Iowa State University Capstones, Theses and Graduate Theses and Dissertations Dissertations
2010 Translation and replication of Rhopalosiphum padi virus RNA in a plant cellular environment Zachary Thomas Regelin Iowa State University
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Recommended Citation Regelin, Zachary Thomas, "Translation and replication of Rhopalosiphum padi virus RNA in a plant cellular environment" (2010). Graduate Theses and Dissertations. 11888. https://lib.dr.iastate.edu/etd/11888
This Thesis is brought to you for free and open access by the Iowa State University Capstones, Theses and Dissertations at Iowa State University Digital Repository. It has been accepted for inclusion in Graduate Theses and Dissertations by an authorized administrator of Iowa State University Digital Repository. For more information, please contact [email protected]. Translation and replication of Rhopalosiphum padi virus RNA in a plant cellular environment
by
Zachary Thomas Regelin
A thesis submitted to the graduate faculty
in partial fulfillment of the requirements for the degree of
MASTER OF SCIENCE
Major: Molecular, Cellular, and Developmental Biology
Program of Study Committee:
Bryony C. Bonning, Co-Major Professor
W. Allen Miller, Co-Major Professor
Steven A. Whitham
Iowa State University
Ames, Iowa
2010 ii
Table of Contents
LIST OF FIGURES iv LIST OF TABLES v ABSTRACT vi CHAPTER 1. GENERAL INTRODUCTION 1 Thesis organization 1 Literature review 1 Introduction 1 Picornavirales 3 Dicistroviruses 5 Rhopalosiphum padi virus 6 Identification of RhPV permissive cell lines and production of an RhPV infectious clone 9 Aphids as major agricultural pests 12 RhPV and other small RNA viruses as biopesticides 13 Translation 17 Canonical translation 17 IRESs as alternatives to canonical translation 20 Dicistrovirus IRESs 26 Goals of thesis research 32 References 37 CHAPTER 2. INVESTIGATION OF TRANSLATION AND REPLICATION OF RHOPALOSIPHUM PADI VIRUS (DICISTROVIRIDAE), IN A PLANT CELLULAR ENVIRONMENT 47 Abstract 47 Introduction 48 Results 51 In vitro translation in wheat germ extract and dual luciferase assays indicate RhPV IGR IRES functions in an in vitro plant translation system 51 In vitro translation in rabbit reticulocyte lysate 59 Translation in oat protoplasts 59 Northern blot hybridization of RNA extracted from oat protoplasts electroporated with RhPV6-1_polyA positive-strand transcript failed to detect negative-strand replication product 48 hours post-electroporation 60
iii
Strand-specific RT-PCR of oat protoplasts electroporated with positive-strand RhPV6-1 transcript suggests replication in protoplasts 62 Discussion 69 Methods 73 Cloning strategy for translation assay constructs 73 Cloning strategy for replication assay constructs 76 In vitro transcription 76 In vitro translation in wheat germ extract 77 SDS polyacrylamide gel electrophoresis of 35S-methionine-labeled wheat germ extract translation product 77 In vitro translation in rabbit reticulocyte lysate 77 Dual luciferase assays 78 Electroporation of oat protoplasts 78 Northern blot hybridization 80 Strand-specific RT-PCR 81 References 82 CHAPTER 3. GENERAL CONCLUSIONS 86 References 91 ACKNOWLEDGEMENTS 93
iv
LIST OF FIGURES
Chapter 1
Figure 1. Organization of Picornavirales genomes. 2
Figure 2. Model of the canonical pathway of eukaryotic translation initiation. 18
Figure 3. Comparisons of eIF requirements of cap-dependent translation and
four IRES groups. 22
Figure 4. Dicistrovirus IGR IRES structure and molecular mimicry. 24
Figure 5. Immunoblot and RT-PCR indicate production of infectous RhPV
virions in oat protoplast electroporated with wildtype RhPV RNA. 34
Chapter 2
Figure 1. RhPV infectious clone and dual luciferase construct maps. 52
Figure 2. IGR IRES activity in WGE. 54
Figure 3. IGR IRES activity in WGE, RRL, and oat protoplasts. 57
Figure 4. SDS-PAGE fractionation of WGE translation products from reactions
with dual luciferase transcripts. 58
Figure 5. Northern blot hybridization of RhPV 6.1 polyA inoculated oat
protoplasts. 61
Figure 6. Schematic of strand-specific RT-PCR using nongenomic tag sequences. 63
Figure 7. Strand-specific RT-PCR detection of positive- and negative-strand
RhPV6.1 RNA from oat protoplasts electroporated with RhPV6.1
positive-strand transcripts demonstrates RhPV6.1 replication in protoplasts. 65 v
LIST OF TABLES
Chapter 1
Table 1. Dicistroviruses, their arthropod hosts, and translation systems
supporting 5’ UTR IRES- and IGR IRES-mediated translation
in vivo and in vitro. 8
Table 2. IGR IRES structural elements and brief descriptions of proposed
functions. 33
Chapter 2
Table 1. RT-PCR primers. 68
Table 2. Primers used for cloning and confirmation of clones. 74
vi
ABSTRACT
Dicistroviruses are small, monopartite, positive-strand RNA viruses that infect arthropods. Unlike other members of the order Picornavirales, viruses in the Dicistroviridae family possess two open reading frames (ORFs), each preceded by a distinct internal ribosome entry site (IRES). Availability of an infectious clone of Rhopalosiphum padi virus (RhPV) provides a useful molecular tool for the investigation of dicistrovirus translation and replication.
Cross-kingdom analysis of translation and replication of this insect virus in a plant cellular environment could elucidate key factors involved in these processes and prove vital in efforts to apply dicistroviruses in transgenically expressed biopesticide management strategies. Intergenic region (IGR) IRES-mediated translation was tested in wheat germ extract (WGE) and oat protoplasts. IGR IRES translation was observed in WGE, but translation in oat protoplasts was insufficient to show IGR IRES function. Replication of RhPV infectious transcripts was assayed in oat protoplasts using strand-specific RT-PCR and northern blot hybridization. Negative- strand replication products were detected 48 hours post-electroporation of protoplasts with strand-specific RT-PCR, but were undetected via northern blot analysis. Though translation was not shown in this study, other preliminary experiments suggest that, with slight protocol modifications, IGR IRES-mediated translation may be detectable in oat protoplasts. Low levels of RhPV replication in planta may be beneficial in a transgenically-expressed biopesticide strategy by supporting virion packaging fidelity while avoiding RNAi silencing. 1
CHAPTER 1. GENERAL INTRODUCTION
Thesis organization
This thesis focuses on the replication and translation of a dicistrovirus of aphids,
Rhopalosiphum padi virus (RhPV) in a plant cellular environment. A literature review is presented in the first chapter. Topics include the taxonomical distinctions of Dicistroviridae and Picornavirales and the biology of RhPV. The identification of RhPV-permissive cell lines, the development of a system for baculovirus-expressed infectious RhPV, and the construction of a full length RhPV infectious clone are also presented. Canonical cap- dependent translation and noncanonical IRES-driven translation are described, with emphasis on translation mediated by the dicistrovirus IGR IRES. At the end of the chapter, the use of small RNA viruses as agricultural biopesticides and future directions in dicistrovirus research are discussed. A justification for the research presented in chapter 2 is provided. Chapter 2 presents work done to investigate translation and replication of RhPV in a plant cellular environment. Last is a general conclusions chapter that describes the implications of this work and future experiments.
Literature review
Introduction
Dicistroviruses are small, positive-strand RNA viruses that infect arthropods. While they share many characteristics with other members of the proposed order Picornavirales, they are distinct in possessing two open reading frames (ORFs), each preceded by an internal ribosome entry site (IRES) (Figure 1A). IRESs are RNA elements that drive translation of an adjacent ORF by a non-canonical, cap-independent mechanism. The intergenic region 2
A. Dicistroviridae Rhopalosiphum padi virus (RhPV) ORF1: nonstructural proteins 5’ UTR IGR ORF2: virion proteins 3’ UTR
SS hel 3A pro pol VP2 VP3 VP1 (A)n VP4
5’ UTR VPg IGR IRES IRES
B. Picornaviridae Poliovirus (PV)
2A IRES VP2 VP3 VP1 2B 2C hel 3A 3C pro 3D RdRp (A)n VP4 pro
5’ VPg cloverleaf
C. Secoviridae Parsnip yellow fleck virus (PYFV)
putative protein CP2 CP3 CP1 hel pro pol
VPg stem loop
D. Secoviridae; subfamily Comovirinae Cowpea mosaic virus (CPMV)
RNA1 RNA2 pro-cofactor hel pro pol (A)n MP CPL CPS (A)n
VPg
nt: 0 1000 2000 3000 4000 5000 6000 7000 8000 9000 10,000
Figure 1. Organization of Picornavirales genomes. (A) The dicistrovirus genome encodes two polyproteins. ORF 1 encodes the nonstructural proteins and ORF2 encodes the virion proteins 1-4 (VP 1-4). The 5’ UTR IRES precedes ORF1 and the IGR IRES precedes ORF2. Genomic RNA is bound by a 3-4 kDa VPg (shown as a circle at the 5’end) and is polyadenylated. Solid vertical lines indicate proteolytic cleavage sites (Liljas et al. 2002; ; Moon et al. 1998; Nakashima and Nakamuru, 2008). A dotted vertical line bisects the 5’ UTR IRES into its minimal active sequences (Groppelli et al., 2007). (B) Picornaviridae genome organization (Kitamura et al., 1981; Racaniello and Baltimore, 1981). (C) Secoviridae genome organization (Turnbull-Ross, et al, 1992; 1993). (D) Secoviridae, subfamily Comovirinae genome organization (Lomonossoff and Shanks, 1983; van Wezenbeek et al., 1983). Abbreviations: SS, silencing supressor domain (in CrPV and DCV); hel, superfamily 3 helicase; 3A, 3A hydrophobic Golgi-ER transport-inhibition membrane protein; VPg, viral protein, genome linked. Found in variable imperfect tandem repeats in dicistroviruses, but not identified in RhPV; pro, chymotrypsin-like cysteine protease; pol, RNA-dependent RNA polymerase; MP, movement protein; CP, Capsid protein; CPL, large capsid protein; CPS, small capsid protein. 3
(IGR) IRES of dicistroviruses is particularly interesting, as it drives the most streamlined form of translation initiation yet known. The IGR IRES-mediated translation occurs independently of eukaryotic initiation factors (eIFs), and translation starts from the ribosomal
A-site in a methionine-independent manner. The activity of the IGR IRES provides a focus for this thesis. Rhopalosiphum padi virus (RhPV) is a dicistrovirus that infects aphids, which are economically important agricultural pests. Research into RhPV is aided by the availability of permissive insect cell lines and a full-length infectious clone. Characteristics of RhPV grant this virus potential for use in the development of agricultural biopesticides.
Future research may not only address dicistrovirus infections of economically important arthropods, but may also shed light on important features of canonical and noncanonical translation.
Picornavirales
Members of the order Picornavirales share several similarities in genome organization
(Figure 1) and virion structure. Member families include Dicistroviridae, Picornaviridae,
Secoviridae (which includes plant viruses formerly classified in the families Comoviridae and Sequiviridae) ,and Marnaviridae, (Le Gall et al., 2008; Sanfaçon et al. 2009). Member genera not assigned to a family include Iflavirus. The proposed order Picornavirales is actually more exclusive than the group less formally referred to as the ―picornavirus-like superfamily‖. Ongoing efforts to organize virus taxonomy may clarify this in the future.
Several criteria distinguish the order Picornavirales. Viruses in the order Picornavirales have small, positive-sense single-stranded RNA (ssRNA) genomes. The 5’ end of a
Picornavirales genomic RNA is bound by a 3-4 kDa VPg (viral protein, genome linked)
(Nakashima & Shibuya, 2006). Most members of the order are polyadenylated at the 3’ end, 4 with the exception of some members of Secoviridae (Figure 1C). While viruses of this order may have segmented genomes (as seen in Secoviridae subfamily Comovirinae) (Figure 1D), only genomic RNAs are translated, rather than utilizing subgenomic RNA. Most have monocistronic RNAs, with the exception of Dicistroviridae (Wilson et al., 2000b) (Figure
1A). ORFs are translated into polyproteins that are subsequently processed autoproteolytically. Proteins are cleaved by the virus’s own chymotrypsin-like cysteine protease (Pro), rather than relying on host proteases (Allaire et al., 1994). Viruses of
Picornavirales also possess a superfamily I RNA-dependent RNA-polymerase (RdRp, or
Pol) and superfamily III helicase (Hel). Genes are distinctively ordered Hel-(VPg)x-Pro-Pol, in which VPgs can occur in imperfect tandem copies (Argos et al., 1984). Virions are about
30 nm in diameter, icosahedral, and non-enveloped. Virions have pseudo-T=3 (p=3) symmetry and are assembled from 60 capsomers, each with three 8-stranded beta barrel
―jelly rolls‖ (Chandrasekar & Johnson, 1998). These capsomers are usually ―pseudo- trimers‖, composed of three separate yet structurally similar virion proteins (VP1-3), about
25 kDa each, and cleaved from the same polyprotein. In Comovirinae, the structural polyprotein is not always cleaved to produce three separate proteins; instead, the three jelly roll domains are in one or two proteins. This still results in p=3 symmetry with the virion assembled from 60 capsomers, rather than T=3 symmetry, in which the virion is assembled from 90 dimers. When a single polyprotein contains structural and nonstructural proteins, structural proteins are located at the N-terminal end. While exceptions exist, members of
Picornavirales generally conform to these criteria.
Some virus families are not included in Picornavirales due to structural, organizational, or sequence divergence, despite being grouped in the picornavirus-like 5 superfamily and sharing a subset of Picornavirales properties, such as virion structure or gene order. These families are Caliciviridae, Hypoviridae, and Potyviridae. Caliciviridae has two to three ORFs, a 12-15 kDa VPg, T=3 symmetry, and 90 kDa structural proteins
(Koopmans et al., 2005). Hypoviridae probably has a double-stranded RNA (dsRNA) genome, is dicistronic, has a superfamily II helicase, and does not code for structural proteins
(Nuss et al., 2005). Potyviridae has a 25 kDa VPg, a different order for nonstructural proteins, and a helical, rather than an icosahedral, virion (Shukla et al., 1994). These characteristics exclude Caliciviridae, Hypoviridae, and Potyviridae families from the proposed order Picornavirales.
Dicistroviruses
Dicistroviruses are members of the recently recognized Dicistroviridae family in the proposed order Picornavirales (Christian et al., 2005a; Le Gall et al., 2008). Dicistroviruses are positive-strand, monopartite RNA viruses. Dicistrovirus virions are approximately 30 nm diameter icosahedrons with pseudo-T=3 (p=3) symmetry, consisting of 60 copies each of the structural proteins, virion protein 1-3 (VP 1-3). Early research with the ―cricket paralysis virus-like‖ arthropod viruses, now designated dicistroviruses, grouped them with picorna-like viruses. This was based on physicochemical properties, virion size and shape, small structural proteins, and characteristic nonstructural proteins (Christian et al., 2000; Moore et al., 1980; Moore & Tinsley, 1982; Scotti, 1985). Sequence data revealed dicistroviruses are distinct from other picorna-like viruses in gene order and separation of nonstructural and structural polyproteins into two ORFs (Govan et al., 2000; Moon et al., 1998; Sasaki &
Nakashima, 1999; 2000; Wilson et al., 2000b). A 5’ UTR IRES drives translation of the nonstructural polyprotein, while structural protein translation is under the control of the IGR 6
IRES (Figure 1A). Observations that nonstructural proteins are present in much lower molar amounts than structural proteins during CrPV or DCV infections of insect cells indicate different activity and mechanisms for the two IRESs (Moore et al., 1980; Wilson et al.,
2000b). This ability to produce a higher molar amount of structural proteins is beneficial to the virus, as a higher amount is needed for the production of new virions.
Dicistroviruses infect several economically important arthropods, including: honeybees, which are hosts of Israeli acute paralysis virus (IAPV), Black queen cell virus
(BQCV), Kashmir bee virus (KBV), and Acute bee paralysis virus (ABPV); aphids, which are hosts of Rhopalosiphum padi virus (RhPV) and Aphid lethal paralysis virus (ALPV); shrimp, which are hosts of Taura syndrome virus (TSV); and fire ants, which are hosts of
Solenopsis invicta virus-1 (SINV-1) (reviewed in Bonning & Miller, 2010). One area of dicistrovirus research currently receiving attention is the investigation of members of the proposed genus Aparavirus. These viruses are of particular interest due to their economic importance. Aparaviruses of bees, most notably IAPV, have been implicated in colony collapse disorder, a phenomenon that decimates commercial honeybee colonies crucial to agriculture (Cox-Foster et al., 2007; Van Engelsdorp et al., 2009; van Engelsdorp et al.,
2008). TSV has a large economic impact on Asian shrimp industries (Lightner et al., 1997;
Overstreet et al., 1997), and SINV-1 may be an important tool for control of invasive red imported fire ants. Work is underway to develop infectious clones to facilitate research of these viruses.
