication and accumulation of these viruses.

1.5.2. The P protein

The SYNV P protein is a component of the viral nucleocapsid core in the virus

particle, and the nucleus-associated polymerase complex during virus replication

(Martins et al., 1998; Wagner and Jackson, 1997). A recent study showed that the SYNV

P protein may also function as a RNA silencing suppressor (Jackson et al., 2005). The

MMV and RYSV P proteins contain putative NLSs at the carboxyl terminus, whereas no

NLS motifs are identified in the P proteins of other plant rhabdoviruses, including MFSV

and SYNV, and vertebrate-infecting rhabdoviruses (Table 1.2). The prediction of the

presence of NLS is confirmed by cellular localization studies, since monomeric DsRed

(mDsRed) fusions of the MMV P protein localizes to the nuclei of N. benthamiana leaf cells (Chapter 4 of this dissertation). In contrast, the SYNV P protein has a karyophilic

region and a nuclear export signal, and GFP fusion of this protein is distributed

throughout the cell in N. benthamiana leaves, suggesting that the SYNV P protein has the

ability to shuttle between the nucleus and cytoplasm (Goodin et al., 2001; 2002; Martins

et al., 1998). However, immunofluorescence localization studies show that the SYNV P

protein localizes to the nuclei of SYNV-infected N. benthamiana protoplasts (Martins et

al., 1998). The nuclear localization of the SYNV P protein is due to the interaction with

other viral proteins (Goodin et al., 2005). The MFSV P protein does not contain a NLS

motif, and YFP fusions of the MFSV P protein is also distributed throughout the cell (Tsai

et al., 2005).

The interaction between rhabdovirus N and P proteins is examined by in planta

colocalization of these two proteins. Several cellular localization studies reveal that

rhabdovirus P proteins follow the distribution of the N proteins when the N and P genes

express simultaneously in plant cells. The YFP-fused MFSV P protein relocalizes to the nucleolus of N. benthamiana leaf cell in the presence of the CFP-fused MFSV N protein

(Tsai et al., 2005), and the DsRed fused SYNV P protein relocalizes to a subnuclear

locale in the presence of the GFP-fused SYNV N protein (Goodin et al., 2002). For MMV,

the mDsRed-fused P protein is redistributed throughout the cell in the presence of the

YFP-fused MMV N protein (Chapter 4 of this dissertation). Thus, the N and P proteins of rhabdoviruses clearly interact resulting in cellular relocalizations. The interaction between the N and P proteins is specific for each virus, because the MFSV P protein does

not relocalize to the subnuclear locale in the presence of the SYNV N protein, and the

SYNV P protein does not relocalize to the subnuclear locale in the presence of the MFSV

N protein (Tsai et al., 2005).

1.5.3. The M protein

The plant rhabdovirus M protein is thought to be responsible for condensing the nucleocapsid of virus particles during maturation and for interacting with the G protein during morphogenesis (Jackson et al., 2005). Similarly to vertebrate-infecting rhabdoviruses (Ahmed et al., 2003; Kopecky and Lyles, 2003), the SYNV M protein inhibits gene expression of the host as evidenced by the observation that expression of the

M gene in plant cells interferes with accumulation of nonviral reporter proteins (Jackson et al., 2005). The M proteins from cytorhabdoviruses and other sequenced vertebrate-infecting rhabdoviruses do not contain NLS motifs, except for two novirhabdoviruses (Redinbaugh and Hogenhout, 2005). However, the MFSV and SYNV

M proteins have NLSs and in planta cellular localization studies show that these proteins localize to the nuclei of N. benthamiana leaf cells (Goodin et al., 2002; Tsai et al., 2005).

1.5.4. The G protein

The plant rhabdovirus G protein is glycosylated and forms spikes on the surface of rhabdovirus virions (Jackson et al., 2005; van Beek et al., 1985). The G protein has a predicted N-terminal signal peptide and endopeptidase cleavage site, glycosylation sites,

and a transmembrane anchor domain, consistent with its role as viral membrane glycoprotein. Immunofluorescence localization studies indicate that the SYNV G protein localizes at the periphery of the nucleus (Martins et al., 1998).

1.5.5. The L protein

The plant rhabdovirus L protein carries conserved motifs characteristic of RNA dependent RNA polymerases, e.g. a catalytic domain, a RNA template-binding site and a metal-binding motif (Choi et al., 1992; Huang et al., 2003). RNA polymerase activity is detected in the nuclei of SYNV-infected plant cells (Choi et al., 1992; Wagner and

Jackson, 1997; Wagner et al., 1996). For vertebrate-infecting rhabdoviruses, it has been demonstrated that the L protein is associated with mRNA 5’ capping, 3’ polyadenylation, and protein kinase activities (Rose and Whitt, 2001; Walker et al., 2000). The L protein forms a complex with the viral N and P proteins that is important for virus genome replication and gene expression (Rose and Whitt, 2001; Wagner and Jackson, 1997). The

L proteins of cytorhabdoviruses are present as active forms in virions (Francki and

Randles, 1973 Francki and Randles 1972), but nucleorhabdovirus L proteins require activation by host factors early in infection (Wagner and Jackson, 1997; Wagner et al.,

1996). The L proteins of MMV, SYNV, RYSV, TaVCV have NLS motifs and are predicted to localize to the nuclei of plant cells by PSORT analysis (Table 1.2). NLS motifs are also present in the L proteins of NCMV and LNYV. It is not surprising because

NLS motifs are also found in the L proteins of BEFV and novirhabdoviruses, which

replicate in the cytoplasm and undergo morphogenesis in the endoplasmic reticulum

(Redinbaugh and Hogenhout, 2005).

1.5.6. Ancillary proteins

Genes located between the P and M genes are found in all plant rhabdoviruses sequenced to date, but not in vertebrate-infecting rhabdoviruses. These genes are most

likely having a role in cell-to-cell and/or systemic movement in plants rather than a role

in vector transmission, because VSIV is capable of replicating in its vertebrate and insect

hosts with just the five core genes (Redinbaugh and Hogenhout, 2005). The secondary

structure prediction of the MFSV P4, MMV P3, RYSV P3, and SYNV sc4 proteins

shows relatedness of these proteins with the 30K superfamily of plant virus movement

proteins (Huang et al., 2005; Melcher, 2000). Furthermore, BLASTP database search and

secondary structure prediction of the cytorhabdovirus LNYV 4b protein reveal a close

similarity to the movement proteins of capillo- and trichoviruses (Family Flexiviridae) that belong to a different group of movement proteins in the 30K superfamily of plant virus movement proteins (Dietzgen et al., 2006). The computer-based predictions were confirmed by complementation and biochemical experiments showing that the RYSV P3 protein functions as a viral cell-to-cell movement protein (Huang et al., 2005). The association of the SYNV sc4 protein with cellular membranes and virion also suggests a role in cell-to-cell movement (Goodin et al., 2002; Scholthof et al., 1994). NCMV, RYSV, and Strawberry crinkle virus (SCV) have genes located between the G and L genes

(Jackson et al., 2005). The RYSV P6 protein is detected in total protein extract of purified virions and viruliferous vector insects by immunoblot analyses, but not in extract of infected rice tissues (Huang et al., 2003). The results suggested that the RYSV P6 protein is associated with virion and may play a role in vector transmission.

1.6. Virus-insect interactions

1.6.1. Insect transmission of rhabdoviruses

Plant rhabdoviruses are transmitted by aphid, leafhopper or planthopper vectors

(Table 1.1), and the viruses replicate in both their insect and plant hosts (Hogenhout et al.,

2003; Jackson et al., 2005). Insects also transmit many vertebrate-infecting rhabdoviruses.

VSIV, Vesicular stomatitis New Jersey virus (VSNJV), and BEFV, which are all

economically important livestock pathogens, are transmitted to their vertebrate hosts by

black flies, sand flies, midges, and mosquitoes (Table 1.1) (Rose and Whitt, 2001). Nault

(1997) proposed that plant rhabdoviruses evolved from an ancestral insect virus and

secondarily adapted to plants for three reasons: 1) Insects appear to be the better-adapted

hosts for plant rhabdoviruses, because no detrimental cytopathological changes have been reported in the rhabdovirus-infected vector insects; 2) Many of these viruses can be

transmitted from virus-infected females to their progeny by transovarial transmission

without the involvement of a plant host; 3) Seed or pollen transmission of plant

rhabdoviruses has not yet been described, and therefore virus transmission is likely to be

dependent on insect acquisition and inoculation for spread among plants.

1.6.2. Persistent propagative transmission and barriers to virus transmission

Insects transmit rhabdoviruses from plant to plant in a persistent propagative manner, i.e. the viruses replicate within the vector insects, and the insects transmit the viruses for the remainder of their lifetimes (Ammar and Nault, 2002; Nault, 1997). For transmission to occur, viruses must first infect the midgut and other organs in order to reach the salivary glands from where to be introduced into plants along with the saliva while the insect feeds (Fig. 1.3). Rhabdoviruses cross at least four cellular membranes in this process. These are: 1) the apical membrane of the microvilli when rhabdoviruses enter the gut lumen upon ingestion of sap from infected plants; 2) the basement membrane at the haemocoel site of the midgut when the viruses enter the hemolymph or other tissues;

3) the basal membrane of the salivary glands; and 4) the apical membrane of the salivary glands when the viruses exit the salivary gland and move with the salivary secretions.

Even though it is generally thought that rhabdovirus particles enter the salivary

glands from the hemolymph, it is perhaps more likely that the salivary glands are infected

via nerve or tracheal cells (Hogenhout et al., 2003). This hypothesis is supported by

recent studies reporting the route of temporal and spatial spread of mammal- and plant-

infecting rhabdoviruses in their vector insects (Drolet et al., 2005; Ammar and Hogenhout,

submitted). The two studies show very similar infection patterns of VSNJV and MMV in

their respective vector insects. VSNJV first accumulates in foregut and midgut cells,

followed by the nerve cells, and subsequently the virus is detected in the hemolymph,

salivary glands, ovaries, and other tissues of the midge vectors Culicoides sonorensis

(Drolet et al., 2005). Similarly, MMV initially accumulates in epithelial cells of insect

midgut and anterior diverticulum, subsequently in tracheae and nerve cells, and then the virus is detected in hemocytes, muscles, and salivary glands and other tissues in the planthopper vector P. maidis (Ammar and Hogenhout, submitted). According to the progress of virus infection and how nerve and trachea connect various insect tissues, rhabdoviruses probably enter insect midgut cells first, and then spread throughout other tissues, including the salivary glands, via the nervous system, trachea, and at a later stage, the hemolymph.

Insect guts are probably important barriers for virus infection that determine the transmission efficiency of plant rhabdoviruses as has been demonstrated in studies with two serologically related rhabdoviruses, MMV and Maize Iranian mosaic virus (MIMV).

MIMV is naturally transmitted by the planthopper Ribautodelphax notabilis, but is transmitted very inefficiently by the MMV vector P. maidis when the virus is acquired orally from MIMV-infected maize (Izadpanah et al., 1983). In contrast, P. maidis transmits MIMV at high efficiency when MIMV is needle-injected into the haemocoel of

P. maidis (Nault and Ammar, 1989). In addition to the midgut barriers, there are probably also other barriers that obstruct rhabdovirus transmission by vector insects. In a laboratory population, only 20% of P. maidis transmits MMV, even though about 40% of

P. maidis is infected with MMV (Ammar and Hogenhout, submitted). In the P. maidis individuals that are infected but do not transmit MMV, MMV either accumulates only in the midgut, or accumulates in several tissues except the salivary glands, suggesting the presence of a midgut-escape and salivary gland infection barriers for MMV invasion in

P. maidis (Ammar and Hogenhout, submitted). Furthermore, it also suggests a midgut

infection barrier for 60% of the P. maidis population that is not MINV-infected at all.

Similar barriers were also reported for aphid-transmitted rhabdoviruses (Sylvester

and Richardson, 1992). Virus injections increase the transmission efficiencies for some

aphid-transmitted rhabdoviruses (Sylvester and Richardson, 1992). However, in other

cases, virus injections may not result in successful transmission by virus-injected insects.

The aphid Hyperomyzus lactucae naturally transmits Sowthistle yellow vein virus

(SYVV), but the aphid Macrosiphum euphorbiae remains an inefficient vector of SYVV even if the virus is injected into the haemocoel of M. euphorbiae (Behncken, 1973).

Electron microscopy reveals SYVV accumulation in various insect tissues, except the salivary glands (Behncken, 1973). The result suggested that the salivary gland basal membranes are barriers for SYVV transmission.

At the subcellular level of virus infection, plant rhabdoviruses are thought to interact

with the cellular receptors exposed to the surface of host cells (see also section 1.6.3) and

to enter cells by pH-dependent receptor-mediated endocytosis in a manner similar to

vertebrate-infecting VSIV and RABV (Lewis and Lentz, 1998; Superti et al., 1987). Thus,

the inability of rhabdoviruses to infect midgut and salivary gland cells in nonvector

insects may be caused by the inability of viruses to bind or activate cellular receptors.

Similarly, the inability of the rhabdoviruses to escape the midgut may be due to the

inability of viruses to recognize receptors of nerve or tracheal cells that allow dispersal of

viruses from gut tissues. However, other possibilities should be taken into account as

well.

1.6.3. Cellular receptor hypothesis of vector specificity

Insect transmission of rhabdoviruses is highly specific; a given virus is transmitted by one or a few insect species. For example, MFSV is specifically transmitted by the leafhopper G. nigrifrons, and MMV by the planthopper P. maidis (Falk and Tsai, 1985;

Redinbaugh et al., 2002). Two strains of Potato yellow dwarf virus (PYDV) are also

transmitted by different species of leafhoppers; one strain is specifically transmitted by

Agallia constricta, and the other strain by Aceratagallia sanguinolenta (Black, 1941). It

has been hypothesized that a rhabdovirus binds specific cellular receptor(s) in its

respective vector insect, allowing successfully transmission of a rhabdovirus by a vector

insect. Recent discovery showed that MFSV replicates in, but is not transmitted by, the

MMV vector P. maidis and other maize-feeding leafhoppers (Todd, Redinbaugh, and

Hogenhout, unpublished data). This result suggested that the nonvector P. maidis may

have cellular receptor(s) for MFSV in their midguts but not in their salivary glands.

Similarly, salivary gland receptor(s) of the nonvector P. maidis may allow transmission of

MIMV when the virus injected into the haemocoel of P. maidis.

Although rhabdovirus receptors have not been identified in insects, putative

receptors have been reported in vertebrate cells. The acetylcholine receptor, neural cell

adhesion molecule, and the low-affinity nerve growth receptor p75NTR serve as receptors

for the vertebrate-infecting rhabdoviruses (Lentz et al., 1984; Thoulouze et al., 1998;

Tuffereau et al., 1998). Considering that receptors of nerve cells are often conserved among vertebrate and invertebrate animals, it would be interesting to determine whether rhabdoviruses also bind analogous proteins in insect vectors.

In vertebrates, the rhabdovirus G protein is essential for interaction with host cell receptors (Langevin et al., 2002). The recombinant viruses in which the RABV G protein is replaced with the Human immunodeficiency virus type I (HIV-1; Family Retroviridae) envelope protein enter cells in a pH-independent manner as HIV-1, and mimic an

HIV-1-like cell tropism. Similarly, when the VSIV G protein is replaced by the G protein of Ebola or Marburg viruses (Family Filoviridae), the cell tropism and virus-induced immunity of these recombinant viruses become similar to those of Ebola or Marburg viruses (Garbutt et al., 2004). The G protein of plant rhabdoviruses is probably involved in determining insect vector specificity. Two strains of PYDV are specifically transmitted by different species of leafhoppers (Black, 1941). Adam and Hsu (1984) reported that the structure difference of the G proteins of PYDV strains might be recognized by different receptors in their respective leafhopper vectors. These results suggested that the G protein is important for the virus uptake and virus specific immunity for vertebrate- and plant-infecting rhabdoviruses.

1.6.4. Other factors affecting vector competence

Vector competence is probably not only determined by cellular receptors of insect organs, but also determined by other factors, for example, the ability of the viruses to replicate in insect cells, to escape from the insect immune system, to prevent or delay apoptosis of host cells, and to prevent degradation in the protease-rich environment of the insect digestive fluids and saliva. Sigma virus is a drosophila-infecting rhabdovirus and is transmitted to the progeny of flies through female and male germinal cells (Teninges,

1999). There are some examples that rhabdoviruses can modulate insect immune responses. Recent microarray experiments indicated that the transcriptional response of

Drosophila melanogaster to Sigma virus infection is much weaker compared to those of

two other drosophila-infecting viruses, Drosophila C virus (DCV; Family Dicistroviridae)

and Drosophila X virus (DXV; Family Birnaviridae) (Chapter 5 of this dissertation).

DCV and DXV usually induce cellular lysis and increase the mortality of drosophila

(Gravot et al., 2000; Teninges et al., 1979; Zambon et al., 2005). A Sigma virus strain that

generally has low titers when it infects drosophila is transmitted at a higher efficiency to

drosophila progeny than another Sigma virus strain that has higher titers (Seecof, 1966).

Further, the low titer strain has lower death rates of drosophila progeny relatively to the

high titer strain (Seecof, 1966). To increase the chances of successful passage to next

generation flies, Sigma virus probably evolved to become less virulent to drosophila

possibly through avoidance of a strong antiviral defense response in drosophila.

The interaction of Potato leafroll virus (PLRV; Family Luteoviridae) with their

aphid vector is another example of a virus trying to escape from the immune defense

responses of its insect host. The aphid Myzus persicae naturally transmits PLRV. PLRV

selectively binds to a homologue of GroEL in virus-infected M. persicae (Hogenhout et

al., 1996). GroEL is abundantly synthesized by Buchnera sp., the primary endosymbiotic

bacteria of M. persicae. The Buchnera GroEL protects PLRV particles against proteolytic

breakdown in aphid hemolymph by avoiding the aphid’s immune system (van den Heuvel

et al., 1994; van den Heuvel et al., 1997).

1.7. Virus-plant interactions

1.7.1. Host range of plant rhabdoviruses

Maize (Zea mays) is the common plant host of MFSV and MMV. In addition to

maize, MFSV also infects barley (Hordeum vulgare), wheat (Triticum aestivum), oat

(Avena sativa), giant foxtail (Seteria faberi), and rye (Secale cereale), whereas MMV infects sorghum (Sorghum bicolor), itchgrass (Rottboellia exaltata), and Setaria vulpiseta

(Redinbaugh et al. 2002; Brunt et al., 1996). The different host range of MFSV and

MMV may be due to the feeding preferences of their respective vector insects, because plant-infecting rhabdoviruses are highly dependent on their vector insects for their transmission from plant to plant (Hogenhout et al., 2003). Plant rhabdoviruses may have a greater plant host range compared to the feeding range of their vector insects (Todd,

Redinbaugh and Hogenhout, unpublished data). Plant host range is also determined by the ability of the viruses to replicate and systemically move in plant hosts.

MFSV and MMV virions accumulate in all cell types of the host plants, including

mesophyll, epidermis, vascular cells, etc. (Bradfute and Tsai, 1983; McDaniel et al., 1985;

Redinbaugh et al., 2002). MFSV symptoms on maize include dwarfing and fine chlorotic

streaks along intermediate and small veins (Redinbaugh et al., 2002). MMV symptoms

on maize include dwarfing and long, even, distinct chlorotic streaks on large or small

veins (Bradfute and Tsai, 1983).

In the cellular level, membrane proliferation around the nucleus has been noted in

the nucleorhabdovirus-infected plant cell (Goodin et al., 2005). SYNV and PYDV

infections enlarge the nuclei of infected cells but the nucleus enlargement does not occur

in other cytoplasmic virus infections, for example Impatients necrotic spot virus (INSV;

Family Bunyaviridae) (Goodin et al., 2005). Goodin et al. (2005) suggested that the cytoplasm membranes are redirected to the nuclei of SYNV and PYDV infected cells.

This membrane redirection is associated with the assembly of new virions.

1.7.2. Molecular response of host plant to rhabdovirus infection

Genetically distinct viruses elicit unique changes of gene expression in virus-

infected plant hosts. Using microarray analysis, two genetically distinct viruses, SYNV and INSV, elicit unique changes in gene expression of their common host N.

benthamiana (Senthil et al., 2005). SYNV and INSV are distinct enveloped viruses within different family. SYNV is a rhabdovirus with a single segmented, negative-sensed

RNA genome, and the virus replicates and assembles in the nucleus of infected plant cell.

INSV is a tospovirus (Family Bunyaviridae) and has a tripartite negative-sensed RNA

genome, and this virus replicates and assembles in the cytoplasm of infected plant cells.

Throughout the infection process, more genes at greater fold changes were up- and

downregulated in INSV-infected plants than SYNV-infected plants (Goodin et al., 2005).

1.8. Research objectives

The research objectives of my Ph.D. studies are to: 1) Complete the genome

sequence of MFSV and MMV; 2) Determine the cellular localization of the MFSV and

MMV proteins in plant and insect cells; 3) Examine whether the nuclear import of MFSV

and MMV proteins is dependent on Importin αs in plant cells; 4) Investigate the

drosophila response to Sigma virus infection using cDNA microarray analysis.

This dissertation consists of five chapters, including this literature review. Chapter

1 is this literature review, introducing the research background and current status of the

studies related to research described in this dissertation and is titled “Maize fine streak

virus and Maize mosaic virus: a good system to study the interactions among rhabdovirus,

plant, and insect vector” Chapter 2 “Complete genome sequence and in planta

subcellular localization of Maize fine streak virus proteins” was published in the Journal

of Virology (Tsai et al., 2005). This chapter describes the completion of the MFSV

genome sequence and subsequent functional analysis of viral proteins. The cellular

localization of MFSV proteins and viral protein interactions are also investigated in this

chapter. Chapter 3 “Shotgun sequencing of the negative-sense RNA genome of the

rhabdovirus Maize mosaic virus”, was published in the Journal of Virological Methods

(Reed et al., 2005). It describes the random shotgun approach method for sequencing the

MMV genome to near completion. It also compares time and cost requirements for shotgun versus primer walking sequencing procedures. Chapter 4 “Subcellular localization and nuclear import of Maize fine streak virus and Maize mosaic virus proteins” describes the cellular localization of MFSV proteins in insect cells and that of

MMV proteins in plant cells. In addition, it demonstrates that the nuclear import of

MFSV proteins is dependent on Importin αs in plant cells. Chapter 5 “Sigma rhabdovirus activates the innate immune response of drosophila” reports on the drosophila innate immune response to Sigma rhabdovirus infection and compares this to other antiviral responses.

1.9. References

Adam, G. and Hsu, H.T. (1984) Comparison of structural proteins from two potato yellow dwarf viruses. J. Gen. Virol., 65, 991-994.

Ammar, E.D. and Nault, L.R. (2002) Virus transmission by leafhoppers, planthoppers and treehoppers (Auchenorrhyncha, Homoptera). Adv. Bot. Res., 36, 141-167.

Barr, J.N., Whelan, S.P. and Wertz, G.W. (2002) Transcriptional control of the RNA-dependent RNA polymerase of vesicular stomatitis virus. Biochim. Biophys. Acta, 1577, 337-353.

Behncken, G.M. (1973) Evidence of multiplication of sowthistle yellow vein virus in an inefficient aphid vector, Macrosiphum euphorbiae. Virology, 53, 405-412.

Black, L.M. (1941) Specific transmission of varieties of potato yellow-dwarf virus by related insects. Am. potato J., 18, 231-233.

Bradfute, O.E. and Tsai, J.H. (1983) Identification of maize mosaic virus in Florida. Plant Dis., 67, 1339-l342.

Brun, G. (1991) Rhabdoviridae. In Adams, J.R. and Bonami, J.R. (eds.), Atlas of invertebrate viruses. CRC Press, Boca Raton, pp. 443-460.

Brunt, A., Crabtree, K., Dallwitz, M., Gibbs, A. and Watson, L. (1996) Viruses of plants: descriptions and lists from the VIDE database. 1484 pp.

Choi, T.J., Kuwata, S., Koonin, E.V., Heaton, L.A. and Jackson, A.O. (1992) Structure of the L (polymerase) protein gene of sonchus yellow net virus. Virology, 189, 31-39.

Dietzgen, R.G., Callaghan, B., Wetzel, T. and Dale, J.L. (2006) Completion of the genome sequence of Lettuce necrotic yellows virus, type species of the genus Cytorhabdovirus. Virus Res., 118, 16-22.

Drolet, B.S., Campbell, C.L., Stuart, M.A. and Wilson, W.C. (2005) Vector competence of Culicoides sonorensis (Diptera: Ceratopogonidae) for vesicular stomatitis virus. J. Med. Entomol., 42, 409-418.

Falk, B.W. and Tsai, J.H. (1985) Serological detection and evidence for multiplication of maize mosaic virus in the planthopper, Peregrinus maidis. Phytopathology, 75, 852-855.

Falk, B.W. and Tsai, J.H. (1983) Physicochemical characterization of maize mosaic virus. Phytopathology, 73, 1536-1539.

Foley, H.D., Otero, M., Orenstein, J.M., Pomerantz, R.J. and Schnell, M.J. (2002) Rhabdovirus-based vectors with human immunodeficiency virus type 1 (HIV-1) envelopes display HIV-1-like tropism and target human dendritic cells. J. Virol., 76, 19-31.

Francki, R.I. and Randles, J.W. (1973) Some properties of lettuce necrotic yellows virus RNA and its in vitro transcription by virion-associated transcriptase. Virology, 54, 359-368.

Garbutt, M., Liebscher, R., Wahl-Jensen, V., Jones, S., Moller, P., Wagner, R., Volchkov, V., Klenk, H.D., Feldmann, H. and Stroher, U. (2004) Properties of replication-competent vesicular stomatitis virus vectors expressing glycoproteins of filoviruses and arenaviruses. J. Virol., 78, 5458-5465.

Goodin, M., Yelton, S., Ghosh, D., Mathews, S. and Lesnaw, J. (2005) Live-cell imaging of rhabdovirus-induced morphological changes in plant nuclear membranes. Mol. Plant Microbe Interact., 18, 703-709.

Goodin, M.M., Austin, J., Tobias, R., Fujita, M., Morales, C. and Jackson, A.O. (2001) Interactions and nuclear import of the N and P proteins of sonchus yellow net virus, a plant nucleorhabdovirus. J. Virol., 75, 9393-9406.

Goodin, M.M., Dietzgen, R.G., Schichnes, D., Ruzin, S. and Jackson, A.O. (2002) pGD vectors: versatile tools for the expression of green and red fluorescent protein fusions in agroinfiltrated plant leaves. Plant J., 31, 375-383.

Heaton, L.A., Hillman, B.I., Hunter, B.G., Zuidema, D. and Jackson, A.O. (1989) Physical map of the genome of sonchus yellow net virus, a plant rhabdovirus with six genes and conserved gene junction sequences. Proc. Natl. Acad. Sci. U.S.A., 86, 8665-8668.

