ABSTRACT

NI, SIHUI. RNA Translocation into Chloroplasts. (Under the direction of Dr. Heike Inge Sederoff).

Plastids and mitochondria are known as semi-autonomous organelles because they can encode and synthesize proteins that are essential for metabolism. They are both derived from

endosymbiotic events. The bacterial ancestor of plastids was cyanobacteria, while the bacterial ancestor of mitochondria was a proteobacteria. During their evolution, most of the ancestral bacterial DNA was translocated into the nuclear genome DNA. The current plastid genome encodes proteins for the photosynthetic apparatus, the transcription/translation system, and various biosynthetic processes. The current mitochondrial genome mainly encodes proteins for electron transport, ATP synthesis, translation, protein import and metabolism. Plastids and mitochondria are not self-sufficient; they import the majority of their proteins, which are synthesized in the cytosol. For plastids, a transit peptide at the N-terminus of protein precursors can be recognized by TOC/TIC complex and mediate their import. Similarly, for mitochondria, a piece of β-signal at the N-terminal of protein precursors can be recognized by TOM/TIM complex and mediate the import of the protein precursors. It worth noting that plastids also encode a suite of transfer RNAs while mitochondria need to import certain types of tRNAs.

In most angiosperms, plastids and mitochondria are inherited maternally and can only be passed down from the maternal plant organs through seed to the next generation. Maternal inheritance has benefits for organellar genetic engineering such as stable inheritance of genes

over generations, absence of out-crossing, absence of contamination by pollen and minimal

pleiotropic effects. The tradition methodology of plastid genomic engineering is biolistic transformation, where DNA covered gold particles parent cell walls of tissue or protoplasts at

high velocity, leading to integration of DNA into chromosome(s) through recombination. However, biolistic based integration of double-stranded DNA is relatively random.

Regeneration of transformed plants is complicated and often require a species-specific protocol

before a homoplasmic plants can be established. The biolistic transformation of mitochondria has not yet been accomplished.

For chloroplast genetic engineering, new methods are needed. The central idea of our

proposed method is to import two components into chloroplasts to engineer the organellar

genomic DNA and to generate homoplasmic plants. First, the reverse transcriptase is needed to generate double-stranded transgene DNA in the chloroplast in vivo. Second, the cas9

endonuclease associated with single guide RNAs is needed to degrade untransformed chromosomes to generate homoplasmic plants. Enzymes can be imported as proteins by being fused to a transit peptide for plastids and a β-signal for mitochondria. While, the mechanism of RNA translocation into organelles is not yet well understood. We tested two candidates, the derived Eggplant Latent Viroid (ELVd) and the eucaryotic initiation factor 4E (eIF4E), which have the potential to mediate the translocation of RNA into chloroplasts. To detect successful translocation, enhanced green fluorescent protein (eGFP) was used as a marker. The chimeric

complementary DNA (cDNA) of the derived ELVd and eGFP was transformed into Nicotiana

benthamiana using agroinfiltration and into Arabidopsis thaliana using floral dip. EGFP was detected in N. benthamiana by eGFP fluorescence and confocal microscopy but not by western blots. EGFP was not detected in A. thaliana either by confocal microscopy or western blots.

The chimeric cDNA containing the eIF4E and eGFP construct was transformed into N. benthamiana using agroinfiltration. EGFP was not detected either by confocal microscopy or western blots. More research, such as in situ hybridization, is needed to determine the cause of the reason for the absence of eGFP expression.

© Copyright 2018 by Sihui Ni

All Rights Reserved RNA Translocation into Chloroplasts

by Sihui Ni

A thesis submitted to the Graduate Faculty of North Carolina State University in partial fulfillment of the requirements for the degree of Master of Science

Plant Biology

Raleigh, North Carolina

2018

APPROVED BY:

______Heike Inge Sederoff Richard L. Blanton Chair of Advisory Committee

______Deyu Xi ii

BIOGRAPHY

Sihui Ni was born in China. The very first motivation for her working in plant biology is to develop a reliable and nutritious food source in Africa using transgenic technologies.