Rhopalosiphum padi virus
Rhopalosiphum padi virus (RhPV) was first identified in laboratory-maintained colonies of Rhopalosiphum padi, the bird cherry-oat aphid (Gildow & D'Arcy, 1988). It is a 7 positive strand RNA virus in the Dicistroviridae family. The icosohedral RhPV virion is 27 nm in diameter and consists of four structural proteins, three of which are surface proteins.
The virus consists of a single messenger-sense RNA with two ORFs. The 5’ ORF encodes the nonstructural proteins, and the 3’ ORF encodes the 4 structural proteins, VP1-4. Each
ORF is translated via a different internal ribosomal entry site (IRES). IRESs are noncoding mRNA sequence that allow for translation of an adjacent ORF without the requirement of a
5’ cap or ribosomal scanning from the 5’ end. The two dicistrovirus IRESs are not related to each other nor to any other known IRESs (Sasaki & Nakashima, 1999).
The 5’ IRES facilitates initiation of cap-independent translation in the absence of eukaryotic initiation factor (eIF) 4E (Woolaway et al., 2001). eIF1, eIF2, and eIF3 are required for 48S complex formation on the 5’ UTR IRES. While not required, eIF1A, eIF4A, and the C-terminal fragment of eIF4G strongly stimulate 48S complex formation on the 5’ UTR IRES (Terenin et al., 2005). The IGR-IRES, however, is able to bypass the initiation phase of translation entirely by forming a complex pseudoknot structure that mimics the structure of tRNA base paired to mRNA in the ribosomal P site (Costantino et al.,
2008; Sasaki & Nakashima, 1999; 2000). This causes the entering ribosome to proceed immediately to the elongation phase of translation on the second ORF. It also allows the virus to bypass host antiviral translation regulation. The 5’ IRES has been shown to function in plant extracts, but this has yet to be shown for the RhPV IGR IRES (Woolaway et al.,
2001) (Table 1).
RhPV has been shown to infect seven economically important aphids: R. padi,
Schizaphis graminum, R. rufiabdominalis, R maidis, Metopolophium dirhodum, Diuraphis noxia, and Sitobion avenae (D'Arcy et al., 1981; Gildow & D’Arcy, 1988; von Wechmar & 8
Table 1. Dicistroviruses, their arthropod hosts, and translation systems supporting 5’ UTR IRES- and IGR IRES-mediated translation in vivo and in vitro.
5' UTR IRES 5' UTR IRES IGR IRES IGR IRES Virus Abbreviation Host Order Function in vivo Function in vitro Function in vivo Function in vitro Genus Cripavirus
Diptera, Drosophila melanogaster, D. melanogaster, Hemiptera, Aedes albopictus, Ae. aegypti, Ae. albopictus, RRL, Cricket paralysis virus CrPV Hymenoptera, Anticarcia gemmatalis, RRL†, 7, 13 Ae. aegypti, T. ni, WGE†, 1, 7, 12, 13 Lepidoptera, Plutella xylostella, S. frugiperda, Bombyx Orthoptera Trichoplusia ni, 7, 13 mori, yeast*, 1, 7, 12, 13
Ae. aegypti, Aphid lethal paralysis virus ALPV Hemiptera ------D. melanogaster, ---- S. frugiperda, B. mori1 Ae. aegypti, Black queen cell virus BQCV Hymenoptera ------D. melanogaster, ---- S. frugiperda, B.mori1 Ae. aegypti, Drosophila C virus DCV Diptera ------D. melanogaster, ---- S. frugiperda, B. mori1 Himetobi P virus HiPV Hemiptera ------Plautia stali intestine virus PSIV Hemiptera ---- S. frugiperda10 ---- RRL, WGE9, 11
D. melanogaster, Rhopalosiphum padi virus RhPV Hemiptera S.frugiperda4, 8, 14 S. frugiperda, S. frugiperda4, 5 RRL4, 5 RRL, WGE4, 8, 14 Xenopus oocytes, Xenopus oocytes, BHK cells, Triatoma virus TrV Hemiptera ---- BHK cells, ---- Ae. albopictus3 Ae. albopictus3 Homalodisca coagulata virus-1 HoCV-1 Hemiptera ------
Proposed genus Aparavirus Acute bee paralysis virus ABPV Hymenoptera ------Taura syndrome virus TSV Decapoda ------B. mori1, 2, 6 RRL, WGE1, 2, 6 Kashmir bee virus KBV Hymenoptera ------B. mori1 ---- Solenopsis invicta virus-1 SINV-1 Hymenoptera ------B. mori1 ---- Israeli acute paralysis virus IAPV Hymenoptera ------
* Note only functions efficiently in yeast with mutations that lower eIF2-GTP/initiator tRNA-met complex levels. † RRL – rabbit reticulocyte lysate, WGE – wheat germ extract.
1 Carter et al., 2008; 2 Cevallos and Sarnow, 2005; 3 Czibener et al., 2005; 4 Domier and McCoppin, 2003; 5 Domier et al., 2000 ; 6 Hatakeyama et al., 2004; 7 Masoumi et al., 2003; 8 Royall et al., 2004; 9 Sasaki and Nakashima; 1999, 10 Shibuya and Nakashima, 2006; 11 Shibuya et al., 2003; 12 Thompson et al., 2001; 13 Wilson et al., 2000b; 14 Woolaway et al., 2001. 9
Rybicki, 1981; Williamson & Rybicki, 1989). Decreased longevity and fecundity result from infection. Gildow and D’Arcy (1988) showed that transovarial vertical transmission from parent aphid to nymph occurs at a rate of 28%, while 87% of adults in an RhPV-infected colony tested positive for RhPV by immunoassay. Horizontal transmission occurs through feeding on plants previously fed on by infected aphids. RhPV circulates through the phloem of the plant without replicating, using it as a passive reservoir (Gildow & D’Arcy, 1988).
Various tests showed horizontal transmission at a rate of about 50%. Infection in the midgut and hindgut tissues results in the release of viruses into the gut and hemocoel. After entering the hemocoel, the virus possibly makes its way to infecting the accessory salivary gland, then the salivary gland (Gildow & D’Arcy, 1988). However, it is not known whether the virus moves through the salivary glands for inoculation into the plant or if inoculation occurs by other means, such as regurgitation. Virions could then be transmitted to the host plant along with saliva while feeding. The fact that plants serve as a reservoir for normal RhPV transmission may prove beneficial for transmission of an engineered aphicidal RhPV virus.
Identification of RhPV permissive cell lines and production of an RhPV infectious clone
Identification of a cell line capable of supporting RhPV replication and viral function was an important step in researching this virus (Boyapalle et al., 2007). From the lepidopteran, dipteran and hemipteran cell lines screened, two hemipteran cell lines that supported RhPV replication were identified. These cell lines were GWSS-Z10 derived from the glassy-winged sharpshooter, Homoladisca coagulata, and Dm1 derived from the corn leafhopper, Dalbulus maidis. After inoculation with a large titer of RhPV wildtype virus
RNA, cytopathic effects (CPE) were observed in the GWSS-Z10 cell line four days post- inoculation (dpi) and 8-10 dpi in Dm1 cells. Northern blot hybrization of RhPV inoculated 10
GWSS cells also showed very small amounts of negative-strand RNA 48 and 96 hours post- inoculation (hpi), but not at 12 hpi, and with diminishing amounts at later time points. Virus- like particles matching the expected size of RhPV (27nm) were detected by electron microscopy within GWSS cells and purified virus preparations. Immunoblot analysis also detected 28 and 30 kDa RhPV coat proteins (but not 29 kDa protein). Virus-like particles were found to be infectious to aphids by membrane feeding.
Further work was done to develop a system for baculovirus-expression of RhPV (Pal et al., 2007). A full length clone of RhPV, along with 121 bases 5’ and 210 bases 3’ of non- viral vector sequence, was inserted into a baculovirus expression vector. Due to a polyhedron promoter and simian virus 40 (SV40) termination/polyadenylation signal, the transcript produced was capped and polyadenylated. Lepidopteran Spodoptera fugiperda
(Sf21) cells were inoculated with the recombinant baculovirus and found to produce RhPV virons in the nuclei. Virions were observed via electron microscopy in cells and virus purifications. Northern blot hybridization and RT-PCR confirmed the presence of RhPV
RNA within these virions. Northern blot hybridization detected RhPV RNA of a size significantly longer than wildtype. Feeding of the baculovirus-expressed RhPV to aphids showed these viruses functioned normally. Thirty-two days after feeding on baculovirus- expressed virions, RhPV6 extracted from the aphids was detected with the 5’ and 3’ vector- derived bases still present. This maintenance of the vector sequence confirms viability of the baculovirus-expressed RhPV, as well as giving an indication that additional sequence can be tolerated by the virus. This opens up the possibility that exogenous genes could be added to the virus for increased pathogenicity by using the virus to deliver a toxin that is aphicidal in the hemocoel but not aphicidal per os (Schmidt et al., 2009). Electron microscopy and 11 immunoblotting 47 dpi also confirmed maintenance of the baculovirus-expressed RhPV in aphids.
To avoid complications of using the baculovirus expression system in the study of
RhPV, an infectious clone was developed from which infectious genomic RNA can be transcribed (Boyapalle et al., 2008). The full RhPV genome, plus 15 non-viral bases at the 5’ end and 5 non-viral bases at the 3’ end, was cloned from the baculovirus-expression vector into vector pGEM3ZF (Promega) to produce RhPV6-1. The genome was inserted between oppositely oriented T7 and SP6 promoters, allowing positive-strand transcription by T7 polymerase and negative strand transcription by SP6 polymerase. Unlike in the baculovirus expression system, these transcripts were not capped or polyadenylated. Furthermore, sequence data showed 119 differences between RhPV6-1 and the previously published RhPV sequence (Moon et al., 1998). Translation in wheat germ extract (WGE) with 35S-methionine radiolabel and a vast abundance of RNA (10 μg) revealed that the infectious clone transcript could be translated to produce the 90 kDa polyprotein expected from translation of ORF2
(Boyapalle et al., 2008). The polyprotein was not proteolytically-cleaved in WGE.
However, translation in WGE occurred at a much lower level for the infectious clone transcript than for wildtype virus. GWSS cells transfected with RhPV6-1 positive-strand transcript showed CPE four dpi. Northern blot hybridization was performed to investigate replication of RhPV transcript in GWSS. RNA extracted from GWSS cells transfected with
RhPV6-1 positive-strand transcript was probed in a northern blot hybridization with a negative strand probe, in order to detect positive-strand RNA. RNA from RhPV6-1 negative-strand transfected cells was probed for detection of negative strand RNA. The observation that full length positive-strand RNA could be detected at later time points than 12 full length negative-strand RNA was offered as evidence for replication on the basis that full- length positive-strand RNA to be detected as late as the 120 hours post-transfection (hpt) timepoint, while a lack of replication meant that full length negative-strand was barely detectable after 48 hpt (Boyapalle et al., 2008). It should be noted that the doublet bands seen in the positive-sense probed (negative-strand detected) northern blot hybridization were far weaker at all time points compared to the negative-sense probed (positive-sense detected) northern blot hybridization.
Strand-specific RT-PCR was attempted to demonstrate a negative-strand replication product in positive-strand transfected GWSS cells (Boyapalle et al., 2008). Amplified fragments indicating detection were seen from RT-PCR at 48, 72, and 96 hpt. Transmission electron microscopy (TEM) revealed 27 nm RhPV-size virus-like particles from virus purifications of GWSS cells four dpt with 20 μg RhPV6-1 positive-sense transcript. This demonstrated that structural proteins could be translated and assembled. TEM of immunohistochemical light microscopy revealed that when these virus-like particles were membrane-fed to aphids, they accumulated in the aphid midgut, as do wildtype RhPV virions.
Aphids as major agricultural pests
Aphids are among the most economically significant insect pests of temperate agriculture (Blackman, 2000). Aphids cause over $1 billion of damage in the US annually through direct damage by phloem-feeding and acting as major plant virus-vectors. The introduction of invasive aphid species, including the soybean aphid, Aphis glycines, has necessitated an increased use of chemical insecticides. Since its introduction to the Midwest in 2000, the invasive soybean aphid has caused yield reductions of 10-40% in soybean fields 13 lacking pest management and has reached population levels as high as >1,000 aphids/plant
(Johnson et al., 2006). As a result, 2.9 million acres were sprayed with an insecticide for soybean aphid in Iowa in 2003 (Pilcher et al., 2005) and more than 1 million acres in 2006.
Problems associated with insecticides include high purchase and application costs, yield loss from driving over fields during application, fossil fuel consumption in production and application, and environmental damage to beneficial insects, including natural predators of insect pests (Pilcher et al., 2005). Also, efficacy of insecticides is likely to go down with the emergence of insecticide-resistant aphids. Development of a sustainable, environmentally benign alternative to chemical pesticides for aphid management has become a priority for the agricultural industry.
RhPV and other small RNA viruses as biopesticides
The use of RhPV as an agricultural biopesticide has been proposed due to the relative simplicity of this small virus, host specificity, and the fact that the virus naturally uses plants as a passive reservoir from which aphid pests acquire infectious viruses (Gildow & D’Arcy,
1988). Advantages of using biopesticides in an integrated pest management strategy as an alternative to chemical pesticides include: a lower cost to growers, preservation of beneficial insects due to the host specificity of the virus, improved safety for humans, and long-term persistence (Ginting & Desmier de Chenon, 1987). Previous research into using small RNA viruses as biopesticides has been done with Helicoverpa armigera stunt virus (HaSV), a tetravirus with a lepidopteran host, and flock house virus (FHV), a nodavirus associated with the grass grub Costelytra zealandica (Christian et al., 2005b; Dasgupta et al., 2001; Gordon et al., 2001). 14
Plants and protoplasts inoculated with FHV supported the synthesis of new infectious virions (Selling, et al., 1990). Barley (Hordeum vulgare), cowpea (Vigna sinensis), chenopodium (Chenopodium hybridum), and tobacco (Nicotiana tabacum and Nicotiana benthamiana) leaves were inoculated with FHV using carborundum, and barley protoplasts were inoculated with FHV using polyethylene glycol. A 100-fold increase in virus production, as well as cell-to-cell and systemic movement of the virus, resulted from FHV infection of transgenic Nicotiana benthamiana expressing movement proteins from tobacco mosaic virus or red clover necrotic mosaic virus (Dasgupta et al., 2001).
In field trials for control of Helicoverpa armigera on sorghum, preparations of HaSV purified from laboratory-infected H. armigera cadavers were found to control the pest as well as the commercial baculovirus biopesticide Helicoverpa zea single-nucleopolyhedrovirus
(HzSNPV, commercial preparation Gemstar) (Christian et al., 2005b). This type of control is easy to implement because insect cadavers can be collected manually after an epizootic, kept at room temperature for years, then ground into a suspension and sprayed on crops.
While this method has been effective with baculoviruses, a lower-cost and more reliable method that does not require infection of caterpillar hosts for virion production has been deemed necessary for small RNA virus biopesticides (Gordon & Waterhouse, 2006).
Suggested alternatives include using a baculovirus expression system or assembly in transgenic plants. Baculovirus expression has not been effective for HaSV, but RhPV produced by baculovirus expression has been found to be infectious (Pal et al., 2007).
Norwalk virus, a calicivirus which can cause gastroenteritis, is a non-plant virus of which capsid proteins have been transgenically expressed and assembled in tomato (Huang et al.,
2005). The assembled virus-like particles (VLPs) mimicked wildtype virus size and 15 symmetry and were highly immunogenic. HaSV genomic RNAs under control of a
Cauliflower mosaic virus 35S promoter was expressed in Nicotiana plumbaginifolia protoplasts (Gordon et al., 2001). When HaSV capsid protein was also expressed from a separate plasmid, infectious virions were produced. This was confirmed by EM and bioassay with host larvae, which demonstrated disease symptoms and HaSV production when larvae were fed VLPs. However, HaSV RNAs were not detected by northern blot hybridization or pulse-labeling. It was concluded that virus assembly could occur in protoplasts independently of replication (Gordon et al., 2001).
After HaSV assembly was demonstrated in protoplasts, attempts were made to produce transgenic tobacco expressing the virus from three separate plasmids (one plasmid expressing RNA1, which includes the replicase, one expressing RNA2, which contains the capsid protein, and one expressing just the capsid protein) (Hanzlik and Gordon, 1998;
Larkin et al. 1996; Schumann et al., 2003). Agrobacterium tumefaciens-mediated transformation produced plants that synthesized infectious HaSV, based on feeding tissue to neonate larvae and observing stunting and detectable HaSV RNA. However, analysis of progeny plants suggested that the transformation was unstable as infectious virions were not produced. Detection of multiple deletions in HaSV genes also raises the question of whether the transformants actually did produce infectious virions, or if there had been contamination with wildtype HaSV. A second strategy to produce transgenic plants expressing HaSV involved transforming the genes individually and then breeding the plants together to produce progeny that would express all three genes (Gordon and Waterhouse, 2006).