Hogenhout, S.A., Verbeek, M., Hans, F., Houterman, P.M., Fortass, M., van der Wilk, F., Huttinga, H. and van den Heuvel, J.F. (1996) Molecular basis of the interactions between luteoviruses and aphids. Agronomie, 16, 167-173.

Hogenhout, S.A., Redinbaugh, M.G. and Ammar, E.D. (2003) Plant and animal rhabdovirus host range: a bug's view. Trends Microbiol., 11, 264-271.

Huang, Y., Zhao, H., Luo, Z., Chen, X. and Fang, R.X. (2003) Novel structure of the genome of Rice yellow stunt virus: identification of the gene 6-encoded virion protein. J. Gen. Virol., 84, 2259-2264.

Huang, Y.W., Geng, Y.F., Ying, X.B., Chen, X.Y. and Fang, R.X. (2005) Identification of a movement protein of rice yellow stunt rhabdovirus. J. Virol., 79, 2108-2114.

Izadpanah, K., Ahmadi, A.A., Parvin, S. and Jafari, S.A. (1983) Transmission, particle size and additional hosts of the rhabdovirus causing maize mosaic in Shiraz, Iran. Phytopathologische Zeitschrift, 107, 283-288.

Jackson, A.O., Dietzgen, R.G., Goodin, M.M., Bragg, J.N. and Deng, M. (2005) Biology of plant rhabdoviruses. Annu. Rev. Phytopathol., 43, 623-660.

Kurath, G., Higman, K.H. and Bjorklund, H.V. (1997) Distribution and variation of NV genes in fish rhabdoviruses. J. Gen. Virol., 78, 113-117.

Langevin, C., Jaaro, H., Bressanelli, S., Fainzilber, M. and Tuffereau, C. (2002) Rabies virus glycoprotein (RVG) is a trimeric ligand for the N-terminal cysteine-rich domain of the mammalian p75 neurotrophin receptor. J. Biol. Chem., 277, 37655-37662.

Lentz, T.L., Wilson, P.T., Hawrot, E. and Speicher, D.W. (1984) Amino acid sequence similarity between rabies virus glycoprotein and snake venom curaremimetic neurotoxins. Science, 226, 847-848.

Lewis, P. and Lentz, T.L. (1998) Rabies virus entry into cultured rat hippocampal neurons. J. Neurocytol., 27, 559-573.

Martins, C.R., Johnson, J.A., Lawrence, D.M., Choi, T.J., Pisi, A.M., Tobin, S.L., Lapidus, D., Wagner, J.D., Ruzin, S., McDonald, K. and Jackson, A.O. (1998) Sonchus yellow net rhabdovirus nuclear viroplasms contain polymerase-associated proteins. J. Virol., 72, 5669-5679.

McDaniel, L.L., Ammar, E.D. and Gordon, D.T. (1985) Assembly, morphology, and accumulation of a Hawaiian isolate of maize mosaic virus in maize. Phytopathology, 75, 1167-1172.

McWilliam, S.M., Kongsuwan, K., Cowley, J.A., Byrne, K.A. and Walker, P.J. (1997) Genome organization and transcription strategy in the complex GNS-L intergenic region of bovine ephemeral fever rhabdovirus. J. Gen. Virol., 78, 1309-1317.

Melcher, U. (2000) The '30K' superfamily of viral movement proteins. J. Gen. Virol., 81, 257-266.

Nault, L.R. (1997) Arthropod transmission of plant viruses: a new synthesis. Ann. Entomol. Soc. Am., 90, 521-541.

Nault, L.R. and Ammar, E.D. (1989) Leafhopper and planthopper transmission of plant viruses. Annu. Rev. Entomol., 503-529.

Randles, J.W. and Francki, R.I. (1972) Infectious nucleocapsid particles of lettuce necrotic yellows virus with RNA-dependent RNA polymerase activity. Virology, 50, 297-300. 27

Redinbaugh, M.G. and Hogenhout, S.A. (2005) Plant rhabdoviruses. Curr. Top. Microbiol. Immunol., 292, 143-163.

Redinbaugh, M.G., Seifers, D.L., Meulia, T., Abt, J.J., Anderson, R.J., Styer, W.E., Ackerman, J., Salomon, R., Houghton, W., Creamer, R., Gordon, D.T. and Hogenhout, S.A. (2002) Maize fine streak virus, a new leafhopper-transmitted rhabdovirus. Phytopathology, 92, 1167-1174.

Reed, S.E., Tsai, C.W., Willie, K.J., Redinbaugh, M.G. and Hogenhout, S.A. (2005) Shotgun sequencing of the negative-sense RNA genome of the rhabdovirus Maize mosaic virus. J. Virol. Methods, 129, 91-96.

Revill, P., Trinh, X., Dale, J. and Harding, R. (2005) Taro vein chlorosis virus: characterization and variability of a new nucleorhabdovirus. J. Gen. Virol., 86, 491-499.

Rose, J.K. and Whitt, M.A. (2001) Rhabdoviridae: the virus and their replication. In Knipe, D.M. and Howley, P.M. (eds.), Fields virology. Lippincott Williams & Wilkins, Philadelphia, pp. 1221-1244.

Scholthof, K.B., Hillman, B.I., Modrell, B., Heaton, L.A. and Jackson, A.O. (1994) Characterization and detection of sc4: a sixth gene encoded by sonchus yellow net virus. Virology, 204, 279-288.

Seecof, R.L. (1966) Low titers in Drosophila that regularly transmit the sigma virus infection. Genetics, 53, 79-83.

Senthil, G., Liu, H., Puram, V.G., Clark, A., Stromberg, A. and Goodin, M.M. (2005) Specific and common changes in Nicotiana benthamiana gene expression in response to infection by enveloped viruses. J. Gen. Virol., 86, 2615-2625.

Superti, F., Seganti, L., Ruggeri, F.M., Tinari, A., Donelli, G. and Orsi, N. (1987) Entry pathway of vesicular stomatitis virus into different host cells. J. Gen. Virol., 68, 387-399.

Sylvester, E.S. and Richardson, J. (1992) Aphid-borne rhabdoviruses - relationship with their vectors. In Harris, K.F. (ed.), Advances in disease vector research. Springer-Verlag, New York, pp. 313-341.

Tanno, F., Nakatsu, A., Toriyama, S. and Kojima, M. (2000) Complete nucleotide sequence of Northern cereal mosaic virus and its genome organization. Arch. Virol., 145, 1373-1384.

Teninges, D. (1999) Sigma rhabdoviruses (Rhabdoviridae). In Granoff, A. and Webster, R.G. (eds.), Encyclopedia of virology. Academic Press, San Diego, pp. 1635-1639.

Tesh, R.B., Chaniotis, B.N. and Johnson, K.M. (1972) Vesicular stomatitis virus (Indiana serotype): transovarial transmission by phlebotomine sandflies. Science, 175, 1477-1479.

Thoulouze, M.I., Lafage, M., Schachner, M., Hartmann, U., Cremer, H. and Lafon, M. (1998) The neural cell adhesion molecule is a receptor for rabies virus. J. Virol., 72, 7181-7190.

Tordo, N., Poch, O., Ermine, A., Keith, G. and Rougeon, F. (1986) Walking along the rabies genome: is the large G-L intergenic region a remnant gene? Proc. Natl. Acad. Sci. U.S.A., 83, 3914-3918.

Tsai, C.W., Redinbaugh, M.G., Willie, K.J., Reed, S., Goodin, M. and Hogenhout, S.A. (2005) Complete genome sequence and in planta subcellular localization of Maize fine streak virus proteins. J. Virol., 79, 5304-5314.

Tuffereau, C., Benejean, J., Blondel, D., Kieffer, B. and Flamand, A. (1998) Low-affinity nerve-growth factor receptor (P75NTR) can serve as a receptor for rabies virus. EMBO J., 17, 7250-7259. van den Heuvel, J.F., Bruyere, A., Hogenhout, S.A., Ziegler-Graff, V., Brault, V., Verbeek, M., van der Wilk, F. and Richards, K. (1997) The N-terminal region of the luteovirus readthrough domain determines virus binding to Buchnera GroEL and is essential for virus persistence in the aphid. J. Virol., 71, 7258-7265. van den Heuvel, J.F., Verbeek, M. and van der Wilk, F. (1994) Endosymbiotic bacteria associated with circulative transmission of potato leafroll virus by Myzus persicae. J. Gen. Virol., 75, 2559-2565.

Wagner, J.D., Choi, T.J. and Jackson, A.O. (1996) Extraction of nuclei from sonchus yellow net rhabdovirus-infected plants yields a polymerase that synthesizes viral mRNAs and polyadenylated plus-strand leader RNA. J. Virol., 70, 468-477.

Wagner, J.D. and Jackson, A.O. (1997) Characterization of the components and activity of Sonchus yellow net rhabdovirus polymerase. J. Virol., 71, 2371-2382.

Walker, P.J., Benmansour, A., Dietzgen, R., Fang, R.X., Jackson, A.O., Kurath, G., Leong, J.C., Nadin-Davies, S., Tesh, R.B. and Tordo, N. (2000) Family Rhabdoviridae. In van Regenmortel, M.H.V., Fauquet, C.M., Bishop, D.H.L., Carstens, E.B., Estes, M.K., Lemon, S.M., Maniloff, J., Mayo, M.A., McGeoch, D.J., Pringle, C.R. and Wickner, R.B. (eds.), Virus taxonomy: classification and nomenclature of viruses. Seventh report of the International Committee on Taxonomy of Viruses. Academic Press, San Diego, pp. 563-583.

Wetzel, T., Dietzgen, R.G. and Dale, J.L. (1994) Genomic organization of lettuce necrotic yellows rhabdovirus. Virology, 200, 401-412.

Whelan, S.P., Barr, J.N. and Wertz, G.W. (2004) Transcription and replication of nonsegmented negative-strand RNA viruses. Curr. Top. Microbiol. Immunol., 283, 61-119.

Genus Type species Hosts Vectors Vesiculovirus Vesicular stomatitis Indiana Mammals, fish Mosquitoes, midges, virus black flies, sand flies

Ephemerovirus Bovine ephemeral fever virus Mammals Mosquitoes

Lyssavirus Rabies virus Humans, mammals None

Nucleorhabdovirus Potato yellow dwarf virus Higher plants Planthoppers, leafhoppers, aphids

Cytorhabdovirus Lettuce necrotic yellows virus Higher plants Planthoppers, leafhoppers, aphids

Novirhabdovirus Infectious hematopoietic Fish None necrosis virus

Table 1.1. Six genera of the Family Rhabdoviridae

Protein SYNV MFSV RYSV MMV TaVCV NCMV LNYV

N + + + + + - -

P - - + + - - -

sc4/P3/4b ------

P4 + -

M + + - - - - -

G + - - + - - -

L + - + + + + +

Virus acronyms: SYNV, Sonchus yellow net virus; MFSV, Maize fine streak virus; RYSV, Rice yellow stunt virus; MMV, Maize mosaic virus; TaVCV, Taro vein chlorosis virus; NCMV, Northern cereal mosaic virus; LNYV, Lettuce necrotic yellows virus.

Abbreviations: +, present; -, absent; N, nucleocapsid protein; P, phosphoprotein; sc4, SYNV sc4 protein; P3, ORF3 protein; 4b, LNYV 4b protein; P4, ORF4 protein; M, matrix protein; G, glycoprotein; L, polymerase.

Table 1.2. Predicted nuclear localization signal of plant rhabdovirus proteins with PSORT

Fig. 1.1. A. Transmission electron micrograph of Maize mosaic virus (MMV) infected maize leave. Note viral particles (V) accumulating in the perinuclear space of parenchyma cells. The inset shows a purified MMV stained with uranyl acetate. (Reproduced from Bradfute and Tsai, 1983; Falk and Tsai, 1983). B. Transmission electron micrograph of Maize fine streak virus (MFSV) infected maize leave. The arrows indicate viral particles budding through the nuclear envelope and accumulating in the perinuclear space of a parenchyma cell. The inset shows a purified MFSV negatively stained with phophotungstic acid. Abbreviations: Nu, nucleus; Cy, cytoplasm; Cl, chloroplast. (Reproduced from Redinbaugh et al. 2002).

Fig. 1.2. A. Schematic diagram of the MMV and MMV genomes. The completed MFSV genome consists of 13,782 nt. The MFSV genome has seven open reading frames (ORFs) on the antigenomic strand. The MMV genome consists of 12,133 nt, and the sequences of the 3’ and 5’ ends were not determined. The MMV genome has six ORFs on the antigenomic strand. B. Representations of the minimal rhabdovirus genome (above) and virion composition (below) (Reproduced from Hogenhout et al, 2003). Abbreviations: N, nucleocapsid protein; P, phosphoprotein; 3, ORF3; 4, ORF4; M, matrix protein; G, glycoprotein; L, polymerase.

Fig. 1.3. Persistent propagative transmission of plant rhabdovirus by a leafhopper. Virus particles are acquired from virus-infected plant cells through the food canal inside the stylet and move from the gut lumen through the epithelial cell layer and visceral muscle cells into the hemolymph and/or nervous system. The viruses spread throughout the insect tissues and infect the salivary glands. At last the viruses are introduced into new plant hosts through the salivary canal during insect feeding. (Reproduced from Hogenhout et al. 2003).

CHAPTER 2

COMPLETE GENOME SEQUENCE AND IN PLANTA SUBCELLULAR

LOCALIZATION OF MAIZE FINE STREAK VIRUS PROTEINS

Chi-Wei Tsai1, Margaret G. Redinbaugh2,3, Kristen J. Willie3, Sharon Reed1,

Michael Goodin4, and Saskia A. Hogenhout1

1Department of Entomology, 2Department of Plant Pathology, Ohio Agricultural Research and Development Center (OARDC), The Ohio State University, Wooster, Ohio

3U.S. Department of Agriculture-Agricultural Research Service, Corn and Soybean

Research, Wooster, Ohio

4Department of Plant Pathology, University of Kentucky, Lexington, Kentucky

2.1. Abstract

The genome of the nucleorhabdovirus Maize fine streak virus (MFSV) consists of

13,782 nucleotides of nonsegmented, negative-sense, single-stranded RNA. The

antigenomic strand consisted of seven open reading frames (ORFs), and transcripts of all

ORFs were detected in infected plants. ORF1, ORF6, and ORF7 had significant

similarities to the nucleocapsid protein (N), glycoprotein (G), and polymerase (L) genes of other rhabdoviruses, respectively, whereas the ORF2, ORF3, ORF4, and ORF5 proteins had no significant similarities. The N (ORF1), ORF4, and ORF5 proteins localized to the nuclei, consistent with the presence of nuclear localization signals (NLSs)

in these proteins. ORF5 likely encodes the matrix protein (M), based on its size, the

position of its NLS, and the localization of fluorescent protein fusions to the nucleus.

ORF2 probably encodes the phosphoprotein (P) because, like the P protein of Sonchus

yellow net virus (SYNV), it was distributed throughout the cell when expressed alone but

was relocalized to a subnuclear locus when coexpressed with the MFSV N protein.

Unexpectedly, co-introduction of the MFSV N and P proteins, but not the orthologous

proteins of SYNV, resulted in accumulations of both proteins in the nucleolus. The N and

P protein relocalization was specific to cognate proteins of each virus. The subcellular

localizations of the MFSV ORF3 and ORF4 proteins were distinct from that of the SYNV

sc4 protein, suggesting different functions. To our knowledge, this is the first comparative

study of the cellular localization of plant rhabdoviral proteins. This study indicated that plant rhabdoviruses are diverse in genome sequence and viral protein interactions.

2.2. Introduction

Members of the family Rhabdoviridae have a broad host range, including humans,

livestock, plants, and insects. Six genera of rhabdoviruses have been described (Walker et

al., 2000). The four genera of animal-infecting rhabdoviruses (Vesiculovirus,

Ephemerovirus, Lyssavirus, and Novirhabdovirus) include the livestock pathogens

Vesicular stomatitis Indiana virus (VSIV), Vesicular stomatitis New Jersey virus

(VSNJV), and Bovine ephemeral fever virus; the human pathogen Rabies virus (RABV);

and the fish pathogen Infectious hematopoietic necrosis virus. The two genera of plant

rhabdoviruses are Cytorhabdovirus (type species, Lettuce necrotic yellows virus [LNYV])

and Nucleorhabdovirus (type species, Potato yellow dwarf virus). The viruses that cause

rabies and fish diseases appear to be confined to vertebrate hosts, whereas vesiculo-,

ephemero-, cyto-, and nucleorhabdoviruses are transmitted to their vertebrate or plant

hosts by insects (Hogenhout et al., 2003; Rose and Whitt, 2001). Plant rhabdoviruses are

particularly interesting because they are able to replicate and systemically spread in very

divergent hosts: plants and insects.

Generally, a rhabdovirus virion is composed of a lipid envelope derived from host

membranes and a ribonucleocapsid core consisting of a nonsegmented, negative-sense, single-stranded RNA bound to complexes of nucleocapsid protein (N), phosphoprotein

(P), and polymerase (L) (Rose and Whitt, 2001). The glycoprotein (G) protrudes from the

exterior of the lipid envelope, and the matrix protein (M) connects the envelope to the

ribonucleocapsid core (Rose and Whitt, 2001). VSIV and VSNJV have the simplest

genomes, encoding just the five structural proteins of the virion in the gene order

3’-N-P-M-G-L-5’, whereas the genomes of other rhabdoviruses harbor additional genes

(Heaton et al., 1989; Kurath et al., 1997; McWilliam et al., 1997; Tanno et al., 2000;

Wetzel et al., 1994).

Although >70 plant rhabdoviruses have been described, the genomes of only a few

plant rhabdoviruses have been sequenced to completion, including LNYV, Strawberry

crinkle virus (SCV), Northern cereal mosaic virus (NCMV), Rice yellow stunt virus

(RYSV), Taro vein chlorosis virus (TaVCV), and Sonchus yellow net virus (SYNV)

(Jackson et al., 2005). The nucleorhabdovirus SYNV is presently the most extensively

characterized plant-infecting rhabdovirus (Jackson et al., 2005). In planta subcellular

localization studies of fluorescent-protein fusions provided an indication of the function

and protein-protein interaction of viral proteins of SYNV (Goodin et al., 2002). The

SYNV N and M proteins both target the plant cell nuclei when expressed individually,

whereas the SYNV P and sc4 proteins do not (Goodin et al., 2002). However, the SYNV

P protein targets subnuclear locale when coexpressed with the SYNV N protein,

suggesting that the N and P proteins interact with each other in SYNV-infected plants

(Goodin et al., 2001; Goodin et al., 2002). The SYNV sc4 gene, located between the P

and M genes, encodes a membrane-associated protein that may be involved in viral

cell-to-cell movement in plants (Melcher, 2000; Scholthof et al., 1994).

Maize fine streak virus (MFSV) was first reported in maize fields in southwestern

Georgia in 1999 and was described as a new plant nucleorhabdovirus (Redinbaugh et al.,

2002). The symptoms caused by MFSV include dwarfing and fine chlorotic streaks along intermediate and small veins (Redinbaugh et al., 2002). MFSV is transmitted by the leafhopper Graminella nigrifrons and is not transmissible by rub inoculation of maize

39 leaves but can be mechanically transmitted by vascular puncture inoculation (VPI)

(Hogenhout et al., 2003; Redinbaugh et al., 2002). Like those of other rhabdoviruses, the

MFSV virion is a bacilliform particle measuring 231 by 71 nm with a lipid envelope, and its genome consists of a nonsegmented, negative-sense, single-stranded RNA

(Redinbaugh et al., 2002; Walker et al., 2000). Purified preparation of MFSV contains three abundant proteins corresponding to the major rhabdovirus structural proteins: the G protein (82 kDa), N protein (50 kDa), and M protein (32 kDa) (Redinbaugh et al., 2002).

To define MFSV genes and to begin to characterize their functions, we determined the complete genomic sequence of the virus and investigated virus gene expression in infected maize. We showed for the first time the localization of coinfiltrated rhabdoviral

N and P proteins in the nucleolus of plant cell and that the N and P proteins of MFSV and

SYNV specifically interact with each other and not with the orthologous proteins of another rhabdovirus. Further, we demonstrated that the MFSV ORF3 and ORF4 proteins had different subcellular localizations than the SYNV sc4 protein.

2.3. Materials and methods

2.3.1. Virus maintenance and purification

MFSV from Georgia was maintained in maize seedlings by serial inoculations with viruliferous insect vectors (G. nigrifrons) or by VPI (Louie, 1995; Redinbaugh et al.,

2002). The virus was purified from maize leaf laminar tissue collected 26 to 40 days after

VPI or vector inoculation, as previously described (Redinbaugh et al., 2002).

2.3.2. MFSV genomic RNA extraction and library construction

For initial cDNA library construction, virus genomic RNA was extracted using a

previously described protocol (Redinbaugh et al., 2002). For all other applications

(reverse transcription [RT]-PCR, 3’ rapid amplification of cDNA ends (RACE), 5’ RACE, and RNA ligase-mediated (RLM) RACE), genomic RNA was extracted from virus pellet

suspensions using a ToTALLY RNA isolation kit (Ambion, Austin, TX.) following the

manufacturer's instructions.

The cDNA synthesis of the MFSV genomic RNA using random hexamers was

carried out with the Superscript Choice system (Invitrogen Corp., Carlsbad, CA)

according to the manufacturer's instructions. The cDNAs were ligated into the

EcoRI-digested, phosphatase-treated pGEM4Z vector (Promega Corp., Madison, WI) or

the pZeroAmp vector (T. Meulia, unpublished results) and transformed into Escherichia

coli TOP10 cells (Invitrogen Corp.). Twelve clones (G2A, G2C, G3A, G3B, G4C, G5C,

G6A, G9B, Z6B, Z11B, Z12B, and Z15B) carrying inserts larger than 1 kb that

hybridized with viral RNA were selected for sequence analysis.

Regions of the MFSV genomic RNA not represented by clones were amplified using

RT-PCR with primers flanking the missing sequence. RT-PCR was carried out using a

Platinum RT-PCR kit (Invitrogen Corp.). RT-PCR products for the 3’ end of the MFSV genome (GAU3’) and MFSV4 (Fig. 2.1) were ligated into the pCR-Blunt II-TOPO vector

(Invitrogen Corp.) for sequencing. Three other RT-PCR products, MFSV5, MFSV6, and

MFSV7 (Fig. 2.1) were sequenced directly, without prior cloning.

2.3.3. MFSV genome sequencing and sequence analysis

Sequencing was carried out in 550- to 700-bp segments by primer walking using a

3700 DNA Sequence Analyzer and BigDye Terminator Cycle Sequencing chemistry

(Applied Biosystems, Inc., Foster City, CA) according to the supplier's instructions. Base calling was done with MacPHRED-MacPHRAP (CodonCode Corp., Dedham, MA), and sequences were assembled by Sequencher (Gene Codes Corp., Ann Arbor, MI).

ORFs were identified with MacVector (Accelrys, San Diego, CA). Putative MFSV protein sequences were compared to the National Center for Biotechnology Information

(NCBI) GenBank database by BLASTP search to identify sequence similarity to other known proteins. Protein sequences were searched for domains and motifs, including transmembrane domains (TMHMM version 2.0 [http://www.cbs.dtu.dk/services/

TMHMM/]) (Sonnhammer et al., 1998), N-terminal signal peptides (SignalP version 3.0

[http://www.cbs.dtu.dk/services/SignalP/]) (Bendtsen et al., 2004), nuclear localization signals (NLSs) (PROSITE [http://us.expasy.org/prosite/] [Falquet et al., 2002] and

PredictNLS [http://cubic.bioc.columbia.edu/predictNLS/] [Cokol et al., 2000]), and glycosylation sites (PROSITE).

2.3.4. Sequencing of the 3’ and 5’ ends of the MFSV genome

The terminal sequences of MFSV were identified by RACE. The viral 5’ trailer region was determined by both 5’ RACE (Invitrogen Corp.) and RLM-RACE (Ambion) following the manufacturers' instructions. For 5’ RACE, the cDNA of the MFSV genomic

RNA was synthesized using a gene-specific primer, 5’ RACE GSP1 (5’-AAATCTCTGT

TGAGCC-3’), and tailed with dCTP using terminal deoxynucleotidyl transferase. The

first amplification was carried out with the abridged anchor primer and a 5’ RACE GSP2

primer (5’-GGTCCATTGCAGAGAGATCAAC-3’). Nested amplification used the

abridged universal amplification primer (AUAP) and a 5’ RACE GSP3 primer

(5’-CTATCCTATCAGATCCCATAATGC-3’). For RLM-RACE, a 45-bp 5’ RACE

adapter was added to the MFSV genomic RNA, and then the cDNA was synthesized

using random decamers. The first amplification was carried out with the 5’ RACE outer

primer and the 5’ RACE GSP2 primer. Nested amplification used the 5’ RACE inner

primer and the 5’ RACE GSP3 primer.

The viral 3’ leader region was determined by 3’ RACE (Invitrogen Corp.). The

MFSV genomic RNA was tailed with ATP using the Poly(A) Tailing kit (Ambion), and cDNA was synthesized using the oligo(dT)-containing adapter primer. The first amplification was carried out with the AUAP and a 3’ RACE GSP1 primer

(5’-CTAAGAATGTCAGGAATAGGTCCTG-3’), and nested amplification was done with the AUAP and a 3’ RACE GSP2 primer (5’-CACCATAGGATAGACATGCA

TTCC-3’). The 5’ RACE and 3’ RACE products were ligated into the pGEM-T Easy vector (Promega Corp.) for sequencing.

Rhabdovirus nucleotide sequences used for interspecies comparison were obtained

from the genome sequences in the NCBI GenBank database for the following viruses:

SYNV (NC 001615), RYSV (NC003746), NCMV (NC 002251), LNYV (L24365 and

L24364) (Wetzel et al., 1994), VSIV (NC 001560), and RABV (NC 001542).

2.3.5. Characterization of transcription start sites of the MFSV G and L genes

The transcription start sites of the MFSV G and L genes were identified by

RLM-RACE. Total RNA from infected maize leaves was extracted with a ToTALLY

RNA isolation kit. Subsequently, mRNA was isolated from the total RNA preparation

using a Dynabead mRNA DIRECT kit (Dynal ASA, Oslo, Norway) following the manufacturer's instructions. The cDNA was synthesized from the adapter-ligated mRNA with random decamers. The first amplification was carried out with either the 5’ RACE outer primer and a G outer primer (5’-GTACTTAGTGGCAATGATGGTGTC-3’) or the

5’ RACE outer primer and an L outer primer (5’-GCTTGTAACAGTGCCCACA

TATC-3’). Nested amplification was carried out with either the 5’ RACE inner primer and a G inner primer (5’-CGATTATCAGTGTCGAGTTGTTC-3’) or the 5’ RACE inner primer and an L inner primer (5’-GTATGTCCCCCATGAGATAGTC-3’). The

RLM-RACE products were ligated into the pGEM-T Easy vector for sequencing.