Fortunately, she was introduced to Dr. Heike Sederoff by Dr. Mari Chinn and therefore had a chance to explore the world of plant biology.

Sihui gained her bachelor’s degree in Functional Materials at Lanzhou University in

China, which helped her build a background in chemistry and physics. Driven by the strong motivation of working in plant biology, she started to work in plant biology labs since the second year in undergraduate school. During this period, she obtained various experimental skills in molecular biology.

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ACKNOWLEDGMENTS

Funding:

NCSU Chancellor’s Innovation Fund

Advisors, Committee, Mentors, Graduate Student Support:

Dr. Heike Sederoff, Professor, Department of Plant & Microbial Biology (PMB) at North Carolina State University (NCSU); Dr. Larry Blanton, Professor, Director of Graduate Programs, Department of PMB at NCSU; Dr. Deyu Xie, Professor, Department of PMB at NCSU; Sue Vitello, Executive Assistant, Department of PMB at NCSU; Dwayne Barnes, Graduate Services Coordinator, Department of PMB at NCSU; Catherine Freeman, Executive Assistant, Department of PMB at NCSU； Dr. Jay Cheng, Professor, Department of Biological and Agricultural Engineering (BAE) at NCSU.

Colleagues & Collaborators:

Dr. Imara Perera, Research Professor, Department of PMB at NCSU; Dr. Eva Johannes, Director of Cellular and Molecular Imaging Facility at NCSU; Dr. Mari Chinn, Professor, Department of BAE at NCSU; Colin Murphree, Graduate student, Department of PMB at NCSU; Jacob Dums, Doctor of Philosophy, Department of PMB at NCSU; Danielle Young, Graduate student, Department of Plant Biology at Michigan State University; Sathya Jali, post doctorate, Department of PMB at NCSU; Christophe La Hovary, post doctorate, Department of Crop and Soil Sciences at NCSU; Eli Hornstein, Melodi Charles, Nathan Wilson and Samuel Acheampong, Graduate students, Department of PMB at NCSU; Brianne Edwards, technician, Department of PMB at NCSU; Avery Ashley and Nikki Khoshnoodi, Undergraduate students, Department of PMB at NCSU

Other moral support:

Juanying Shen, Jianhua Ni, Guangming Wang, Xiting Liu, Lora J. Gary, Jing Wu, Wenbin Zhou, Yue Zhu, Huangchao Yu, Tao Jiang and Katherine Speight.

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TABLE OF CONTENTS

LIST OF TABLES ...... v LIST OF FIGURES ...... vi LIST OF ABBREVIATIONS ...... vii

CHAPTER 1: RNA Transport into Organelles ...... 1 1.1 Organellar Genomes in Plant Cells ...... 2 1.2 Plastids Genome Engineering ...... 14 1.3 RNA Translocation into Plastids ...... 20 1.4 Mitochondrial Genome Editing ...... 23 1.5 RNA Translocation into Mitochondria ...... 24 REFERENCES ...... 28

CHAPTER 2: RNA Transport into Chloroplasts Mediated by ELVd ...... 39 Introduction ...... 40 Methods and Materials ...... 45 Results ...... 50 Plasmid Constructions ...... 50 Tobacco Transformation ...... 52 EGFP Was Not Detected in Arabidopsis ...... 54 Discussion ...... 60 REFERENCES ...... 65

APPENDICES ...... 69 Appendix A: Recipes and Protocols ...... 70 Appendix B: DNA Segments Sequences and Primers ...... 73

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LIST OF TABLES

Table 1. Successful cases of plastid gene transformation using biolistics. …………………..15

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LIST OF FIGURES

CHAPTER 1 Figure 1.1 The predicted evolutionary stages of the mitochondrion, the nucleus, and the plastid...... 5 Figure 1.2 Maps of the chloroplast and mitochondrion of Arabidopsis thaliana...... 12 Figure 1.3 The design of our Cas9-ligation chloroplast genomic DNA engineering process...... 18 Figure 1.4 The structural models and replication pathways of the Pospiviroidae family and the Avsunviroidae family...... 21 CHAPTER 2