Transformants expressing the structural protein alone were shown to produce assembled
HaSV particles, as confirmed by EM, but were only able to maintain this expression for a 16 short time. No genomic RNA was detected from plants transformed with genomic RNA1 or
RNA2 constructs, although a small amount of capsid protein was detected from RNA2 transformants. Gordon and Waterhouse (2006) suspected RNA silencing was responsible for the lack of HaSV RNA expression in plants, and suggested the use of RNA interference
(RNAi) mutant Arabidopsis for future attempts to produce transgenic plants expressing the virus for basic research (Gordon & Waterhouse, 2006). RNA silencing is a major antiviral defense mechanism in plants and insects (van Rij and Andino, 2006). In RNA silencing, dsRNA or dsRNA secondary structural features of ssRNA are cleaved by a Dicer endoribonuclease into discrete-sized fragments characteristic of the particular Dicer involved in cleavage (reviewed in Baulcombe, 2004; Ding and Voinnet, 2007, Aliyari and Ding,
2009). Short interfering RNAs (siRNAs) are derived from perfectly base-paired dsRNAs, whereas microRNAs (miRNAs), involved in regulation of cellular mRNAs, can be derived from imperfectly base-paired dsRNA. siRNAs derived from viruses are termed viRNAs.
Following cleavage by Dicer, fragments are incorporated into an effector complex termed the
RNA induced silencing complex (RISC) along with Argonaute (Ago) protein. Ago1 recruits miRNA and Ago2 recruits siRNA. After the dsRNA fragment is loaded into the RISC, one strand is degraded and the remaining strand serves as a guide-strand to target complementary
RNA for degradation by the RISC. Host RdRps can amplify siRNAs. Viral suppressors of
RNA silencing (VSRs) can target various components of the RNA silencing pathway. Like most known VSRs, the silencing suppressor of Drosophila C virus DCV-1A binds dsRNA, preventing Dicer from cleaving it into siRNA (van Rij et al., 2006). The CrPV silencing suppressor CrPV-1A binds and inhibits the activity of Ago2, but not Ago1, thereby suppressing siRNA-mediated silencing and leaving miRNA-mediated silencing unaffected 17
(Nayak, et al. 2010). Dicistroviruses with silencing suppressors may be more resistant to silencing in plants than HaSV, which does not have a silencing suppressor that can function in plants (Gordon and Waterhouse, 2006). It is also possible that RNAi-pathway mutant plants could be used for transgenic small RNA virus production, since Dicer-like (Dcl) 2, 3, and 4 mutants only show mild phenotypes , though they are less able to respond to environmental stresses and more susceptible to virus infection (Borsani et al., 2005; Xie et al., 2005).
Translation
Canonical translation
Most eukaryotic translation occurs through a cap-dependent scanning mechanism
(Figure 2). This process involves over 1 MDa of various eukaryotic translation initiation factors (Pestova et al., 2007). The process begins with the recruitment of eukaryotic initiation factor 4F (eIF4F). eIF4F contains the 5’ 7-methyl guanosine cap-binding protein eIF4E, the helicase eIF4A, and the scaffolding protein eIF4G, which serves to bind other initiation factors. With eIF4G binding eIF4E and poly(A) binding protein, the mRNA is
Met circularized and the 40S subunit, primed with eIF2–GTP–Met-tRNA i ternary complex and other factors to form the 43S complex, is recruited to the mRNA (Jackson et al., 2010;
Sonenberg & Hinnebusch, 2009). Scanning then proceeds in an ATP-dependent manner until an AUG start codon in a favorable context (i.e. GCC(A/G)CCAUGG) is encountered
(Kozak, 1991; Pestova & Kolupaeva, 2002). The start codon placed in the P-site of the
Met ribosome binds the anticodon loop of the initiator tRNA i, eIF5 facilitates the hydrolysis of
GTP attached to eIF2, and eIF2-GDP is released (Algire et al., 2005). Then, eIF5B facilitates the hydrolysis of GTP to recruit the large ribosomal subunit, and various factors 18
Figure 2. Model of the canonical pathway of eukaryotic translation initiation. (A) Canonical translation initiation begins with the recruitment eIF4F to the 5’ end of the mRNA. eIF4F is composed of cap-binding protein eIF4E, scaffolding protein eIF4G, and helicase eIF4A. (B) Binding of eIF4G to eIF4E and polyA binding protein (PABP) circularizes the mRNA. mRNA is activated by ATP-dependent unwinding of the cap-proximal region by eIF4F and eIF4B. (C) The 43S complex attaches to the activated mRNA. The 43S pre-initiation complex includes the 40S ribosomal subunit, eIF1, eIF1A, Met eIF3, eIF–GTP–Met-tRNA i and probably eIF5. (D) The 43S complex scans the 5’ UTR in the 5’ to 3’ direction. Recognition of the initiation codon leads to 48S initiation complex formation, with displacement of eIF1 to allow eIF5-mediated hydrolysis of eIF2-bound GTP and Pi release. (E) Joining of the 60S subunit and displacement of eIF2 – GDP, eIF1, eIF3, eIF4B, eIF4F, and eIF5, is mediated by eIF5B. After GTP-hydrolysis by eIF5B and the release of eIF1A and GDP-bound eIF5B, the 80S ribosome is ready to begin elongation (See Jackson et al., 2010 for details). 19
5’ cap A 4E 4B 4A AUG 3’ m7GpppN AAAAAAAAAA 4G PABP PABP eIF4F = eIF4E + eIF4G + eIF4A 4E 4B 4A AUG m7GpppN B 4G
PABP PABP 3’ AAAAAAAAAA eIF2 C met-tRNA GTP 43S complex = eIF5 40S + eIF1 + 1 UAC 1A eiF1A +eIF3 + eIF3 E P A eIF2–GTP–Met- Met 40S tRNA i + eIF5 4E 4B m7GpppN 4A AUG 4G
PABP PABP 3’ AAAAAAAAAA D eIF2 met-tRNA GDP Pi eIF5 UAC 4E 4B 1 AUG 1A 48S complex m7GpppN 4A eIF3 E P A 4G 40S E PABP PABP eIF5 eIF2 3’ AAAAAAAAAA GDP
4A eIF3 1 60S 4B 1A
met-tRNA 4G 4E UAC AUG 3’ m7GpppN AAAAAAAAAA E P A Figure 2. (continued) 40S PABP PABP 20 are released from the complex, releasing the ribosome to proceed into elongation (Unbehaun et al., 2004). Elongation proceeds when the next encoded amino acid is brought to the A-site by eukaryotic elongation factor 1a-GTP (eEF1a-GTP) (Valle et al., 2003a). Following hydrolysis of GTP, the aminoacylated-tRNA is bound in the A-site. The peptidyl transferase reaction is catalyzed by the large ribosomal subunit and the growing peptide chain is attached to the tRNA in the A-site (Wintermeyer et al., 2004). The deacylated-tRNA in the P-site is bound by eukaryotic elongation factor 2-GTP (eEF-2-GTP) and ribosomal protein L1 (rpL1) of the L1 stalk, making the tRNA take on a hybrid state (Dorner et al., 2006; Fei et al., 2008;
Moazed & Noller, 1989). Following hydrolysis of GTP from eEF-2, translocation occurs, moving the E-site tRNA out of the ribosome, the P-site tRNA to the E-site, and the A-site tRNA to the P-site, leaving the A-site ready to accept the next cognate aminoacylated-tRNA
(Spirin, 1985; Valle et al., 2003b).
IRESs as alternatives to canonical translation
Alternative forms of translation initiation exist, allowing cap-independent, end- independent translation in the absence of a full complement of initiation factors. Internal ribosome entry sites (IRESs) are RNA structural elements that recruit the ribosome near the start of an ORF, independently of a 5’ cap and the canonical scanning mechanism (Jang et al., 1988; Pelletier et al., 1988). This mechanism was first reported for poliovirus (PV) and encephalomyocarditis virus (EMCV), both picornaviruses which lack a 5’ cap, use a cap- independent, end-independent translation mechanism (Jang et al., 1988; Pelletier et al., 1988;
Pelletier & Sonenberg, 1988). The cell can regulate the production of proteins through alterations to the translation initiation machinery. Cap-dependent translation efficiency can be lowered during mitosis, apoptosis, and cellular stress via several mechanisms. These 21 include changes in phosphorylation, competitive binding, and capsase cleavage of initiation factors (Ali et al., 2001; Gradi et al., 1998a; Gradi et al., 1998b; Pyronnet, 2000; Pyronnet et al., 2001; Spriggs et al., 2005; Svitkin et al., 1999). Cellular and viral IRESs can efficiently recruit the ribosome despite such changes. This allows for certain cellular mRNAs to remain functional even when canonical translation is suppressed. Viral IRESs can allow translation of viral proteins when cap-dependent translation initiation efficiency is lowered by host antiviral responses or viral inhibition of cap-dependent translation. At least 85 cellular
IRESs and 39 viral IRESs have been identified (Baird et al., 2006).
IRESs are divided into four groups, distinguished by structural complexity and protein factor requirements (Kieft, 2008) (Figure 3). Group 1 IRESs are the most structurally
Met complex and do not require protein factors or initiator methionyl-tRNA i to bind the ribosome and activate translation (Jan & Sarnow, 2002; Sasaki & Nakashima, 1999; 2000;
Wilson et al., 2000a; Wilson et al., 2000b). Each subsequent group of IRESs is progressively less complex and requires a greater number of protein factors to function. Group 1 consists of the intergenic region (IGR) IRESs of dicistroviruses. These IRESs start translation from the A-site rather than the P-site, and independently of an AUG start codon (Sasaki &
Nakashima, 1999; Wilson et al., 2000a). Binding of the cognate aminoacylated tRNA to the first translated codon occupying the A-site leads to pseudotranslocation, in which translocation occurs without a preceding peptidyl transferase reaction. Of all the IRES groups, the most is known about the 3-dimensional structure of the Group 1 IRESs
(Costantino & Kieft, 2005; Costantino et al., 2008; Kieft, 2008; 2009; Pfingsten et al., 2010;
Pfingsten et al., 2006). The structure includes three pseudoknots making up two globular domains: (1) a ribosomal binding domain that recruits the ribosome by binding both subunits 22
eIF2 A met-tRNA GTP eIF5 1 UAC 1A 5’ 4E 4B AUG m7GpppN 4A eIF3 E P A 4G 40S Canonical cap- dependent initiation PABP PABP 3’ AAAAAAAAAA
B eIF2 met-tRNA GTP eIF5 Group 4 IRES 1 UAC 1A (examples: PV, HAV) and 4B 4A AUG 5’ eIF3 E P A Group 3 IRES 4G 40S (examples: FMDV, EMCV) ITAF PABP PABP 3’ AAAAAAAAAA
C eIF2 Group 2 IRES met-tRNA GTP eIF5 (examples: HCV, UAC 4A AUG CSFV) 5’ 3’ eIF3 E P A 40S
D 40S 5’ Group 1 IRES (Dicistroviruses)
3’ E P A
60S
Figure 3. Comparisons of eIF requirements of cap-dependent translation and four IRES groups. (A) Canonical cap-dependent translation, requiring all eIFs and Met- Met Met tRNA i (B) Group 4 IRESs require some canonical eIFs, Met-tRNA i, and ITAFs, and require HeLa cell extract to translate in RRL. Group 3 IRESs require some canonical Met eIFs, Met-tRNA i, and ITAFs. (C) Group 2 IRESs require a subset of canonical eIFs Met Met and Met-tRNA i. (D) Group 1 IRESs require no eIFs nor Met-tRNA i. 23 at the E-site, and (2) a P-site domain that mimics the molecular interactions and structure of an mRNA-bound tRNA and docks in the P-site (Figure 4). The crystal structure of the ribosomal binding domain of Plautia stali intestine virus (PSIV) has been solved to 3.1 Å, and the P-site domain of cricket paralysis virus (CrPV) has been solved to 2.4 Å (Costantino et al., 2008; Pfingsten et al., 2006). An understanding of this structure has hinted at the mechanism by which it functions (discussed in Dicistrovirus IRESs section below).
Met Group 2 IRESs require eIF3, eIF2, and initiator methionyl-tRNA i to function (Kieft et al.,
2001; Otto & Puglisi, 2004). Examples of Group 2 IRESs can be found in hepatitis C virus
(HCV) (family Flaviviridae, genus Hepacivirus), classical swine fever virus (CSFV) (family
Flaviviridae, genus Pestivirus), and porcine teschovirus (family Picornaviridae, genus
Teschovirus). These IRESs are not globular like Group 1 IRESs. Their extended conformation complicates crystal formation and NMR, necessitating a divide-and-conquer strategy of solving secondary structural elements individually (Kieft et al., 2002). Cryo-EM has been used to solve the structure of a ribosome-bound HCV IRES to 20 Å (Spahn et al.,
2001). Like Group 1 IRESs, Group 2 IRES RNA binds the ribosome directly. A number of
HCV stem loops contact the ribosome directly at the E-site, and the bent conformation of the unbound IRES is retained by the ribosome-bound IRES (Otto et al., 2002). When recruiting the ribosome, the HCV IRES first binds the 40S subunit. This complex then recruits eIF3
Met and eIF2-met-tRNA i-GTP ternary complex before recruiting the large ribosomal subunit
(Hellen & Pestova, 1999; Pestova & Hellen, 1999).
Comparisons of Group 1 and 2 IRES structures show no structural similarities, with the overall architecture of the first being globular and the architecture of the second extended. Nevertheless, similarities in IRES functions exist. In both Group 1 and 2 IRESs, 24
A Class 1 B Class 2 Ribosome Binding SL IV Ribosome Binding SL IV Shorter Domain Domain 5’ P2.2 5’ PK III P1.1 PK III P1.1 P2.2 SL V SL V P2.2 L1.1 L2.1 PK II L2.1 PK II L1.1 Longer P-site P-site L1.1 Domain Domain SL III Additional stem loop 3’ 3’ PK I PK II C D SL V tRNA-mRNA P-site Domain PK III SL IV 5’
P1.1 PK II tRNA anticodon loop 3’ mRNA codon L1.1 L1.2
Figure 4. Dicistrovirus IGR IRES structure and molecular mimicry. Secondary structural features of a Class 1 IGR IRES. Boxes surround the ribosomal binding domain and P-site domain as indicated. Structural features are labeled. (B) Secondary structure diagram of a Class 2 IGR IRES. Structural features are labeled and differences with Class 1 IRESs are indicated in gray shaded text. (C) Crystal structure of PSIV IGR IRES ribosome binding domain with structural features labeled. (D) The crystal structure of tRNA bound to an mRNA codon (left) and the crystal structure of the CrPV IGR IRES P-site domain (right), illustrating that nucleotide orientation of a tRNA bound to mRNA is mimicked by the P-site domain. Some bases have been removed for clarity. Figure adapted from (Pfingsten el al., 2008; Pfingsten et al., 2010; Kieft, 2009) 25 binding at the E-site near ribosomal protein (rp) S5 results in a change in 40S subunit conformation, demonstrating active manipulation of the translation machinery (Pfingsten et al., 2010; Pfingsten et al., 2006; Spahn et al., 2001). However, it should be noted that while
Group 1 IRESs are positioned within the intersubunit space, Group 2 IRESs can be found along the solvent-accessible side of the 40S subunit. IRESs of both groups also contact the
L1 stalk of the large ribosomal subunit, though at different places. The binding of this highly mobile element suggests movement may be important in IRES function, either through the ribosome moving the IRES or the IRES manipulating the ribosome. Lastly, in both IRES groups, the most compact regions bind the ribosome, but this binding is not sufficient for
IRES function, and an additional domain is required.
Less is known about the structures of Group 3 and 4 IRESs. Group 3 IRESs require a
Met subset of eIFs, met-tRNA i, and protein called IRES trans-activating factors (ITAFs)
(Spriggs et al., 2005). Translation starts at the 3’ end of the IRES. Examples of Group 3
IRESs can be found in EMCV, foot-and-mouth disease virus (FMDV), and Theiler Murine
Encephalomyelitis virus (TMEV) (genus Picornavirus) (Jang et al., 1988; Kuhn et al., 1990).
Met Group 4 IRESs require a subset of canonical eIFs, met-tRNA i,, ITAFs and supplementation with HeLa cell extract to translate in rabbit reticulocyte lysate (RRL)
(Hellen & Sarnow, 2001). Translation initiates at an AUG downstream of the IRES.