2.3.6. Northern blot hybridization analysis

Total RNA (10 μg) from infected and healthy maize was denatured using glyoxal sample loading dye (Ambion) and separated on a 1.2% agarose gel in 1x BPTE electrophoresis buffer {10 mM PIPES [piperazine-N,N’-bis(2-ethanesulfonic acid)], 30

mM Bis-Tris, 1 mM EDTA, pH 6.5}, transferred to a positively charged BrightStar-Plus

nylon membrane (Ambion) with 20× SSC (3 M NaCl, 0.3 M sodium citrate, pH 7.0), and

cross-linked to the membrane by exposure to UV light (UV Transilluminator; Fisher

Scientific, Pittsburgh, PA) for 2 min. Probes were prepared by PCR amplification of DNA

fragments corresponding to each gene from the cDNA clones and subsequent

incorporation of [32P]dCTP (Amersham Biosciences Corp., Piscataway, NJ) (Feinberg

and Vogelstein, 1983). Hybridization of probes to Northern blots was carried out as

previously described (Redinbaugh et al., 1988). Northern blots were washed three times

for 10 min each in 2× SSC and 0.1% sodium dodecyl sulfate (SDS) and four times for 10

min each in 0.1× SSC and 0.1% SDS at 65°C and then exposed to the Storage Phosphor

Screen (Molecular Dynamics, Sunnyvale, CA) for 24 h. Images were captured using

ImageQuant software (Molecular Dynamics), converted to TIFF for export, and

processed in Photoshop version 7.0 (Adobe, San Jose, CA).

2.3.7. Construction of pGD derivatives for in planta subcellular localization

Construction of the binary pGDG and pGDR vectors, the pGDG construct for in

planta synthesis of the Arabidopsis thaliana Fib1 (AtFib1)-green fluorescent protein

(GFP), GFP-SYNV N, and the pGDR construct for in planta synthesis of DsRed-SYNV

P were described previously (Goodin et al., 2002). The pGDB and pGDY vectors were constructed by modification of pGD (Goodin et al., 2002). The cyan fluorescent protein

(CFP) and yellow fluorescent protein (YFP) genes were amplified from pECFP-C1 and

pEYFP-C1 (BD Biosciences, Palo Alto, CA), respectively. The PCR products were subcloned into the pGD vectors in a manner that reconstituted the multiple cloning site of pGD, since BglII, HindIII, and XhoI restriction sites were incorporated into the 3’ ends of

the PCR products. The CFP and YFP genes in the new vectors were verified by DNA

sequencing and expression in plant cells. The complete predicted MFSV ORFs (i.e., from

the putative start codon to the first stop codon) were expressed as fusions to the C termini

of fluorescent proteins. The MFSV ORFs were amplified from corresponding cDNA

clones by PCR, and primers with overhanging restriction sites were introduced to

facilitate directional cloning into the binary vector pGDB, pGDG, pGDR, or pGDY as

previously described (Goodin et al., 2002). PCR was performed using the high-fidelity

DynazymeEXT polymerase (Finnzymes, Espoo, Finland). PCR products were cloned

directly into the pCRII vector (Invitrogen Corp.) using topoisomerase-mediated cloning.

The DNA sequences of the full-length clones for the MFSV N, P, 3, 4, and M genes in

pGD derivatives were verified prior to transformation into Agrobacterium tumefaciens.

2.3.8. Agroinfiltration procedures

A. tumefaciens was infiltrated into leaves of Nicotiana benthamiana as previously described (Goodin et al., 2002). Briefly, suspensions of transformed C58C1 agrobacteria were adjusted to an optical density at 600 nm of 0.6 in agroinfiltration buffer (10 mM

MgCl2, 10 mM MES [morpholineethanesulfonic acid], pH 5.6), and acetosyringone was

added to a final concentration of 150 μM (Goodin et al., 2002). An agrobacterial

suspension was infiltrated into the mesophyll of leaves using a 1-ml disposable syringe.

Following infiltration, the leaves were examined by epifluorescence microscopy between

40 and 90 h postinfiltration.

2.3.9. DAPI staining of plant nuclei

DAPI (4’-6-diamidino-2-phenylindole dihydrochloride; 15 μg/ml) in agroinfiltration

buffer was infiltrated into leaves as described by Goodin et al. (Goodin et al., 2002).

Following infiltration, the plants were incubated in the dark for 1 to 2 h before

examination of leaf sections by epifluorescence microscopy.

2.3.10. Epifluorescence microscopy

Epifluorescence micrographs were acquired using an Axiocam MR monochromatic digital camera mounted on a motorized Axioplan2 microscope (Carl Zeiss Microimaging

Inc., Thornwood, NY). Camera and microscope settings were controlled by Axiovision

software version 4.1. False colors for differentiating DAPI, CFP, GFP, and DsRed2

fluorescences were assigned using color settings in the Axiovision software. Filter sets

that permitted viewing of the relevant fluorescences were purchased from Chroma

Technology Corp. (Rockingham, VT) and included the following filter sets. (i) Filter set

31001 for viewing GFP; this set consisted of a D470/40× excitation (Ex) filter, a 505

DCLP dichroic, and a D540/40× emission (Em) filter. (ii) Filter set 31000 was used for

viewing DAPI-stained nuclei. This set consisted of a D360/40× Ex filter, a 400 DCLP

dichroic, and a D460/50 M Em filter. (iii) Filter set 310044 V2, used for capturing CFP

fluorescence, consisted of a D436/20× Ex filter, a 455DCLP dichroic, and a D480/40 M

Ex filter. (iv) YFP fluorescence was viewed using a 41028 filter set that consisted of an

HQ500/20× Ex filter, a Q5151LP dichroic, and a HQ535/30 M Em filter. (v) For viewing

fluorescence from DsRed2, a 41035 filter set, consisting of a HQ546/12× Ex filter, a

Q560LP dichroic, and a HQ650/75 M Em filter, was used. The lenses used in this study

included the Plan Neofluar 10×/NA 0.3; the Plan Neofluar 25×/NA 0.8 multiple

immersion lens, used primarily in the water immersion setting; and a Plan Apochromat

100×/NA 1.4 oil immersion lens. Sections of plant tissue were mounted in water on

standard glass slides and covered with no. 1 glass coverslips (Corning Inc., Corning, NY).

Leaves were mounted so that the abaxial surface was viewed. Micrographs were exported

from the Axiovision software as TIFF files. All subsequent cropping and image

manipulations were carried out in Photoshop version 7.0 (Adobe Systems Inc., San Jose,

CA) and Canvas version 8.0 (Deneba Software, Miami, FL).

2.3.11. Confocal laser scanning microscopy

Confocal laser scanning micrographs were acquired on a TCS SP2-AOBS

microscope (Leica Microsystems, Bannockburn, IL.). GFP variants were excited

simultaneously using the 488-nm laser line. Fluorescence emissions from CFP, GFP, and

YFP could be distinguished using the spectral imaging capability provided by the prism

spectrophotometer of the microscope. The ability to unambiguously differentiate these three fluors when coexpressed in leaf epidermal cells allowed assignment of the subnuclear locales in which the N and P proteins accumulate.

2.3.12. Nucleotide sequence accession number

The sequence of the MFSV genome was deposited in GenBank under accession

number AY618417.

2.4. Results

2.4.1. Nucleotide sequence of MFSV

The MFSV genome sequence was obtained by assembling sequences derived from

the overlapping sequences of 12 randomly primed cDNA clones, five RT-PCR products, and two RACE products (Fig. 2.1). The MFSV genome consisted of 13,782 nucleotides

(nt) and seven ORFs on the antigenomic strand.

The sequence of the 3’ end of the MFSV genome was determined by 3’ RACE, and

that of the 5’ end was determined by 5’ RACE and RLM-RACE. The 3’ RACE PCR

products were cloned, and inserts of all six clones were identical, indicating that the 3’

end of the MFSV genome was 3’-UGUGUGGUUUUUCCCACUGC · · · -5’. The

sequences of inserts of four 5’ RACE and RLM-RACE clones were identical, indicating

that the 5’ end of the MFSV genome was 3’- · · · GCAGUAAAAAAACGGACACA-5’.

Identical 5’- and 3’-end sequences using either poly(A) or poly(C) adapters were

obtained at the University of Wisconsin (P. Flanary and T. L. German, personal

communication). These results led to the conclusion that the MFSV genomic RNA had a

184-nt 3’ leader region preceding the leader-N intergenic sequence and a 145-nt 5’ trailer

region following the L-trailer intergenic sequence.

Comparison of the 3’ and 5’ ends of the MFSV genomic RNA revealed that 19 of 30

nucleotides were complementary and might give rise to a putative panhandle structure

(Fig. 2.2). Similar structures were reported for SYNV, LNYV, and NCMV (Choi et al.,

1994; Tanno et al., 2000; Wetzel et al., 1994). In addition, the first 30 nt of the 3’ leader sequence had a high U residue content (47%), similar to those described for SYNV (53%),

RYSV (37%), NCMV (37%), and LNYV (60%). Although the plant rhabdoviruses share complementarity of their 3’ and 5’ ends and have similar nucleotide biases, these sequences did not have significant sequence identity among rhabdoviruses (Fig. 2.2).

2.4.2. Gene junctions of the MFSV genome

The MFSV ORFs were separated by gene junctions with the consensus sequence

3’-UUUAUUUUGUAGUUG-5’(Fig. 2.3A). This sequence was broadly conserved

among plant and animal rhabdoviruses (Fig. 2.3B) and was divided into three distinct motifs: the sequences corresponding to the 3’ ends of mRNAs, the intergenic sequences, and the sequences corresponding to the 5’ ends of mRNAs (Rose and Whitt, 2001) (Fig.

2.3). The corresponding sequences of MFSV mRNA 3’ ends were A/U rich and 8 nucleotides in length, and only the ORF7-5’ trailer junction had 2 nucleotide differences

from the consensus sequence. The intergenic sequence GAUG was conserved in all gene

junctions of MFSV; however, this sequence was not conserved among rhabdoviruses (Fig.

2.3B). The transcription start site consisted of 3 nucleotides, and only the ORF4 mRNA start site had a single-nucleotide change relative to the consensus sequence, UUG. The

MFSV transcription start site was identical to those of other rhabdoviruses (SYNV, RYSV,

VSIV, and RABV), except for the two cytorhabdoviruses NCMV and LNYV (Fig. 2.3B).

2.4.3. Transcription start sites of the MFSV G and L genes

RLM-RACE was used to confirm the sequences of the putative transcription start

sites for the MFSV G and L genes (Fig. 2.4). The sequences of six RLM-RACE products

for the G gene were identical (Fig. 2.4A). The sequences of four RLM-RACE products

for the L gene were identical, while the sequences of two others differed from these at

position 3 (Fig. 2.4B). These data are consistent with the putative UUG transcription start

sites identified in Fig. 2.3A. However, the 5’ ends of transcripts contained a nucleotide A

that does not correspond to the viral genomic sequence (Fig. 2.4). Whether the

single-nucleotide change at the third nucleotide found in two L gene RLM-RACE clones

was due to true variability in the L gene mRNAs is not known.

2.4.4. Analysis of the MFSV ORF sequences

The predicted proteins encoded by the seven MFSV ORFs were examined for

sequence similarity to other proteins in the NCBI GenBank nonredundant database using

BLASTP. The deduced protein sequence of ORF1 had significant similarity to the N

proteins of RYSV (expected [E] value = 8e-21), SYNV (E value = 1e-19), and NCMV (E

value = 5e-6). The ORF6 protein sequence had significant similarity to those of the G proteins of RYSV (E value = 5e-22) and SYNV (E value = 1e-16), and the ORF7 protein

sequence had significant similarity to those of the L proteins of SYNV (E value = 0.0),

RYSV (E value = 0.0), and other nonsegmented, negative-sense RNA viruses. Thus,

based on sequence similarity, MFSV ORF1, ORF6, and ORF7 likely encode the N, G,

and L proteins, respectively. The ORF2, ORF3, ORF4, and ORF5 protein sequences did

not show any significant similarities to other rhabdovirus proteins or other sequences in

GenBank.

The protein sequences of MFSV and other nucleorhabdoviruses were searched for

domains and motifs, including NLSs, glycosylation sites, N-terminal signal peptides, and

transmembrane domains. Putative NLSs were found at amino acid positions 436 to 452

(KRSSDGTGNVSKKKSRK) of the N protein, at positions 17 to 33

(RKALTKASKALFKGKIK) of the ORF4 protein, and at positions 195 to 211

(KKEDKAEKATTEKRKRQ) of the ORF5 protein, whereas no NLSs were identified in

the MFSV ORF2, ORF3, G, and L proteins (Table 2.1). The SYNV N and M and RYSV

N proteins also had putative NLSs in the carboxyl regions, but no putative NLS was

identified in the RYSV M protein (Redinbaugh and Hogenhout, 2005).

Similar to other rhabdovirus G proteins, the putative MFSV G protein apparently

has abundant glycosylation signals. Eight potential glycosylation signals (N-[P]-S/T-[P]-)

were located at amino acid positions 64 to 67, 131 to 134, 132 to 135, 139 to 142, 204 to

207, 325 to 328, 438 to 441, and 494 to 497. The MFSV G protein also had an N-terminal

signal peptide sequence (MMARLVPCFTLALLLHLTEC) and a putative cleavage site

(C/A) between amino acid positions 20 and 21. Furthermore, a transmembrane domain

(FIIKLVIGFTVGTIMLYISWIII) was identified at amino acid positions 529 to 551.

Other predicted proteins of MFSV did not have abundant glycosylation sites, signal

peptide sequences, or transmembrane domains.

2.4.5. Detection of MFSV ORF transcripts in plants

To test whether all the identified ORFs in the MFSV genome are transcribed,

Northern blots of total RNAs from healthy and MFSV-infected maize were hybridized

with probes corresponding to each ORF (Fig. 2.1). Transcripts of expected sizes that

corresponded to each of the seven ORFs were detected in MFSV-infected maize (Fig.

2.5), whereas no hybridization occurred with RNA from healthy maize (data not shown).

Several RNAs hybridized to the ORF7 probe, with the size of the largest matching the

expected size of the ORF7 transcript (Fig. 2.5A, lane L). The predicted length of gene 3 is 357 nt, and a transcript of ~400 bp hybridized to the ORF3 probe (Fig. 2.5A, lane 3).

The predicted length of gene 4 is 1,185 nt, and a transcript of ~1,200 bp hybridized to the

ORF4 probe (Fig. 2.5A, lane 4). To exclude the possibility of RNA degradation, a blot that was hybridized with the ORF4 probe was subsequently probed with the ORF3 probe.

As expected, the resulting blot showed two distinct hybridized bands (Fig. 2.5B). These

results demonstrated that seven distinct transcripts were present in MFSV-infected maize and that these transcripts corresponded to the seven putative genes identified in the

MFSV genome.

2.4.6. Subcellular localization of the MFSV proteins

NLSs were identified in three of the seven MFSV ORFs, and nuclear localization

was demonstrated for the SYNV N and M proteins (Goodin et al., 2002). To determine

the subcellular localization of the MFSV proteins, we used in planta localization and

colocalization of fluorescent-protein fusions. Full-length ORF sequences were introduced

into binary pGD derivatives for A. tumefaciens-mediated agroinfiltration of N.

benthamiana leaves (Goodin et al., 2002) and subsequent in planta production of MFSV proteins fused at the N terminus to CFP, YFP, and GFP.

Agroinfiltration of fluorescent-protein fusions of MFSV N, ORF2, ORF3, ORF4,

and ORF5 proteins showed that CFP-MFSV N, GFP-MFSV 4, and GFP-MFSV 5

accumulated in the nuclei (Figs. 2.6 and 2.7B and C), whereas YFP-MFSV 2 was

distributed throughout the cell (Fig. 2.6) and YFP-MFSV 3 accumulated in punctate loci

in the cytoplasm (Fig. 2.7A). These results were consistent with the prediction of NLSs in

the MFSV N, ORF4, and ORF5 proteins.

When leaves were coinfiltrated with mixtures of C58C1 agrobacteria harboring

CFP-MFSV N and YFP-MFSV 2 fusions, both fusions colocalized to the subnuclear

locale (Fig. 2.8A). However, unlike the SYNV N and P proteins, the MFSV N and ORF2

proteins localized to the nucleolus (Fig. 2.8A). To confirm the nucleolar localization, we

coexpressed CFP-MFSV N and YFP-MFSV 2 with the AtFib1-GFP that localizes to the

nucleolus in A. thaliana and N. benthamiana epidermal cells (Barneche et al., 2000;

Goodin et al., 2002). This result confirmed that CFP-MFSV N and YFP-MFSV 2 targeted the same subnuclear locale as AtFib1-GFP, which is in the nucleolus (Fig. 2.8B).

Finally, we determined whether coinfiltration of fluorescent-protein fusions of the

MFSV N protein and the SYNV P protein allowed redirection of cytosolic SYNV P

protein to the subnuclear locale, similar to co-introduction of the SYNV N and P proteins.

Coinfiltrated CFP-MFSV N and DsRed-SYNV P did not interact to target a subnuclear locale but looked very similar to DsRed-SYNV P infiltrated alone (Fig. 2.9). Similar results were obtained when GFP-SYNV N was coinfiltrated with YFP-MFSV 2 (Fig. 2.9).

These data suggest that the interaction of N and P proteins leading to subnuclear targeting

is virus specific.

2.5. Discussion

We have mapped and sequenced the complete genome of the leafhopper-transmitted

maize pathogen MFSV and showed that all seven genes included in its genome were

expressed during infection of maize. Furthermore, in planta subcellular localization

studies of fluorescent-protein fusions showed that the MFSV N, ORF4, and ORF5

proteins targeted to the nuclei and that coinfiltrated MFSV N and ORF2 proteins interacted, resulting in targeting of both proteins to the nucleolus. We also showed that the nuclear targeting of coinfiltrated N and P proteins was virus specific and that the protein products of the two additional genes (3 and 4) of MFSV had different subcellular localizations than the SYNV sc4 protein.

Assignment of functions for the MFSV N (ORF1), G (ORF6), and L (ORF7) proteins could be made largely on the basis of sequence similarities to SYNV genes and proteins. However, nucleotide and deduced protein sequences of ORF2, ORF3, ORF4, and ORF5 showed few similarities to other rhabdovirus genes or sequences in GenBank.

We used in planta subcellular localization of fluorescent-protein fusions, a useful technique in protein subcellular targeting studies and for elucidation of protein function

(Goodin et al., 2002; Hanson and Kohler, 2001; Stewart, 2001), to begin to define the functions of the MFSV ORFs.

The most likely gene order of the MFSV genome is 3’-N-P-3-4-M-G-L-5’.

Sequence analysis and in planta localization results suggest that ORF2 of MFSV encodes the P protein because (i) P is located adjacent to N in all rhabdoviruses sequenced so far,

(ii) the predicted size of the ORF2 protein is similar to those of P proteins of other rhabdoviruses, and (iii) similar to the SYNV N and P proteins (Goodin et al., 2002), coinfiltration of the MFSV N protein with the MFSV ORF2 protein resulted in nucleolar localization of both proteins, whereas the MFSV N protein targeted the whole nucleus and the MFSV ORF2 protein was distributed throughout the cell when the proteins were infiltrated alone. For ORF5 of MFSV, sequence analysis and in planta localization results indicate that this ORF probably encodes the M protein because (i) the position of the

NLS and the size of the ORF5 protein are most comparable to the M protein of SYNV

(Redinbaugh and Hogenhout, 2005) and (ii) the ORF5 protein of MFSV targeted the nucleus when expressed alone, similar to the SYNV M protein (Goodin et al., 2002). The

M protein sequences are not conserved among plant rhabdoviruses, whereas they are conserved among animal- or fish-infecting rhabdoviruses. The M proteins are likely

55 prone to considerable adaptive evolution because these proteins play important roles in the suppression of host transcription (Ahmed et al., 2003; Kopecky and Lyles, 2003) and determine budding sites (Harty et al., 1999; Jayakar and Whitt, 2002).

The MFSV N-P protein complex appears to target a different subnuclear locale than the SYNV N-P protein complex. This suggests that there may be differences in the infection mechanisms for the two viruses. However, N. benthamiana is a host of SYNV, whereas we have not been able to infect N. benthamiana with MFSV by rub inoculation or insect transmission so far (J. C. Todd, S. A. Hogenhout, and M. G. Redinbaugh, unpublished results). More work is needed to determine whether the differential targeting is associated with the abilities of the two viruses to replicate in N. benthamiana.

The hypothesis that the infection mechanisms of MFSV and SYNV in plant cells are different is supported by the observation that the subcellular distributions of ORF3 and

ORF4 proteins of MFSV were different from the subcellular distribution of the SYNV sc4 protein. The SYNV sc4 protein is a membrane-associated protein, although it does not have a transmembrane domain or an NLS (Scholthof et al., 1994). The SYNV sc4 protein has been predicted to be a member of the 30K superfamily of plant virus movement proteins, related to the 30-kDa Tobacco mosaic virus (TMV) movement protein (Melcher, 2000). All of the sequenced plant rhabdovirus genomes encode a protein of unknown function similar in mass to the 36.7-kDa sc4 protein between the P and M genes, except for NCMV, which has four small proteins with unknown functions.

This unique protein includes the MFSV ORF4 protein, the RYSV 3 protein (Chen et al.,

1998), and the LNYV 4b protein (GenBank accession no. AF209034). Although one is tempted to speculate that, based on similar genome locations and sizes, these additional

genes have similar functions, our results show that this may not be the case, because the

SYNV sc4 protein is cytosolic (Goodin et al., 2002) while the MFSV ORF4 protein is

nuclear.

The interaction of the N and P proteins is specific for MFSV and SYNV; that is, co-introduction of the MFSV N protein did not redirect the SYNV P protein to the nucleus, and co-introduction of the SYNV N protein did not redirect the MFSV P protein to the nucleus. This is not surprising, given the lack of conservation among plant rhabdovirus P proteins. The P protein acts as a transcription factor and aids in N protein encapsidation of RNA for rhabdoviruses (Gupta et al., 2003). It remains to be investigated whether the interactions between N and P proteins are specific for other rhabdoviruses as well.

The organization of the MFSV genome is distinct from those of other rhabdoviruses

described so far because MFSV has two additional genes between the P and M genes

compared to other nucleorhabdoviruses (Table 2.2). However, the cytorhabdovirus

NCMV has four genes between the P and M genes (Tanno et al., 2000), and the

nucleorhabdovirus RYSV has two additional genes, with one located between the P and

M genes and one between the G and L genes (Huang et al., 2003) (Table 2.2). Thus, the

locations and numbers of genes, in addition to the basic gene order (3’-N-P-M-G-L-5’)

(Walker et al., 2000), vary extensively among insect-transmitted plant rhabdoviruses. All

seven ORFs of MFSV were transcribed during the infection of maize, and gene junctions

flanked all ORFs. Gene junctions are conserved among plant and animal rhabdoviruses

and are important for transcription termination and reinitiation of the rhabdovirus

polymerase complex (Rose and Whitt, 2001).

The MFSV gene junctions suggest that the MFSV genes use UUG as a consensus transcription start site. The RLM-RACE data for the MFSV G and L genes confirmed this transcription start site. However, a nonviral nucleotide was present at the 5’ ends of

MFSV mRNAs. Heterogeneous nucleotides were found at the 5’ ends of viral mRNAs for

RYSV and LNYV (Luo and Fang, 1998; Wetzel et al., 1994) and were thought to be derived from host cellular RNAs (Kurath et al., 1997). The 3-nucleotide transcription start sites of animal rhabdoviruses and nucleorhabdoviruses are identical, but those of cytorhabdoviruses are clearly different (CUU/A rather than UUG). It is not known whether there is a biological significance in transcription initiation associated with the sequence difference in cytorhabdoviruses.

With the accumulation of completed genome sequences of rhabdoviruses, it is now

becoming clear that plant rhabdoviruses are diverse. In this study, we showed for the first

time that rhabdoviruses are diverse not only in genome sequence, but also in the subcellular localization of proteins, and hence in the interaction with host factors. It is

therefore not surprising that rhabdovirus proteins specifically interact with each other and

not with orthologous proteins of another rhabdovirus, as shown here for the N and P

proteins of MFSV and SYNV. This information will help to determine factors essential to

the host and vector specificities of rhabdoviruses.

2.6. Acknowledgments

The project was supported by the National Research Initiative of the USDA

Cooperative State Research, Education and Extension Service, grant 2002-35302-12653,

and the OARDC Research Enhancement and Competitive Grants Program.

We thank T. Meulia from the Molecular and Cellular Imaging Center of the OARDC for providing us with a pZeroAmp vector; T. L. German and P. Flanary from the

Department of Entomology, University of Wisconsin, for confirming our 3’ and 5’ RACE

MFSV sequence results; J. von der Heiden from Leica Microsystems for his excellent technical assistance with acquisition of the confocal micrographs; and R.G. Dietzgen from the Department of Primary Industries and Fisheries, Queensland Government,

Australia, for critical reading of the manuscript.

2.7. References

Ahmed, M., McKenzie, M.O., Puckett, S., Hojnacki, M., Poliquin, L. and Lyles, D.S. (2003) Ability of the matrix protein of vesicular stomatitis virus to suppress beta interferon gene expression is genetically correlated with the inhibition of host RNA and protein synthesis. J. Virol., 77, 4646-4657.

Barneche, F., Steinmetz, F. and Echeverria, M. (2000) Fibrillarin genes encode both a conserved nucleolar protein and a novel small nucleolar RNA involved in ribosomal RNA methylation in Arabidopsis thaliana. J. Biol. Chem., 275, 27212-27220.

Bendtsen, J.D., Nielsen, H., von Heijne, G. and Brunak, S. (2004) Improved prediction of signal peptides: SignalP 3.0. J. Mol. Biol., 340, 783-795.

Chen, X.Y., Luo, Z.L. and Fang, R.X. (1998) Structure analysis of the rice yellow stunt rhabdovirus gene 3. Chinese Sci. Bull., 43, 745-748.

Choi, T.J., Wagner, J.D. and Jackson, A.O. (1994) Sequence analysis of the trailer region of sonchus yellow net virus genomic RNA. Virology, 202, 33-40.

Cokol, M., Nair, R. and Rost, B. (2000) Finding nuclear localization signals. EMBO Rep., 1, 411-415.

Falquet, L., Pagni, M., Bucher, P., Hulo, N., Sigrist, C.J., Hofmann, K. and Bairoch, A. (2002) The PROSITE database, its status in 2002. Nucleic Acids Res., 30, 235-238.

Feinberg, A.P. and Vogelstein, B. (1983) A technique for radiolabeling DNA restriction endonuclease fragments to high specific activity. Anal. Biochem., 132, 6-13.

Gupta, A.K., Shaji, D. and Banerjee, A.K. (2003) Identification of a novel tripartite complex involved in replication of vesicular stomatitis virus genome RNA. J. Virol., 77, 732-738.

Hanson, M.R. and Kohler, R.H. (2001) GFP imaging: methodology and application to investigate cellular compartmentation in plants. J. Exp. Bot., 52, 529-539.