Figure 2.1 Proposed chloroplast genomic DNA engineering through reverse transcription and homologous recombination...... 44 Figure 2.2 Vector map...... 52 Figure 2.3 Confocal microscopy of N. benthamiana transgenic plant leaf tissue...... 53 Figure 2.4 Western blot for full-length eGFP of N. benthamiana...... 54

Figure 2.5 Confocal microscopy of A. thaliana T1 transgenic plant leaf tissues...... 55 Figure 2.6 Lambda scan of the same leaf tissue of ELVd::eGFP transgenic line from figure 2.5...... 56 Figure 2.7 PCR amplification gel electrophoresis...... 58 Figure 2.8 RT-PCR for two ELVd::eGFP transgenic lines...... 59 Figure 2. 9 Western blot for full-length eGFP...... 60 Figure 2.10 The secondary structure of ELVd (AJ536613) predicted by Mfold...... 61

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LIST OF ABBREVIATIONS

5’ UTR 5’ untranslated region aaRS Aminoacyl-tRNA synthetase A. thaliana Arabidopsis thaliana ADP Adenosine diphosphate ATP Adenosine triphosphate bbp Β-barrel-membrane protein biolistic Biological and ballistic CaMV Cauliflower mosaic virus cDNA Complementary DNA CFP Cyan fluorescence protein cm Centimeter(s) CRISPR Clustered regularly interspaced short palindromic repeats CTAB Cetyl trimethylammonium bromide diH2O Deionized water DNA Deoxyribonucleic acid EDTA Ethylenediaminetetraacetic acid eGFP Enhanced green fluorescent protein eIF4E Eucaryotic translation initiation factor 4E ELVd Eggplant latent viroid g Gram(s) GFP Green fluorescent protein GUS Β-glucuronidase Gyr Billion years hr Hour(s) indel Insertion/deletion mutation LB Lysogeny broth LSC Large single copy region mGFP Modified green fluorescent protein mitomiR Mitochondrial microRNA mL Milliliter(s) mRNA Message RNA

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MRP Mitochondrial RNA processing MTS Mitochondrial targeting sequence mV Millivolt(s) N. benthamiana Nicotiana benthamiana ncRNA Noncoding RNA NUMT Nuclear mitochondrial DNA sequences NUPT Nuclear plastid DNA-like sequence min Minute(s) PCR Polymerase chain reaction PEG Polyether polyethylene glycol PRAT Preprotein and amino acid transporter RNA Ribonucleic acid rRNA Ribosomal RNA RT Reverse transcriptase RT-PCR Reverse transcription polymerase chain reaction sec Second(s) SAM sterile-a-motif Sarkosyl N-Laurosylsacosine sgRNA Single guidance RNA SSC Small single copy region TALEN Transcription activator-like effector nuclease TIM Translocase of the inner membrane TOM Translocase of the outer mitochondrial membrane TRiC Tailless complex polypeptide 1 ring complex tRNA Transport RNA µFD Microfarad µL Microliter(s) µs Microsecond(s) VDAC Voltage-dependent anion channel ZFN Zinc finger nuclease

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CHAPTER 1

RNA Transport into Organelles

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1.1 Organellar Genomes in Plant Cells

Eukaryotic plant cells contain various subcellular organelles, including the nucleus,

plastids, mitochondria, and vacuoles. Plastids and mitochondria are unique in that they

maintain their own genomic DNAs separate from the nuclear DNA. The presence of a genome and other features suggests that they might have once been free-living organisms. The origin and development of these genome-carrying organelles is important for understanding mechanisms of RNA translocation into organelles.