Examples of Group 4 IRESs are found in PV and rhinovirus (Borman & Jackson, 1992;
Pelletier et al., 1988). Groups 3 and 4 might operate by binding initiation factors and ITAFs, which then interact with the ribosome, rather than the IRES RNA binding the ribosome directly. 26
Dicistrovirus IRESs
The 5’ UTR IRES has indistinct boundaries, and partial deletions to the IRES result only in partial reductions in activity (Woolaway et al., 2001). This suggests that the ribosome is recruited by multiple redundant domains, and it is believed that ribosome recruitment relies on long, unstructured poly-U stretches, rather than relying on a structured
IRES (Terenin et al. 2005). The 5’ UTR IRES facilitates initiation at an AUG codon (Royall et al., 2004; Woolaway et al., 2001). In contrast, the IGR IRES was found to initiate translation independently of a methionine at a non-AUG codon. The N-terminal amino acid of dicistrovirus structural polyprotein is not methionine (Johnson & Christian, 1998; Sasaki
& Nakashima, 1999). Methionyl-peptidase treatment showed there is no N-terminal methionine for the peptidase to remove. Also, translation was not inhibited in the presence of edeine, which inhibits the recognition of AUG start codons by ribosomal scanning, indicating AUG-independent translation (Kozak, 1989; Wilson et al., 2000a). Most dicistroviruses code for an alanine in the N-terminal position, except for PSIV, which codes for a glutamine (Sasaki & Nakashima, 2000; Wilson et al., 2000b). Through site-directed mutagenesis, Shibuya et al. (2003) found that any codon can serve as the first decoded codon by IGR IRES-mediated translation.
The structure of the Dicistrovirus IGR IRES allows the RNA-only recruitment of the ribosome in the absence of initiation factors and drives translation from a non-AUG codon
Met independently of methionyl- tRNA i,. This is likely achieved through molecular mimicry of tRNA-ribosome interactions (Figure 4D) and a coordinated series of events coupling IGR
IRES structural changes to ribosome structural changes. A great deal of research has contributed to the current understanding of IGR IRES structure and function. 27
IGR IRES structure (Figure 4) was determined through disruptive and compensatory mutagenesis, as well as phylogenetic comparisons showing conserved RNA structure rather than conserved primary sequence (Kanamori & Nakashima, 2001). IGR IRES structure was also demonstrated by chemical and enzymatic probing (Jan et al., 2001; Nishiyama et al.,
2003). These studies revealed a complex structure with three pseudoknots (PK), with PKIII nested within PKII, along with other well conserved structural elements (Figure 4A, B).
Most IGR sequences are made up exclusively by these structural elements, with the IGR
IRESs of HiPV, TrV and ALPV actually overlapping the 3’ end of ORF1. One can speculate about a timing mechanism by which IGR IRES translation in these viruses is most efficient only when eIF scarcity due to infection prevents 5’ UTR IRES-driven translation. In this hypothetical timing mechanism, 5’ UTR IRES translation would result in IGR IRES disruption by the ribosome, and translation of ORF2 would be most efficient when infection lowers the availability of eIFs.
RhPV is unique in possessing an IGR (533 nt) that is much larger than the IGR IRES
(175 nt) that it contains. Site-directed mutagenesis revealed that all pseudoknots are necessary for IGR IRES activity, and stem loops IV and V (SLIV, SLV) are necessary for the recruitment of the 40S ribosomal subunit (Costantino & Kieft, 2005; Jan & Sarnow, 2002;
Nishiyama et al., 2003). Loop 1.1 (L1.1) was shown to be required for 60S ribosomal subunit recruitment, but not for recruitment of the small ribosomal subunit (Pfingsten et al.,
2006).
Analysis of IGR IRESs also shows that they diverge into two classes (sometimes called types in the literature) (Nakashima & Uchiumi, 2009) (Table 1) (Figure 4A, B).
Dicistroviruses in genus Cripavirus (RhPV, PSIV, CrPV, ALPV, BQCV, DCV, HiPV, TrV, 28 and HoCV-1) have Class I IGR IRESs, and viruses in proposed genus Aparavirus (IAPV,
SINV-1, ABPV, KBV, and TSV) have Class II IGR IRESs. Class II IGR IRESs have longer
L1.1 segments and shorter P2.2 segments compared to Class I IGR IRESs. They also possess an additional stem loop, SLIII, in region 3, which is necessary for translation by Class II IGR
IRESs (Cevallos & Sarnow, 2005; Hatakeyama et al., 2004). Finally, Class I IRESs have a conserved UUAC sequence between SL IV and P2.2. Hydroxy-radical probing and footprinting of unbound and 40S ribosomal subunit bound IGR IRES RNA has shown that the tightly packed structure is preformed before binding of ribosomal subunits (Costantino &
Kieft, 2005; Jan & Sarnow, 2002; Nishiyama et al., 2003). Global structure remains constant, while local changes occur when subunits bind.
Primer extension inhibition (toeprinting) assays were used to reveal where the ribosome binds the IRES. It was shown that the ―initiation triplet‖ of PKI, comprised of the
3’ most IGR IRES nucleotides, occupied the P-site of the assembled ribosome; the first decoded codon occupied the A-site (Jan et al., 2003; Pestova & Hellen, 2003). Also, cyclohexamide was shown to arrest IGR IRES-driven translation after two elongation cycles
(when the first A-site tRNA reached the E-site), demonstrating that translation started from the A-site (Pestova & Hellen, 2003). Eukaryotic elongation factors eEF1A and eEF2, required for canonical translocation, were also required for pseudotranslocation in IGR
IRES-driven translation (Jan et al., 2003; Pestova & Hellen, 2003). Cryo-EM of IGR IRES bound by the 40S ribosomal subunit or the 80S ribosome solved the IGR IRES to 20 Å and
17.5 Å respectively (Spahn et al., 2004). X-ray crystallography was used to solve the ribosome binding domain of PSIV to 3.1 Å and the P-site domain of CrPV to 2.4 Å
(Costantino et al., 2008; Pfingsten et al., 2006) (Figure 4D). 29
Knowledge of the IGR IRES structure has led to an understanding of the mechanism by which it functions. The dicistrovirus IGR IRES bypasses translation initiation, avoiding regulation of this complex process, and proceeds straight into elongation (Kieft, 2009). The
IGR IRES is composed of three regions, each with a pseudoknot (Figure 4). Regions 1 and 2 form one compact body, the ribosome-binding domain, and contain pseudoknots PKII and
PKIII respectively (Costantino & Kieft, 2005; Kanamori & Nakashima, 2001; Pfingsten et al., 2006). PKIII is nested within PKII. Region 2 binds the 40S ribosomal subunit at the E- site by projecting stem-loops SL IV and SL V in the same direction in position to interact with ribosomal protein rpS5 (Jan & Sarnow, 2002; Nishiyama et al., 2003). SLIV has also been shown to bind ribosomal protein rpS25 (Nishiyama et al., 2007). Upon binding the small ribosomal subunit, there is a conformational change in the 40S-IRES structure that allows binding of the large ribosomal subunit (Pfingsten et al., 2010). Loop L1.1 of region 1 binds the 60S subunit at the L1 stalk (Nishiyama et al., 2003; Pfingsten et al., 2010;
Pfingsten et al., 2006). With the ribosome formed around the IGR IRES, region 3 is positioned in the ribosomal P-site. Region 3, which makes up the P-site domain and contains
PKI, structurally mimics a hybrid state tRNA bound to mRNA (Costantino et al., 2008;
Yamamoto et al., 2007). With the initiation triplet of PKI in the P-site, the 3’ proximal codon in the A-site is the first decoded when eukaryotic elongation factor 1 (eEF1) shuttles in aminoacylated-tRNA to bind it (Jan et al., 2001; Wilson et al., 2000a). This codon need not code for methionine (Sasaki & Nakashima, 1999; 2000; Wilson et al., 2000a). Then, eEF2 facilitates a pseudo-translocation event, in which the tRNA-bound codon is moved to the P-site without peptide bond formation occurring. Elongation of the viral polyprotein then proceeds as normal. 30
Though the IGR IRES maintains generally the same global architecture in unbound,
40S subunit bound, and 80S ribosome bound forms, local changes that occur upon binding are likely important to IGR IRES function. Cryo-EM and crystallography show several flexible regions, with the exterior features SL IV, SL V, P1.1, L1.1, and L1.2 being the most dynamic (Pfingsten et al., 2006; Spahn et al., 2004). Changes in flexibility in these regions upon 40S subunit and 80S ribosome binding were examined through selective 2’-hydroxyl acylation analyzed by primer extension (SHAPE) (Pfingsten et al., 2010). Briefly, SHAPE can be used to analyze the degree to which an RNA backbone is constrained by base-pairing or other interactions (Wilkinson et al., 2006). This method is based on the principle that more flexible nucleotides sample more conformations that enhance nucleophilicty of the 2’ hydroxyl. This makes unconstrained nucleotides more reactive with electrophiles such as N- methylisatoic anhydride (NMIA). This reaction with NMIA forms a 2’-O-adduct which stops extension of a radiolabeled primer in a subsequent reverse-transcription reaction.
These stops are detected when product cDNAs are electrophoretically separated on a high resolution gel, providing single nucleotide resolution. SHAPE showed that SL IV and SL V were flexible in the unbound IGR IRES, and less flexible when the IGR IRES is bound to the
40S subunit (Pfingsten et al., 2010). This is consistent with previous conclusions that these stem loops bind the 40S subunit directly. SL IV and SL V were also shown to become more flexible upon binding of the 60S subunit, possibly indicating a conformation change and/or partial release from the 40S subunit following the shape change this subunit undergoes upon formation of the 80S ribosome. Helix P1.1 was also found to be disordered in the unbound state by SHAPE, cyro-EM, and crystallography (Pfingsten et al., 2010; Pfingsten et al., 2006;
Spahn et al., 2004). To analyze the importance of P1.1, mutants were generated. Two 31 mutants were generated in which helix formation would be abolished. This was done by exchanging one half of the helix for its Watson-Crick compliment. A double mutant was also generated in which helix formation was restored. Native polyacrylamide gel electrophoresis (PAGE) of unbound mutant and wildtype IGR IRESs revealed structural differences between wildtype and single-mutant IRESs. Structure was restored to wildtype in the double mutant. This implies that while the helix is not stable in the unbound form, it most likely ―breathes‖ until interaction with the 40S ribosomal subunit. By pre-initiation complex assembly assays with mutant and wildtype IRESs, Pfingsten (2010) also showed
P1.1 helix formation is necessary for 60S subunit binding. This is most likely due to P1.1 constraining the adjacent L1.1 in a way that lets it become structured upon interaction with the 60S subunit. The structuring of P1.1 upon 40S subunit binding may therefore trigger the ability of L1.1 to bind the 60S subunit. SHAPE also showed that L1.1 becomes structurally constrained upon 60S subunit binding. Interestingly, mutants that differed in only two nucleotides also differed in ability to bind the 60S subunit. Mutation of C6065 and U6066
(of the CrPV IGR IRES, while not distorting unbound IRES structure, resulted in an inability to bind the large ribosomal subunit. The same was seen for U6015 and U6017. These four nucleotides are very well conserved in Class I IGR IRESs, but not in Class II IGR IRESs.
The differing ability to bind the 60S subunit based on the identity of these nucleotides suggests a specificity of interaction with the ribosome based on nucleotide identity rather than structure alone. Finally, SHAPE also confirmed L1.2 is dynamic. Insertion mutants and deletion mutants of L1.2 were generated because length is the most conserved feature of
L1.2. Deletion mutants were less flexible than wildtype, but could bind the 60S subunit equally or less well, depending on the position of the deletion. All insertion mutants were 32 less able to form the 80S ribosome. However, all mutants, especially the deletion mutant that bound the 60S subunit as wildtype levels, were greatly deficient in driving translation in a dual-luciferase construct in RRL. This shows there is a functional defect post 80S ribosome formation. Based on the position of P1.2 flanking PK2, the pseudoknot linking the ribosomal binding domain to the P-site domain, this defect may be due to improper placement of the P- site domain. It is suggested that P1.2 may play a role in propagating the allosteric change of
L1.1 becoming structured upon ribosomal binding to the P-site domain, correctly positioning the tRNA-mRNA mimic upon ribosome formation.
In summary, the dynamic nature of several structural elements in the IGR IRES suggests a mechanism by which structural changes of the IGR IRES couple with structural changes with the ribosome in order to achieve ribosome recruitment and translation initiation
(Table 2). In solution, the global structure of the IGR IRES is already formed, folding compactly around P2.2 at the core. This under-wound helix, along with PK III, positions SL
IV and SL V parallel and adjacent to each other, in position to bind the 40S ribosomal subunit. Upon binding of the small ribosomal subunit, P1.1 becomes more structured, constraining the adjacent L1.1. This readies L1.1 for binding with the 60S ribosomal subunit, allowing the loop to become more locally structured. This allosteric change may then be propagated through L1.2 and PKII to properly position the P-site domain for the start of translation.
Goals of thesis research
In preliminary experiments performed by Narinder Pal, oat protoplasts electroporated with wildtype RhPV positive-sense RNA were shown by immunoblot to produce RhPV structural proteins detectable 48 hours post-inoculation (Figure 5A). Aphids fed an 33
Table 2. IGR IRES structural elements and brief descriptions of proposed functions.
Structural Brief description of function element Dynamic. Breathes before 40S is bound. P1.1 Becomes structured at 40S binding, and stabilizes L1.1 for 60S binding Dynamic; structure changes after 40S subunit is bound. L1.1 Binds 60S subunit L1 stalk, gaining local structure Dynamic. Important in providing flexibility or positioning L1.2 P-site domain after ribosome is bound.
PK II Pseudoknot connecting region 1 to region 3. Dynamic
PK III Pseudoknot providing structure for region 2
SL IV Co-planar, project on exterior of IRES, bind 40S subunit at rpS5, rpS25 SL V
Structural core stabilizing region 2, positions SL IV, V P2.2 co-planar in position to bind 40S subunit Pseudoknot that mimicks mRNA-bound PK I tRNA in P-site domain 34
Figure 5. Immunoblot and RT-PCR indicate production of infectious RhPV virions in oat protoplasts electroporated with wildtype RhPV RNA. (A) 28, 29, and 30 kDa structural proteins are detected by immunoblot from oat protoplasts electroporated with 10 μg wildtype RhPV RNA 48 hours post-inoculation. Structural proteins were not detected in mock-electroporated oat protoplasts. (B) RT-PCR amplification of a 1539 bp RhPV fragment or 500 bp BYDV fragment from R. padi aphids inoculated by overnight membrane feeding with oat protoplast-derived virus preparations, then raised on healthy plants for two weeks. Virus preparations were obtained 48 hours post inoculation from oat protoplasts electroporated with wildtype RhPV RNA or Barley yellow dwarf virus RNA. Lane 1 shows 1 kb ladder molecular weight standard. Lanes 2-3 show amplified RhPV fragments from RT-PCR using wildtype RhPV RNA as template. Lanes 4-5 show amplified RhPV fragments from RT-PCR using RNA from aphids fed virus preparations from oat protoplasts electroporated with wildtype RhPV RNA. Lane 6 shows a faint amplified RhPV fragment from RT-PCR using RNA from wildtype RhPV-infected aphids. Lane 7 shows an amplified BYDV fragment from RT-PCR using RNA from aphids fed virus preparations from oat protoplasts electroporated with BYDV RNA. Lane 8 shows the absence of amplified RhPV fragments from RT-PCR using RNA from mock-infected aphids. Arrowheads indicate molecular weight. 35
A. Immunoblot RhPV Mock inoculated inoculated protoplast protoplasts s 30 kDa
29 kDa
28 kDa
B. RT-PCR RhPV BYDV Mock 1kb RhPV protoplast RhPV protoplast infected std virus aphids aphids aphids aphids Lane 1 2 3 4 5 6 7 8
1.6 kb 1.5 kb 1.0 kb
0.5 kb 0.5 kb
Figure 5. Immunoblot and RT-PCR indicate production of infectous RhPV virions in oat protoplast electroporated with wildtype RhPV RNA. 36 enrichment of virus-like particles obtained from these protoplasts 48 hours post-inoculation became infected, as shown by RT-PCR (Figure 5B). However, Northern blot hybridization of the oat protoplasts (not shown) did not detect viral RNA in the first attempts, and it was decided that these tests would be repeated. For production of infectious RhPV virions in transgenic plants for use in aphid management, it is important to address whether negative- strand RhPV RNA is produced in planta. Production of negative-strand RNA, and specifically the intermediate dsRNA would trigger the plant RNA silencing mechanism
(Ding & Voinnet, 2007), as occurred on attempted plant expression of HaSV (Gordon &
Waterhouse, 2006). While high level expression of RhPV in transgenic plants could overcome silencing, this would decrease the efficiency of RhPV production within the plant.
Hence, the first goal of research described in this thesis was to address whether negative- strand RhPV RNA is produced in a plant cellular environment. We found that in oat protoplasts, a small amount of negative-strand RhPV RNA was detectable by strand-specific
RT-PCR 48 hours post-inoculation with positive-strand RhPV infectious transcript.