Harty, R.N., Paragas, J., Sudol, M. and Palese, P. (1999) A proline-rich motif within the matrix protein of vesicular stomatitis virus and rabies virus interacts with WW domains of cellular proteins: implications for viral budding. J. Virol., 73, 2921-2929.

Hogenhout, S.A., Redinbaugh, M.G. and Ammar, E.D. (2003) Plant and animal rhabdovirus host range: a bug's view. Trends Microbiol., 11, 264-271.

Jackson, A.O., Dietzgen, R.G., Goodin, M.M., Bragg, J.N. and Deng, M. (2005) Biology of plant rhabdoviruses. Annu. Rev. Phytopathol., 43, 623-660.

Jayakar, H.R. and Whitt, M.A. (2002) Identification of two additional translation products from the matrix (M) gene that contribute to vesicular stomatitis virus cytopathology. J. Virol., 76, 8011-8018.

Kopecky, S.A. and Lyles, D.S. (2003) Contrasting effects of matrix protein on apoptosis in HeLa and BHK cells infected with vesicular stomatitis virus are due to inhibition of host gene expression. J. Virol., 77, 4658-4669.

Krug, R.M. (1981) Priming of influenza viral RNA transcription by capped heterologous RNAs. Curr. Top. Microbiol. Immunol., 93, 125-149.

Kurath, G., Higman, K.H. and Bjorklund, H.V. (1997) Distribution and variation of NV genes in fish rhabdoviruses. J. Gen. Virol., 78, 113-117.

Louie, R. (1995) Vascular puncture of maize kernels for the mechanical transmission of maize white line mosaic virus and other viruses of maize. Phytopathology, 85, 139-143.

Luo, Z.L. and Fang, R.X. (1998) Structure analysis of the rice yellow stunt rhabdovirus glycoprotein gene and its mRNA. Arch. Virol., 143, 2453-2459.

Melcher,U. (2000) The '30K' superfamily of viral movement proteins. J. Gen. Virol., 81, 257-266.

Redinbaugh, M.G. and Hogenhout, S.A. (2005) Plant rhabdoviruses. Curr. Top. Microbiol. Immunol., 292, 143-163.

Redinbaugh, M.G., Wadsworth, G.J. and Scandalios, J.G. (1988) Characterization of catalase transcripts and their differential expression in maize. Biochim. Biophys. Acta, 951, 104-116.

Rose, J.K. and Whitt, M.A. (2001) Rhabdoviridae: the virus and their replication. In Knipe, D.M. and Howley, P.M. (eds.), Fields virology. Lippincott Williams & Wilkins, Philadelphia, pp. 1221-1244.

Scholthof, K.B., Hillman, B.I., Modrell, B., Heaton, L.A. and Jackson, A.O. (1994) Characterization and detection of sc4: a sixth gene encoded by sonchus yellow net virus. Virology, 204, 279-288.

Sonnhammer, E.L., von Heijne, G. and Krogh, A. (1998) A hidden Markov model for predicting transmembrane helices in protein sequences. Proc. Int. Conf. Intell. Syst. Mol. Biol., 6, 175-182.

Stewart, C.N., Jr. (2001) The utility of green fluorescent protein in transgenic plants. Plant Cell Rep., 20, 376-382.

Tanno, F., Nakatsu, A., Toriyama, S. and Kojima, M. (2000) Complete nucleotide sequence of Northern cereal mosaic virus and its genome organization. Arch. Virol., 145, 1373-1384.

Wetzel, T., Dietzgen, R.G. and Dale, J.L. (1994) Genomic organization of lettuce necrotic yellows rhabdovirus. Virology, 200, 401-412.

ORF Calculated TM (position in NLS (position Putative function no. mass (kDa) protein)a in protein)a 1 51.6 ND + (C-term) Nucleocapsid protein (N) 2 38.4 ND ND Phosphoprotein (P) 3 10.7 ND ND Unknown 4 37.2 ND + (N-term) Unknown 5 28.5 ND + (C-term) Matrix protein (M) 6 67.0 + (C-term) ND Glycoprotein (G) 7 223.5 ND ND Polymerase (L) a TM: transmembrane domain; NLS: nuclear localization signal; ND: Not detected; N-term: N-terminal half of protein; C-term: C-terminal half of protein; +, present.

Table 2.1. Features of the encoded proteins of the MFSV genome

Virus Genome organization Vectora Hosta

Nucleorhabdovirus

MFSV 3’ N P 3 4 M G L 5’ L M

SYNV 3’ N P sc4 M G L 5’ A D

RYSV 3’ N P 3 M G 6 L 5’ L M

Cytorhabdovirus

LNYV 3’ N P 4b M G L 5’ A D

NCMV 3’ N P 3 4 5 6 M G L 5’ P M a A: aphid; L: leafhopper; P: planthopper; D: dicot; M: monocot.

Table 2.2. Genome organization of plant rhabdoviruses

Fig. 2.1. Schematic diagram of the MFSV genome organization. The open reading frames (block arrows) are derived from the antigenomic sequence. The asterisks indicate gene junctions (Fig. 2.3). Bold lines indicate locations of probes A, B, C, D, E, F, and G used for Northern blot hybridization of mRNAs of ORF1, -2, -3, -4, -5, -6, and -7, respectively (Fig. 2.5).

Fig. 2.2. Sequences of the 3’ and 5’ termini of plant rhabdovirus genomes. Sequences are shown in the genomic sense. Vertical lines indicate complementary nucleotides between leader and trailer sequences. Overhangs in leader sequences are underlined.

Fig. 2.3. Comparison of rhabdovirus gene junctions. A. Gene junctions of the MFSV genome. The three motifs are indicated, corresponding to the 3’ ends of the mRNAs (column 1), intergenic sequences (column 2), and the 5’ ends of the mRNAs (column 3). B. Consensus sequences of gene junctions of plant and animal rhabdoviruses. All sequences are presented in genomic sense in the 3’-to-5’ orientation, and nucleotide differences are underlined. Abbreviations: 3’ le, 3’ leader sequence; 5’ tr, 5’ trailer sequence; (N)n, variable number of nucleotides

Fig. 2.4. RLM-RACE of the MFSV G and L mRNAs. The sequences of six clones, each derived from the G gene and L gene mRNAs, were determined. Nucleotides of mRNAs complementary to the transcription start sites in the conserved gene junction sequences of the viral genome (vg) are boxed.

Fig. 2.5. Detection of MFSV gene transcripts in infected maize by Northern blot analysis. A. Blots of total RNA isolated from healthy (data not shown) or MFSV-infected maize hybridized to probes A, B, C, D, E, F, and G, which are located within the MFSV N, 2, 3, 4, 5, G, and L genes, respectively. B. Blot hybridized to probe D and subsequently to probe C. Figure 2.1 shows the localizations of the probes. The asterisks indicate migration of the genomic RNA of MFSV, and the arrows indicate hybridization to mRNAs. Gel images of 25S rRNA were used as loading controls (below).

Fig. 2.6. Epifluorescence microgaphs of subcellular localizations of fluorescent-protein fusions of the MFSV N and ORF2 proteins. The DNA selective dye DAPI was used to determine the position of the nuclei in plant cells. Infiltrations of unfused CFP and YFP were included as negative controls. Cellular views of localizations of CFP-MFSV N to the nucleus (top row), and YFP-MFSV 2 throughout the cell (bottom row) are shown. Bars = 5 μm.

Fig. 2.7. Epifluorescence micrographs of subcellular localizations of fluorescent-protein fusions of the MFSV ORF3, ORF4, and ORF5 proteins in the cellular (top rows) and nuclear (bottom rows) views. The DNA selective dye DAPI was used to determine the position of the nuclei in plant cells. A. Accumulation of YFP-MFSV 3 in punctate loci in the plant cell cytoplasm. B. Nuclear localization of GFP-MFSV 4. C. Nuclear localization of GFP-MFSV 5. Bars = 5 μm.

Fig. 2.8. Subcellular localization of fluorescent-protein fusions of the MFSV N and ORF2 proteins and the SYNV N and P proteins. The DNA selective dye DAPI was used to determine the position of the nuclei in plant cells. A. Epifluorescence microgaphs of GFP-SYNV N infiltrated by itself targeting the nucleus (top row); coinfiltrated GFP-SYNV N and DsRed-SYNV P targeting the subnuclear locale (middle row); and coinfiltrated CFP-MFSV N and YFP-MFSV 2 targeting the nucleolus (bottom row). B. Confocal micrographs of the colocalization of AtFib1-GFP, CFP-MFSV N, and YFP-MFSV 2 in the nucleolus of a plant cell. Single-plane optical sections (0.3 mm thick) for each channel were taken through the largest area of fluorescence within the nucleolus. Bars = 5 μm.

Fig. 2.9. Epifluorescence microgaphs of infiltrated DsRed-SYNV P alone (top row), coinfiltrated CFP-MFSV N and DsRed-SYNV P (middle row), and coinfiltrated GFP-SYNV N and YFP-MFSV 2 (bottom row). The DNA selective dye DAPI was used to determine the position of the nuclei in plant cells. The fluorescence of GFP-SYNV N and YFP-MFSV 2 could not be distinguished by epifluorescence microscopy. Bars = 5 μm.

CHAPTER 3

SHOTGUN SEQUENCING OF THE NEGATIVE-SENSE RNA GENOME OF

THE RHABDOVIRUS MAIZE MOSAIC VIRUS

Sharon E. Reed1, Chi-Wei Tsai1, Kristen J. Willie3, Margaret G. Redinbaugh2,3 and

Saskia A. Hogenhout1

1Department of Entomology, 2Department of Plant Pathology, Ohio Agricultural Research and Development Center (OARDC), The Ohio State University, Wooster, Ohio

3U.S. Department of Agriculture-Agricultural Research Service, Corn and Soybean

Research, Wooster, Ohio

3.1. Abstract

The maize-infecting nucleorhabdovirus, Maize mosaic virus (MMV), was sequenced to near completion using the random shotgun approach. Sequences of 102 clones from a cDNA library constructed from randomly-primed viral RNA were compiled into a 12,133 nucleotide (nt) contig containing six open reading frames. The contig consisted of 97 sequences averaging 660 bp in length. The average sequence coverage was six-fold, and

93% of the contig had sequence reads covering both strands. The remaining sequence was derived from single (5%) or multiple (2%) reads on the same strand. Three of the six

ORFs showed significant similarities to the deduced protein sequences of the nucleocapsid, glycoprotein and polymerase sequences of other rhabdoviruses. The

predicted gene order of the MMV genome was 3’-N-P-3-M-G-L-5’. Shotgun sequencing

of the MMV genome took approximately 127 h and cost $ 0.38 per nt (including labor),

whereas the primer walking approach for sequencing the 13,782-nt MFSV genome [Tsai,

C.W., Redinbaugh, M.G., Willie, K.J., Reed, S., Goodin, M., and Hogenhout, S.A., 2005.

Complete genome sequence and in planta subcellular localization of Maize fine streak

virus proteins. J. Virol. 79, 5304–5314] took about 217 h and cost $ 0.50 per nt. Thus, the

shotgun approach gave good depth of coverage for the viral genome sequence while

being significantly faster and less expensive than the primer walking method. This

technique will facilitate the sequencing of multiple rhabdovirus genomes.

3.2. Introduction

Rhabdoviruses are economically important pathogens that infect animals, humans,

and plants (Hogenhout et al., 2003). The family Rhabdoviridae is comprised of six genera:

Lyssavirus (contains Rabies virus); Vesiculovirus (Vesicular stomatitis Indiana virus);

Ephemerovirus (Bovine ephemeral fever virus); Novirhabdovirus (Infectious hematopoietic necrosis virus); Cytorhabdovirus (Lettuce necrotic yellows virus); and

Nucleorhabdovirus (Sonchus yellow net virus). Currently, 15 plant-infecting members of the Rhabdoviridae are listed in the Universal Database of the International Committee of

Taxonomy of Viruses (ICTVdb; www.ncbi.nlm.nih.gov/ICTVdb/index.htm), and 88 possible plant rhabdoviruses are listed in the Virus Identification Data Exchange (VIDE) database (www.image.fs.uidaho.edu/vide). At this time, only six plant rhabdovirus genomes have been partially or completely sequenced. The lack of virus genome sequences has slowed the development of modern detection techniques and research on plant rhabdoviruses.

Current methods of genome sequencing of viruses, such as primer walking analysis

of large cDNA clones, can be time consuming and hence costly in comparison to the

more rapid shotgun sequencing methods that are currently used to sequence bacterial and

eukaryotic genomes to completion. The random shotgun sequencing method involves the

cloning and sequencing of ~1,000 bp genomic DNA fragments, and subsequent assembly

of sequences into contigs based on sequence overlaps (Kaiser et al., 2003). Originally,

Fleischmann et al. (1995) sequenced Haemophilus influenzae at 48 cents/bp using the

shotgun method, but technological advances have lowered costs to 3–4 cents/bp for 99%

of the genome sequence and 8–9 cents/bp for the complete genome (Fraser et al., 2002).

Here, we outline a procedure for the shotgun sequencing of the genome of Maize

mosaic virus (MMV), a negative-sense RNA virus of the family Rhabdoviridae. MMV is a maize-infecting virus that is found in Central America, the Caribbean, India, Mauritius,

Tanzania, and the United States (Jackson et al., 1981). It is transmitted in a persistent propagative manner by the planthopper Peregrinus maidis (Ammar and Nault, 2002; Falk and Tsai, 1985). MMV was classified as a nucleorhabdovirus because virions show the classic bullet-shape of rhabdoviruses and electron micrographs indicate that MMV viral particles bud through the perinuclear membrane (Ammar and Nault, 1985; Ammar, 1994;

McDaniel et al., 1985). Three structural proteins were apparent after separation of MMV virion proteins by SDS-PAGE similarly to other members of the family Rhabdoviridae

(Falk and Tsai, 1983). Results of this study showed that shotgun sequencing of a rhabdoviral genome is not only possible but is also an affordable and quick method relative to the more traditional primer walking method.

3.3. Materials and methods

3.3.1. MMV and RNA isolation

MMV was maintained on maize seedlings by serial inoculations using viruliferous P.

maidis as previously described (Falk and Tsai, 1985). Leaf laminar tissue (60 g) was collected from symptomatic maize plants approximately six weeks post inoculation and immediately homogenized in 3 vol. of extraction buffer containing 0.1 M sodium

phosphate, pH 7.2 and 10% sucrose (Creamer, 1992; Skaf and Carroll, 1995). The homogenate was filtered through cheesecloth and miracloth then 5% (w/v) activated charcoal was added. After stirring for 20 min and repeating the filtration step, the homogenate was centrifuged at 3,300 × g for 10 min at 4 °C. The resulting supernatant was centrifuged at 27,500 × g for 30 min at 4 °C. The resulting pellet was resuspended in

0.1 vol. extraction buffer, filtered through polyester fiber (Fairfield Corp., Danbury, CT), layered onto 25% (w/v) sucrose in 0.1 M sodium phosphate, pH 7.2, and centrifuged at

99,400 × g for 30 min at 4 °C. The pellet was resuspended in 6 ml 0.1 M sodium phosphate, pH 7.2, layered onto 10–40% (w/v) sucrose gradients in 0.1 M sodium phosphate, pH 7.2 (McDaniel et al., 1985), and then centrifuged at 112,700 × g for 25 min at 4 °C. The virus-containing milky band was extracted from the sucrose gradient, and diluted to 200 ml with 0.1 M sodium phosphate, pH 7.2. Virus particles were collected by centrifugation at 104,800 × g for 3 h at 4 °C. The pellet was resuspended in

500 μl PBS (0.1 M NaCl, 2.0 mM KCl, 10 mM Na2HPO4, 1.3 mM KH2PO4, pH 7.4).

RNA was isolated from the MMV virion preparations using the ToTALLY RNA isolation kit (Ambion, Austin, TX) and concentrations were measured using the

RiboGreen RNA quantitation assay (Molecular Probes Inc., Eugene, OR) according to manufacturers’ directions. Prior to library construction, the RNA was brought to 0.4

μg/ml glycogen (Invitrogen Corp., Carlsbad, CA) and precipitated using sodium acetate and ethanol.

3.3.2. cDNA library construction

The SuperScript Choice System for cDNA Synthesis (Invitrogen Corp.) was used to construct a cDNA library from viral RNA following the manufacturer's instructions.

Briefly, first strand cDNA was prepared from 4.4 μg total RNA by random priming with

125 ng random hexamers. After second strand cDNA synthesis, the double-stranded cDNA was ligated to 10 μg of EcoRI adapters, separated by column chromatography, and cloned into the EcoR1 site of calf intestinal phosphatase-treated pGEM4Z (Promega

Corp., Madison, WI). Ligations were introduced into Top Ten One Shot F’ Chemically

Competent E. coli cells (Invitrogen Corp.). Colonies were grown in 5 ml of Luria-Bertani broth containing 50 μg/ml ampicillin (Sigma, St Louis, MO). Plasmids were isolated using the QIA Miniprep spin kit (Qiagen, Valencia, CA), and digested with EcoR1 and

HindIII to determine insert presence and size. DNA concentrations were measured by

PicoGreen dsDNA quantitation assay (Molecular Probes Inc.) according to the manufacturer’s directions.

3.3.3. Sequencing and sequence analysis

To initially evaluate library quality, ten colonies were selected to determine insert presence, and eight of these clones were sequenced from both directions. Subsequently, plasmids from an additional 100 clones were digested to determine the average insert size in the library. Of these, 94 plasmid inserts were sequenced from both directions.

Sequencing was performed at the Plant-Microbe Genomics Facility at The Ohio State

University (Columbus, OH) using an ABI 3700 DNA Sequence Analyzer and Big Dye

Terminator Cycle Sequencing chemistry (Applied Biosystems Inc., Foster City, CA).

Sequence quality scores were obtained using McPhred/McPhrap (CodonCorp Corp.,

Dedham, MA). Removal of low quality and vector sequence, and contig assembly were

conducted with Sequencher (Gene Codes Corp., Ann Arbor, MI). At the 5’ end, sequences

were trimmed until the first 15 bases contained less than three bases with confidence

levels less than 20. At the 3’ end, sequences were trimmed until the last 15 bases

contained less than three bases with confidence levels less than 20. The average length

was calculated for good quality reads of greater than 100 bp. Subsequently these reads

were assembled into contigs using the Dirty Data algorithm with a minimum match of

85% and a minimum overlap of twenty. The average length of good quality reads that assembled into contigs was calculated.

Open reading frame (ORF) analysis was performed using both MacVector 6.5.3

(Oxford Molecular Group, Oxford, UK) and the ORF Finder database at the NCBI

website. The six ORFs identified in the MMV genome were compared, using BLASTx

database, to the non-redundant (nr) protein database of GenBank. The MMV genome

sequence was submitted to GenBank under accession number AY618418.

3.4. Results and discussion

3.4.1. cDNA library construction and analysis of sequences

MMV virions from three independent isolations totaling 2.4 mg protein were

combined for RNA isolation generating 11 μg of total RNA. Of this, 4.4 μg were used for

randomly primed cDNA library construction. The cDNA library yielded 9,065 clones

with an average insert size of 1,500 [standard deviation (S.D.) = 780 bp] based on EcoR1 and HindIII digestions of 100 plasmids (Table 3.1). Initially, inserts of eight plasmids were sequenced from both directions using SP6 and T7 primers, and of these, six insert sequences had similarities to genome sequences from other rhabdoviruses. Subsequently,

94 more clones from the MMV cDNA library were sequenced from the forward (SP6 primer) and reverse (T7 primer) directions.

After removal of low quality sequence and vector sequences, 163 of the 204

sequences (80%) contained reads of more than 100 bp, and were used for further analysis

(Table 3.1). Thus, only 20% of the sequence reads were below 100 bp after trimming the

vector sequences. The average length of a read was 704 (S.D. = 194 bp).

Assembly of the insert sequences of more than 100 bp resulted in four contigs made

up of 158 sequences and five singletons. The largest contig of 12,133 bp consisted of 97

sequence reads with an average length of 660 bp (S.D. = 180 bp). A BLASTx analysis of

this contig against the nr protein database of GenBank revealed significant similarity to

the polymerase (L) protein of the plant rhabdoviruses: Rice yellow stunt virus (RYSV)

(E-value: 0.00), Sonchus yellow net virus (SYNV) (E-value: e-168) and Northern cereal

mosaic virus (NCMV) (E-value: e-130), indicating that the contig represented the MMV

genome sequence. This 12,133 nt sequence of the MMV genome had a six-fold average

coverage, and both strands were sequenced for 93% of the contig, while 5.0% of the sequence was derived from single reads of one strand and 2.0% was derived from multiple sequences on the same strand (data not shown).

Five singlet sequences remained after contig formation. BLASTx analysis of the

nucleotide sequences against GenBank nr database revealed that three singlets (635, 669

and 732 bp) were most similar to a ferritin like protein (E-value: 8e-69), an E. coli

hypothetical protein (E-value: 9e-35), and the L protein of MMV (E-value: 2e-78),

respectively, while the remaining two singlets had no significant similarities to the database. BLASTx database searches on three other smaller contigs (1,587, 1,700, and

2,968 bp) indicated a high level similarity with an rRNA intron encoding an endonuclease of Oryza sativa (E-value: 3e-24), MMV L protein (E-value: 0.0), and cytochrome P450

monooxygenase of Zea mays (E-value: e−130), respectively. These three contigs were

assembled from 62 sequences (37%) of the 163 sequences of more than 100 bp in length.

Thus, one singlet and one small contig contained fragments of the MMV genome

sequence. To investigate why these two were not assembled into the larger contig, these

sequences were examined further and revealed that the singlet was derived from a

chimeric insert composed of a segment of the MMV genome and a plant gene, and the

contig sequence was most likely derived from a misassembly as it was missing a

fragment corresponding to nucleotides 11,171 to 11,600 of the 12,133 bp contig.

3.4.2. Annotation of the MMV genome sequence

Six ORFs located on the antigenomic strand were identified in the 12,133 bp MMV

contig (Fig. 3.1) and showed a genome organization similar to those of other

rhabdoviruses (Table 3.2). The ORF sequences were compared with the GenBank

non-redundant (nr) protein database using BLASTx. The sequence of the largest ORF

(ORF6), located at the most distal portion of the genome, was most similar to the L

protein of RYSV and SYNV (E-values: 0.0), and ORF5 was most similar to the

glycoproteins (G) of RYSV (E-value: 2e-22). ORF1 was most similar to the nucleocapsid proteins (N) of RYSV (E-value: 5e-48) and Maize fine streak virus (MFSV) (E-value:

3e-30). The deduced protein sequences of the remaining three ORFs were not closely

related to any proteins in GenBank. However, based on gene order in other rhabdoviruses

and ORF size (Redinbaugh and Hogenhout, 2005), ORF2 likely encodes the

phosphoprotein (P) and ORF4 possibly encodes the matrix protein (M). Thus, the most

likely gene order of the MMV genome was 3’-N-P-3-M-G-L-5’ (Fig. 3.1). Nucleotide

lengths of these MMV ORFs were 1,344 nt for N, 810 nt for the tentative P, 861 nt for

ORF 3, 708 nt for the tentative M, 1,776 nt for G, and 5,769 nt for L. The calculated

molecular weights of the proteins based on the deduced protein sequences would be ~60

kDa for the N protein, ~36 kDa for the P protein, ~38 kDa for the ORF3 protein, ~31 kDa

for the M protein, ~80 kDa for the G protein, and ~259 kDa for the L protein.

The basic gene order of a rhabdovirus genome is 3’ leader, N, P, M, G, L, and 5’

trailer (Hogenhout et al., 2003). Thus, MMV had an additional gene, ORF3, located between the P and M genes. The genome structure of MMV most closely matched those

of SYNV and Lettuce necrotic yellows virus (LNYV), which also have an additional ORF

between the P and M genes (Table 3.2). In SYNV, this additional gene is known as sc4,

which is involved in cell-to-cell movement in plants (Scholthof et al., 1994), and this may

also be the role of the ORF3 protein product of MMV.

Interestingly, the genome organization of MMV was different from those of two

other cereal-infecting nucleorhabdoviruses, MFSV and RYSV (Table 3.2). MFSV contained two extra genes, 3 and 4, which were expressed in MFSV-infected plants (Tsai et al., 2005). The RYSV genome sequence also contained an additional ORF, gene 3, between the P and M genes, and in addition had a second additional ORF, gene 6, located between the G and L genes (Chen et al., 1998; Huang et al., 2003). The RYSV P6 is a phosphorylated structural protein that is expressed in insects, but not in plants (Huang et al., 2003) suggesting a role of this protein in insect transmission. Insect specific

expression of other plant rhabdovirus sequences remains to be tested.

Clones corresponding to the 3’ and 5’ termini of the MMV genome were not

sequenced with the shotgun approach. By comparison to other nucleorhabdoviruses

(RYSV, SYNV, and MFSV), it is likely that the MMV genome sequence lacks 20–80 nt

of the leader sequence at the 3’end and 30–100 nt of trailer sequence at the 5’ end. The

inefficiency of cloning the genomic termini may not be surprising as other plant

rhabdoviruses have potentially stable secondary structures at the 5’ and 3’ ends of the

genomic RNA (Choi et al., 1994; Tanno et al., 2000; Tsai et al., 2005; Wetzel et al., 1994).

We will determine the terminal sequences of MMV by RACE (Tsai et al., 2005).

The gene junction sequences between rhabdovirus ORFs contain a conserved region

that correspond to the polyadenylation signature at the end of a gene sequence, the intergenic region and the transcriptional start site of the next gene. For MMV, the conserved gene junction sequence was 3’-AUUCUUUUUGGGUUG-5’ (Fig. 3.2A), and was similar to those of animal and other plant rhabdoviruses (Fig. 3.2B). Among

rhabdoviruses, the polyadenylation signatures consist of 8–9 nucleotides of (U)n

interspersed with an occasional A or a C, the intergenic regions are most variable in

length and sequence but usually contain one or more Gs, and the transcription start sites

contain the three consensus nucleotides, UUG, except for the two cytorhabdoviruses

(NCMV and LNYV), which have CUA and CUU transcription start sites (Fig. 3.2B). As

expected, the MMV gene junction sequence was most similar to those of RYSV and

SYNV (Fig. 3.2B).

3.4.3. Time and cost analysis of the shotgun and primer walking sequencing

procedures

To determine the effectiveness of the shotgun sequencing of a viral genome, the time

and cost of the shotgun sequencing procedure of the MMV genome was compared to that of the slightly larger MFSV genome (Tables 3.3 and 3.4), which was recently sequenced using the traditional primer walking method (Tsai et al., 2005). Shotgun sequencing of the MMV genome took a total of 127 h, whereas primer walking sequencing of the

MFSV genome took 217 h (Table 3.3). For cost analysis, personnel costs were calculated

at $ 15.86/h including employee benefits as 30% of salary. Supply costs were derived

from a lab supply budget of $ 10,000 per 260-day (2,080 h) work year multiplied by work

hours, and did not include computer programs or large equipment used in the project

since none were purchased for this purpose. Further, costs of the cDNA library kit, the cloning vector, and the competent E. coli cells were calculated separately under the item

“cDNA library costs” that were the same for the shotgun and primer walking methods.