Although plastids and mitochondria share some features with the nucleus such as DNA and a double membrane, the origins of plastids and mitochondria differ from the origin of the nucleus. It is widely accepted that the nucleus came from an autogenous origin, which is

described as the nucleus arising originally in the ancestral protoeukaryote as a structure

modification within a single evolving lineage (Baum, 2015). Some biologists still hold the

view that the nucleus formed from an endosymbiotic origin. The endosymbiotic origin is

described as the nucleus formed when a nuclear endosymbiont, which might be archaeon,

spirochaete or membrane-bound virus, engulfed by a cytoplasmic host, an archaeon,

proteobacterium or Planctomycetes-Verrucomicrobia-Chlamydiae bacterium (Forterre, 2013;

Horiike et al., 2004; Margulis et al., 2000; Takemura, 2001; Gupta & Golding, 1996; Lake &

Rivera, 1994).

Unlike the nucleus, the origins of plastids and mitochondria have less debate and both

are generally accepted as having an endosymbiotic origin. Plastids formed through an

endosymbiotic event when a cyanobacterium engulfed by a non-plastid bearing cell formed a

stable intercellular coexistence and resulted in the integration of the majority of the

cyanobacterial genome into the nuclear genome (Martin et al., 2015). This endosymbiotic

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origin theory of plastids has been specified multiple times since Richard Altmann first claimed the idea of coexistence of bioblasts in eukaryotic cells (Altmann, 1894; Martin et al., 2015).

However, Altmann did not suggest anything about the endosymbiotic theory. The first

thorough argument of the endosymbiotic theory was not clearly defended until Constantin

Mereschkowsky, a Russian botanist, published in his paper in 1905 that “Chromatophores,”

another name known as chloroplasts at that time, “are foreign organisms that invaded the

colourless plasm of the cell and entered into a symbiotic coexistence with it” (See the translated

version in English: Martin & Kowallik, 1999).

Similarly, the endosymbiotic origin theory of mitochondria is described as mitochondria

formed when a proteobacterium engulfed by an anaerobic cell forming a stable intercellular coexistence and resulted in the integration of the proteobacterial genome into the nuclear genome (Martin et al., 2015). The idea of the endosymbiotic origin of mitochondria was not

elaborated until Ivan Emanuel Wallin reviewed prior works and summarized that

“mitochondria are bacterial organisms symbiotically combined with cells of all higher

organisms” (Wallin, 1927).

Numerous forms of evidence have been put forth over the years to confirm the endosymbiotic origin of plastids and mitochondria. Mereschkowsky proposed his results based on four pieces of evidence using the most common technique available at that time, an optical microscope. First, chromatophores do not arise de novo. Instead, they preexist in egg cells and

are passed down to the offspring from there. Second, chromatophores are highly independent

of the nucleus and can live and/or even divide either with or without the nucleus. Third, chromatophores behave similarly to Zoochlorellae, a respective structure in animals that they divide from pre-existing individuals independently to the nucleus. Four, chromatophores are

4

highly similar to a free-living organism, Cyanophyceae (cyanobacteria), on all aspects (See the translated version in English: Martin & Kowallik, 1999).

In addition, modern techniques further support the endosymbiotic origin of both plastids and mitochondria. New evidence includes the sequence analysis and phylogenetic comparisons of the genomes of plastids and mitochondria. It indicates that the closest extant relative of the plastid is a cyanobacterium (Martin et al., 2002, 2003), while the closest extant relative of the mitochondrion is a proteobacterium (Andersson et al., 1998; Douglas & Raven, 2003). It worth noting that, according to phylogenetic analyses, the order of the diverging time of the three

DNA-carrying organelles should be first the mitochondrion at around 2 billion years (Gyr) ago

(Abhishek et al., 2011), then the nucleus at around 1.45 Gyr ago (Javaux et al., 2001), and finally the plastids at 1600-600 million years ago (McFadden & Van Dooren, 2004) (Figure

1.1). Plastids and cyanobacteria have the same protein components in metabolic pathways such as photosystem I and II (Ben-Shem et al., 2003). Mitochondria and proteobacteria have the

Krebs cycle in common (Walden, 2002). The outer membranes of plastids and mitochondria contain β-barrel-membrane proteins (bbps) that are coded by the nuclear genome and synthesized in the cytosol but also exist in the proteobacterial ancestor of mitochondria

(Paschen et al., 2003). The molecular origin of plant plastids and mitochondria continues to be an area of active research.