The second goal was to address whether the RhPV IGR IRES is active in a plant cellular environment. We tested the RhPV IGR IRES in in vitro and in vivo plant translation systems. While many dicistrovirus IGR IRESs have demonstrated in vivo and in vitro activity in a variety of insect, mammalian, and yeast translation systems, and several dicistroviruses IGR IRESs have demonstrated in vitro activity in WGE, no IGR IRES has yet been shown to function in an in vivo plant translation system (Table 1). We found that the
RhPV IGR IRES was functional in WGE, but failed to detect translation in oat protoplasts.
37
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Williamson, C. & Rybicki, E. P. (1989). A comparative study on the cell-free translation of the genomic RNAs of two aphid picorna-like viruses. Arch Virol 109, 59-70.
Wilson, J. E., Pestova, T. V., Hellen, C. U. & Sarnow, P. (2000a). Initiation of protein synthesis from the A-site of the ribosome. Cell 102, 511-520.
Wilson, J. E., Powell, M. J., Hoover, S. E. & Sarnow, P. (2000b). Naturally occurring dicistronic cricket paralysis virus RNA is regulated by two internal ribosome entry sites. Mol Cell Biol 20, 4990-4999.
Wintermeyer, W., Peske, F., Beringer, M., Gromadski, K. B., Savelsbergh, A. & Rodnina, M. V. (2004). Mechanisms of elongation on the ribosome: dynamics of a macromolecular machine. Biochem Soc Trans 32, 733-777. 46
Woolaway, K. E., Lazaridis, K., Belsham, G. J., Carter, M. J. & Roberts, L. O. (2001). The 5' untranslated region of Rhopalosiphum padi virus contains an internal ribosome entry site which functions efficiently in mammalian, plant, and insect translation systems. J Virol 75, 10244-10249.
Xie, Z., Allen, E., Wilken, A. & Carrington, J. C. (2005). DICER-LIKE 4 functions in trans-acting small interfering RNA biogenesis and vegetative phase change in Arabidopsis thaliana. Proc Natl Acad Sci USA 102, 12984–12989.
Yamamoto, H., Nakashima, N., Ikeda, Y. & Uchiumi, T. (2007). Binding mode of the first aminoacyl-tRNA in translation initiation mediated by Plautia stali intestine virus internal ribosome entry site. J Appl Biol Chem 282, 7770-7776.
47
CHAPTER 2. INVESTIGATION OF TRANSLATION AND
REPLICATION OF RHOPALOSIPHUM PADI VIRUS
(DICISTROVIRIDAE), IN A PLANT CELLULAR ENVIRONMENT
Zachary Regelin, W. Allen Miller, Bryony Bonning
Abstract
Dicistroviruses are small, monopartite viruses with two open reading frames, each preceded by an internal ribosome entry site (IRES). Availability of an infectious clone of
Rhopalosiphum padi virus (RhPV) provides a useful molecular tool for the investigation of dicistrovirus translation and replication. Cross-kingdom analysis of translation and replication of this insect virus in a plant cellular environment could elucidate key factors involved in these processes and prove vital in efforts to apply dicistroviruses in transgenically expressed biopesticide management strategies. Dual luciferase constructs incorporating the intergenic region (IGR) IRES were constructed to test IRES function in wheat germ extract (WGE) and oat protoplasts. IGR IRES translation was observed in WGE, but translation in oat protoplasts was insufficient to show IGR IRES function. Replication of capped and polyadenylated RhPV infectious clone transcripts electroporated into oat protoplasts was assayed. Negative-strand replication products were detected 48 hours post- electroporation of protoplasts with strand-specific RT-PCR, but were undetected via northern blot hybridization.
48
Introduction
Dicistroviruses are positive-sense monopartite RNA viruses with two open reading frames (ORFs) that encode two polyproteins (Nakashima & Uchiumi, 2009). Translation of these polyproteins is driven by two distinct internal ribosome entry sites (IRESs) (Moon et al., 1998; Wilson et al., 2000b). IRESs are RNA elements that recruit and activate ribosomes in proximity of the start of an ORF, in a fashion differing from the canonical 5’ cap- dependent scanning mechanism (Garrey et al., 2010; Jackson, 2005; Kieft, 2009). A highly unstructured, U-rich 5’ IRES drives translation of nonstructural proteins from the first ORF of the dicistrovirus genome (Terenin et al., 2005; Woolaway et al., 2001). Translation of capsid proteins is driven by the well-characterized dicistrovirus intergenic region IRES (IGR
IRES) (Jan, 2006; Kanamori & Nakashima, 2001). This highly structured element contains two tightly-folded modular domains, the ribosome binding domain and the P-site domain, which recruit the ribosome and initiate translation from the A-site of the ribosome in a methionine-independent manner without the need for eukaryotic initiation factors or ternary complex (Kieft, 2009; Sasaki & Nakashima, 2000).
Most eukaryotic translation occurs through a cap-dependent scanning mechanism, which involves over 1 MDa of various eukaryotic translation initiation factors (Pestova
2007). The dicistrovirus IGR IRES bypasses translation initiation, and proceeds straight into elongation thereby avoiding regulation of this complex process, which can be part of the host’s antiviral response (Sasaki & Nakashima, 1999; 2000; Wilson et al., 2000a; Wilson et al., 2000b). The IGR IRES is composed of three regions, each with a pseudoknot. Regions 1 and 2 form one compact body, the ribosome-binding domain, and contain 49 pseudoknots PKII and PKIII respectively (Costantino & Kieft, 2005; Kanamori &
Nakashima, 2001; Pfingsten et al., 2006). PKIII is nested within PKII. Region 2 binds the
40S ribosomal subunit at the E-site by projecting stem-loops SL IV and SL V in the same direction and in position to interact with ribosomal protein rpS5 (Jan & Sarnow, 2002;
Nishiyama et al., 2003). SL IV has also been shown to bind ribosomal protein rpS25
(Nishiyama et al., 2007). Upon binding the small ribosomal subunit, there is a conformational change in the 40S-IRES structure that allows binding of the large ribosomal subunit (Pfingsten et al., 2010). Loop L1.1 of region 1 binds the 60S subunit at the L1 stalk
(Nishiyama et al., 2003; Pfingsten et al., 2010; Pfingsten et al., 2006). With the ribosome formed around the IGR IRES, region 3 is positioned in the ribosomal P-site. Region 3, which makes up the P-site domain and contains PKI, structurally mimics a hybrid state tRNA bound to mRNA (Costantino et al., 2008; Yamamoto et al., 2007). With the initiation triplet of PKI in the P-site, the 3’ proximal codon in the A-site is the first decoded when eukaryotic elongation factor 1 (eEF1) shuttles in aminoacylated-tRNA to bind it (Jan et al., 2001;
Wilson et al., 2000a). This codon need not code for methionine. Then, eEF2 facilitates a pseudo-translocation event, in which the tRNA-bound codon is moved to the P-site without peptide bond formation occurring. Elongation of the viral polyprotein synthesis then proceeds as normal.
Dicistroviruses infect several economically important arthropods, including: honeybees, which are hosts of Israeli acute paralysis virus (IAPV), Black queen cell virus
(BQCV), Kashmir bee virus (KBV), and Acute bee paralysis virus (ABPV); aphids, which are hosts of Rhopalosiphum padi virus (RhPV) and Aphid lethal paralysis virus (ALPV); 50 shrimp, which are hosts of Taura syndrome virus (TSV); and fire ants, which are hosts of
Solenopsis invicta virus-1 (SINV-1). (reviewed in Bonning & Miller, 2010)
The use of transgenically-expressed dicistroviruses in plants for the management of pestiferous insects has been proposed due to the relative simplicity of the dicistrovirus genome and the fact that some dicistroviruses, including RhPV, are naturally vectored by plants (Bonning & Miller, 2010; Gildow & D’Arcy, 1988). Previous attempts to transgenically express an insect virus, Helicoverpa armigera stunt virus, as a biopesticide in plants encountered problems with RNAi silencing (Gordon & Waterhouse, 2006). Work with flock house virus has shown promise, as this insect virus has been shown to replicate and disperse through inoculated plants expressing movement proteins (Dasgupta et al., 2001)
(Discussed in greater detail in Chapter 1, RhPV and other small RNA viruses as biopesticides).
Before attempting to transgenically express dicistroviruses in plants as biopesticides, more must be investigated about dicistrovirus function in a plant cellular environment.
Practical applications aside, basic questions on virus function and cross-kingdom translational mechanisms will also be addressed through this line of inquiry. Elucidating whether dicistrovirus IGR IRESs are functional and competitive in plants would indicate whether fundamental factors required for IGR IRES-driven translation in animal cells are also present and comparable in plant cells. If not, noting differences between plant and animal factors could help elucidate important factors involved in IGR IRES mediated translation. In this study, we assessed the potential for RhPV replication in a plant cellular environment by using RT-PCR and northern blot analysis. Evaluation of whether RhPV can replicate in planta provides insight into the potential for plant expression of RhPV without 51 induction of the plant silencing system through production of RhPV dsRNA. We also evaluated the RhPV IGR IRES activity in dual luciferase transcripts in WGE and oat protoplasts. This will further our understanding of cross-kingdom IRES activity, which is crucial for the potential of dicistrovirus translation functioning in transgenic plants.
Comparisons between the efficiencies of wildtype IGR IRES and a mutant IGR IRES, produced with a disrupted PK1, suggest a significant level of RhPV IGR IRES activity in
WGE. Tests in oat protoplasts lacked sufficiently robust translation to show IGR IRES function.
Replication of previously described RhPV infectious clone transcripts (Boyapalle et al., 2008), modified by the addition of a 60 bp polyA tail, was investigated in oat protoplasts using strand-specific RT-PCR. Strand-specific RT-PCR was able to detect low levels of negative strand replication product in oat protoplasts 48 hours post-electroporation with capped positive-strand RhPV6-1_polyA transcripts. Northern blot hybridization was also attempted to monitor RhPV6-1_polyA transcript replication in oat protoplasts. Northern blot hybridization failed to detect negative-strand replication intermediate in oat protoplasts 48 hours post-electroporation with capped positive-strand RhPV6-1 polyA transcripts.
Results
In vitro translation in wheat germ extract and dual luciferase assays indicate RhPV
IGR IRES functions in an in vitro plant translation system
In vitro translations of dual luciferase transcripts were carried out in wheat germ extract to evaluate function of the IGR IRES in a plant in vitro translation system. Dual luciferase transcripts contained a 5’ rluc sequence with translation dependent on ribosomal scanning, followed by a polylinker region and a 3’ fluc sequence (Figure 1). The RF 52
A. RhPV 6.1 B. RhPV 6.1_polyA
5’ UTR IGR IRES 3’ UTR SmaI T7 T7 5’ UTR IGR IRES 3’ UTR SalI ORF1 ORF2 ORF1 ORF2 EcoRI SP6 EcoRI polyA SP6 C. RF D. RIF
T7 PmlI HpaI T7 IGR IRES PmlI HpaI Rluc Fluc AmpR Rluc Fluc AmpR
E. RImF F. RsF
T7 mut IGR IRES PmlI HpaI T7 PmlI HpaI Rluc Fluc AmpR Rluc Fluc AmpR Stop
G. RsIF H. RsImF
T7 IGR IRES PmlI HpaI T7 mut IGR IRES PmlI HpaI Rluc Fluc AmpR Rluc Fluc AmpR Stop Stop I. RsFa J. RsIFa
T7 NotI T7 IGR IRES NotI Rluc Fluc AmpR Rluc Fluc AmpR Stop polyA Stop polyA
K. RsImFa L. RFa
T7 mut IGR IRES NotI T7 NotI Rluc Fluc AmpR Rluc Fluc AmpR Stop polyA polyA Figure 1. Infectious clone and dual luciferase construct maps. (A) RhPV 6.1 infectious clone construct. (B) RhPV 6.1 infectious clone construct with additional 60 bp polyA sequence added. (C-L) Dual luciferase constructs. (C) Dual luciferase construct p2luci in (Grentmann et al, 1998), renamed RF for consistency. (D) RIF construct. (E) RImF construct. (F) RsF construct. (G). RsIF construct. (H) RsImF construct. (I) RsFa construct. (J) RsIFa construct. (K) RsImFa construct. (L) RFa construct. R in the construct name or Rluc on the figure denotes a Renilla
luciferase gene; F / Fluc = Firefly luciferase; I / IGR IRES = RhPV IGR IRES; Im / mut IGR IRES = RhPV IGR IRES with initiation triplet mutated to UGC, disrupting PKI; s / Stop = stop codon introduced to polylinker in Renilla luciferase reading frame; a / polyA = 60 bp polyA; T7 = T7 promoter; SP6 = SP6 promoter; AmpR = ampicilin resistance gene. Relevant restriction sites are shown in italics. 53 transcript (named p2luci in Grentzmann et al.(1998), but renamed here for consistency) contains an 18 codon polylinker region devoid of in-frame stop codons (Figure 1C).
Therefore translation results in a rluc-fluc fusion protein that can serve as a positive control for rluc and fluc assays. The RIF construct (Figure 1D) was produced by cloning the 175 bp wild-type IGR IRES sequence of RhPV into the polylinker region of RF, in-frame and two codons upstream of the fluc gene. Translation of fluc is therefore dependent on IGR IRES function. The RImF construct (Figure 1E) was produced as a negative control with the CCU initiation codon of the IRES mutated to UGC. This mutation disrupts the formation of pseudoknot I (PKI) of Domain 3 and has been shown to drastically decrease IGR IRES- driven translation (Sasaki & Nakashima, 2000). A dual luciferase translation system was used because both proteins can be assayed independently in the same reaction mixture and ribosomal scanning dependent translation of rluc serves as an internal positive control. In
IGR IRES containing transcripts, the fluc/rluc ratio can be used as the normalized measure of
IRES activity, with a higher ratio indicating a higher proportion of IGR IRES driven translation. Also, the IGR IRES is positioned between two ORFs, in an analog of its natural context in a virus.
Linearized constructs were templates for in vitro transcription of positive-sense RNA using T7 RNA polymerase. Three wheat germ extract translation replicates were set up for each transcript. After 1 hour of translation, fluc/rluc activities were measured (Figure 2A).
The fluc/rluc ratio was significantly higher for the RIF translation products compared to
RImF translation products (p-value = 0.0076; ~3-fold higher). This suggests that the IGR
IRES drives translation in wheat germ extract. Translation products from RIF and RImF transcript had a far smaller fluc/rluc ratio than the activity ratio produced by the fusion 54
WGE Fluc/Rluc Ratios
0.05 e A B C 0.04
0.03
0.02 c f
fluc /fluc rluc ratio d f 0.01 a a b a 0.00 RsIFRIF RImFRImF RF RsIFRIF + RImFRImF + RFRF + RsIF RsImFRsImF RsF Mutant Mutant Mutant IRES IGR IRES No IGR IRES No IGR IRES No IGR IRES IGR IRES IGR IRES Only in Only in Only in Only in Stop codons No No Yes Yes Yes IRES IRES IRES IRES 3' UTR 10 nt 10 nt 10 nt 161 nt 161 nt 161 nt 161 nt 161 nt 161 nt
Figure 2. IGR IRES activity in WGE. Ratios of fluc/rluc activity from translation reactions in WGE for different transcripts. Presence of RhPV IGR IRES, mutant IGR IRES, or no IRES is indicated, as is the presence of rluc-frame stop codons in the polylinker and the length of each transcript’s 3’ UTR (A)
Translations of RIF, RImF, and RF with short 3’ UTRs. These transcripts lack rluc-frame stop codons in the polylinker between rluc and the IGR IRES. (B)
Translations of RIF, RImF, and RF with longer 3’ UTRs. (C) Translations of RsIF, RsImF, and RsF with long 3’UTRs. These transcripts have rluc-frame stop codons in the polylinker between rluc and the IGR IRES. Error bars show standard deviation. Different letters above bars indicate statistically significant difference by paired t-test. 55 protein product of RF (RF ratios were roughly five times those of RIF), indicating that cap- dependent translation greatly exceeded IRES-driven translation in this system.
To produce constructs with increased IGR IRES function, the polylinker regions of
RIF and RImF were manipulated to move two stop codons into the rluc reading frame between rluc and the IGR IRES. This would prevent the ribosome from translating into the
IGR IRES sequence and disrupting the formation of helix P1.1, which is required for the formation of the IRES-80S ribosome complex. These constructs were named RsIF and
RsImF (Figure 1G, H). The same change was made to the RF construct to introduce two stop codons between the rluc and fluc genes and move fluc out of the rluc reading frame. This was done to produce RsF, a negative control construct in which no fluc translation is expected (Figure 1F). Translation and dual luciferase assays were repeated with RNA transcribed from these new constructs, as well as RNA from the previous constructs for comparison (Figure 2A, C). The fluc/rluc ratios were greater in RsIF than in RIF, as expected (p-value = 0.0401; ~38% higher ratio). Stop codons added to the polylinker region in RsF lowered the fluc/rluc ratio but did not eliminate fluc activity as expected (p-value =
0.0018; ~4.2 fold decrease of RsF vs. RF fluc/rluc ratios). Again, fluc/rluc activity ratios indicated that translation of fluc driven by a wild type IGR IRES was 3.7 times the translation driven by the mutant IGR IRES (p-value = 0.0038). However, fluc/rluc ratios were nearly as high for RsF transcripts as they were for RsIF transcripts (p-value = 0.1024).