The supplies costs for MFSV also included $ 350 for primers and $ 50 for PCR cloning

materials. Sequencing costs included costs for 204 reactions and was higher for the

MFSV genome, because the price for individual sequence reaction is $ 10/reaction as

opposed to $ 8/reaction for a 96 well plate. The total cost to sequence the 12,133-nt

MMV and 13,782-nt MFSV genomes was $ 0.38 and 0.51, respectively (Table 3.4).

In summary, in this manuscript we show that shotgun sequencing of a rhabdovirus

genome is possible while being an affordable and quick method in comparison to the

traditional primer walking method and, in addition, gives a good depth of genome

coverage. The shotgun sequencing allowed for quick and almost complete coverage of

the MMV genome although MMV is difficult to purify and MMV preparations had

significant plant contamination. At this time, only six of the ~90 plant rhabdovirus

genomes have been partially or completely sequenced. The shotgun sequencing technique

may facilitate the sequencing of more rhabdovirus genomes. This genome sequence

information will lead to a better understanding of rhabdovirus evolution and factors that

determine the host range of these viruses. This is important, because despite their small

genomes, rhabdoviruses are pathogens of humans, animals, insects and plants

(Hogenhout et al., 2003). Further, arthropods, such as mosquitoes, flies and plant-feeding insects, transmit many rhabdoviruses and may initiate new outbreaks by introducing rhabdoviruses among vertebrate and plant hosts.

3.5. Acknowledgements

We thank Angela Strock and John Abt for maintaining P. maidis populations and

MMV-infected plant material. This project was supported by the National Research

Initiative of the USDA Cooperative State Research, Education and Extension Service, grant number 2002-35302-12653.

3.6. References

Ammar, E.D. (1994) Propagative transmission of plant and animal viruses by insects: factors affecting vector specificity and competence. In Harris, K.F. (ed.), Advances in Disease Vector Research. Springer-Verlag, New York, pp. 289-331.

Ammar, E.D. and Nault, L.R. (2002) Virus transmission by leafhoppers, planthoppers and treehoppers (Auchenorrhyncha, Homoptera). Adv. Bot. Res., 36, 141-167.

Ammar, E.D. and Nault, L.R. (1985) Assembly and accumulation sites of maize mosaic virus in its planthopper vector. Intervirology, 24, 33-41.

Chen, X.Y., Luo, Z.L. and Fang, R.X. (1998) Structure analysis of the rice yellow stunt rhabdovirus gene 3. Chinese Sci. Bull., 43, 745-748.

Choi, T.J., Wagner, J.D. and Jackson, A.O. (1994) Sequence analysis of the trailer region of sonchus yellow net virus genomic RNA. Virology, 202, 33-40.

Creamer, R. (1992) Purification and protein characterization of sorghum stunt mosaic rhabdovirus. Phytopathology, 82, 1473-1476.

Falk, B.W. and Tsai, J.H. (1985) Serological detection and evidence for multiplication of maize mosaic virus in the planthopper, Peregrinus maidis. Phytopathology, 75, 852-855.

Falk, B.W. and Tsai, J.H. (1983) Physicochemical characterization of maize mosaic virus. Phytopathology, 73, 1536-1539.

Fleischmann, R.D., Adams, M.D., White, O., Clayton, R.A., Kirkness, E.F., Kerlavage, A.R., Bult, C.J., Tomb, J.F., Dougherty, B.A. and Merrick, J.M. (1995) Whole-genome random sequencing and assembly of Haemophilus influenzae Rd. Science, 269, 496-512.

Fraser, C.M., Eisen, J.A., Nelson, K.E., Paulsen, I.T. and Salzberg, S.L. (2002) The value of complete microbial genome sequencing (you get what you pay for). J. Bacteriol., 184, 6403-6405.

Hogenhout, S.A., Redinbaugh, M.G. and Ammar, E.D. (2003) Plant and animal rhabdovirus host range: a bug's view. Trends Microbiol., 11, 264-271.

Jackson, A.O., Milbrath, J.M. and Jedlinski, H. (1981) Rhabdovirus diseases of the Graminea. In Gordon, D.T., Knoke, J.K. and Scott, G.E. (eds.), Virus and Virus like Diseases of Maize in the United States. Southern Cooperative Series Bulletin 247. Ohio Agricultural Research and Development Center, Wooster, Ohio, pp. 51-76.

Kaiser, O., Bartels, D., Bekel, T., Goesmann, A., Kespohl, S., Puhler, A. and Meyer, F. (2003) Whole genome shotgun sequencing guided by bioinformatics pipelines - an optimized approach for an established technique. J. Biotechnol., 106, 121-133.

McDaniel, L.L., Ammar, E.D. and Gordon, D.T. (1985) Assembly, morphology, and accumulation of a Hawaiian isolate of maize mosaic virus in maize. Phytopathology, 75, 1167-1172.

Redinbaugh, M.G. and Hogenhout, S.A. (2005) Plant rhabdoviruses. Curr. Top. Microbiol. Immunol., 292, 143-163.

Scholthof, K.B., Hillman, B.I., Modrell, B., Heaton, L.A. and Jackson, A.O. (1994) Characterization and detection of sc4: a sixth gene encoded by sonchus yellow net virus. Virology, 204, 279-288.

Skaf, J.S. and Carroll, T.W. (1995) Purification of barley yellow streak mosaic virus and detection by DAS-ELISA and ISEM using polyclonal antibodies. Plant Dis., 79, 1003-1007.

Tanno, F., Nakatsu, A., Toriyama, S. and Kojima, M. (2000) Complete nucleotide sequence of Northern cereal mosaic virus and its genome organization. Arch. Virol., 145, 1373-1384.

Wetzel, T., Dietzgen, R.G. and Dale, J.L. (1994) Genomic organization of lettuce necrotic yellows rhabdovirus. Virology, 200, 401-412.

Number of clones in the MMV cDNA library 9,065 Average insert size (bp)a 1,500 ± 780 Number of inserts sequenced from both directions 102 Number of good quality sequence reads 163 Number of failed reactions 41 Average good quality sequence read length (bp) 704 ± 194 Total number of contigs 4 Sequence reads assembled into the MMV contig 97 Sequence reads that assembled into other contigs 62 Sequence reads that did not assemble into contigs 5 MMV contig length after assembly (bp) 12,133 MMV genome depth of coverage Six-fold a The average size of the insert was calculated based on fragments released after digestion of plasmids with EcoRI and HindIII. Data presented are the mean ± S.D.

Table 3.1. Results of shotgun library construction, sequencing, and assembly

Virus Gene order VSIV 3’ N P M G L 5’ Nucleorhabdovirus MMV 3’ N P 3 M G L 5’ MFSV 3’ N P 3 4 M G L 5’ SYNV 3’ N P sc4 M G L 5’ RYSV 3’ N P 3 M G 6 L 5’

Cytorhabdovirus LNYV 3’ N P 4b M G L 5’ NCMV 3’ N P 3 4 5 6 M G L 5’

Abbreviations: VSIV, Vesicular stomatitis Indiana virus; MMV, Maize mosaic virus; MFSV, Maize fine streak virus; SYNV, Sonchus yellow net virus; RYSV, Rice yellow stunt virus; LYNV, Lettuce necrotic yellows virus; and NCMV, Northern cereal mosaic virus.

Table 3.2. Comparison of the genome organizations of the livestock pathogen VSIV, and the plant infecting nucleorhabdoviruses and cytorhabdoviruses

Shotgun (MMV)a Primer walking (MFSV)ab Virus isolation 24 24 cDNA library construction 25.5 25.5 First round of colony picking and library quality analysis 7.5 7.5 First round of sequence analysis 4 20 Colony picking and plasmid isolation for the 96-well plates 46 NA Sequence and assembly of sequences from 96-well plates 20 NA Design of primers for five rounds of primer walking steps NA 25 Cloning of PCR products NA 15 Sequence analysis and assembly of sequences of a total of 12 cDNA NA 100 clones and 4 PCR products in the five primer walking steps Total hours 127 217 a Time is indicated in hours. b The MFSV genome sequence is described in Tsai et al., 2005. J. Virol. 79, 5304–5314. NA, not applicable.

Table 3.3. Time comparison of the shotgun and primer walking sequencing procedures

Shotgun (MMV) Primer walking (MFSV) Personnel cost, including 30% benefits ($ 15.86/h) ($) 2,014 3,442 Cost of supplies ($ 10,000 for 260 work days (2,080 h per year) ($) 611 1,043 Cost of cDNA library constructiona ($) 265 265 Cost of sequencingb ($) 1,696 1,670 50 Primers ($) NA 350 PCR cloning materials ($) NA 50 Total cost ($) 4,586 6,820 Sequencing cost per basec ($) 0.38 0.50 a Includes the costs of one reaction of the cDNA library kit, the cloning vector, and the competent E. coli cells. b The sequence costs for MMV includes 96 inserts sequenced from both directions (192 reactions) at $ 8/reaction for the 96 well plate and eight inserts from both directions (16 reactions) at $ 10/reaction, and the sequence costs for MFSV includes 167 reactions (i.e. 132 good quality reads, and 35 unsuccessful sequence reactions) at $ 10/reaction. c Genome lengths excluding the 5’ and 3’ ends were 12,133 nt for MMV (this study) and 13,782 nt for MFSV (Tsai et al., 2005. J. Virol. 79, 5304–5314.); NA, not applicable.

Table 3.4. Cost comparison of the shotgun and primer walking sequencing procedures

Fig. 3.1. Schematic representation of the genome organization of Maize mosaic virus. Figure details were derived from MacVector using the sequences assembled with Sequencher. Abbreviations: N, nucleocaspid protein; P, phosphoprotein; 3, deduced protein of open reading frame 3; M, matrix protein; G, glycoprotein; and L, polymerase protein; nt, nucleotides; MW, molecular weight.

Fig. 3.2. Comparative sequence analysis of gene junctions of rhabdovirus genomes. A. Gene junctions of the MMV genome. The three motifs are indicated and correspond to the 3’ end of the mRNAs (1), intergenic sequences (2), and the 5’ end of the mRNAs (3). B. Consensus sequences of gene junctions of plant and animal rhabdoviruses. All sequences are presented in genomic sense in the 3’ to 5’ orientation, and nucleotide differences are underlined. Abbreviations: 3’ le, 3’ leader sequence; 5’ tr, 5’ trailer sequence; (N)n, nucleotide type and number may vary. RABV, Rabies virus.

CHAPTER 4

SUBCELLULAR LOCALIZATION AND NUCLEAR IMPORT OF MAIZE FINE

STREAK VIRUS AND MAIZE MOSAIC VIRUS PROTEINS1

1The manuscript is in preparation: Chi-Wei Tsai, Valdir R. Correa, Margaret G.

Redinbaugh, and Saskia A. Hogenhout. Subcellular localization and nuclear import of

Maize fine streak virus and Maize mosaic virus proteins.

4.1. Abstract

Maize fine streak virus (MFSV) and Maize mosaic virus (MMV) are

insect-transmitted nucleorhabdoviruses. These two viruses do not only replicate in their

maize hosts but also replicate in their vector insects. In both hosts, nucleorhabdoviruses replicate in the nucleus and assemble at the inner nuclear membrane, and therefore

nuclear import of viral proteins is critical to complete morphogenesis. The Importin α and

β dependent nuclear import machinery is highly conserved from yeast to higher plants

and mammals, so we hypothesized that plant nucleorhabdoviruses interact with

conserved nuclear import pathways of plant and insect cells enabling these viruses to

infect both hosts. To begin with addressing this hypothesis, the role of Importin αs in

nuclear import of MFSV and MMV proteins in plant cells was studied by virus-induced

gene silencing (VIGS) combined with in planta cellular localization experiments. The

results showed that silencing of Importin αs in plant cells inhibited the nuclear

localization of the MFSV N protein and the MFSV N-P complex in plant cells,

suggesting that the MFSV N protein and the MFSV N-P complex are dependent on

Importin αs for nuclear import in plants. The cellular localization of the MMV proteins in

plant cells was studied as well. These results showed that the MMV N protein was

distributed throughout the cell, whereas the MMV P protein targeted the nucleus.

Surprisingly, the nuclear targeting of the MMV P protein was not dependent on Importin

αs. Co-introduction of the MMV N and P proteins resulted in the distribution of both

proteins throughout the cell. Thus, the MMV P protein followed the distribution of the

MMV N protein. To determine how the distribution of the MFSV N and P proteins in

plant cells compared to that in insect cells, fluorescent protein fusions were introduced into drosophila S2 cells. This showed that the MFSV N protein and the MFSV N-P complex also targeted the nuclei of insect cells, consistent with the hypothesis that the

MFSV proteins interact with conserved nuclear import pathway of plants and insects.

Studies to determine whether the Importin αs are involved in nuclear import of the MFSV

N protein and the MFSV N-P complex into the nuclei of drosophila cells are ongoing.

4.2. Introduction

Many rhabdoviruses are economically important pathogens of humans, livestock,

fish, and crops, of which Rabies virus is probably the most well known species (Rose and

Whitt, 2001). The rhabdoviruses that cause rabies and fish diseases appear to be confined

to vertebrate hosts, whereas vesiculo-, ephemero-, cyto-, and nucleo-rhabdoviruses are

transmitted to their vertebrate or plant hosts by insects (Ammar and Nault, 1985;

Hogenhout et al., 2003; Jackson et al., 2005; Rose and Whitt, 2001). Rhabdoviruses also infect and replicate in their insect vectors. Thus, most rhabdoviruses have two natural

hosts: either insects and plants, or insects and vertebrates. Plant-infecting rhabdoviruses

are transmitted by aphid, leafhopper or planthopper vectors, and vertebrate-infecting

rhabdoviruses are transmitted to animal hosts by black flies, sand flies, midges, and

mosquitoes (Hogenhout et al., 2003; Jackson et al., 2005; Rose and Whitt, 2001).

Plant-infecting rhabdoviruses are detected in all tissues of the insect and plant hosts

(Hogenhout et al., 2003; Jackson et al., 2005; McDaniel et al., 1985). There are two

genera of plant rhabdoviruses, Nucleorhabdovirus and Cytorhabdovirus, depending on

the site of replication and morphogenesis in plant cells (Jackson et al., 2005; Walker et al.,

2000). Nucleorhabdoviruses replicate in the nucleus, bud through the inner nuclear

membrane, and accumulate in the perinuclear space of plant cell. Cytorhabdoviruses

replicate in the cytoplasm and undergo morphogenesis from the endoplasmic reticulum, and accumulate in the cytoplasm of plant cell, similarly to vertebrate-infecting rhabdoviruses. Maize fine streak virus (MFSV) and Maize mosaic virus (MMV) are insect-transmitted nucleorhabdoviruses (Jackson et al., 2005; Redinbaugh et al., 2002).

Nucleorhabdoviruses replicate in the nucleus and assemble at inner nuclear membrane, and therefore nuclear import of viral proteins is critical to complete morphogenesis

(Walker et al., 2000). We hypothesized that plant rhabdoviruses interact with conserved nuclear import pathways of plant and insect cells enabling these viruses to infect diverse hosts.

Nuclear import of plant rhabdovirus proteins is mediated by Importin α/β (Goodin et

al., 2001; Jackson et al., 2005). Two key cellular processes are separated by the nuclear

envelope: transcription takes place in the nucleus, whereas proteins are synthesized in the

cytoplasm. This implies that many macromolecules must be exchanged constantly

between the nucleus and the cytoplasm. Nuclear import of nuclear localization signal

(NLS)-containing proteins is initiated by the specific recognition of the NLS of a

karyophilic protein by the NLS receptor (also called adapter) Importin α in the cytoplasm

(Adam and Gerace, 1991; Gorlich et al., 1994). The nuclear import receptor Importin β

binds co-operatively to the cargo protein-Importin α complex in the absence of Ran-GTP

(Gorlich et al., 1995ab). The ternary complex (cargo protein-Importin α-Importin β) then

docks to the nuclear pore complex via Importin β (Gorlich et al., 1995ab; Moroianu et al.,

1995; Radu et al., 1995). After translocation into the nucleus, the ternary complex is dissociated by the action of nuclear GTPase Ran-GTP, which is present in high concentrations in the nucleus (Izaurralde et al., 1997; Kutay et al., 1997b). This leads to the release of Importin α and the import cargo protein into the nucleoplasm. Importin α and β are recycled back to the cytoplasm in a complex with the Exportin CAS and

Ran-GTP (Izaurralde et al., 1997; Kutay et al., 1997ab). Importin α and β are then ready for a new round of nuclear import. In some cases, Importin β can also interact directly with NLS-containing proteins (Palacios et al., 1997; Tiganis et al., 1997).

This Importin α and β dependent nuclear import machinery is highly conserved from yeast to higher plants and mammals (Goldfarb et al., 2004). Our first objective is to determine the cellular localization of MFSV and MMV proteins in plant and insect cells.

Our second objective is to examine whether nuclear import of MFSV proteins is dependent on Importin α. Virus-induced gene silencing (VIGS) has been used widely in plants for analysis of gene function (Baulcombe, 1999; Lu et al., 2003). RNA interference

(RNAi) has been widely applied to knockdown genes in model organisms for functional analysis (Boutros et al., 2004). We employed VIGS to study the role of Importin αs in nuclear import of MFSV and MMV proteins in plant cells.

4.3. Materials and methods

4.3.1. cDNA cloning for cellular localization

Construction of different fluorescent versions of the binary pGD vectors was described previously (Goodin et al., 2002; Tsai et al., 2005). For cloning MFSV and

100

MMV genes in the pGD derivatives, each full-length gene was RT-PCR amplified from

virus infected maize total RNA, and primers with overhanging restriction sites were

introduced to facilitate directional cloning into the binary vector pGDY or pGDmR as

previously described (Goodin et al., 2002). RT-PCR was performed using the

MasterAMP High Fidelity RT-PCR kit (Epicentre, Madison, WI). The recombinant

plasmids were transformed into electro-competent Agrobacterium tumefaciens strain

GV3101. Successful cloning was verified by DNA sequencing with vector specific

primers.

To allow production of YFP fused MFSV N protein and mDsRed fused MFSV P

protein in drosophila S2 cells, the fragments corresponding to YFP-MFSV-N and

mDsRed-MFSV-P were amplified from the pGDY:MFSV-N and pGDmR:MFSV-P

plasmids with primers annealing at the 5’ termini of YFP and mDsRed and 3’ termini of

the MFSV N and P genes, and cloned into the pMT/V5-His-TOPO vector (Invitrogen,

Carlsbad, CA). To serve as negative controls, the fragments corresponding to YFP and mDsRed were amplified from the pGDY and pGDmR plasmids with primers annealing at the 5’ and 3’ termini of YFP and mDsRed, respectively. Similarly, to obtain a positive control for nuclear localization in S2 cells, the portion corresponding to YFP fused simian virus 40 large T-antigen NLS (SV40 NLS) was amplified from pGDY:SV40 NLS plasmid

(Kanneganti et al., in press) with primers annealing at the 5’ terminus of YFP and 3’ terminus of YFP-SV40 NLS. PCR was performed using the ProofStart DNA polymerase

(Qiagen, Valencia, CA). Successful cloning was verified by DNA sequencing with vector specific primers.

101

4.3.2. Tobacco rattle virus (TRV)-induced gene silencing

Construction of the pTV00:NbImp-α1 and pTV00:NbImp-α2 for TRV-induced gene silencing of Nicotiana benthamiana were described previously (Kanneganti et al., in press). A. tumefaciens strain GV3101 carrying the binary TRV RNA1 construct

(pBINTRA6) and the TRV RNA2 construct (pTV00, pTV00:PDS [phytoene desaturase], pTV00:NbImp-α1 and pTV00:NbImp-α2 )were introduced into N. benthamiana by agroinfiltration as previously described (Goodin et al., 2002). Briefly, suspensions of transformed agrobacteria were adjusted to an optical density at 600 nm (OD600) of 1.0 in agroinfiltration buffer (10 mM MgCl2, 10 mM MES [morpholineethanesulfonic acid], pH

5.6), and acetosyringone was added to a final concentration of 150 μM. The cultures were maintained at room temperature for 3 h to allow for induction of the vir gene. Before agroinfiltration, the agrobacterial suspensions were mixed at a 1:1 ratio (RNA1:RNA2).

The agrobacterial suspension was infiltrated into leave cells of 3-week old plants using a

1-ml disposable syringe. Following agroinfiltration, plants were maintained in a greenhouse at 25 °C and 16 h photoperiod. The empty pTV00 vector was used as a negative control to monitor phenotypes resulting from TRV infection. The pTV00:PDS construct was used as a positive control to monitor and visualize the onset of gene silencing as evidenced by a severe photobleaching phenotype resulting from PDS silencing (Ratcliff et al., 2001). This phenotype was usually seen between 6-10 days after infiltration. At two weeks after agroinfiltration with the TRV constructs, leaves from all plants in the experiment were collected for total RNA isolation.

102

4.3.3. RT-PCR confirmation of gene silencing

At two weeks after agroinfiltration with the TRV constructs, the fourth leaf above

the infiltrated leaves in silenced and non-silenced N. benthamiana plants were collected

for total RNA isolation using the Trizol reagent (Invitrogen) according to the

manufacturer instructions. RT-PCR amplifications were carried out with 20 ng total RNA using the Qiagen (Valencia, CA) OneStep RT-PCR kit and primer pairs NbImp-α1-F

(5’-GCGctcgagccATGTCGCTGAGGCCGAATTCGAGAAC-3’) and NbImp-α1-R

(5’-GCGggatccGGGGACACACTCCAGCTTCAATAACAGC-3’) for the NbImp-α1

gene, and NbImp-α2- F (5’-GCGctcgagccATGTCTCTGAGACCAAGTGCTAG

GACGG-3’) and NbImp-α2-R (5’-GCGggatccAGGAGACGGATGCATGAGGAGCTC

AACCA-3’) for the NbImp-α2 gene. These primers anneal outside the region targeted for

silencing to ensure that the endogenous gene was tested. Integrity of total RNA was

controlled with primers Tub-F1 (5’-ATCGCATCCGAAAGCTTGCAG-3’) and Tub-R1

(5’-ACATCAACATTCAGAGCTCCATC-3’), which were specific for the constitutively

expressed tomato tubulin factor 1 gene. Because tubulin factor 1 gene was highly expressed in leaves, RT-PCR amplifications of this gene were carried out with 2 ng RNA templates each. PCRs were performed for 30 cycles.

4.3.4. Agroinfiltration for in planta cellular localization

The agroinfiltration procedures were the same as described in TRV-induced gene

silencing section, except for using agrobacterial suspension at OD600 of 0.6. For

co-infiltration of different fluorescent fused protein constructs, equal volumes of each

103

agrobacterial suspension were mixed prior to infiltration and infiltrated as described

above. For in planta cellular localization analysis, non-silenced, NbImp-α1 and α2

silenced N. benthamiana plants of the same age were used. The leaves to be infiltrated

were at similar growth stages as those of the PDS-silenced leaves with the

photobleaching phenotype. Leaves were examined by laser-scanning confocal

microscopy between 40 h and 90 h post-infiltration. Experiments were conducted three times and each construct was tested in a total of 10 plants.

4.3.5. S2 cell culture and transfection

Maintenance and transient transfection of drosophila S2 cells were performed

following the instructions provided in the Drosophila Expression System manual

(Invitrogen). S2 cells were cultured and passaged every 3 or 4 days in Sf-900 II SFM

medium (Invitrogen) without Fetal bovine serum supplement. Drosophila S2 cells were

transfected with 10 μg Drosophila expression vector pMT/V5-His harboring YFP,

mDsRed, YFP-SV40NLS, YFP-MFSV N gene and mDsRed-MFSV P gene using the

calcium phosphate method. To induce protein synthesis under the control of the

metallothionein promoter, CuSO4 was added to the medium at a final concentration of

500 μM at 48 h after transfection. Cells were analyzed for fluorescence by laser-scanning

confocal microscopy between 24 h and 48 h post-induction.

4.3.6. Laser-scanning confocal microscopy

N. benthamiana leaves and drosophila S2 cells were examined under a Leica TCS

SP spectral laser-scanning confocal microscope. Drosophila S2 cells were fixed in 4%

104

paraformaldehyde with 0.1% Triton X-100 for 30 min at room temperature. Specimens

were subsequently stained the actin filaments with 6.6 μM of Alexa Fluor 633 Phalloidin

(Invitrogen) for 30 min at room temperature. YFP were excited at 488 nm, mDsRed was

excited at 568 nm, and Phalloidin was excited at 633 nm. The emitted fluorescence was

filtered through the trichronic 488/543/633, and detected at the following wavelengths:

510-550 nm for YFP, 575-620 nm for mDsRed, and 640-690 nm for Phalloidin. Images

were captured using Leica TCSNT software, and processed in Photoshop CS2 (Adobe,

San Jose, CA).

4.4. Results

4.4.1. Prediction of NLSs and cellular localization of the MFSV and MMV proteins

The presence of NLSs and cellular localization of the MFSV and MMV proteins were predicted by PSORT (http://psort.hgc.jp/) (Table 4.1). Although MFSV and MMV both undergo morphogenesis at inner nuclear membranes, the distribution of NLSs in both viral genomes was different. The N, P4, and M proteins of MFSV harbored NLSs, and the MFSV P, P3, G, and L proteins did not. In contrast, the N, P, G and L proteins of

MMV contain NLSs, and the MMV M and P3 proteins did not. The cellular localization

for the MFSV N, P, P3, P4, and M proteins are confirmed by in planta protein expression

using agroinfiltration (Tsai et al., 2005). The experimental results confirmed the PSORT

predictions. However, PSORT predicted that MFSV P locates to mitochondria, whereas

this protein is distributed throughout the cell. Even though the MMV G protein harbors

105

an NLS, PSORT predicted that the MFSV and MMV G proteins are plasma membrane

bound proteins because they have characteristic N-terminal signal peptides,

endopeptidase cleavage sites, and transmembrane anchor domains.

4.4.2. Cellular localization of the MMV proteins in plant cells

Because the cellular localizations of the MFSV proteins have already been studied

(Tsai et al., 2005), this section focuses solely on the localization of the MMV proteins. To

this end, the MMV genes were cloned into the pGD binary vector that creates fluorescent

protein fusion of the MMV proteins at the N-terminus. Agrobacteria carrying the pGDY:MMV-N, pGDY:MMV-M, pGDmR:MMV-P, or pGDmR:MMV-3 were infiltrated into the mesophyll of N. benthamiana leaves. As negative controls, agrobacteria carrying the empty vectors (pGDY and pGDmR) were used for agroinfiltration. The results showed that the YFP-MMV N, YFP-MMV M, and mDsRed-MMV P3 were distributed throughout the plant cells, (Fig. 4.1A). The mDsRed fused MMV P protein localized to the nuclei of plant cells (Fig. 4.1A). Thus, the nuclear targeting of the MMV proteins in plant cells was consistent with the presence of a NLS in the MMV P protein, and the absence of NLS in the MMV P3 and M proteins by PSORT. However, the two overlapping monopartite NLSs predicted by PSORT in the MMV N protein were apparently not functional as this N protein did not specifically target the nucleus in this study.