5

A B

Figure 1.1 The predicted evolutionary stages of the mitochondrion, the nucleus, and the plastid (Martin et al., 2015). (A) A proteobacterium (in blue) engulfed by an anaerobic cell (in red) forming a stable intercellular coexistence, and the the nucleus formed autogenously and the proteobacterial genome integrated into the nuclear genome. (B) A cyanobacterium (in green) engulfed by a non-plastid bearing cell formed a stable intercellular coexistence and resulted in the integration of the majority of the cyanobacterial genome into the nuclear genome The genomes in plastids and mitochondria are much smaller than those in their free- living ancestors indicating that genes were transferred from the endosymbionts to the nucleus of the host cells during evolution (Weeden, 1981). The identification of translocated organellar genes into the nuclear genomes is based on the sequence similarities between the genomes of ancestral organism and eukaryotic nuclei (Timmis et al., 2004). The term nuclear plastid DNA- like sequences (NUPTs) is used to describe the transpositions of all types of ancestral cyanobacterial DNA into the nuclear genome in eukaryotic organisms (Smith et al., 2011).

Taking Arabidopsis thaliana as an example, there are 17 insertions with a total of 11 kilobase pairs (kb) in the nuclear genome maintained from the cyanobacterial ancestor of plastids (The

6

Arabidopsis Genome Initiative, 2000). Correspondingly, the term nuclear mitochondrial DNA

sequences (NUMTs) is used to describe the transpositions of all types of ancestral proteobacterial DNA into the nuclear genome in eukaryotic organisms (Lopez et al., 1994).

There is one large insertion of around 620 kb plus 13 small insertions with a total of 7 kb in nuclear genomes maintained from the proteobacterial ancestor of mitochondria (Stupar et al.,

2001; The Arabidopsis Genome Initiative, 2000).

Nevertheless, organelles have conserved part of the genomes from their endosymbiotic ancestors. The current genomes in plastids and mitochondria are double-stranded, circular molecules with multiple copy numbers, with some exceptions in algae and some higher plants showing leaner architecture (Bock, 2007; Taanman, 1999). Interestingly, mitochondria in most plant species sequenced so far possess plasmids in addition to their genomic DNA. The gene number in plastids varies from 7 in Pilostyles hamiltonii to 193 in Pelargonium endlicherianum

(NCBI, 2004; Weng et al., 2017; Bellot & Renner, 2015). In contrast, in mitochondria, the gene number varies from none on a mitochondrial plasmid in Polytomella piriformis to 221 in the mitochondrial genome in Capsicum annuum (Smith et al., 2010; NCBI, 2004; Jo et al.,

2011).

The reverse phenomenon that nuclear genes transfer back to organellar genomes also exists but is very rare. Only one putative case was reported for a nuclear gene from the MutS homolog gene family in a Coral species, Sarcophyton glaucum, that might have been transposed in the mitochondrial genome (Pont-Kingdon et al., 1998). No nuclear to plastid gene transposition have been reported so far.

As the eukaryotic species evolved in different environments with different phenotypes, proteins encoded by plastid and mitochondrial genomes adapted to the organism’s needs. In

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modern plant species, plastids contain genes encoding proteins functioning in three different metabolic systems: the photosynthetic apparatus, the transcription/translation system and the

biosyntheses (Wakasugi et al., 2001). These genes are generally organized in one large single

copy region (LSC), one small single copy region (SSC), and two inverted repeat regions (IRA

and IRB) separating the single copy regions (Bock, 2007). Genes in IRs usually have higher

recombination frequency and lower mutation frequency than the genes in other positions on

the plastid genomes (Bock, 2007; Fan & Mower, 2016) (Figure 1.2A). Correspondingly, the

mitochondrial genes encode proteins functioning in electron transport, ATP synthesis,

translation, protein import and metabolisms (Fan & Mower, 2016). Mitochondrial genomes do

not form similar physical structures as plastid genomes do, and they have much lower

recombination frequency than the plastid genomes (Barr et al., 2005) (Figure 1.2B).