Comparison between transcripts also showed that a longer 3’ UTR in the transcripts from
HpaI-linearized constructs resulted in greater fluc translation, compared to transcripts cut with the PmlI site added for linearization in Grentzmann et al. (1998) ( p-value = 0.0013,
0.0076 , 0.0056 for RIF, RImF, and RF comparisons, respectively) (Figure 2A, B). 56
Next, we tested constructs with a 3’ 60 nt polyA tail, because RhPV RNA is polyadenylated. Constructs RsIFa, RsImFa , RsFa, and RFa (Figure 1I-L) were linearized and used to transcribe 7-methyl guanosine-capped positive-sense transcripts. When these transcripts were translated in wheat germ extract, rluc and fluc luciferase assays gave readings roughly one order of magnitude higher than was observed with uncapped, non- polyadenylated transcripts, but otherwise showing the same patterns in fluc/rluc ratios
(Figure 3A). Activity ratios from RsIFa translation product and RsFa were similar and higher than the fluc/rluc ratio from RsImFa (RsIFa to RsImFa p-value = 0.1002; RsFa to
RsImFa (p-value = 0.0187). The greatest fluc/rluc ratio was seen from the fusion protein product of RFa.
To discern why the RsF and RsFa transcripts produced a translation product with fluc activity in the absence of an IRES, we determined the size of the translation products by polyacrylamide gel electrophoresis (PAGE). Following PAGE, phosphorimager scans
35 revealed S methione-radiolabeled translation products of expected sizes for RsIFa, RsImFa,
RFa, and firefly luciferase control translation reactions (Figure 4). Scans also revealed that translation products from RsFa contained rluc, fluc, and rluc-fluc fusion protein.
Furthermore, fluc/rluc ratios for RsIFa and RsFa are similar according to peak, average, trace, Gaussian peak, Gaussian trace, and relative quantity analyses of the images. It is unclear how RsFa transcripts can be transcribed to produce rluc/fluc fusion proteins, despite the presence of rluc frame stop codons in the polylinker, and the fact that rluc and fluc do not share a frame.
57
A B
WGE Fluc/Rluc Ratios RRL Fluc/Rluc Ratios 0.010 0.010 a
c 0.008 0.008 a,b b 0.006 0.006 b
0.004 0.004
fluc / rluc ratio / rluc fluc fluc / rluc / ratio rluc fluc
a 0.002 0.002
0.000 0.000
RsIFa RsImFaRsImFa RsFa RsIFa RsImFaRsImFa RsFa
C Oat Protoplast Fluc/Rluc Ratios 0.00020 b 0.00015
0.00010 a
0.00005 a fluc / rluc / ratio rluc fluc
0.00000
-0.00005
RsIFa RsIRsImFamFa RsFa
Figure 3. IGR IRES activity in WGE, RRL, and oat protoplasts. Ratios of fluc/rluc activity from translation reactions in (A) WGE, (B) RRL, and (C) oat protoplasts for capped and polyadenylated transcripts of RsIFa, RsImFa, and RsFa. Error bars indicate standard deviation. Different letters above bars indicate statistically significant difference by paired t-test. 58
Firefly kDa RsIFa RsImFa RsFa RFa control 250 -
150 -
100 - 100 fusion 75 - 61 Fluc 50 -
37- 36 Rluc
25 - 20 - 15 -
10 -
Figure 4. SDS-PAGE fractionation of WGE translation products from reactions with dual luciferase transcripts. Phosphorimager scan of radiolabeled translation products from WGE translation reactions with RsIFa, RsImFa, RsFa, RFa, and firefly control transcripts. Translation products were separated by SDS-PAGE. Precision plus protein standard is shown to the left with molecular weights (kDa) indicated. Arrowheads indicate the expected sizes of fluc-rluc fusion protein, fluc, and rluc. 59
In vitro translation in rabbit reticulocyte lysate
Because RhPV IGR IRES function was demonstrated previously in rabbit reticulocyte lysate (Domier & McCoppin, 2003; Domier et al., 2000), capped positive-sense transcripts from RsIFa, RsImFa, RsFa, and RFa constructs were tested in this system (Figure 3B).
Translation products were used in dual luciferase assays. Translation products from RsIFa transcript had significantly higher fluc/rluc ratios than the RsImFa and RsFa translation product (RsIFa to RsImFa p-value = 0.0057; RsIFa to RsFa p-value = 0.0061; RsFa to RsImFa p-value = 0.0102). Fluc/rluc ratios were roughly twice as high in RRL as they were in WGE.
The fusion protein from RFa translation gave a fluc/rluc ratio roughly 18 times higher than that of RIFa translation product.
Translation in oat protoplasts
7-methyl guanosine capped positive-sense transcripts of RsIFa, RsImFa, RsFa, and
RFa, as well as uncapped fluc single luciferase transcript, were electroporated into oat protoplasts. Mock electroporations without transcript were also performed as negative controls. Following 4hr translation reactions, the luciferase readings from lysates for each transcript were measured (Figure 3C). Luciferase levels were 2-3 orders of magnitude lower than readings from in vitro translations. fluc levels expressed from RsIFa, RsImFa, and RsFa were near those from lysates from mock electroporations, making it difficult to draw conclusions about IGR IRES function in oat protoplasts. While RsFa transcripts gave fluc readings significantly greater than the fluc readings of the mock electroporation protoplast lysates (some rluc-fluc fusion protein and fluc may be translated in protoplasts, as they were in wheat germ extract and rabbit reticulocyte lysate), there was no statistically significant difference between mock electroporation fluc levels and fluc levels from RsIFa and RsImFa 60 protoplast lysates. This could indicate that the RhPV IGR IRES is not functional in plant protoplasts, or that it is not competitive for the ribosome when canonical translation is active.
Northern blot hybridization of RNA extracted from oat protoplasts electroporated with
RhPV6-1_polyA positive-strand transcript failed to detect negative-strand replication product 48 hours post-electroporation
Replication of viral RNA is a sensitive test for the ability of the RNA to be translated, because translation of the viral replication genes is required for replication. To determine whether RhPV replicates in oat protoplasts, we first attempted to detect accumulation of viral
RNA by northern blot hybridization. Protoplasts were electroporated with 10 ug of RhPV6-
1_polyA positive-strand capped transcript (Figure 1B), RhPV6-1_polyA negative strand transcript, PAV6 Barley yellow dwarf virus infectious clone positive-sense transcript, or mock transfected with water. RNA was extracted from protoplasts electroporated with
RhPV6-1_polyA positive-strand capped transcript 1 hour, 24 hour, and 48 hours post- electroporation. RNA from remaining protoplasts was extracted 48 hours post- electroporation. Northern blot hybridization using positive-strand radiolabled RNA probe for the detection of negative-strand RhPV6-1_polyA RNA detected fragments only for samples electroporated with negative-strand RhPV6-1_polyA transcript and for RhPV6-1_polyA negative strand transcript (Figure 5, lanes 11-13, 22). A lack of detectable full-length negative-strand RNA from RhPV6-1_polyA-electroporated protoplasts does not necessarily rule out replication because a small amount of negative-strand RNA may not be detectable when large amounts of positive-strand RNA are present. An increase in positive-strand RNA was not detectable due to the massive amount of inoculum used.
61
6.1 polyA2 T7 capped transcript PAV6 T7 capped transcript 6.1 polyA2 PAV T7 6.1 polyA2 SP6 transcript 6.1 polyA2 T7 capped pos. sense SP6 capped H2O 1 hr 24 hr 48 hr 48 hr 48 hr 48 hr 1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22
Figure 5. Northern blot hybridization of RhPV 6.1_polyA inoculated oat protoplasts. Northern blot hybridization with RhPV 6.1 positive-sense probe for detection of negative- sense RhPV RNA. (Lanes 2-10) Northern blot hybridization of RNA extracted from oat protoplasts 1, 24, or 48 hr after electroporation with capped RhPV 6.1_polyA positive- sense transcript. (Lanes 11-13) RNA extracted from oat protoplasts 48 hr after electroporation with RhPV 6.1_polyA negative-sense transcript. (Lanes 14-16) RNA extracted from oat protoplasts 48 hr after electroporation with capped PAV6 positive- sense transcript. (Lanes 17-19) RNA extracted from oat protoplasts 48 hr after mock electroporation with water. (Lane 20) Capped RhPV 6.1_polyA positive-sense transcript. (Lane 21) Capped PAV6 positive-sense transcript. (Lane 22) RhPV 6.1_polyA negative- sense transcript. Lanes 2-19 show a phosphorimager scan from a screen exposed for 24 hrs. Lanes 20-22 show a 1 minute exposure. 62
Strand-specific RT-PCR of oat protoplasts electroporated with positive-strand
RhPV6-1 transcript suggests replication in protoplasts
We next attempted RT-PCR to detect negative strand genomic RNA accumulation because it is far more sensitive than northern blot hybridization. RT-PCR employing a tag system for strand specificity, as adapted from Lin (2002) and Plaskon (2009) (Figure 6), was used to separately detect positive- and negative-sense RhPV viral RNA sequence in inoculated oat protoplasts. Protoplasts were electroporated with positive-sense transcripts from infectious clone RhPV6-1 and incubated for 1, 24 or 48 hr before extracting RNA.
Protoplasts were also electroporated with positive-sense transcript from clone
RhPV62A_DA, which harbors a lethal frameshift mutation in the RNA-dependent RNA polymerase coding region, as a negative control for replication. RNA was extracted from
RhPV62A_DA -inoculated protoplasts 48 hr after electroporation. As positive controls for strand-specific RT-PCR, reactions were carried out using positive- or negative-strand transcripts (Figure 7A). Reactions designed to detect negative-strand RNA used tag1-ZF5 primer for reverse transcription and tag1 primer and ZR6 in PCR (Table 1). Reactions designed to detect positive-strand RNA used tag1-ZR6 primer for reverse transcription and
ZF5 primer and tag1 in PCR. Accurate strand-specific detection of positive-sense and negative-sense transcripts was confirmed by the presence of a 1 kb band with known transcripts as template (Figure 7A, lanes 2 and 3). This band was absent when the template was positive transcript and the primers were designed for negative-strand detection, and vice- versa (Figure 7A, lanes 1 and 4). To rule out DNA contamination of transcripts or mispriming, reactions were carried out without an RT step, without reverse transcriptase, or without primer in the RT step (Figure 7B, lanes 1-3, 4-7, and 8-11). Lack of any product 63
Figure 6. Schematic of strand-specific RT-PCR using nongenomic tag sequences. The figure illustrates how a non-genomic tag sequence can be utilized in RT-PCR to ensure strand-specificity. In the described system, RT-PCR can amplify a product when one strand is present, but no product can be produced from that strand’s complement. For comparison, standard RT-PCR not utilizing a nongenomic tag can lead to false-positive detection of a strand if mispriming leads to amplification from the complementary strand. (A) Hypothetical viral sequence with forward primer, forward primer complement, reverse primer, and reverse primer complement sequences indicated. (B) A primer with a 5’ nongenomic sequence and a 3’ viral sequence (tag-viral primer) primes first strand synthesis from complementary viral RNA in reverse transcription. A cDNA with a 5’ nongenomic tag is produced. Following exonuclease I and RNAse H treatment, PCR with tag sequence primer (no 3’ viral sequence on this primer) and viral sequence complement primer produces fragments of an expected size. (C) A mispriming event with a tag-viral primer will not lead to PCR amplification. If the tag-viral primer is able to prime first strand synthesis of cDNA from its same-sense RNA strand, rather than its complement, the resulting cDNA will not be amplified by subsequent PCR. In PCR with tag sequence primer and viral sequence complement primer, neither primer will be complementary to the cDNA sequence, and PCR priming will not occur. Therefore, RT-PCR using a tag-viral primer in reverse transcription is strand-specific, and will not result in a false-positive indication of a strand’s presence if its complement is misprimed. (D) For comparison, a mispriming event in RT-PCR using standard genomic primers can lead to a false-positive in the form of a PCR product of the expected size. When a viral sequence primer primes first-strand synthesis from its same-sense RNA, and the false-priming occurs downstream of the region intended to be amplified and near enough that the cDNA includes it, subsequent PCR can amplify the intended region and result in an amplified fragment of the expected size. 64
A (+) sense (-) sense
B
C
D
Figure 6. Schematic of strand-specific RT-PCR using nongenomic tag sequences. 65
Figure 7. Strand-specific RT-PCR detection of positive- and negative- strand RhPV6.1 RNA from oat protoplasts electroporated with RhPV6.1 positive-strand transcripts demonstrates RhPV6.1 replication in protoplasts. RT-PCR using tag1-ZF5 as the primer for reverse transcription (first strand synthesis) and primers tag1 and ZR6 for PCR detected negative-strand RhPV 6.1 RNA. RT-PCR using tag1-ZR6 as the primer for reverse transcription and primers ZF5 and tag1 for PCR detected positive-strand RhPV 6.1 RNA. (A) Lanes 1-4 show RT-PCR with RhPV 6.1 T7 or RhPV 6.1 SP6 transcripts. Primer sets with tag1-ZF5 for first strand synthesis and tag1, ZR6 for PCR showed bands only for negative strand RNA (SP6 transcripts). Primer sets with tag1-ZR6 for first strand synthesis and ZF5, tag1 for PCR showed bands only for positive- strand RNA (T7 transcripts). Lanes 5-7 show PCR using RhPV 6.1 plasmid as template. Presence of plasmid only results in a band from PCR with ZF5 and ZR6 primer in the PCR positive control in lane 5, and not in the PCR reactions with tag1 and ZR6 in lane 6 or ZF5 and tag1 in lane 7. (B) Lane 1 shows a negative control for PCR with water and ZF5 and ZR6 primer (no DNA or RNA sample added). Lanes 2 and 3 show negative PCR controls in which 6.1 T7 or SP6 transcripts are added directly to PCR with ZF5 and ZR6 primers (no RT step). Lanes 4-7 show negative RT-PCR controls in which 6.1 T7 or SP6 transcripts were added to first strand reverse transcription reactions with tag1- ZF5 primers or tag1-ZR6 primers, but with reverse transcriptase absent. First strand reaction products were then used in PCR reactions with the appropriate primer pair: tag1 and ZR6 or ZF5 and tag1. Lanes 2-7 demonstrated that there was no DNA contamination of the 6.1 T7 or 6.1 SP6 transcripts. Lanes 8-11 show RT-PCR reactions in which no first-strand primer was used in order to confirm that tag1 will not prime PCR reactions that lack first strand PCR product. (C) Lanes 1-6 show RT-PCR with RNA extracted from oat protoplasts 0, 24 or 48 hours post-electroporation with RhPV 6.1 T7 transcript. Bands in lanes 2, 4, and 6 show detection of positive-strand RNA at all time points. A faint band in lane 5, but no bands in lanes 1 or 3 shows detection of negative- strand RNA at the 48 hr timepoint only. Lanes 7 and 8 show RT-PCR with RNA extracted from oat protoplasts 48 hours post-electroporation with RhPV DA mutant T7 transcript. Positive-sense but not negative-sense RNA is detected. (D) Lanes 1-8 show RT-PCR reactions with RNA extracted from oat protoplasts as described above, but without reverse transcriptase. These lanes rule out false- priming in PCR with oat protoplast extracted RNA. All reactions shown were done concurrently and all images shown are from the same agarose gel. 66
PCR PCR control for Figure 7 (continued). RT-PCR with transcripts Pos false priming AA control RNA 6.1 T7 6.1 SP6 Plasmid tag1- tag1- tag1- tag1- FS Primer ---- ZF5 ZR6 ZF5 ZR6 Detected (-) (+) (-) (+) ---- Strand PCR tag1, ZF5, tag1, ZF5, ZF5, tag1, ZF5, Primers ZR6 tag1 ZR6 tag1 ZR6 ZR6 tag1
Lane 1 2 3 4 5 6 7
PCR neg Test for DNA contamination of RNA Test for false-priming B control 6.1 RNA H O 6.1 T7 6.1 T7 6.1 SP6 6.1 T7 6.1 SP6 2 SP6 tag1- tag1- tag1- tag1- FS Primer No RT Step No Primer, RT Present ZF5 ZR6 ZF5 ZR6 Detected ---- No RT --- Strand PCR tag1, ZF5, tag1, ZF5, tag1, ZF5, tag1, ZF5, ZF5, ZR6 Primers ZR6 tag1 ZR6 tag1 ZR6 tag1 ZR6 tag1 FigureLane 8 1 2 3 4 5 6 7 8 9 10 11 67
C Figure 7 (continued).