Because the interaction between the N and P proteins of MFSV and Sonchus yellow

net virus (SYNV) leads to subnuclear localization in plant cells (Goodin et al., 2002; Tsai

106

et al., 2005), we investigated the relocalization of the MMV N and P proteins by

coinfiltration of pGDY:MMV-N and pGDmR:MMV-P into the mesophyl of N. benthamiana leaves. The results suggest that small amount of the MMV P protein was distributed throughout the cell (Fig. 4.1B). Thus, the N and P proteins of MMV apparently interact as well, similarly to the N and P proteins of MFSV and SYNV. These results also indicate that the MMV P protein follows the distribution of the MMV N protein, and not the opposite (i.e. the N protein follows the distribution of the P protein).

Finally, we discovered that the cellular localization pattern of MMV proteins was totally different from that of MFSV and SYNV proteins in plant cells.

4.4.3. Nuclear import of the MFSV and MMV proteins is dependent on Importin αs

in N. benthamiana.

Analysis of the N. benthamiana EST database resulted in the identification of two genes with similarities to Importin αs of tomato, pepper, and Arabidopsis, and these two genes were targeted for silencing in N. benthamiana plants using the TRV-based VIGS system (Kanneganti et al., in press). This experiment was repeated with the ultimate goal to study whether the nuclear import of the MFSV and MMV proteins with NLSs are dependent on Importin αs for nuclear import. To verify the silencing of the N.

benthamiana Importin αs, RT-PCR was conducted using the same primers as described in

Kanneganti et al. (in press). RT-PCR results showed significantly lower levels of

amplification for NbImp-α1 and α2 transcripts in the TRV-NbImp-α1 and TRV-NbImp-α2

treated leaves than in control leaves infiltrated with empty pTV00 and pBINTRA6 and

107

non-infiltrated leaves. The amplification levels of the tubulin transcripts were similar in all samples (Fig. 4.2). Therefore, these RT-PCR results suggest that the NbImp-α1 and α2 genes were silenced in N. benthamiana leaves.

To examine whether nuclear import of the MFSV proteins is dependent on Importin

αs, NbImp-α1 and α2 silenced and control N. benthamiana leaves were infiltrated with A. tumefaciens cultures carrying the pGDY:MFSV-N or pGDmR:MFSV-P. The YFP fused

MFSV N protein was distributed throughout the cells in NbImp-α1 and α2 silenced leaves, but not in control leaves (Fig. 4.3A), indicating that the nuclear import of the MFSV N protein into the nuclei of plant cells is dependent on NbImp-α1 and α2. The distributions of mDsRed fused MFSV P protein were similar in NbImp-α1 and α2 silenced and control

N. benthamiana leaves (Fig. 4.3A), which was expected as the MFSV P protein does not specifically target the nuclei of plant cells (Tsai et al., 2005).

Coinfiltration of the MFSV N and P constructs results in relocalization of both proteins to the nucleolus of both proteins (Tsai et al., 2005). To examine whether the nuclear transport is also dependent on Importin αs, leaves of NbImp-α1 and α2 silenced and control N. benthamiana plants were coinfiltrated with A. tumefaciens carrying the pGDY:MFSV-N and pGDmR:MFSV-P. The YFP fused MFSV N protein and mDsRed fused MFSV P protein were distributed throughout the cells in NbImp-α1 and α2 silenced, but not control, N. benthamiana leaves (Fig. 4.3B). Thus, transport of the MFSV N-P complex into the nuclei of plant cells is dependent on Importin α1 and α2.

Similar experiments were conducted to assay for Importin αs dependence of nuclear import of the MMV proteins. The mDsRed fused MMV P protein localized in the nuclei

108

of cells in both NbImp-α1 and α2 silenced and control N. benthamiana plants (Fig. 4.4).

As expected, the YFP fused MMV N protein was distributed throughout the cells in both

NbImp-α1 and α2 silenced and control N. benthamiana plants (Fig. 4.4). This result

showed that nuclear import of the MMV P protein is not dependent on Importin α1 and

α2.

4.4.4. Cellular localization of the MFSV proteins in insect cells

MFSV and MMV also infect insects and bud from nuclear membrane in cells of

most tissues (Ammar and Nault, 1985; Hogenhout et al., 2003). To investigate whether

the proteins of these viruses also target the nuclei of insect cells, the localization of

fluorescent protein fusions of the MFSV N and P proteins, YFP-MFSV-N and

mDsRed-MFSV-P, were examined in drosophila S2 cells. To this end,

DES:YFP-MFSV-N or DES:mDsRed-MFSV-P along with the DES:YFP or

DES:mDsRed controls were transfected into drosophila S2 cells. The results showed that

the mDsRed fused MFSV P protein distributed throughout the insect cells, whereas the

YFP fused MFSV N protein localized to the nuclei of insect cells (Fig. 4.5). When the

YFP fused MFSV N proteins and the mDsRed fused MFSV P protein were transfected together into drosophila cells, the YFP fused MFSV N protein and mDsRed fused MFSV

P protein colocalized to the nuclei of drosophila cells (Fig. 4.5). Hence, the localization of MFSV proteins is similar between plant and insect cells suggesting that rhabdovirus proteins use a mechanism conserved among plants and animals for nuclear import.

109

4.5. Discussion

We discovered that the presence of NLSs is consistent with in vivo nuclear

localization data, and nuclear import of the MFSV N protein and the MFSV N-P complex

is dependent on Importin αs in plant cells by VIGS and in planta cellular localization

assay. Comparing the cellular localization of MFSV and MMV proteins, we showed that

the cellular localization of proteins of these two nucleorhabdoviruses is distinct even

though all viral proteins require import into the nuclei of infected cells for virus morphogenesis at the nuclear membrane.

MFSV and MMV are both nucleorhabdoviruses and undergo morphogenesis at the

inner nuclear membrane of plant cell, however the distribution and type of the NLSs are

different among the MFSV and MMV proteins. The presence of NLSs is consistent with

the phylogenetic relationships of five fully sequenced nucleorhabdoviruses: MFSV and

SYNV form a subgroup, and Rice yellow stunt virus (RYSV), MMV, and Taro vein

chlorosis virus (TaVCV) form the other subgroup. NLSs are classified into three

categories, which are SV40-like (monopartite), bipartite and Mat α2-like (reviewed in

Hicks and Raikhel, 1995). The MFSV and SYNV N proteins have a bipartite NLS at the

carboxyl termini of the proteins (Goodin et al., 2005; Tsai et al., 2005). The MMV and

TaVCV N proteins have monopartite NLSs and are predicted to localize in the cytoplasm

by PSORT with a certainty of 0.45. The M proteins of SYNV and MFSV have NLSs, but

those of RYSV, MMV, and TaVCV do not. The P proteins of MMV and RYSV contain

NLSs, but those of MFSV, SYNV and TaVCV do not. Therefore, it is likely that these

viruses also use different strategies for import of proteins into the nuclei of cells.

110

All nucleorhabdovirus structure proteins need to be imported into the nuclei of

infected cells for virus morphogenesis, except for the G protein. Synthesis of the

rhabdovirus G protein occurs at the rough endoplasmic reticulum, is processed for

glycosylation in the Golgi apparatus, and then is transported to nuclear membrane

(Jackson et al., 2005; Rose and Whitt, 2001). How do these viral proteins transport into

the nuclei of infected cells if they do not have NLSs? It seems most likely that viral

protein without their own NLS may enter the nuclei by co-transport with other viral

proteins that have NLSs. Co-introduction of the MFSV N and P proteins that both proteins localized to the nuclei of plant and insect cells is an example.

We show evidence that nucleorhabdoviruses use conserved mechanisms for

importing viral proteins into the nuclei of cells. The MFSV N and P proteins behave

similarly regarding nuclear import in N. benthamiana and drosophila cells; the MFSV N

protein localizes to the nucleus independently of the P protein, whereas the MFSV P

protein localizes specifically to the nucleus only when the P gene is coexpressed with the

N gene. Although most plant viruses replicate in the cytoplasm of host cells, several

groups of viruses replicate in the nuclei, such as nucleorhabdoviruses, geminiviruses, and

caulimoviruses (Krichevsky et al., 2006). For these nucleus-replicating viruses, nuclear

import is a crucial stage in the infection cycle. Several studies show that geminiviruses also use conserved mechanisms for importing viral proteins and genomes into the nuclei of plant and insect cells (Kunik et al., 1998; Liu et al., 1999; Sanderfoot and Lazarowitz,

1995; Sanderfoot et al., 1996).

111

Our results showed that nuclear import of the MFSV N protein and the MFSV N-P complex is dependent on Importin αs. The Importin α/β dependent nuclear import

machinery is highly conserved from yeast to higher plants and mammals (Goldfarb et al.,

2004). The Importin α gene family has undergone considerable expansion along the

eukaryotic evolution. Importin α2 and α3 groups only occur in metazoan animals and

perform cell and tissue specific roles in development and differentiation. Drosophila

melanogaster contain a single representative of each Importin α1, α2, and α3 group

(Goldfarb et al., 2004). N. benthamiana genome contains at least two copies of each

Importin α gene, NbImp-α1 and NbImp-α2 (Kanneganti et al., in press). These two genes

belong to α1 group of Importin αs and are paralogs of drosophila DmImp-α1 (Goldfarb et

al., 2004). Thus, it is likely that nuclear import of the MFSV N protein and the MFSV

N-P complex in drosophila S2 cells is dependent on DmImp-α1. Our future experiments

will be targeted toward the silencing of DmImp-α1 expression in order to determine

whether nuclear import of the MFSV N protein and the MFSV N-P complex is dependent

on DmImp-α1.

The nuclear import of the MFSV N protein and N-P complex is dependent on

Importin αs in plant cells. However, the nuclear import of the MMV P protein and N-P complex was not dependent on Importin αs in plant cells. Therefore, The MMV P protein must use another karyopherin adaptor or karyopherin for its nuclear import. Recent coimmunoprecipatation experiment indicated that the SYNV P protein interacts with yeast Impotin β (M. Deng and A.O. Jackson, personal communication). These results suggest that the MMV P protein may use Importin β for its nuclear import. Indeed, in

112

some cases, Importin β or Importin β-like proteins interact directly with NLS-containing proteins, such as hnRNP A1, snRNPs, and T-cell protein tyrosine phosphatase (Palacios et al., 1997; Pollard et al., 1996; Tiganis et al., 1997).

4.6. Acknowledgments

The project was supported by the National Research Initiative of the USDA

Cooperative State Research, Education and Extension Service Grant 2002-35302-12653, and the OARDC Research Enhancement and Competitive Grants Program.

113

4.7. References

Adam, S.A. and Gerace, L. (1991) Cytosolic proteins that specifically bind nuclear location signals are receptors for nuclear import. Cell, 66, 837-847.

Ammar, E.D. and Nault, L.R. (1985) Assembly and accumulation sites of maize mosaic virus in its planthopper vector. Intervirology, 24, 33-41.

Baulcombe, D.C. (1999) Fast forward genetics based on virus-induced gene silencing. Curr. Opin. Plant Biol., 2, 109-113.

Boutros, M., Kiger, A.A., Armknecht, S., Kerr, K., Hild, M., Koch, B., Haas, S.A., Paro, R., Perrimon, N. and Heidelberg Fly Array Consortium. (2004) Genome-wide RNAi analysis of growth and viability in Drosophila cells. Science, 303, 832-835.

Goldfarb, D.S., Corbett, A.H., Mason, D.A., Harreman, M.T. and Adam, S.A. (2004) Importin α: a multipurpose nuclear-transport receptor. Trends Cell Biol., 14, 505-514.

Gorlich, D., Kostka, S., Kraft, R., Dingwall, C., Laskey, R.A., Hartmann, E. and Prehn, S. (1995a) Two different subunits of importin cooperate to recognize nuclear localization signals and bind them to the nuclear envelope. Curr. Biol., 5, 383-392.

Gorlich, D., Prehn, S., Laskey, R.A. and Hartmann, E. (1994) Isolation of a protein that is essential for the first step of nuclear protein import. Cell, 79, 767-778.

Gorlich, D., Vogel, F., Mills, A.D., Hartmann, E. and Laskey, R.A. (1995b) Distinct functions for the two importin subunits in nuclear protein import. Nature, 377, 246-248.

Hicks, G.R. and Raikhel, N.V. (1995) Protein import into the nucleus: an integrated view. Annu. Rev. Cell Dev. Biol., 11, 155-188.

Hogenhout, S.A., Redinbaugh, M.G. and Ammar, E.D. (2003) Plant and animal rhabdovirus host range: a bug's view. Trends Microbiol., 11, 264-271.

114

Izaurralde, E., Kutay, U., von Kobbe, C., Mattaj, I.W. and Gorlich, D. (1997) The asymmetric distribution of the constituents of the Ran system is essential for transport into and out of the nucleus. EMBO J., 16, 6535-6547.

Jackson, A.O., Dietzgen, R.G., Goodin, M.M., Bragg, J.N. and Deng, M. (2005) Biology of plant rhabdoviruses. Annu. Rev. Phytopathol., 43, 623-660.

Kanneganti, T., Bai, X., Tsai, C.W., Win, J., Meulia, T., Goodin, M., Kamoun, S., and Hogenhout, S.A. 2006. A functional genetic assay for nuclear trafficking in plants. Plant J., In press.

Krichevsky, A., Kozlovsky, S.V., Gafni, Y. and Citovsky, V. (2006) Nuclear import and export of plant virus proteins and genomes. Mol. Plant Pathol., 7, 131-146.

Kunik, T., Palanichelvam, K., Czosnek, H., Citovsky, V. and Gafni, Y. (1998) Nuclear import of the capsid protein of tomato yellow leaf curl virus (TYLCV) in plant and insect cells. Plant J., 13, 393-399.

Kutay, U., Bischoff, F.R., Kostka, S., Kraft, R. and Gorlich, D. (1997a) Export of importin α from the nucleus is mediated by a specific nuclear transport factor. Cell, 90, 1061-1071.

Kutay, U., Izaurralde, E., Bischoff, F.R., Mattaj, I.W. and Gorlich, D. (1997b) Dominant-negative mutants of importin-β block multiple pathways of import and export through the nuclear pore complex. EMBO J., 16, 1153-1163.

Liu, H., Boulton, M.I., Thomas, C.L., Prior, D.A., Oparka, K.J. and Davies, J.W. (1999) Maize streak virus coat protein is karyophyllic and facilitates nuclear transport of viral DNA. Mol. Plant Microbe Interact., 12, 894-900.

Lu, R., Martin-Hernandez, A.M., Peart, J.R., Malcuit, I. and Baulcombe, D.C. (2003) Virus-induced gene silencing in plants. Methods, 30, 296-303.

McDaniel, L.L., Ammar, E.D. and Gordon, D.T. (1985) Assembly, morphology, and accumulation of a Hawaiian isolate of maize mosaic virus in maize. Phytopathology, 75, 1167-1172.

Moroianu, J., Hijikata, M., Blobel, G. and Radu, A. (1995) Mammalian karyopherin α1β and α2β heterodimers: α1 or α2 subunit binds nuclear localization signal and β subunit interacts with peptide repeat-containing nucleoporins. Proc. Natl. Acad. Sci. U.S.A., 92, 6532-6536.

Palacios, I., Hetzer, M., Adam, S.A. and Mattaj, I.W. (1997) Nuclear import of U snRNPs requires importin β. EMBO J., 16, 6783-6792.

115

Pollard, V.W., Michael, W.M., Nakielny, S., Siomi, M.C., Wang, F. and Dreyfuss, G. (1996) A novel receptor-mediated nuclear protein import pathway. Cell, 86, 985-994.

Radu, A., Blobel, G. and Moore, M.S. (1995) Identification of a protein complex that is required for nuclear protein import and mediates docking of import substrate to distinct nucleoporins. Proc. Natl. Acad. Sci. U.S.A., 92, 1769-1773.

Ratcliff, F., Martin-Hernandez, A.M. and Baulcombe, D.C. (2001) Tobacco rattle virus as a vector for analysis of gene function by silencing. Plant J., 25, 237-245.

Redinbaugh, M.G. and Hogenhout, S.A. (2005) Plant rhabdoviruses. Curr. Top. Microbiol. Immunol., 292, 143-163.

Rose, J.K. and Whitt, M.A. (2001) Rhabdoviridae: the virus and their replication. In Knipe, D.M. and Howley, P.M. (eds.), Fields virology. Lippincott Williams & Wilkins, Philadelphia, pp. 1221-1244.

Sanderfoot, A.A., Ingham, D.J. and Lazarowitz, S.G. (1996) A viral movement protein as a nuclear shuttle. The geminivirus BR1 movement protein contains domains essential for interaction with BL1 and nuclear localization. Plant Physiol., 110, 23-33.

Sanderfoot, A.A. and Lazarowitz, S.G. (1995) Cooperation in viral movement: The geminivirus BL1 movement protein interacts with BR1 and redirects it from the nucleus to the cell periphery. Plant Cell, 7, 1185-1194.

Tiganis, T., Flint, A.J., Adam, S.A. and Tonks, N.K. (1997) Association of the T-cell protein tyrosine phosphatase with nuclear import factor p97. J. Biol. Chem., 272, 21548-21557.

Protein MFSV MMV Nucleocapsid protein (N) 436 KR SSDGTGNVSK KKSRK 438 PAPKKTR 440 PKKTRSG 0.82 nucleus 0.45 cytoplasm Phosphoprotein (P) None 263 KRPR 154 PSIKRKA 0.64 mitochondrial matrix 0.30 nucleus/microbody ORF3 protein (P3) None None 0.64 microbody (peroxisome) 0.45 cytoplasm ORF4 protein (P4) 17 RK ALTKASKALF KGKIK 0.76 nucleus Matrix protein (M) 207 KRKR None 195 KK EDKAEKATTE KRKRQ 0.91 nucleus 0.65 cytoplasm Glycoprotein (G) None 432 RKKP 0.46 plasma membrane 0.46 plasma membrane Polymerase (L) None 375 KK NPRQSVLDEI RRQFK 0.70 plasma membrane 0.76 nucleus

Table 4.1. Predicted nuclear localization signal and cellular localization of Maize fine streak virus (MFSV) and Maize mosaic virus (MMV) proteins with PSORT

117

Fig. 4.1. A. Cellular localization of fluorescent protein fusions of the MMV N, P, P3 and M proteins in N. benthamiana leaves. Infiltrations of pGDY (YFP) and pGDmR (mDsRed) were included as negative controls. B. Cellular localization of fluorescent protein fusions of the MFSV N and P proteins in N. benthamiana leaves. Bar = 20 μm. All micrographs were taken at identical magnifications. Representative of more than three independent experiments is presented.

118

Fig. 4.2. RT-PCR analysis confirming the silencing of two NbImp-α homologs of N. benthamiana. NbImp-α silencing was performed by Tobacco rattle virus (TRV)-induced gene silencing. Products amplified from total RNA of non-infiltrated (non-silenced) plants, NbImp-α1-silenced plants, NbImp-α2-silenced plants, and TRV-infiltrated (non-silenced) plants were shown. Gene-specific primers that amplify both NbImp-α1 and α2 were used. As a control for RNA amount, the mRNA levels of tubulin factor 1 were examined. Representative of more than three independent experiments is presented.

119

Fig. 4.3. A. Cellular localization of the MFSV N and P proteins in the NbImp-α1 and α2 silenced N. benthamiana leaves. Nuclear import of the YFP fused MFSV N protein was NbImp-α1 and α2 dependent in N. benthamiana leaves. B. Cellular localization of the MFSV N-P complex in the NbImp-α1 and α2 silenced N. benthamiana leaves. Upper panel shows the overlay of YFP and DsRed signals. Lower panel shows the YFP signal. Transport of the MFSV N-P complex into the nucleus/nucleolus of plant cells is dependent on NbImp-α1 and α2 in N. benthamiana leaves. Bar = 20 μm. All micrographs were taken at identical magnifications. Representative of more than three independent experiments is presented.

120

Fig. 4.4. Cellular localization of the MMV N and P proteins in the NbImp-α1 and α2 silenced N. benthamiana leaves. Nuclear import of the MMV P protein was NbImp-α1 and α2 independent in N. benthamiana leaves. Bar = 20 μm. All micrographs were taken at identical magnifications. Representative of more than three independent experiments is presented.

121

Fig. 4.5. Cellular localization of fluorescent protein fusions of the SV40 NLS and the MFSV N and P proteins in drosophila S2 cells. Transfection of pGDY (YFP) and pGDmR (mDsRed) were included as negative controls. Panels A to D, expression pattern of YFP; D, overlay of micrographs B and C; Panels E to H, expression pattern of YFP:SV40 NLS; H, overlay of micrographs F and G; Panels I to L, expression pattern of YFP:MFSV-N; L, overlay of micrographs J and K; Panels M to P, expression pattern of mDsRed; P, overlay of micrographs N and O; Panels Q to T, expression pattern of mDsRed:MFSV-P; T, overlay of micrographs R and S; Panels U to X, expression pattern of coexpressed YFP:MFSV-N and mDsRed:MFSV-P; X, overlay of micrographs V and W. Bars = 10 μm. All micrographs were taken at identical magnifications. Representative of two independent experiments is presented.

122

CHAPTER 5

SIGMA RHABDOVIRUS ACTIVATES THE INNATE IMMUNE

RESPONSE OF DROSOPHILA1

1 The manuscript is in preparation: Chi-Wei Tsai, Elizabeth A. McGraw, Ralf G.

Dietzgen, and Saskia A. Hogenhout. Sigma rhabdovirus activates the innate

immune response of drosophila.

123

5.1. Abstract

Sigma rhabdovirus only infects Drosophila flies and is transmitted to the progeny of

flies through germinal cells. Rhabdoviruses are generally neurotropic and frequently

induce lethal diseases, for example Rabies virus mainly spreads through the central nervous system of humans and animals and causes slow painful deaths. Sigma virus also

spreads mainly through the nervous system of its drosophila host. However, compared to

healthy flies, Sigma virus-infected flies do not display disease symptoms in natural

environments, but become permanently paralyzed when exposed to high concentration of

CO2. Therefore, we hypothesized that drosophila mounts a potent immune response to restrict excessive rhabdovirus replication and spread. To test this hypothesis, hybridization experiments with RNAs from virus-infected and virus-free drosophila adults and cDNA microarrays that contain whole drosophila genome were conducted.

The Imd signaling pathway was identified as the main component of the drosophila anti-Sigma virus response, whereas there were no indications of activation of the Toll

pathway. This is in contrast with Drosophila C virus (DCV) and Drosophila X virus

(DXV), which mainly activate the Jak-STAT and Toll pathways, respectively. The different antiviral responses among these viruses were expected because data herein and other evidences suggested that Sigma virus inhibits the Toll pathway activation. Further,

Sigma virus has a lipid envelope with exposed glycoproteins, whereas DCV and DXV are

naked viruses with only protein capsids. Thus, viruses can induce different immune

pathways in drosophila, similarly to Gram-negative bacteria and Gram-positive bacteria,

which predominantly activate the Imd and Toll pathways, respectively. Nerve cell

developmental genes were also upregulated in Sigma virus-infected versus healthy flies.

124

This may enhance spread of viruses in its host and change the host’s behavior. These

findings and future studies of Sigma virus-drosophila interactions with will help to

elucidate innate immune response pathways against enveloped viruses in vertebrates, and elucidate how rhabdoviruses manipulate nerve cells to enhance spread.

5.2. Introduction

Sigma virus is a symbiont of Drosophila melanogaster and belongs to the family

Rhabdoviridae, which contains many economically important human, livestock, fish, and

plant pathogens (Teninges, 1999; Tordo et al., 2005). Rabies virus is arguably the most

well-known rhabdovirus and causes lethal disease in humans and animals. Even though

Rabies virus is not insect-transmitted, many rhabdoviruses are. Black flies, sand flies, and mosquitoes transmit Vesicular stomatitis virus (VSV) and Bovine ephemeral fever virus

(BEFV), and all plant rhabdoviruses have insect vectors, mostly aphids, leafhoppers, and planthoppers (Hogenhout et al., 2003; Rose and Whitt, 2001; de Mattos et al., 2001).

Insects are not only vectors but also replication hosts for rhabdoviruses. In addition, some rhabdoviruses are restricted to insect hosts. Sigma virus only infects drosophila and is transmitted to the progeny of flies through female and male germinal cells (Teninges,

1999). Many plant rhabdoviruses are also passed on to the progeny of their insect vectors

(Jackson et al., 2005; Nault, 1997).

Rhabdovirus infection is generally neurotropic in their vertebrate and invertebrate

hosts. Rabies virus spreads throughout the human and animal body via the central

nervous system and also infects other tissues (e.g. salivary glands) (de Mattos et al.,

2001). This results in behavioral changes of hosts, such as aggressive behavior and

125

increased salivation. Similarly, Sigma virus replicates in the nervous system of drosophila

as well as other tissues (Teninges, 1999), and plant rhabdoviruses also infect the nervous

system of their insect vectors (Ammar et al., 2004; Hogenhout et al., 2003; Ammar and

Hogenhout, Submitted). Nerve cell infection accounts for the characteristic CO2 sensitivity of Sigma virus-infected drosophila. Sigma virus-infected flies remain irreversibly paralyzed and die after CO2 anesthetization, whereas virus-free flies recover

(Brun, 1991; Teninges, 1999). Similarly, vesiculoviruses also confer CO2 sensitivity to

their black fly hosts (Brun, 1991).

Most of our knowledge on the drosophila innate immune response is based on

antibacterial and antifungal responses. Insects have several innate immune responses triggered by complex signaling pathways to combat invading microbes, including encapsulation, blood coagulation, melanization, phagocytosis, and antimicrobial peptide

(AMP) production (Leclerc and Reichhart, 2004; Steiner, 2004). The Toll and immune

deficiency (Imd) pathways regulate expression of genes encoding these AMPs through

activation of NF-κB family transcription factors (Reviewed in Hoffmann, 2003; Leclerc

and Reichhart, 2004; Royet et al., 2005). Fungal and most Gram-positive bacteria activate the Toll pathway that controls the gene expression of AMP genes via translocation of Dif

(NF-κB family) to the nucleus. In contrast, Gram-negative bacteria mainly activate the

Imd pathway through the peptidoglycan recognition protein (PGRP)-LC receptor

resulting in the expression of another set of AMP genes via translocation of cleaved

Relish (NF-κB family) to the nucleus.