Although plastids and mitochondria each have the ability to synthesize some of the

proteins contained in these organelles, they are not self-sufficient, because most of the plastids

and mitochondrial genomes have transferred genes into the nucleus from their endosymbiotic

ancestors. The majority of organellar proteins are encoded in the nuclear genome, translated in

the cytosol and imported into organelles under the guidance of an N-terminal targeting signal.

Thses targeting sequences consist of 20 to 60 amino acids (Herrmann & Neupert, 2000; Li &

Chiu, 2010), and are called transit peptides in plastid translocations and β-signals in mitochondrial translocations. These peptide domains enable protein transport across the membrane mediated by TOC/TIC complexes (translocon on the outer/inner chloroplast membrane) in plastids and TOM/TIM complexes (translocase of the outer/inner membrane) in mitochondria, and are cleaved off the protein after transport and refolding (Herrmann &

Neupert, 2000; Li & Chiu, 2010; Schleiff & Becker, 2011). While the plastid genomes encode

8 an entire suite of transfer RNAs (tRNAs) and ribosomal RNAs (rRNAs) required for translation (Sugiura et al., 1998), the mitochondrial genome only encodes a subset of the tRNAs and therefore has to import selected tRNAs (Taanman, 1999). The number of tRNA species that are lacking in the mitochondrial genome and need to be imported, varies between plant species from three in Marchantia polymorpha to at least 16 in A. thaliana (Akashi et al.,

1998; Duchene & Marechal-Drouard, 2001).

So why do organelles need to retain genomes at all? One of the hypotheses is the

“hydrophobicity–importability” therory that states that the hydrophobicity of the organellar proteins may interfere with the efficiency of their targeting to and importing into organelles, and results in misstargeting to the endoplasmic reticulum, if they are synthesized in the cytosol

(Daley & Whelan, 2005; Leister & Kleine, 2011). Therefore, the genes of these proteins tend to be retained in the organelles. However, the hydrophobicity hypothesis has two major flaws.

First, not all the organellar proteins are hydrophobic, such as the large subunit of ribulose 1,5- bisphosphate carboxylase/oxygenase (Rubisco) in chloroplasts, which is hydrophilic but encoded by the chloroplast genome (Leister & Kleine, 2011). Second, some hydrophobic proteins are nuclear encoded but can be imported into organelles. For example, the light- harvesting complex (LHC) II and I, are encoded by the nuclear goneme and imported into chloroplasts (Allen, 2003).

Another hypothesis developed more recently, is know as the co-location for redox regulation (CoRR) hypotheis, which states that “mitochondria and chloroplasts contain genes whose expression is required to be under the direct regulatory control of the redox state of their gene products, or of electron carriers with which their gene products interact” (Allen, 2003).

To be more specific, the primary function of chloroplasts and mitochondria is energy

9 transduction that proced through the vectorial electron transfer to ATP synthesis, which is a redox chemistry interaction. Unlike the hydrophobicity hypothesis, the shared feature of organelle-encoded proteins is that they are all involved in electron transport, such as photosynthesis in chloroplasts and respiration in mitochondria (Allen, 1993, 2003). Being encoded in the organelles, these proteins can confer a selective advantage. One direct factor of evidence is that the D1 subunit of photosystem II, which is needed during photo-oxidative damage, is encoded by the chloroplast genome. Being nuclear encoded results in continuous synthesis and translocation to each chloroplast. In comparason, being encoded by chloroplasts might be more energy efficient (Allen & Martin, 2016). However, there is no direct experimental evidence for the CoRR hypothesis observed in mitochondira. Some chloroplast encoded subunits cannot function without being combined with additional nuclear-encoded subunits. This is in conflict with the CoRR hypothesis that offers immediate advantages in terms of protein functions, when they are encoded within chloroplasts (Daley & Whelan,

2005).