RNA 0 hr 6.1 24 hr 6.1 48 hr 6.1 48 hr DA tag1- tag1- tag1- tag1- tag1- tag1- tag1- tag1- FS Primer ZF5 ZR6 ZF5 ZR6 ZF5 ZR6 ZF5 ZR6 Detected (-) (+) (-) (+) (-) (+) (-) (+) Strand PCR tag1, ZF5, tag1, ZF5, tag1, ZF5, tag1, ZF5, Primers ZR6 tag1 ZR6 tag1 ZR6 tag1 ZR6 tag1 Lane 1 2 3 4 5 6 7 8
D Test for false-priming
RNA 0 hr 6.1 24 hr 6.1 48 hr 6.1 48 hr DA
FS Primer No Primer, RT Present Detected --- Strand PCR tag1, ZF5, tag1, ZF5, tag1, ZF5, tag1, ZF5, Primers ZR6 tag1 ZR6 tag1 ZR6 tag1 ZR6 tag1 Lane 1 2 3 4 5 6 7 8 68
Table 1. RT-PCR primers. tag1 sequence is shown in bold
Primer Length Sequence (5'-3‘) ZF5 24 CAGCCGCAAAACTCTTATTGACTG ZR6 22 CACCATCTCTATCGCCACGAAG tag1 25 GGCAGAGATTTCGATGACCTGATGC tag1-ZF5 45 GGCAGAGATTTCGATGACCTGATGCCGCAAAACTCTTATTGACTG tag1-ZR6 44 GGCAGAGATTTCGATGACCTGATGCCATCTCTATCGCCACGAAG 69 from these reactions confirms that there is no DNA contamination or false-priming that would lead to false-positives in RT-PCR with transcripts. RT-PCR reactions carried out without primer in the RT step also confirmed oat protoplasts were free of RhPV construct
DNA contamination (Figure 7D).
In RT-PCR products from inoculated protoplasts, the 1 kb product was seen for all positive-sense detection reactions (Figure 7C, lanes 2, 4, 6, and 8). RNA from RhPV6-1– electroporated protoplasts incubated for 48 hours was the only protoplast RNA template from which negative-sense detecting RT-PCR produced a product (Figure 7C, lane 5). RT-PCR with RNA from shorter incubations, or from protoplasts electroporated with RhPV62A_DA
RNA-dependent RNA polymerase mutant transcript, did not detect negative-strand RNA.
Lack of negative-strand detection accompanied by robust positive-sense detection of the replication mutant RhPV62A_DA supports the notion that negative-sense detection from
RhPV6-1– inoculated protoplasts indicates authentic replication. These findings suggest that
RhPV6-1 infectious clone transcript is replicative in oat protoplasts. The faintness of the band from RT-PCR with RhPV6-1 electroporated protoplasts incubated for 48 hours compared to the band from RT-PCR with 1.5 pg of negative-sense transcript implies that the amount of single-stranded negative-strand RNA available for reverse transcription is low at
48 hours post electroporation. We conclude that RhPV6-1 may replicate at very low levels in oat protoplasts.
Discussion
Translation assays in wheat germ extract indicate the RhPV IGR IRES functions in a plant translation system because the wildtype IRES out-performs mutant IRES in driving translation. However, unexpected translation from transcripts lacking an IRES, as seen in 70 luciferase assays as well as visualized from radiolabeled translation products, lowers the reliability of these tests. It is also unclear how fluc is translated from transcript lacking any
IGR IRES structure (mutant or wildtype) in levels equaling those of the wildtype IGR IRES- containing transcript’s levels and exceeding those of the mutant IGR IRES-containing transcript. Fluc and rluc-fluc fusion protein were not observed from translations with a similar negative-control construct used by Grentzmann et al. (1998), in which rluc and fluc were out of frame by a single base pair deletion in a dual luciferase transcript (Compare to
RsF and RsFa, which have fluc in the rluc +1 frame, as well as two stop codons in the rluc frame). However, the presence of these proteins may have been obfuscated by the high background of their gels. Repeating translation experiments in a less problematic dual luciferase construct may be advisable. Also, longer translation incubations may enable detection of fluc in protoplasts.
No negative-strand replication product was detected by northern blot hybridization from positive strand electroporated protoplasts extracted at 1, 24, and 48 hr time points. This may simply indicate that such products were not present in adequate quantities for detection at these time points, and does not necessarily mean that replication cannot occur. Negative- strand can be very difficult to detect owing to the massive excess of positive strand RNA to which it may anneal during extraction (Ahlquist, 2002; Buck, 1996). Advantages of northern blot analysis include providing the size of detected strands, which can distinguish full length
RNAs from degradation products. Also, the lower sensitivity of this analysis makes it less likely to return false positive results. The drawback is that it may not be sensitive enough to detect the low amount of negative-strand produced, especially when positive strand is present in such abundance. The more sensitive RT-PCR using a tagged-viral first-strand primer 71 proved to be a reliable system for strand-specific RNA detection from RNA obtained from protoplasts electroporated with RhPV6-1 transcript. Numerous controls support the reliability of this test. While false-positive detection of a strand when only its complement was present often occurred when using an RT-PCR system with only viral sequence primers, the incorporation of nongenomic tags (Lin et al., 2002; Plaskon et al., 2009) in the system used here overcame these problems. The use of tag-viral primers does not add to the time or difficulty of the RT-PCR procedure, thus the added reliability of this system makes it advisable to use for future strand-specific RNA detection.
The dicistrovirus IGR IRES drives the most streamlined form of translation initiation known to date. As such, it may be an important molecular tool for elucidating key steps and factors in translation. An understanding of IGR IRES functionality in a plant cellular environment could be important in comparing fundamentals of translation in plant and animal cells. An understanding of dicistrovirus translation and replication in a plant cellular environment could prove vital for the application of transgenically produced insect specific virus as a crop biopesticide. While IGR IRES-driven translation was observed in a wheat germ extract translation system, any potential IGR IRES-driven translation in oat protoplasts was below detectable levels. Similarly, while RT-PCR results suggest replication of RhPV infectious clone in oat protoplasts, the amount of negative sense replication product was below detectable levels via northern blot analysis. While low levels of replication may indicate functionality of viral processes including translation, minimal levels of dsRNA resulting from replication may be crucial for avoiding RNAi silencing in plants (Gordon &
Waterhouse, 2006). 72
Our results support previous demonstrations of IGR IRESs from other dicistroviruses driving translation in wheat germ extract (Domier et al., 2000; Shibuya et al., 2003; Wilson et al., 2000a). These previous works examined IGR IRES function by using SDS PAGE to compare IRES-driven fluc translation with cap-dependent translation of green fluorescent protein (GFP) or chloramphenicol acetyl transferase (CAT), precluding a quantitative comparison to our dual luciferase assay results. However, in in vivo translation in
Spodoptera frugiperda (Sf9) insect cells wildtype RhPV IGR IRES was four times more active in driving translation than a PK I-disruption mutant, which is comparable to the three- to 3.7-fold activity increase we saw between wildtype and PK I-disruption mutant IGR IRES- driven translation. Previous insights (Pfingsten et al., 2010) have suggested that the IGR
IRES functions by manipulating the ribosome through universally conserved key steps in translation. Thus, the conservation of IGR IRES functionality in both animal and plant systems supports this notion.
RhPV is naturally transmitted via plant reservoirs, but the virus does not replicate or persist indefinitely in plant tissue (Gildow & D’Arcy, 1988). However, the stability of RhPV in plants and ability of plants to serve as reservoirs for the virus from which aphid insect pests acquire infectious virus, suggest the virus may be useful if transgenically expressed in crop plants to act as an insect specific biopesticide. Previous attempts to express the insect virus HaSV, a tetravirus, in transgenic plants encountered problems due to RNAi silencing of the virus (Gordon & Waterhouse, 2006). The low levels of replication by the RhPV infectious clone transcript may prove beneficial in avoiding similar obstacles. 73
Methods
Cloning strategy for translation assay constructs
To make construct RIF (Figure 1D), a dual luciferase construct in which translation of the firefly luciferase (fluc) is driven by the RhPV IGR IRES, nucleotides 6935-7109 of the
RhPV sequence (the RhPV IGR IRES) were amplified by PCR from the RhPV62A construct with forward primer IGR_IRES_XmaIf (5’ AATT C^CCGGG AGTGTTGTGTGATCTTGC
3’), with XmaI restriction site underlined, and reverse primer IGR_IRES_SacIr (5’ AATT
GAGCTC AGGTCATATATAAGGATA 3’) with SacI restriction site underlined (Table 2).
To produce construct RImF (Figure 1E), a negative control construct in which formation of pseudoknot I (PKI) of Domain 3 is disrupted, reverse primer UGC_mut_IGR_IRES_SacIr (
5’ AAT TGAGCTC GCA TCATATATAAGGATA 3’) was used in PCR amplification, resulting in a mutation of the CCU initiation triplet to an unrelated UGC triplet (UGC reverse-complement italicized). The amplified product was double digested with XmaI and
SacI, and ligated into an XmaI and SacI double-digested RF construct (p2luci in Grentzmann
(1998), but renamed RF here for consistency) (Figure 1C). This inserted the IGR IRES (or
UGC mutant-IGR IRES) sequence with the initiation triplet in frame with and two codons upstream of the initiation methionine codon of fluc. Correct insertion was screened by restriction digest profiling and confirmed by sequencing.
Additional constructs, RsIF, RsImF, and RsF (Figure 1F-H), where made by manipulating the polylinker region between the Renilla luciferase (rluc) and the IGR IRES to introduce two stop codons, a TAG C and a TAG G, into the rluc reading frame. This was done by SalI-HF digesting RF, RIF, and RImF constructs, then filling in the 4 base 5’ overhangs with DNA Polymerase I, Large (Klenow) fragment, and ligating the blunt ends 74
Table 2. Primers used for cloning and confirmation of clones.
Purpose Oligonucleotide Name Nucleotide sequence (5'-->3') AAT TCC CGG GAG TGT TGT GTG IGR_IRES_XmaIf ATC TTG C Cloning RhPV IGR IRES AAT TGA GCT CAG GTC ATA TAT into dual luciferase IGR_IRES_SacIr AAG GAT A constructs AAT TGA GCT CGC ATC ATA TAT UGC_mut_IGR_IRES_SacIr AAG GAT A Confirm correct reading GCA AGA AGA TGC ACC TGA TGA p2IGf frame and sequence of AAT GG added IGR IRES, check GGG CGT ATC TCT TCA TAG CCT p2IGr polylinker (stop) TAT GC Addition of polyA to dual PmlI_polyA_NotI GTG (A×60)GC luciferase constructs NotI_polyT_PmlI GGC CGC (T×60) CAC Addition of polyA to RhPV KpnI_polyA_SalI C(A×60)G 6.1 SalI_polyT_KpnI TCG AC(T×60) GGT AC Confirm addition of polyA p2polyAf GTT GTT GTT TTG GAG CAC G to dual luciferase constructs p2polyAr CAC TGC ATT CTA GTT GTG G Confirm addition of polyA PV-6214f TGG TTT TAA CCG TGT TAT TG to RhPV 6.1 polyA R-1 reverse M13 stock primer CAG GAA ACA GCT ATG ACC 75 together. This process added 4 bases to the length of the polylinker, thereby moving two stop codons into the rluc reading frame. The process also eliminated the SalI restriction site and introduced a PvuI restriction site. Following ligation, SalI-HF digestion was carried out again to eliminate any constructs that had re-ligated without having undergone Klenow fill- in, prior to transforming the ligation product into chemically-competent Top 10 E.coli
(Invitrogen, Carlsbad, CA) by heat shock method. Constructs were confirmed by restriction digest profiling and sequencing (Table 2).
A final round of constructs was made in which a 60 bp poly A tail sequence was added for in vivo translation in protoplasts, producing RsIFa, RsImFa, RsFa, and RFa (Figure
1I-L). 5’-phosphorylated oligonucleotides PmlI_polyA_NotI and NotI_polyT_PmlI
(Invitrogen, Carlsbad, CA) (Table 2) were resuspended to 500 μM in duplex buffer (100 mM potassium acetate, 30 mM HEPES, pH 7.5) and duplexed together by heating at 94°C for 2 minutes, then allowing to cool to room temperature. This duplex was inserted between the
PmlI and NotI sites of double-digested and calf intestinal phosphatase-treated (NEB, Ipswich,
MA) RsF construct. The RsIFa and RsImFa constructs were made using the RsF construct as vector and the inserts used were the same PCR products used for making the RIF and RImF constructs. To make RFa, the gel-purified (Qiagen, QiaSpin Gel extraction kit) KpnI, EcoRI
2,207 bp restriction fragment (nucleotides 62–2269) of RF was used as insert and the gel- purified 3,720 bp KpnI, EcoRI restriction fragment (nucleotides 2448–62) of RsIFa was used as vector. Restriction digest profiling and sequencing were used to check the polyA region and polylinker regions (including the presence or absence of IGR IRES, mutant IGR IRES, and rluc frame stop codons) (Table 2). 76
Cloning strategy for replication assay constructs
A 60 bp polyA tail sequence was added to the RhPV6-1 construct (Boyapalle et al.,
2008). 5’-phosphorylated KpnI_polyA_SalI (5’ CA(60)G 3’) and SalI_polyT_KpnI (5’
TCGACT60GGTAC) oligonucleotides (Table 2) were duplexed as above and inserted between the KpnI and SalI sites of double digested and gel purified RhPV6-1 construct (bp 5-
22). Construct was verified by restriction digest profiling and sequencing (Table 2).
In vitro transcription
Constructs RF, RIF, and RImF (Figure 1C-E) were linearized by digestion with PmlI, cutting 10 bp downstream of fluc, as done in Grentzmann et al. (1998). For DNA that could be used to produce transcripts with longer 3’UTRs, HpaI was used to cut RF, RIF, RImF,
RsF, RsIF, and RsImF constructs (Figure 1C-H) 161 bp downstream of the fluc stop codon.
Constructs RFa, RsFa, RsIFa, and RsImFa (Figure 1I-L) were linearized with NotI. RhPV6-1 was linearized with SmaI and RhPV6-1_polyA was linearized with SalI for positive-strand transcription. RhPV6-1 and RhPV6-1_polyA were linearized with EcoRI for negative-strand transcription. For non-polyA constructs, uncapped positive-strand transcripts were transcribed with T7 Polymerase using a T7 MegaScript kit (Ambion, Austin, TX). For polyA containing constructs, 7-methyl guanosine capped positive-strand transcripts were transcribed with mMessage mMachine T7 MegaScript kits (Ambion, Austin, TX). Negative- strand transcripts were transcribed using an SP6 MegaScript kit (Ambion, Austin, TX).
Transcripts were DNAse treated with Turbo DNAse for 15 minutes at 37°C, then purified by lithium chloride precipitation or phenol/chloroform extraction and stored in aliquots at -
80°C. Concentrations were determined using a Nanodrop spectrophotometer (Thermo
Scientific, Rockford, IL). 77
In vitro translation in wheat germ extract
Non-isotopically labeled in vitro translations were carried out using 190 ng (0.2 pmole) of RNA in 10 μl translation reactions containing: 50% wheat germ extract (Wheat
Germ Extract kit, Promega, Madison, WI), 40 μM methionine, 40 μM leucine, 80 μM of all other amino acids, 8 units RNasin, and 40 mM potassium acetate. 35S-Methionine-labeled translations were done as above but with 5 pmol 1000 Ci/mmol 35S-methionine (0.5 μM final concentration) and 80 μM of all other amino acids. Translations were carried out for one hour at 25 °C.
SDS polyacrylamide gel electrophoresis of 35S-methionine-labeled wheat germ extract translation product
Five μl of each translation reaction product was added to 20 μl of LDS running buffer (Invitrogen, Carlsbad, CA) + β-mercaptoethanol and denatured at 70°C for 15 minutes. Samples were loaded onto a 4-12% Tris-Glycine gel (Invitrogen, Carlsbad, CA) buffered with 1X SDS MOPS buffer (Invitrogen, Carlsbad, CA) in an XCell SureLock Mini-
Cell (Invitrogen, Carlsbad, CA) and ran for 30 minutes at 250 V. Gels were fixed in 50% methanol, 10% glacial acetic acid for 30 minutes, then soaked in 7% glacial acetic acid, 7% methanol, 1% glycerol for 5 minutes prior to drying gel at 80°C under vacuum. The gel was then used to expose an Imaging Screen-K (Kodak) phosphorimage screen for 8 hours prior to imaging. Quantity One (Bio-Rad) was used to analyze images to determine peak, average, trace, Gaussian peak, Gaussian trace, and relative quantity of bands.
In vitro translation in rabbit reticulocyte lysate
Non-isotopically labeled in vitro translations were carried out using 190 ng (0.2 pmole) of RNA in 10 μl translation reactions containing: 70% Rabbit Reticulocyte Lysate 78
(Promega, Madison, WI), 10 μM methionine, 10 μM leucine, 20 μM of all other amino acids, and 8 units RNasin. Translations were carried out for one hour at 30 °C.