Drosophila C virus (DCV, Family Dicistroviridae) and Drosophila X virus (DXV,

Family Birnaviridae) are two drosophila-infecting viruses that have been used for

126

molecular studies of virus-host interactions (Dostert et al., 2005; Roxstrom-Lindquist et

al., 2004; Zambon et al., 2005). Microarray studies have shown DCV infection induces

components of the Toll, Imd, and Janus kinase-signal transducer and activator of

transcription (Jak-STAT) signaling pathways (Dostert et al., 2005). The Toll and Imd

pathways are both activated in response to DXV infection (Zambon et al., 2005). Further,

genetic screens of drosophila immunity mutants identify the Toll pathway as an essential

component of the drosophila anti-DXV responses (Zambon et al., 2005). DCV and DXV

have RNA genomes encapsidated in non-enveloped icosahedral particles (Jousset et al.,

1977; Zambon et al., 2005). In contrast, Sigma virus is an enveloped virus and has one

type of transmembrane glycoprotein (G) protruding from the exterior of the lipid

envelope (Rose and Whitt, 2001). The host-derived lipid bilayer surrounds the

nucleocapsid core consisting of the negative-sense, single-stranded RNA genome,

nucleocapsid protein (N), polymerase-associated phosphoprotein (P), and multifunctional

polymerase (L) (Rose and Whitt, 2001). The matrix protein (M) connects the envelope to

the ribonucleocapsid core (Rose and Whitt, 2001). Because of these morphological

differences and neurotropism, we hypothesize that drosophila elicits different immune

responses to Sigma virus infection versus DCV and DXV infection.

Virion structures of Sigma virus and other rhabdoviruses are highly similar, and the majority of the rhabdoviruses are neurotropic. Because innate immunity is conserved from insects to mammals, drosophila response to infectious agents has been used as a model for studying innate immune systems in various organisms. We identified the Imd signaling pathway as a main component of the drosophila anti-Sigma virus responses.

The results presented will further assist to identify genes involved in innate immunity in

127

drosophila, and will lead to a better understanding of innate immune responses to enveloped viruses. In addition, this may help in the further to develop novel control strategies against insect borne viruses.

5.3. Materials and methods

5.3.1. Drosophila stocks and crossings

Fe strain (Sigma virus-infected) and CST strain (Sigma virus-free) D. melanogaster were used as starting stocks. To minimize genetic background effects, CST females were crossed with Fe males, and then the progeny females of each generation were backcrossed against Fe males for four generations to create a BC4 strain with 97% Fe background. Small portion of BC4 flies remained infected with Sigma virus because

paternal transmission is possible although it is less efficient than maternal transmission

(Teninges, 1999). Sigma virus-infected BC4 flies were removed from the population

using the CO2 sensitivity assay (described below). Fe strain (Sigma virus-infected) and

BC4 strain (Sigma virus-free) drosophila were used throughout this study. All drosophila

stocks were maintained at 25˚C, 70% humidity, and a 12 h light/dark cycle on standard

cornmeal/yeast media.

5.3.2. Sigma virus detection

Presence of Sigma virus in drosophila was determined by CO2 sensitivity assay and

N gene-specific reverse transcription-polymerase chain reaction (RT-PCR). For the CO2

sensitivity assay, flies were treated with CO2 gas and kept on ice for 10 min. Next, CO2

128

was replaced with fresh air, and flies were kept at room temperature for 10 min. The

Sigma virus-infected drosophila remained irreversibly paralyzed and died. In contrast, the

virus-free drosophila recovered from anesthetization after returning to fresh air and room

temperature. For RT-PCR detection of Sigma virus the primer pair SigV 33F

(5’-AGACACCAAGACCGAGTTCG-3') and SigV 527R (5’-TGCGTTGAAAAT

GGAGCGAG-3’) was designed to amplify a 500-bp fragment of the N gene using a

Qiagen (Valencia, CA, USA) One Step RT-PCR kit according to the manufacturer's

instructions. This RT-PCR assay detected Sigma virus infection in individual drosophila

flies.

5.3.3. RNA extractions and hybridizations

For microarray experiments, 3-day-old BC4 adult flies were treated with CO2 to remove Sigma virus-infected flies from virus-free flies. To prevent artifacts of CO2 treatment, Sigma virus-infected Fe adult flies with the same age were also treated with

CO2. Then, equal numbers of males and females of the virus-free BC4 and Sigma

virus-infected Fe flies were snap frozen in liquid nitrogen. Subsequently, the heads were

separated from the bodies (thorax and abdomen) by vortexing with glass beads and

passage through a 0.71 mm standard sieve. RNA was extracted using Trizol (Invitrogen,

Carlsbad, CA, USA) following the manufacturer’s instructions with an additional

chloroform extraction before isopropanol precipitation. The integrity and concentration of

the RNA was determined spectrophotometrically using a NanoDrop and associated

software version 1000 (NanoDrop Technologies, Wilmington, DE, USA). Prior to

labeling and hybridization an additional step of mRNA isolation was carried out using the

129

GeneElute mRNA mini prep kit (Sigma-Aldrich, St. Louis, MO, USA) according to the manufacturer’s instructions. A total of 4.0 µg of mRNA from paired Sigma virus-infected and uninfected lines was labeled using random hexamers and oligo dT and direct incorporation of fluorescently labeled (Cy3 and Cy5) nucleotides (GE Healthcare,

Chalfont St. Giles, UK) as previously described (Grimmond et al., 2000). Material was then hybridized to whole genome D. melanogaster arrays (White et al., 1999) containing replicate spots for 13,098 genes. Three replicate hybridizations were carried out for heads

only samples and bodies only samples.

5.3.4. Microarray analysis

Microarray slides were scanned with a GMS418 scanner (Genetic MicroSystems,

Woburn, MA, USA). Annotation was completed using SoftWorx Tracker (Applied

Precision, Issaquah, WA, USA), and microarray analysis was performed in GeneSpring

version 7 (Silicon Genetics Inc., Agilent Technologies, Palo Alto, CA, USA). A Lowess

normalization was applied to each chip. Spots were filtered by the presence of flags and

minimum expression level. Only profiles of genes meeting the criteria in all replicate

hybridizations are reported. An uncorrected p-value of 0.05 was used to determine

significance. This approach allowed for the identification of a broad class of candidate

genes whose expression could then be confirmed by additional methods [Northern blot

hybridization, quantitative reverse transcription real-time polymerase chain reaction

(qRT-PCR)]. All gene annotations are from FlyBase (http://flybase.bio.indiana.edu).

130

5.3.5. Northern blot hybridization

Total RNA (20 µg) from Sigma virus-infected and virus-free drosophila was

denatured using Glyoxal sample loading dye (Ambion, Austin, TX, USA), separated on a

1.2% agarose gel in 1x BPTE electrophoresis buffer {10 mM PIPES

[piperazine-N,N’-bis(2-ethanesulfonic acid)], 30 mM Bis-Tris, 1 mM EDTA, pH 6.5}, transferred to a positively charged BrightStar-Plus nylon membrane (Ambion) with 20x

SSC, and cross-linked to the membrane by exposure to UV light (UV Transilluminator;

Fisher Scientific, Pittsburgh, PA, USA). Probes were prepared by RT-PCR amplification

(primers used are listed in Table 5.1) of DNA fragments corresponding to each target

gene from drosophila total RNA using Qiagen OneStep RT-PCR kit and subsequently

incorporation of [32P] dCTP (PerkinElmer, Boston, MA, USA) using Ready-To-Go DNA

Labeling Beads (GE Healthcare) according to the manufacturers’ instructions.

Membranes were prehybridized in modified Church Buffer [0.5 M sodium phosphate (pH

7.0), 1 mM EDTA, and 7% SDS] at 65 oC for 16-18 hours. Next, membranes were

hybridized with heat-denatured, radiolabeled probes in modified Church Buffer at 65 oC

for another 16-18 hours. After hybridization, blots were washed three times for 10 min

each in 2x SSC and 0.1% SDS, four times for 10 min each in 0.1x SSC and 0.1% SDS at

65°C, and then exposed to the Storage Phosphor Screen (Molecular Dynamics,

Sunnyvale, CA, USA) for 24 hours. Images were captured and analyzed using Storm 840

Gel and Blot Imaging System and ImageQuant software (Molecular Dynamics),

converted to TIFF for export, and processed in Photoshop CS2 (Adobe, San Jose, CA,

USA).

131

5.4 Results

5.4.1. Overview of transcript changes in response to Sigma virus infection

Genome-wide transcript changes in response to Sigma virus infection were obtained

from transcription profile comparisons between virus-infected and virus-free drosophila.

Because Sigma virus is neurotropic, we decided to analyze transcript changes in the drosophila head, which is predominantly neural tissue, and the rest of the drosophila body

(thorax and abdomen) separately. In the heads only experiments, a total of 56 genes (out of 13,098 genes on the microarray) were upregulated, and 32 genes were downregulated in Sigma virus-infected versus uninfected drosophila (Table 5.2). In the bodies only

experiments, a total of 113 genes were upregulated, and 12 genes were downregulated in

Sigma virus-infected versus uninfected drosophila (Table 5.2). Only 13% of the

upregulated genes and 5% of the downregulated genes were shared between the heads

and bodies experiments (Table 5.2). Thus, most of these differential expressed genes were

head-specific or body-specific. Overall, Sigma virus induced more genes in bodies than in

heads, but more genes were suppressed in heads than in bodies. The repression of gene

expression in drosophila brain is consistent with the finding that the predominant effect

of Rabies virus infection is the repression of gene expression in mouse CNS (Prosniak et

al., 2001). When the heads and bodies data were pooled, approximately 25% of the

transcript changes were related to the defense response of drosophila, including immunity,

protease, and protease inhibitor genes (Fig 5.1). The results suggest that Sigma virus

infection affects drosophila gene expression, and most of the differentially expressed genes are involved in the defense response.

132

5.4.2. Confirmation of differential expressed genes

To validate Sigma virus infection induced immunity gene expressions, Northern

blots of total RNAs from Sigma virus-infected and virus-free drosophila were hybridized

with probes corresponding to drosophila immunity genes. Drosocin, Attacin, and

Diptericin showed strong upregulations in Sigma virus-infected drosophila, and

PGRP-SB1 and SD showed week upregulations (Fig 5.2). However, the differential expression of Jun-related antigen (Jra), Cabut, and PGRP-SC1 in the microarray experiments were not confirmed by Northern analysis (Fig 5.2). The expression of Toll pathway controlled Defensin, Drosomycin, and Metchnikowin were variable in Sigma virus-infected drosophila (data not shown). This may explain the nonsignificant differences of these gene expressions in the microarray experiments. These data suggest that Sigma virus infection mainly activates the Imd signaling pathway because the expression of Drosocin, Attacin, and Diptericin is under the control of the Imd pathway.

5.4.3. Comparison of immune response of drosophila infected with DCV, DXV, and

Sigma virus

Signaling pathways controlling drosophila humoral defense have been well

described, and recently, the antiviral responses to two drosophila viruses, DCV and DXV,

were analyzed (Dostert et al., 2005; Roxstrom-Lindquist et al., 2004; Zambon et al.,

2005). Unlike Sigma virus, DCV and DXV are not enveloped by a lipid bilayer and do

not have an exposed G protein. Hence, we investigated how the drosophila immune

system reacts to enveloped viruses (Sigma virus) versus non-enveloped viruses (DCV and

DXV).

133

Genes upstream of the Toll and Imd pathways were similarly regulated in Sigma

virus and DCV infections (Table 5.3). PGRP-SD was upregulated in Sigma virus-infected

drosophila, whereas PGRP-SA and Spatzle were upregulated after DCV infection (Table

5.3). PGRP-SA and SD are pattern recognition receptors detecting Gram-positive bacteria and triggering the activation of the Toll pathway involving the proteolytically cleaved

Spatzle (Bischoff et al., 2004; Michel et al., 2001; Royet et al., 2005). Neither Sigma virus nor DCV upregulated the expression of transmembrane receptors of the Toll and

Imd pathways, Toll and PGRP-LC, although Sigma virus infection upregulated

components of the Imd pathway, and DCV infection upregulated components of the Toll,

Imd, and Jak-STAT pathways (Dostert et al., 2005).

Sigma virus also upregulated a number of other PGRP genes, including PGRP-SB1,

SC1a and SC1b, that were not upregulated in DCV-infected drosophila (Table 5.3).

Sequence analysis categorizes the drosophila PGRPs into two subgroups, catalytic and

recognition PGRPs (Royet et al., 2005). PGRP-SB1, SB2, SC1 and SC2 belong to the

catalytic subgroup harboring zinc-binding residues that account for amidase activity

against peptidoglycan. PGRP-SA and SD belong to the recognition subgroup lacking

zinc-binding residues but retain peptidoglycan-binding ability. PGRP-SC1b cleaves

peptidoglycan to suppress the immune reaction suggests a possible immunomodulatory

role mediated by the amidase of these catalytic PGRPs (Mellroth et al., 2003). None of

the genes encoding known components of the RNA interference machinery were

upregulated by Sigma virus infection. Sigma virus may employ the immunomodulatory

function of the catalytic PGRPs to conquer the antiviral response in drosophila for both

the Imd and Toll pathways.

134

Comparison of upregulated genes that are components of the Imd signaling cascades

show that Sigma virus slightly upregulated Jra and Cabut (Table 5.3). These genes are part of the c-Jun N-terminal Kinase (JNK) pathway that forms a branch in the Imd signaling cascade through the activation of JNK, and were not upregulated in

DCV-infected drosophila. The involvement of the Imd pathway in anti-Sigma virus response was also suggested by the upregulation of Imd pathway modulated AMP genes, including Drosocin, Diptericin, Diptericin-B, Attacin-A, B, and C (Table 5.3). DCV infection also activates the Imd pathway modulated AMP genes, although mostly different AMP genes are upregulated, including Diptericin-B, Attacin-A, B, C, D, and

Cecropin-A1, A2, B, C (Table 5.3). Tested by qRT-PCR, DXV infection slightly activates

the Imd pathway modulated AMP genes, including Drosocin, Diptericin, Attacin, and

Cecropin after 2 and 24-hour infections, although most fold changes are below two folds

(Zambon et al., 2005).

Sigma virus did not upregulate genes that are components of the Toll pathway. DCV

activates the NF-κB family transcription factor Dorsal and downstream AMP genes

Drosomycin, Defensin, and Metchnikowin of the Toll pathway. qRT-PCR results reveal

that DXV infection dramatically upregulates (greater than 60 folds) at least two AMP

genes, Drosomycin and Metchnikowin (Zambon et al., 2005).

In addition to the Toll and Imd pathways, we also found a Jak-STAT pathway

regulated gene, CG9080, upregulated in Sigma virus-infected drosophila (Table 5.3).

Therefore, the Jak-STAT signaling pathway may be involved in anti-Sigma virus response

of drosophila as well. The Jak-STAT pathway is also involved in the anti-DCV response

of drosophila (Dostert et al., 2005).

135

In summary, Sigma virus infection triggers different transcriptional responses in

drosophila than those triggered by DCV and DXV infections. First, Sigma virus infection

mainly activates the Imd pathway of drosophila immune system, whereas the Toll and

Jak-STAT signaling pathways only slightly respond to Sigma virus infection. In contrast,

DCV and DXV appear to mainly activate the Jak-STAT and Toll pathways, respectively.

Secondly, drosophila response to Sigma virus infection is much less pronounced, and this

may be due to the interactions of Sigma virus with catalytic PGRPs. The differential

expression between Sigma virus, DCV, and DXV was expected because Sigma virus has a

lipid envelope with exposed G proteins, but DCV and DXV do not.

5.4.4. Differential expression of defense-related genes

Sigma virus infection also upregulated more genes encoding proteases and serine-type endopeptidase inhibitors (serpins) in drosophila than those activated by DCV infection (Fig 5.3). A number of transcript changes correspond to protease and protease

inhibitors. A total of ten protease genes were upregulated and five protease genes were

downregulated in Sigma virus-infected versus uninfected drosophila (Figs 5.3 and 5.4).

Most of these proteases were serine-type endopeptidases. Interestingly, all differential expressed protease genes were specific to the heads or bodies. Four serpins were

upregulated only in the bodies experiments. Serine proteases and serpins regulate several

invertebrate defense responses, including blood coagulation, AMP synthesis, and

melanization (Reichhart, 2005; Ross et al., 2003). In addition, three lysozyme genes were

136

downregulated (Fig 5.4). Lysozyme hydrolyzes the polysaccharide component of the cell

wall to reduce the negative charge and facilitate phagocytosis of bacteria (Masschalck

and Michiels, 2003).

5.4.5. Differential expression of cell fate-related genes

Sigma virus infection upregulated ten and downregulated four cell fate-related genes

(Fig 5.5). Cell fate-related genes include genes required for cell morphogenesis,

proliferation, and apoptosis. Most upregulated genes were related to nervous system, embryo, and muscle development. For example, stripe, astray, and off-track were related

to nervous system development and axon guidance, and they were activated in response

to Sigma virus infection (Fig 5.5). Among downregulated cell fate-related genes, a gene

related to cell proliferation, CG8319, was greatly downregulated in both heads and bodies

experiments. These results suggest that Sigma virus infection stimulates nerve

development enhancing spread of viruses and interferes with cell proliferation and

drosophila development.

5.4.6. Differential expression of metabolism and structural genes

Sigma virus infection upregulated 17 and downregulated seven metabolism genes

(Fig 5.6). Most upregulated metabolism genes were related to carbohydrate and lipid metabolisms and included many lipases. Interestingly, there were no downregulated lipid metabolism genes in Sigma virus-infected flies. Upregulation of carbohydrate and lipid metabolisms provide energy, lipid, and amino acid supporting virus morphogenesis.

137

Another interesting observation was that Sigma virus infection strongly suppressed five

cuticle synthesis genes in heads only experiments (Fig 5.7). It is not clear whether these

genes are biologically relevant to the survival of the viruses and infected flies.

5.4.7. Differential expression of sensing genes

Six genes related to vision, gustatory, and olfaction were upregulated with Sigma

virus infection, and only one sensing gene was downregulated (Fig 5.8). These

upregulated genes were Ggamma30A, trp-like, dpr13, and odorant-binding protein 49a,

56a, and 99b. One downregulated gene was Gustatory receptor 98a. These results may imply the possibility that Sigma virus affects drosophila behavior. Rhabdovirus infection induces behavior changes have been described for Rabies virus-infected animals as they

generally become more aggressive and have increased salivation (de Mattos et al., 2001).

5.5. Discussion

Sigma virus infection triggers different immune responses in drosophila compared to

the other drosophila viruses, DCV and DXV. Sigma virus mainly activates the Imd

signaling pathway, whereas DCV and DXV appear to mainly activate the Jak-STAT and

Toll pathways, respectively (Dostert et al., 2005; Zambon et al., 2005). Gram-negative

and Gram-positive bacteria also trigger different immune pathways in drosophila.

Gram-negative bacteria induce mainly the Imd pathway through interaction of the

bacterial peptidoglycan and the transmembrane receptor PGRP-LC, whereas

Gram-positive bacteria induce mainly the Toll pathway through interaction among the

bacterial peptidoglycan with the circulating protein PGRP-SA and GNBP-1 (Leclerc and

138

Reichhart, 2004; Royet et al., 2005; Steiner, 2004). Thus, our findings demonstrate that

viruses induce different immune pathways in drosophila, similarly to Gram-negative and

Gram-positive bacteria. Just as there are distinct signaling pathways activated by different types of bacteria, there may be some capacity for recognition and response to diverse viral types as well.

The finding that the drosophila immune response can differentiate between Sigma

virus versus DCV and DXV may not be surprising as these viruses are different in virion

structures. Sigma virus is an enveloped virus. Its RNA genome is surrounded by a protein

core (form a nucleocapsid) and a lipid envelope with one type of transmembrane G

protein protruding from the exterior (Teninges, 1999). In contrast, DCV and DXV are

non-enveloped viruses, and their RNA genomes are encapsidated solely in a protein core

(Jousset et al., 1977; Zambon et al., 2005). It has been shown that different types of

peptidoglycans on the surface of the Gram-negative and Gram-positive bacteria are

discriminated by various PGRPs and trigger the Imd and Toll pathways (Steiner, 2004).

Hence, it is most likely that the Sigma virus lipid envelop and the G protein, which are

absent from DCV and DXV, are mainly responsible for activating the observed immune

responses in drosophila.

In addition to activate the Imd pathway, Sigma virus appears to also inhibit

drosophila immune responses. Sigma virus infection upregulates several catalytic PGRP

genes, including PGRP-SB1, SC1a, and SC1b demonstrating to suppress induction of drosophila immune pathways. For example, PGRP-SC1b has amidase activity and cleaves peptidoglycans to make peptidoglycans unable to elicit immune response

(Mellroth et al., 2003). Interestingly, the rhabdovirus G protein is synthesized in the host

139

cells, and their glycan chains are derived from the host cell in the Golgi apparatus.

Therefore, it is very possible that the host-derived glycans of the Sigma virus G protein are important for inhibition of drosophila immune response. Further, none of the genes encoding known components of the RNA interference machinery were upregulated in

Sigma virus infected drosophila. It has been suggested that the Sonchus yellow net rhabdovirus (SYNV) P protein functions as a silencing suppressor in plants (Jackson et al., 2005). Similarly, the Sigma virus P protein may be function as a silencing suppressor in virus-infected drosophila. In summary, our results indicate that Sigma virus inhibits antiviral immune responses in drosophila.

There is more evidence that Sigma virus modulates drosophila immune responses.

At least six drosophila genes confer resistance to Sigma virus infection in natural

drosophila population (Teninges, 1999). Among these, ref(2)P is most extensively

investigated. The Ref(2)P protein shares conformation-dependent epitopes with the Sigma

virus N protein and forms a complex with the Sigma virus P protein (Wyers et al., 1993;

Wyers et al., 1995). Interestingly, Ref(2)P interacts with the atypical protein kinase C that

induces Dif transcriptional activity in the Toll signaling cascade (Avila et al., 2002; Royet

et al., 2005). Thus, Sigma virus may actively inhibit the transcription induction in the Toll

pathway by interacting with the atypical protein kinase C. Evidence that Sigma virus

interferes with induction of the Toll pathway through interactions with catalytic PGRPs

and Ref(2)P also confirms our results that Sigma virus does not upregulate transcripts

corresponding to the AMP Drosomycin, which is typically induced upon activation of the

Toll pathway (Royet et al., 2005).

140

Overall, the number of genes differentially regulated in Sigma virus-infected

drosophila versus non-infected flies is low. Only 1.5% of drosophila genes are 1.5 – 3.0 fold up- or downregulated during Sigma virus infection. Sigma virus is exclusively vertically transmitted to progeny flies (Teninges, 1999). For vertically transmitted parasite, a balance typically evolves between maximizing parasite transmission and minimizing virulence to hosts (Messenger et al., 1999; Stewart et al., 2005). Therefore, it is beneficial to Sigma virus transmission to minimize pathogenic effects to its host. DCV and DXV are transmitted horizontally, and are more virulence as observed by cytopathological effects in DCV and DXV-infected flies (Gravot et al., 2000; Teninges et al., 1979).

The drosophila Toll receptor is most similar to the Toll-like receptor TLR 4 of

mammals and humans. In mammalian immune system, 13 Toll-like receptors (TLRs) are

now recognized, and the expression of 10 is known in humans (Pandey and Agrawal,

2006). Rabies virus also activates the innate immune response in mammals through the

upregulation of TLRs in the CNS, especially TLR 2 and 3 (McKimmie et al., 2005). TLR

2 is involved in the recognition of viral G proteins, bacterial peptidoglycans, and fungal

components, and TLR 3 recognizes dsRNA (Alexopoulou et al., 2001; Bieback et al.,

2002; Pandey and Agrawal, 2006; Takeuchi et al., 1999; Underhill et al., 1999).

Interestingly, like Sigma virus in drosophila, Rabies virus also suppresses the immune

response in the nervous system of its host (Lafon, 2005). Rabies virus infection mainly

downregulates the gene expression, including immune genes, in mouse brain (Prosniak et

141

al., 2001). The street strain of Rabies virus evades virus-mediated apoptosis and host

immune response (Lafon, 2005; Wang et al., 2005). Thus, the innate immune responses of

vertebrate and invertebrate animals to rhabdovirus infections are comparable.

Our study also reveals that several nervous system development genes are upregulated in Sigma virus-infected flies, such as stripe, astray, and off-track that are involved in nervous system development and axon guidance. Sigma virus may traffic

through the nervous system of drosophila and is accumulated in cephalic and thoracic ganglia (Teninges, 1999). The paralysis that occurs upon exposure of Sigma virus-infected drosophila to CO2 is related to the virus infection in the CNS (Teninges,

1999). Other insect- transmitted rhabdoviruses are also neurotropic in their insect vectors,

including Vesicular stomatitis New Jersey virus (VSNJV) in the midge vector Culicoides

sonorensis and plant rhabdovirus Maize mosaic virus in the planthopper vector

Peregrinus maidis (Drolet et al., 2005; Ammar and Hogenhout, Submitted). Further,

Rabies virus completes its infection by traveling from one neuron to the next, along the spinal cord to the brain and the salivary glands. The establishment of new connections between neurons may be an element of the spread of Rabies virus in the CNS (Prosniak et

al., 2001; Prosniak et al., 2003). Interestingly, Sigma virus and Rabies virus may both

actively upregulate nerve developmental genes that enhance spread of these viruses

through the animal body. Rabies virus induces some obvious behavioral changes in

infected animals, such as increased aggression and salivation that increase the likelihood of transmission of this virus to other hosts. It remains to be determined whether Sigma virus infection of the drosophila nervous system initiates any transmission-enhancing changes in the behavior of flies.

142

5.6. Acknowledgments

This work was supported by the Australian Research Council Linkages International

Grant LX0452397. We thank Dr. S. L. O’Neill from the School of Integrative Biology,

The University of Queensland, for his advice in planning the described experiments.

143

5.7. References

Alexopoulou, L., Holt, A.C., Medzhitov, R. and Flavell, R.A. (2001) Recognition of double-stranded RNA and activation of NF-κB by Toll-like receptor 3. Nature, 413, 732-738.

Ammar, E.D., Meulia, T., Ozbek, E. and Hogenhout, S.A. (2004) Assembly and accumulation sites of Maize mosaic virus (Rhabdoviridae) in plant host and insect vector using transmission electron and confocal laser scanning microscopy. In Mendez-Vilas, A. and Labajos-Broncano, L. (eds.), Current issues on multidisciplinary microscopy research and education. Formatex, Badajoz, Spain, pp. 56-64.

Avila, A., Silverman, N., Diaz-Meco, M.T. and Moscat, J. (2002) The Drosophila atypical protein kinase C-ref(2)p complex constitutes a conserved module for signaling in the toll pathway. Mol. Cell. Biol., 22, 8787-8795.

Bieback, K., Lien, E., Klagge, I.M., Avota, E., Schneider-Schaulies, J., Duprex, W.P., Wagner, H., Kirschning, C.J., Ter Meulen, V. and Schneider-Schaulies, S. (2002) Hemagglutinin protein of wild-type measles virus activates Toll-like receptor 2 signaling. J. Virol., 76, 8729-8736.