In comparison, the hypothesis of Muller’s ratchet, explains the organelles retaining at least partial genomes in the long-term evolution (Bock, 2007; Muller, 1964; W. Martin &

Herrmann, 1998). Muller’s ratchet is described as the non-recombining inheritance of mutant genes will cause an irreversible retrogression because most mutations are deleterious, while sexual reproduction can efficiently correct mutations by recombination or gene conversion.

Plastid and mitochondrial genomes have a high degree of polyploidy and a lower mutation rate

(Wolfe et al., 1987), while they typically follow maternal inheritance, which efficiently conserves these organellar genomes during nuclear sexual recombination and prevents deleterious mutations. This explains why plastids and mitochondria maintain some genes from

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their ancestral proteobacterial DNA. This theory also reveals some clues about the maternal

inheritance of the organellar genomes, although there are several cases of biparental or paternal inheritance reported (Barr et al., 2005; Ralph Bock, 2007). The dominant maternal inheritance leads to a relatively stable inheritance of genes that only pass from the maternal plants to the next generation and will not be affected by pollen. For the same reason, there will not be gene dispersal via pollen.

Another feature of plastids and mitochondria that encourages biologists to design strategies to edit their genomes is the high DNA copy number. There are usually hundreds to thousands of plastids and mitochondria in a mature leaf cell and each plastid or mitochondrion contains hundreds to thousands copy of organellar DNA (Alberts et al., 2002). High DNA copy number can potentially result in a high protein expression level and accordingly, a high yield of proteins. In addition, as organelles compartmentalize proteins inside, they may accumulate

toxic proteins by preventing the adverse interactions with cytoplasmic proteins, and therefore

minimize the pleiotropic effects of the transgenesis (Adem et al., 2017).

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Figure 1.2 Maps of the chloroplast and mitochondrion of Arabidopsis thaliana. Maps were drawn using the complete genome sequence (GenBank: NC_001284.2 and NC_000932.1) with version 1.1.1 of the OrganellarGenomeDRAW software tool (Lohse et al., 2007). (A) A. thaliana chloroplast genome map. Genes are mainly organized in one LSC and one SSC encoding proteins for the photosynthetic apparatus, the transcription/translation system and the biosyntheses. (B) A. thaliana mitochondrial genome map. Genes encode proteins for electron transport, ATP synthesis, translation, protein import and metabolisms.

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A

13

B

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1.2 Plastid Genome Engineering

Modifying the plastid genome has been a challenge for many research groups. The first

successful engineering of a chloroplasts genome was achieved in 1988 in a unicellular alga,

Chlamydomonas reinhardtii, using biolistic transfer (Boynton et al., 1988). Biolistic

transformation uses a gene gun to deliver DNA fragments coated on metal particles. The gene gun is uses microprojectiles coated with DNA fragments and accelerates them to a high speed to penetrate the plasma membrane, as well as the double membranes of the organelles, and thereby reach an organellar genome target. A screen placed between the microprojectiles and plant tissues blocks most of the microprojectiles to size select them and to reduce the damage

to the cells. The most common microprojectiles are made from gold or tungsten. Historically,

the first gene gun accelerated microprojectiles using a gunpowder charge (Klein et al., 1987).

Soon after, helium gas was used to accelerate microprojectiles, and instead of being set loosely in the chamber, microprojectiles were placed on a microcarrier (Johnston, 1990). The helium gas model of gene gun continuous to be the most commonly used.

The first successful biolistic transformation of a plastid genome in vascular plants was achieved in Nicotiana tabacum (tobacco) in 1990 (Svab et al., 1990). Since then, the plastid

genomes of several plant species have successfully been modified, including Marchantia polymorpha, Brassica oleracea (cauliflower, cabbage), Glycine max (soybean), Solanum

tuberosum (potato) (Reviewed by Bock, 2015).