Dual luciferase assays
Dual luciferase assays were performed using a Dual Luciferase Assay kit (Promega,
Madison, WI) and a 20/20 GloMax luminometer (Promega, Madison, WI). One μl of translation product, 40 μl of LARII reagent, and 40 μl of Stop and Glo were used for each sample.
Electroporation of oat protoplasts
Electroporation of oat protoplasts with dual luciferase of infectious clone transcripts was done by a method adapted from Rakotondrafara et al. (2007). Briefly, oat suspension culture in liquid MS Medium was started from oat callus culture maintained on MS Medium plates. Oat suspension culture was maintained at 25°C in a shaker incubator set at 220 rpm.
Oat suspension culture was subcultured every 7 days, adding 10 mL culture to 40 mL fresh
MS Medium under asceptic conditions.
Oat protoplasts were made from 3- to 7-day-old suspension culture. Fifty mL cultures were split into two 50-mL conical tubes. Cells were left to settle, and then the supernatant was drawn off with a serological pipette. Cells were resuspended in 25 mL of fresh oat protoplast enzyme solution (0.1% w/v Driselase (Sigma), 0.175% w/v Cellulase
(Onozuka RS, Yakult Pharmaceuticals), 0.8% w/v hemicellulase (Sigma) in 1:1 ASW/0.6 M mannitol, pH 5.6. ASW solution is 311 mM sodium chloride, 18.8 mM magnesium sulfate,
6.8 mM calcium chloride, 10 mM 2-(N-morpholino)ethanesulfonic acid, 6.9 mM potassium chloride, 16.7 mM magnesium chloride, 1.75 mM sodium bicarbonate. Tubes were sealed with parafilm and covered with aluminum foil, then placed on a gyratory shaker for 14 hr at 79
50 rpm at 30°C. Digested cells were checked with a light microscope to ensure protoplasts appeared healthy without cell clumps or over-digested protoplasts. Protoplasts were centrifuged at 100 x g, 4°C, supernatant was removed, and protoplasts were gently resuspended in 10 mL ASW/0.6 M mannitol. This wash step was repeated 2 more times and protoplasts were resuspended in 10 mL oat protoplast electroporation buffer (0.15 mM potassium dihydrogen phosphate, 0.43 mM sodium phosphate dibasic, 130 mM sodium chloride, 0.2 M mannitol, 3.2 mM calcium chloride, 0.2 mM spermidine, pH 7.2).
Protoplasts were again centrifuged at 100 x g, 4°C, supernatant was removed, and protoplasts were resuspended in electroporation buffer at a concentration of ~6 106 protoplasts/mL, as checked with a hemacytometer. Protoplasts were checked under a light microscope at each resuspension to ensure they remained intact.
Aliquots of 0.4 mL of protoplasts in oat protoplast electroporation buffer were transferred to 4 mm electroporation cuvettes (BTX, Holliston, MA) and kept on ice 5-30 minutes. Ten μg of transcript RNA were added to each sample, which was then gently mixed by inversion and electroporated in an Electro Square Porator T820 (BTX, Holliston, MA) with one pulse for 6 msec at 300V. For replication assays, 1hr, 24 hr, and 48 hr samples were collected. For translation assays, one hr samples were incubated on ice before they were used for TRIzol (Invitrogen, Carlsbad, CA) RNA extraction or protein extraction.
Twenty-four hr and 48 hr samples were added to 5 mL of MS Medium with 0.4 M mannitol in 6 well plates, sealed with parafilm and incubated at room temperature in the dark for the appropriate time before Trizol RNA extractions or lysis. Lysis was done prior to dual luciferase assays by resuspending protoplasts in 150 μL of passive lysis buffer and lysing protoplasts via 3 freeze/thaw cycles. 80
Northern blot hybridization
Trizol-extracted total RNA from 1 hour, 24 hour, and 48 hour incubations of oat protoplasts electroporated with capped RhPV6-1_polyA positive-strand T7 transcripts, as well as 48 hour incubations of oat protoplasts electroporated with capped RhPV6-1_polyA negative-strand SP6 transcripts, PAV6 positive-strand T7 transcripts, or negative controls electroporated with water, was used for Northern blotting. Capped RhPV6-1_polyA positive-strand T7 transcripts, PAV6 positive-strand T7 transcripts, and RhPV6-1_polyA negative strand SP6 transcripts, were also run as controls.
RNA was denatured in glyoxal buffer (Ambion, Austin, TX) at 50°C for 1 hr and size fractionated in a 10 mM sodium phosphate-buffered 1.8% SeaKem GTG agarose gel
(FMC, Rockland, ME) with 0.01M iodoacetic acid (Sigma, St. Louis, MO) for 4.5 hours at
4°C. RNA was transferred to a Hybond XL membrane (GE Healthcare /Amersham
Biosciences, Piscataway, NJ) with 20x SSC (3 M NaCl, 0.3 M sodium citrate, pH 7.0) transfer buffer by downward capillary transfer (Palani & Lin, 2007) for 16 hours. RNA was then crosslinked to the membrane with 220 MJ of UV light emitted by 312 nm bulbs in a
Stratalinker 2400 (Stratagene, La Jolla, CA). RNA was de-glyoxalated by incubating the membrane in 20 mM Tris-Cl pH 8 at 65°C for 30 minutes. The membrane was then prehybridized in prehybridization buffer (Palani & Lin, 2007) for 1 hour at 65°C. The membrane was then hybridized for 16 hours in fresh prehybridization buffer plus RhPV6-1
T7 transcribed positive-sense α-32P CTP-labeled probe (Palani & Lin, 2007) at 65°C.
Following hybridization, a low stringency wash was carried out for 1.5 hours in 2X SSC,
0.2% SDS, followed by two 1 hour high stringency washes in 0.2X SSC, 0.2% SDS. The membrane was then wrapped in plastic wrap and used to expose an Imaging Screen-K 81
(Kodak)( Bio-Rad, Hercules, CA) phosphorimage screen in an amplifying cassette. After scanning the phosphorimage screen with a PharosFX Plus Molecular Imager (Bio-Rad,
Hercules, CA), the membrane was stripped in boiling 0.1% SDS until probe was removed.
Prehybridization, hybridization, and imaging was repeated using RhPV6-1 negative strand
SP6 transcript.
Strand-specific RT-PCR
Strand-specific RT-PCR using a tagged primer system was used to test for replication in oat protoplasts electroporated with positive-sense RhPV6-1 infectious clone transcript.
Total RNA was extracted 1hr, 24 hr and 48hr post-electroporation, and 48 hr post electroporation from RhPV62A_DA RdRp-deficient mutant negative control noninfectious clone. Reverse-transcription reactions with 290 ng TRIzol-extracted oat protoplast RNA, 1
μM final concentration of first-strand tagged primer (ZF5-tag1 for negative-strand detection or ZR6-tag1 for positive-strand detection) (Table 1), and 50 units of SuperScript III reverse transcriptase (Invitrogen, Carlsbad, CA) for each 10 μl reaction, synthesized cDNA with the non-genomic tag sequence incorporated at the 5’end. Reverse-transcriptions were carried out at 55°C for 50 minutes and heat inactivated at 70°C for 15 minutes. This was followed by exonuclease I (NEB, Ipswich, MA) treatment for the removal of unincorporated primers, then RNAse H (NEB, Ipswich, MA) treatment for 20 minutes at 37°C to digest RNA hybridized to DNA. This was followed by PCR with tag1 and ZR6 primers for negative- strand detection or tag1 and ZF5 primers for positive-strand detection (Table 1). Each 30 μl
PCR reaction used 3 μl reverse-transcription product, 1 unit Ex Taq DNA Polymerase
(Takara, Otsu, Shiga, Japan), and 0.5 μM final concentration of each primer. The PCR 82 program was: 95°C for 3 minutes, followed by 35 cycles of 94°C for 45 seconds, 58 °C for
45 seconds, 72°C for 1 minute, followed by 5 minutes at 72°C and a 4°C hold.
To demonstrate strand-specificity, strand specific RT-PCR was also carried out for positive and negative sense RhPV6-1 transcripts, using only 1.5 pg (=0.46 attomole,
300,000 molecules RNA) transcript for the reverse transcription. Reactions without first- strand primer in the reverse transcription were done to test for false-priming to ensure strand specificity. Reactions without reverse transcriptase were set up to test for DNA contamination. A PCR reaction with RhPV6-1 plasmid DNA was done as a positive control for PCR.
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CHAPTER 3. GENERAL CONCLUSIONS
Dicistroviruses are small, positive-strand RNA viruses that infect arthropods. They are distinguished from other members of the order Picornavirales in possessing two ORFs each preceded by a distinct IRES. The dicistrovirus IGR IRESs are currently the only IRESs known to drive translation independently of eIFs. Long term goals of our lab include the development of transgenic plants employing modified RhPV as an insect-specific biopesticide. Implementation of this project requires detailed knowledge of RhPV translation and replication in a plant cellular environment. To these ends, the work described in this thesis tested RhPV IGR IRES-mediated translation in plant in vitro and in vivo systems. The
IGR IRES was found to be capable of driving translation in WGE. IGR IRES-mediated translation was not detected in oat protoplasts. Replication of RhPV infectious transcripts in oat protoplasts was also a focus of this thesis work. Small amounts of replication were detected by RT-PCR 48 hours post-electroporation with infectious transcript, but this was below the detection level of northern blot hybridization. Detection of IGR IRES-mediated translation in WGE is an encouraging finding for future utilization of this IRES in plants.
The lack of a detectable level of translation in oat protoplasts transfected with RhPV IGR
IRES dual luciferase transcripts does not necessarily mean that the IGR IRES is not functional in a plant in vivo environment. It may mean that in the in vivo environment, the
IRES was not able to compete with the cellular mRNAs for the ribosome strongly enough to drive a detectable amount of translation in the 4 hour incubation. In fact, preliminary work done in our lab by Narinder Pal showed that when oat protoplasts were electroporated with large amounts (10 μg) of wildtype RhPV viral RNA, structural proteins could be detected by immunoblot 48 hours post-inoculation. Differences between these experiments include the 87 amount of incubation time and using a dual luciferase transcript vs. wildtype virus RNA. A longer incubation time may be beneficial for detection. For comparison, studies investigating
RhPV virion production in aphids or insect cell culture typically looked for virions several days after inoculation, with virions reaching their highest levels after 7 days. (Boyapalle et al., 2008; Boyapalle et al., 2007; D'Arcy et al., 1981). For comparison with other insect small RNA viruses translated in plants, Dasgupta et al. (2001) monitored plants for FHV production 12-14 days post-inoculation and Gordon et al. (Gordon et al., 2001) monitored protoplasts for HaSV production 3-5 days post-inoculation. Poor health and longevity of protoplasts at the beginning of our investigation led us to choose a shorter incubation time for translations. However, recent optimization of protoplast digestion and handling protocols, resulting in better health and longevity of protoplasts, will allow for longer incubations, which may be advisable. Also, due to unexpected translation activity in a dual luciferase construct expected to be a negative control for firefly luciferase translation, it may be advisable to use different dual luciferase constructs. Dual luciferase constructs containing an
RhPV IGR IRES, produced by Domier & McCoppin (2003) or Carter et al. (2008), should be adaptable for such use.
Strand-specific RT-PCR detected a small amount of replication in oat protoplasts 48 hours post-inoculation with capped infectious RhPV transcript. Northern blot hybridization of protoplasts inoculated with capped and polyadenylated infectious RhPV transcript did not detect negative-strand replication product. This is not surprising, as northern blot hybridization is less sensitive and an abundance of positive-strand RNA may interfere with the detection of negative-strand RNA. 88
Regarding prospects for use as a transgenically-expressed biopesticide in crops, the literature on HaSV expression in plants would suggest that an inability to replicate in plants is preferred (Gordon & Waterhouse, 2006). This is because dsRNA generated by replication triggers RNAi silencing of the virus genome. This is what Waterhouse and Gordon (2006) believe happened with HaSV plant expression. While it is generally believed that replication is an absolute requirement for virion assembly and packing, Annamalai & Rao (2005) showed with Brome mosaic virus (BMV) replication mutants that, although replication helped packing fidelity, both viral genomic RNA and cellular mRNA were packaged into virions in the absence of replication. If translation and assembly occur in plants — as suggested by Narinder Pal’s preliminary data — then it may be possible to get enough properly packaged virions in transgenic plants expressing modified RhPV to infect aphids, even though improperly packed virions may also be present. Once aphids acquire the virions, the virus would be in an environment where it can replicate. Aphids would then be able to produce modified RhPV and transmit it to other aphids through plant passive reservoirs or transovarially, essentially introducing a new ―natural enemy‖. This may be especially useful against invasive species of aphids that have escaped their natural enemies.
Little to no replication may be preferable to strong replication in plants because low packaging fidelity may be preferable to RNAi-mediated silencing. However, even a small amount of dsRNA from replication may lead complete silencing due to amplification of silencing (Ding & Voinnet, 2007).
Expanding on our labs’ work with RhPV and ALPV, members of the dicistrovirus genus Cripavirus, we are currently in the process of beginning research on members of the proposed genus Aparavirus. These viruses are of particular interest due to their economic 89 importance. Aparaviruses of bees, most notably IAPV, have been implicated in colony collapse disorder, a phenomenon that decimates commercial honeybee colonies crucial to agriculture (Cox-Foster et al., 2007; van Engelsdorp et al., 2009; van Engelsdorp et al.,
2008). TSV has a large economic impact on Asian shrimp industries (Lightner et al., 1997;
Overstreet et al., 1997), and SINV-1 may be an important tool for control of invasive red imported fire ants. Work is underway to develop infectious clones to facilitate research on these viruses. Aparaviruses are also of particular interest because they possess Class II IGR
IRESs. While the stem loop unique to this class, SL III, has been shown to be necessary for
Class II IGR IRES function, the mechanism by which it operates is unknown (Cevallos &
Sarnow, 2005; Hatakeyama et al., 2004). Recently, Jang and Jan (2010) constructed chimeric IGR IRESs with the ribosomal binding domain of one class and the P-site domain of the other. These observations show that the separate domains are modular in function.
While answering some questions, other questions are raised about these varying IGR IRES classes. If SL III is not required in a chimeric IGR IRES, why is it necessary in wildtype
Class II IGR IRESs? Several highly-conserved nucleotides in the Class I IGR IRESs are required for IGR IRES activity, but are not conserved in Class II IGR IRESs (Pfingsten et al.
2010). Are there similarly important nucleotides in corresponding locations of the Class II
IGR IRES structure, and what would this mean for the function of such nucleotides?
Another area of interest is examining the dynamic nature of the IGR IRESs, and investigating the initiation mechanism based on a carefully controlled series of coupled structural changes to the IRES and ribosome. Comparing this mechanism to other IRES mechanisms and canonical translation could result in interesting insights into the crucial steps of translation initiation. Hernandez (2008) postulates that IRES-driven translation is an 90 evolutionary predecessor of canonical cap-dependent translation. When translation is viewed through this lens, what does an understanding of IRES translational mechanisms tell us about the development of cap-dependent translation? What aspects of IRES-driven translation would have been consequential to a developing mechanism of cap-dependent translation if canonical translation was the hijacker of IRES-driven translation, rather than the other way around? Finally, new information about IRES structure and function could be useful for the identification of additional viral and cellular IRESs. RNAs translated via an IRES have the potential to be expressed in conditions less suitable for canonical translation, and in ways outside the control of conventional regulatory methods. Therefore, such RNAs could be important in times of cell stress or infection, and knowledge of them could shed light on unknown areas of cell response and regulation. Dicistroviruses provide a unique platform for the investigation of translation, as well as potential tools for the application of an insect virus as a transgenically-expressed biopesticide.
91
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ACKNOWLEDGEMENTS
I would like to thank my family, friends, and coworkers who made my Masters studies possible. My parents always encouraged my interests in the sciences. I would like to thank Nikki Krueger for helping me learn various protocols and always showing an interest in helping my research to progress. Thanks to Jimena Carrillo-Tripp. With your cooperative nature, attention to details, and efficiency, you have become something of a science role model to me. Thanks to Vijaya Palani, Karin Werner, and Julie Meyer for their assistance with northern blot hybridization techniques. Special thanks to Julie Meyer for her patience and hard work helping me to prepare this thesis. Thank you to Jelena Kraft and Zhaohui
Wang for allowing me to benefit from your research experience. I would also like to thank
Alice Hui and Mariko Peterson-Gordon for their help in troubleshooting and contributing to the positive energy of the lab. I would like to thank Dr. Steven Whitham for serving on my committee. Finally, I would like to thank my major professors, Dr. Bryony C. Bonning and
Dr. W. Allen Miller for their guidance and the opportunity they have given me to pursue research in their labs.