Bischoff, V., Vignal, C., Boneca, I.G., Michel, T., Hoffmann, J.A. and Royet, J. (2004) Function of the drosophila pattern-recognition receptor PGRP-SD in the detection of Gram-positive bacteria. Nat. Immunol., 5, 1175-1180.

Brun, G. (1991) Rhabdoviridae. In Adams, J.R. and Bonami, J.R. (eds.), Atlas of invertebrate viruses. CRC Press, Boca Raton, pp. 443-460. de Mattos, C.A., de Mattos, C.C. and Rupprecht, C.E. (2001) Rhabdoviruses. In Knipe, D.M. and Howley, P.M. (eds.), Fields virology. Lippincott Williams & Wilkins, Philadelphia, pp. 1245-1277.

Dostert, C., Jouanguy, E., Irving, P., Troxler, L., Galiana-Arnoux, D., Hetru, C., Hoffmann, J.A. and Imler, J.L. (2005) The Jak-STAT signaling pathway is required but not sufficient for the antiviral response of drosophila. Nat. Immunol., 6, 946-953.

Drolet, B.S., Campbell, C.L., Stuart, M.A. and Wilson, W.C. (2005) Vector competence of Culicoides sonorensis (Diptera: Ceratopogonidae) for vesicular stomatitis virus. J. Med. Entomol., 42, 409-418.

Gravot, E., Thomas-Orillard, M. and Jeune, B. (2000) Virulence variability of the Drosophila C virus and effects of the microparasite on demographic parameters of the host (Drosophila melanogaster). J. Invertebr. Pathol., 75, 144-151.

144

Grimmond, S., Van Hateren, N., Siggers, P., Arkell, R., Larder, R., Soares, M.B., de Fatima Bonaldo, M., Smith, L., Tymowska-Lalanne, Z., Wells, C. and Greenfield, A. (2000) Sexually dimorphic expression of protease nexin-1 and vanin-1 in the developing mouse gonad prior to overt differentiation suggests a role in mammalian sexual development. Hum. Mol. Genet., 9, 1553-1560.

Hoffmann, J.A. (2003) The immune response of Drosophila. Nature, 426, 33-38.

Hogenhout, S.A., Redinbaugh, M.G. and Ammar, E.D. (2003) Plant and animal rhabdovirus host range: a bug's view. Trends Microbiol., 11, 264-271.

Jackson, A.O., Dietzgen, R.G., Goodin, M.M., Bragg, J.N. and Deng, M. (2005) Biology of plant rhabdoviruses. Annu. Rev. Phytopathol., 43, 623-660.

Jousset, F.X., Bergoin, M. and Revet, B. (1977) Characterization of the Drosophila C virus. J. Gen. Virol., 34, 269-283.

Lafon, M. (2005) Modulation of the immune response in the nervous system by rabies virus. Curr. Top. Microbiol. Immunol., 289, 239-258.

Leclerc, V. and Reichhart, J.M. (2004) The immune response of Drosophila melanogaster. Immunol. Rev., 198, 59-71.

Masschalck, B. and Michiels, C.W. (2003) Antimicrobial properties of lysozyme in relation to foodborne vegetative bacteria. Crit. Rev. Microbiol., 29, 191-214.

McKimmie, C.S., Johnson, N., Fooks, A.R. and Fazakerley, J.K. (2005) Viruses selectively upregulate Toll-like receptors in the central nervous system. Biochem. Biophys. Res. Commun., 336, 925-933.

Mellroth, P., Karlsson, J. and Steiner, H. (2003) A scavenger function for a Drosophila peptidoglycan recognition protein. J. Biol. Chem., 278, 7059-7064.

Messenger, S.L., Molineux, I.J. and Bull, J.J. (1999) Virulence evolution in a virus obeys a trade-off. Proc. Biol. Sci., 266, 397-404.

Michel, T., Reichhart, J.M., Hoffmann, J.A. and Royet, J. (2001) Drosophila Toll is activated by Gram-positive bacteria through a circulating peptidoglycan recognition protein. Nature, 414, 756-759.

Nault, L.R. (1997) Arthropod transmission of plant viruses: a new synthesis. Ann. Entomol. Soc. Am., 90, 521-541.

Pandey, S. and Agrawal, D.K. (2006) Immunobiology of Toll-like receptors: emerging trends. Immunol. Cell Biol., 84, 333-341.

145

Prosniak, M., Hooper, D.C., Dietzschold, B. and Koprowski, H. (2001) Effect of rabies virus infection on gene expression in mouse brain. Proc. Natl. Acad. Sci. U.S.A., 98, 2758-2763.

Prosniak, M., Zborek, A., Scott, G.S., Roy, A., Phares, T.W., Koprowski, H. and Hooper, D.C. (2003) Differential expression of growth factors at the cellular level in virus-infected brain. Proc. Natl. Acad. Sci. U.S.A., 100, 6765-6770.

Reichhart, J.M. (2005) Tip of another iceberg: Drosophila serpins. Trends Cell Biol., 15, 659-665.

Rose, J.K. and Whitt, M.A. (2001) Rhabdoviridae: the virus and their replication. In Knipe, D.M. and Howley, P.M. (eds.), Fields virology. Lippincott Williams & Wilkins, Philadelphia, pp. 1221-1244.

Ross, J., Jiang, H., Kanost, M.R. and Wang, Y. (2003) Serine proteases and their homologs in the Drosophila melanogaster genome: an initial analysis of sequence conservation and phylogenetic relationships. Gene, 304, 117-131.

Roxstrom-Lindquist, K., Terenius, O. and Faye, I. (2004) Parasite-specific immune response in adult Drosophila melanogaster: a genomic study. EMBO Rep., 5, 207-212.

Royet, J., Reichhart, J.M. and Hoffmann, J.A. (2005) Sensing and signaling during infection in Drosophila. Curr. Opin. Immunol., 17, 11-17.

Steiner, H. (2004) Peptidoglycan recognition proteins: on and off switches for innate immunity. Immunol. Rev., 198, 83-96.

Stewart, A.D., Logsdon, J.M.,Jr and Kelley, S.E. (2005) An empirical study of the evolution of virulence under both horizontal and vertical transmission. Evolution, 59, 730-739.

Takeuchi, O., Hoshino, K., Kawai, T., Sanjo, H., Takada, H., Ogawa, T., Takeda, K. and Akira, S. (1999) Differential roles of TLR2 and TLR4 in recognition of gram-negative and gram-positive bacterial cell wall components. Immunity, 11, 443-451.

Teninges, D. (1999) Sigma rhabdoviruses (Rhabdoviridae). In Granoff, A. and Webster, R.G. (eds.), Encyclopedia of virology. Academic Press, San Diego, pp. 1635-1639.

Teninges, D., Ohanessian, A., Richard-Molard, C. and Contamine, D. (1979) Isolation and biological properties of Drosophila X vrus. J. Gen. Virol., 42, 241-254.

146

Tordo, N., Benmansour, A., Calisher, C., Dietzgen, R.G., Fang, R.X., Jackson, A.O., Kurath, G., Nadin-Davis, S., Tesh, R.B. and Walker, P.J. (2005) Family Rhabdoviridae. In Fauquet, C.M., Mayo, M.A., Maniloff, J., Dessenberger, U. and Ball, L.A. (eds.), Virus taxonomy: classification and nomenclature of viruses. Eighth report of the International Committee on Taxonomy of Viruses. Elsevier Academic Press, San Diego, pp. 623-644.

Wang, Z.W., Sarmento, L., Wang, Y., Li, X.Q., Dhingra, V., Tseggai, T., Jiang, B. and Fu, Z.F. (2005) Attenuated rabies virus activates, while pathogenic rabies virus evades, the host innate immune responses in the central nervous system. J. Virol., 79, 12554-12565.

White, K.P., Rifkin, S.A., Hurban, P. and Hogness, D.S. (1999) Microarray analysis of Drosophila development during metamorphosis. Science, 286, 2179-2184.

Wyers, F., Dru, P., Simonet, B. and Contamine, D. (1993) Immunological cross-reactions and interactions between the Drosophila melanogaster ref(2)P protein and sigma rhabdovirus proteins. J. Virol., 67, 3208-3216.

Wyers, F., Petitjean, A.M., Dru, P., Gay, P. and Contamine, D. (1995) Localization of domains within the Drosophila Ref(2)P protein involved in the intracellular control of sigma rhabdovirus multiplication. J. Virol., 69, 4463-4470.

Zambon, R.A., Nandakumar, M., Vakharia, V.N. and Wu, L.P. (2005) The Toll pathway is important for an antiviral response in Drosophila. Proc. Natl. Acad. Sci. U.S.A., 102, 7257-7262.

147

Gene Primer name Primer sequence Drosocin Dro_For 5’ TTCCTGCTGCTTGCTTGCG Dro_Rev 5’ CTGTCTTTCGTGTGTTTATTGC Diptericin Dpt_For 5’ CCATTGCCGTCGCCTTAC Dpt_Rev 5’ GACCCACCAGCCTCTGTC Attacin Att_For 5’ CCCACAACAGGACCCATTC Att_Rev 5’ CGATGACCAGAGATTAGCAC PGRP-SB1 PGRP-SB1_For 5’ AGTGCTTTGCTGCTTAGCTC PGRP-SB1_Rev 5’ CGAAGTTGCCAATGAAGACG PGRP-SC1 PGRP-SC1_For 5’ CGTCTATGTCGTCTCCAAG PGRP-SC1_Rev 5’ TTGTAGTTGCCCAGGAAGC PGRP-SD PGRP-SD_For 5’ TCTCGGACATTGGCTACCAC PGRP-SD_Rev 5’ ACATTTCTTCGGACCAGTTG Jra Jra_For 5’ TCAGAATGCTGGCAGTTCC Jra_Rev 5’ GGAGTTAGTGTGAAGATTGTG Cabut Cbt_For 5’ ATGTCCGAGGAGAAACTAAC Cbt_Rev 5’ GCTGATGTGTTGTTGTTGG Actin88F Act_For 5’ ACACCGTGCCCATCTATGAG Act_Rev 5’ TGGTTGCTGCCTTTGAAGAG

Table 5.1. Primers used in this study for amplifying Northern blot probes

148

Upregulated genes Downregulated genes Head Body Head & Head Body Head & Category specific specific body specific specific body Immunity 3 14 4 1 2 1 Protease 1 9 0 2 3 0 Protease inhibitor 0 4 0 0 0 0 Cell fate 4 5 1 2 1 1 Metabolism 3 11 3 5 2 0 Sensing 1 3 2 1 0 0 Structural gene 1 2 0 6 0 0 Others 10 15 3 6 1 0 Unknown 14 31 6 7 1 0 Total 37 94 19 30 10 2

Table 5.2. Overview of differentially regulated genes in response to Sigma virus infection in drosophila

149

Virus-insect host Sigma virus- DCV- DCV- DXV- drosophilaa drosophilab drosophilac drosophilad Pathway Fold change Fold change Fold change Fold change Upstream gene & receptor PGRP-SA Toll - - 3.2 PGRP-SD Toll 2.8 - - GNBP-1 Toll - - - Spatzle Toll - - 3.0 Toll Toll - - - PGRP-LC Imd - - -

Signaling cascade My88D Toll - - - Cactus Toll - - - Dif Toll - - - Imd Imd - - - Relish Imd - - 3.5 Cabut JNK 1.9 - - Jra JNK 1.6 - - CG31764 (Vir-1) Jak-STAT - - 6.4 CG9080 Jak-STAT 2.4 - 5.4 CG12780 Jak-STAT - - 6.5

Antimicrobial peptide Drosomycin Toll - 3.0 2.1 70 Defensin Toll 1.8 - + 4.8 Metchnikowin Toll - - 3.0 60 Drosocin Imd 4.3 - - 3.2 Diptericin Imd 4.0 - - 3.2 Diptericin-B Imd 2.1 - 5.0 Attacin-A Imd 2.4 6.3 8.7 2.5 Attacin-B Imd 5.5 - 4.7 Attacin-C Imd 3.9 - 2.7 Attacin-D Imd - - + Cecropin-A1 Imd - 2.6 - 1.8 Cecropin-A2 Imd - 3.3 - Cecropin-B Imd - - + Cecropin-C Imd - - +

Others PGRP-SB1 2.6 - - PGRP-SC1a 2.7 - - PGRP-SC1b 2.3 - - a Data from this study. Symbol: -, no different expressions between control and virus-infected drosophila. b Data from: Roxstrom-Lindquist et al. (2004) EMBO Rep. 5:207-212. DCV, Drosophila C virus. c Data from: Dostert et al. (2005) Nat. Immunol. 6:946-953. Symbol: +, not express in control but express in DCV-infected drosophila. d Data from: Zambon et al. (2005) Proc. Natl. Acad. Sci. U.S.A. 102:7257-7262. DXV, Drosophila X virus.

Table 5.3. Comparison of upregulated immunity genes of virus-infected drosophila

150

Upregulated genes Downregulated genes

Fig. 5.1. Total up- and downregulated genes in response to Sigma virus infection in drosophila. Each pie chart represents the total number of genes categorized by the process in which these genes are involved.

151

Fig. 5.2. Northern blot analysis of immunity genes expressing in Sigma virus-infected and virus-free drosophila. Each lane contains 20 μg total RNA isolated from virus free (-) and virus-infected (+) drosophila. The blots were analyzed using the 32P-labeled cDNA probes corresponding to each gene. The blots were probed with Actin88F as an internal loading control. Numbers below diagrams show the results of densitometric analysis to determine the relative expression level of each gene after correcting by Actin88F expression. Representative of three independent experiments is presented.

152

Immunity genes Head Body Gene name Gene ID Pathway Fold p-value Fold p-value Dro CG10816 Imd 2.85 0.003 4.26 0.003 AttB CG18372 Imd 2.34 0.013 5.49 0.041 PGRP-SB1 CG9681 1.83 0.006 2.63 0.002 Dpt CG12763 Imd 1.71 0.019 3.99 0.037 CG9080 Jak-STAT 1.57 0.024 2.41 0.016 AttA CG10146 Imd 2.44 0.038 - - DptB CG10794 Imd 2.14 0.009 - - CG13422 1.61 0.034 - - AttC CG4740 Imd - - 3.85 0.001 PGRP-SD CG7496 Toll - - 2.79 0.042 PGRP-SC1a CG14746 - - 2.72 0.009 IM23 CG15066 - - 2.36 0.014 PGRP-SC1b CG8577 - - 2.31 0.044 IM18 CG10332 - - 2.26 0.001 Cbt CG4427 JNK - - 1.87 0.001 Def CG1385 Toll - - 1.81 0.022 CG6676 - - 1.63 0.003 Gli CG3903 - - 1.60 0.031 Jra CG2275 JNK - - 1.60 0.028 L(2)efl CG4533 - - 1.57 0.011 CG10433 - - 1.56 0.006 Ag5r CG9538 - - 1.53 0.027

Protease genes Head Body Gene name Gene ID Type Fold p-value Fold p-value CG8528 Serine 1.70 0.042 - - Jon65Aiii CG6483 Serine - - 2.06 0.000 CG8299 Serine - - 2.06 0.021 CG10472 Serine - - 2.01 0.007 CG4053 Serine - - 1.92 0.011 Jon66Ci CG7118 Serine - - 1.77 0.013 Jon99Fi CG18030 Trypsin - - 1.67 0.014 Jon44E CG8579 Trypsin - - 1.62 0.002 CG11911 Serine - - 1.55 0.011 CG6733 Metallo- - - 1.51 0.006 peptidase

Protease inhibitor genes Head Body Gene name Gene ID Type Fold p-value Fold p-value CG16712 Serpin - - 2.35 0.0170 CG7219 Serpin - - 1.55 0.0024 CG6663 Serpin - - 1.52 0.0474 CG9460 Serpin - - 1.51 0.0190

>1.5 >2 >4

Fig. 5.3. Upregulated defense-related genes in response to Sigma virus infection in drosophila. The fold change color code is indicated at the bottom.

153

Immunity genes Head Body Gene name Gene ID Type Fold p-value Fold p-value LysE CG1180 Lysozyme 0.40 0.015 0.37 0.002 CG5999 0.49 0.009 - - LysB CG1179 Lysozyme - - 0.50 0.003 LysD CG9118 Lysozyme - - 0.36 0.019

Protease genes Head Body Gene name Gene ID Type Fold p-value Fold p-value CG1497 Serine 0.50 0.020 - - Jon74E CG6298 Trypsin 0.42 0.000 - - CG3739 Serine - - 0.46 0.003 CG11529 Serine - - 0.42 0.011 Ser12 CG17240 Serine - - 0.34 0.033

<0.25 <0.5

Fig. 5.4. Downregulated defense-related genes in response to Sigma virus infection in drosophila. The fold change color code is indicated at the bottom.

154

Fig. 5.5. Up- and downregulated cell fate-related genes of Sigma virus-infected drosophila.

155

Fig. 5.6. Up- and downregulated metabolism genes of Sigma virus-infected drosophila

156

Fig. 5.7. Up- and downregulated structural genes of Sigma virus-infected drosophila

157

Fig. 5.8. Up- and downregulated sensing genes of Sigma virus-infected drosophila

158

BIBLIOGRAPHY

Adam, G. and Hsu, H.T. (1984) Comparison of structural proteins from two potato yellow dwarf viruses. J. Gen. Virol., 65, 991-994.

Adam, S.A. and Gerace, L. (1991) Cytosolic proteins that specifically bind nuclear location signals are receptors for nuclear import. Cell, 66, 837-847.

Alexopoulou, L., Holt, A.C., Medzhitov, R. and Flavell, R.A. (2001) Recognition of double-stranded RNA and activation of NF-kappaB by Toll-like receptor 3. Nature, 413, 732-738.

Ammar, E.D. and Nault, L.R. (2002) Virus transmission by leafhoppers, planthoppers and treehoppers (Auchenorrhyncha, Homoptera). Adv. Bot. Res., 36, 141-167.

Ammar, E.D. and Nault, L.R. (1985) Assembly and accumulation sites of maize mosaic virus in its planthopper vector. Intervirology, 24, 33-41.

159

Barr, J.N., Whelan, S.P. and Wertz, G.W. (2002) Transcriptional control of the RNA- dependent RNA polymerase of vesicular stomatitis virus. Biochim. Biophys. Acta, 1577, 337-353.

Baulcombe, D.C. (1999) Fast forward genetics based on virus-induced gene silencing. Curr. Opin. Plant Biol., 2, 109-113.

Behncken, G.M. (1973) Evidence of multiplication of sowthistle yellow vein virus in an inefficient aphid vector, Macrosiphum euphorbiae. Virology, 53, 405-412.

Bendtsen, J.D., Nielsen, H., von Heijne, G. and Brunak, S. (2004) Improved prediction of signal peptides: SignalP 3.0. J. Mol. Biol., 340, 783-795.

Bieback, K., Lien, E., Klagge, I.M., Avota, E., Schneider-Schaulies, J., Duprex,W.P., Wagner, H., Kirschning, C.J., Ter Meulen, V. and Schneider-Schaulies, S. (2002) Hemagglutinin protein of wild-type measles virus activates toll-like receptor 2 signaling. J. Virol., 76, 8729-8736.

Black, L.M. (1941) Specific transmission of varieties of potato yellow-dwarf virus by related insects. Am. potato J., 18, 231-233.

Bradfute, O.E. and Tsai, J.H. (1983) Identification of maize mosaic virus in Florida. Plant Dis., 67, 1339-l342.

Brun, G. (1991) Rhabdoviridae. In Adams, J.R. and Bonami, J.R. (eds.), Atlas of invertebrate viruses. CRC Press, Boca Raton, pp. 443-460.

Brunt, A., Crabtree, K., Dallwitz, M., Gibbs, A. and Watson, L. (1996) Viruses of plants: descriptions and lists from the VIDE database. 1484.

Chen, X.Y., Luo, Z.L. and Fang, R.X. (1998) Structure analysis of the rice yellow stunt rhabdovirus gene 3. Chinese Sci. Bull., 43, 745-748.

160

Choi, T.J., Kuwata, S., Koonin, E.V., Heaton, L.A. and Jackson, A.O. (1992) Structure of the L (polymerase) protein gene of sonchus yellow net virus. Virology, 189, 31-39.

Choi, T.J., Wagner, J.D. and Jackson, A.O. (1994) Sequence analysis of the trailer region of sonchus yellow net virus genomic RNA. Virology, 202, 33-40.

Cokol, M., Nair, R. and Rost, B. (2000) Finding nuclear localization signals. EMBO Rep., 1, 411-415.

Creamer, R. (1992) Purification and protein characterization of sorghum stunt mosaic rhabdovirus. Phytopathology, 82, 1473-1476. de Mattos, C.A., de Mattos, C.C. and Rupprecht, C.E. (2001) Rhabdoviruses. In Knipe, D.M. and Howley, P.M. (eds.), Fields virology. Lippincott Williams & Wilkins, Philadelphia, pp. 1245-1277.

Dietzgen, R.G., Callaghan, B., Wetzel, T. and Dale, J.L. (2006) Completion of the genome sequence of Lettuce necrotic yellows virus, type species of the genus Cytorhabdovirus. Virus Res., 118, 16-22.

Drolet, B.S., Campbell, C.L., Stuart, M.A. and Wilson, W.C. (2005) Vector competence of Culicoides sonorensis (Diptera: Ceratopogonidae) for vesicular stomatitis virus. J. Med. Entomol., 42, 409-418.

Falk, B.W. and Tsai, J.H. (1985) Serological detection and evidence for multiplication of maize mosaic virus in the planthopper, Peregrinus maidis. Phytopathology, 75, 852-855.

Falk, B.W. and Tsai, J.H. (1983) Physicochemical characterization of maize mosaic virus. Phytopathology, 73, 1536-1539.

Falquet, L., Pagni, M., Bucher, P., Hulo, N., Sigrist, C.J., Hofmann, K. and Bairoch, A. (2002) The PROSITE database, its status in 2002. Nucleic Acids Res., 30, 235-238.

Feinberg, A.P. and Vogelstein, B. (1983) A technique for radiolabeling DNA restriction endonuclease fragments to high specific activity. Anal. Biochem., 132, 6-13.

Fleischmann, R.D., Adams, M.D., White, O., Clayton, R.A., Kirkness, E.F., Kerlavage, A.R., Bult, C.J., Tomb, J.F., Dougherty, B.A. and Merrick, J.M. (1995) Whole- genome random sequencing and assembly of Haemophilus influenzae Rd. Science, 269, 496-512.

161

Francki, R.I. and Randles, J.W. (1973) Some properties of lettuce necrotic yellows virus RNA and its in vitro transcription by virion-associated transcriptase. Virology, 54, 359-368.

Fraser, C.M., Eisen, J.A., Nelson, K.E., Paulsen, I.T. and Salzberg, S.L. (2002) The value of complete microbial genome sequencing (you get what you pay for). J. Bacteriol., 184, 6403-5; discusion 6405.

Garbutt, M., Liebscher, R., Wahl-Jensen, V., Jones, S., Moller, P., Wagner, R., Volchkov, V., Klenk, H.D., Feldmann, H. and Stroher, U. (2004) Properties of replication- competent vesicular stomatitis virus vectors expressing glycoproteins of filoviruses and arenaviruses. J. Virol., 78, 5458-5465.

Goldfarb, D.S., Corbett, A.H., Mason, D.A., Harreman, M.T. and Adam, S.A. (2004) Importin α: a multipurpose nuclear-transport receptor. Trends Cell Biol., 14, 505-514.

Gorlich, D., Kostka, S., Kraft, R., Dingwall, C., Laskey, R.A., Hartmann, E. and Prehn, S. (1995) Two different subunits of importin cooperate to recognize nuclear localization signals and bind them to the nuclear envelope. Curr. Biol., 5, 383-392.

Gorlich, D., Prehn, S., Laskey, R.A. and Hartmann, E. (1994) Isolation of a protein that is essential for the first step of nuclear protein import. Cell, 79, 767-778.

Gorlich, D., Vogel, F., Mills, A.D., Hartmann, E. and Laskey, R.A. (1995) Distinct functions for the two importin subunits in nuclear protein import. Nature, 377, 246-248.

162

Gupta, A.K., Shaji, D. and Banerjee, A.K. (2003) Identification of a novel tripartite complex involved in replication of vesicular stomatitis virus genome RNA. J. Virol., 77, 732-738.

Hanson, M.R. and Kohler, R.H. (2001) GFP imaging: methodology and application to investigate cellular compartmentation in plants. J. Exp. Bot., 52, 529-539.

Hicks, G.R. and Raikhel, N.V. (1995) Protein import into the nucleus: an integrated view. Annu. Rev. Cell Dev. Biol., 11, 155-188.

Hoffmann, J.A. (2003) The immune response of Drosophila. Nature, 426, 33-38.

Hogenhout, S.A., Verbeek, M., Hans, F., Houterman, P.M., Fortass, M., van der Wilk, F., Huttinga, H. and van den Heuvel, J.F.J.M. (1996) Molecular basis of the interactions between luteoviruses and aphids. Agronomie, 16, 167-173.

Hogenhout, S.A., Redinbaugh, M.G. and Ammar, E.D. (2003) Plant and animal rhabdovirus host range: a bug's view. Trends Microbiol., 11, 264-271.

Huang, Y.W., Geng, Y.F., Ying, X.B., Chen, X.Y. and Fang, R.X. (2005) Identification of a movement protein of rice yellow stunt rhabdovirus. J. Virol., 79, 2108-2114.

163

Izaurralde, E., Kutay, U., von Kobbe, C., Mattaj,I.W. and Gorlich, D. (1997) The asymmetric distribution of the constituents of the Ran system is essential for transport into and out of the nucleus. EMBO J., 16, 6535-6547.

Jackson, A.O., Goodin, M., Moreno, I., Jackson, J. and Lawrence, D.M. (1999) Plant rhabdoviruses. In Granoff, A. and Webster, R.G. (eds.), Encyclopedia of virology. Academic Press, San Diego, CA, pp. 1531-1541.

Jackson, A.O., Dietzgen, R.G., Goodin, M.M., Bragg, J.N. and Deng, M. (2005) Biology of plant rhabdoviruses. Annu. Rev. Phytopathol., 43, 623-660.

Jousset, F.X., Bergoin, M. and Revet, B. (1977) Characterization of the Drosophila C virus. J. Gen. Virol., 34, 269-283.

Kurath, G., Higman, K.H. and Bjorklund, H.V. (1997) Distribution and variation of NV genes in fish rhabdoviruses. J. Gen. Virol., 78, 113-117.

Kutay, U., Bischoff, F.R., Kostka, S., Kraft, R. and Gorlich, D. (1997a) Export of importin α from the nucleus is mediated by a specific nuclear transport factor. Cell, 90, 1061-1071.

164

Lafon, M. (2005) Modulation of the immune response in the nervous system by rabies virus. Curr. Top. Microbiol. Immunol., 289, 239-258.

Leclerc, V. and Reichhart, J.M. (2004) The immune response of Drosophila melanogaster. Immunol.Rev., 198, 59-71.