There are many copies of plastids in a single plant cell, and it is important to obtain a

plant where all plastid chromosomes in all plastids are identically equivalent to homozygous

plants, these plants are called homoplastomic plants and do not show segregation of the plastid genomes. Generally, two selection systems have been utilized to obtain homoplastomic plants.

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Typically, before a biolistic transformation, plant tissues will first be treated with hormones to

obtain callus. Then after bombardment, the callus will be transferred to an antibiotic-containing regeneration medium. With the antibiotic resistant gene introduced, transformed callus will

grow while untransformed sectors will die. The antibiotic resistance expression in segregating cells is not cell autonomous, which means nontransformed sectors may show antibiotic resistance (Svab et al., 1990). Therefore, a second selection system is necessary. The second selection system is usually a visible marker, such as β-glucuronidase (GUS) or green

fluorescent protein (GFP) (Maliga, 2004). After the second selection, transformed tissues are

transferred to regeneration medium to induce shoot development. Transplastomic shoots will

then be transferred to another medium to induce roots. The shoot/root induced regeneration

will be repeated for several rounds until a homoplastomic plant is obtained (Bock, 2015).

Table 1. Successful cases of plastid gene transformation using biolistic. These are the cases which stable homoplasmic plants have been achieved using the primary selectable maker shown in the table. Selectable Reference Species Tissue marker Chlamydomonas photosynthetic (Kindle et al., Cells reinhardtii marker 1991) Suspension culture (Chiyoda et al., Marchantia polymorpha aadA cells 2007) Brassica oleracea Leaves aadA (Liu et al., 2007) (cauliflower, cabbage) Embryogenic (Dufourmantel et Glycine max (soybean) aadA callus tissue al., 2004) Nicotiana tabacum Leaves and aadA (Langbecker, 2004) (tobacco) suspension cells Solanum tuberosum (Sidorov et al., Leaves aadA (potato) 1999) Solanum lycopersicum Leaves aadA (Ruf et al., 2001) (tomato) (Kanamoto et al., Lactuca sativa (lettuce) Leaves aadA 2006)

There are several disadvantages to biolistic transformation of plastid genomes. First,

several parameters need to be optimized to target the right tissue depth, such as the velocity

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and the particle size needed for each tissue and species. Second, selection and regeneration of homoplastomic plants is a slow labor-intensive process, and protocols for regeneration from callus is species- and tissue-specific. New protocols must be developed for each species. Third, biolistic transformation is a random event and transformation efficiencies are low compared to

Agrobacterium-mediated transformation. Biolistic transformation is also more expensive due to the cost of the gold particles and limited reuse of microcarriers.

Because only a few species and actually individual plants that have successfully been plastome engineered in the past three decades, alternative methodologies should be considered that could improve the range and ease of plastid transformation.

A new methodology derived from a bacterial defense system now used for “gene editing” has been developed to engineer nuclear genomes (Barrangou, 2015). This technology is known as known as CRISPR/Cas9, an abbreviation for “Clustered Regularly Interspaced Short

Palindromic Repeats”, which uses a nuclease (Cas9) that is guided by a short single stranded

RNA (sgRNA) to induce a double strand cut in a target genome. The nuclear repair mechanism is imperfect and randomly induces or deletes nucleotides leading to insertion/deletion mutations (indels) and thereby creates sequence specific, targeted modification of the genome

(Sander & Joung, 2014). This methodology has inspired us to propose a new strategy to engineer plastid genomes. However, plastids lack enzymes to repair the DNA double strand breaks (DSBs). Therefore, non-homologous end joining (NHEJ) repair ligases, associated with heterodimers need to be imported into chloroplasts. In our strategy, Cas9 would first cleave at a desired site on the plastid genome to create DSBs under the guidance of sgRNAs. Then a heterodimer factor, such as Ku, will bind to the terminus of a DSB, so that a DNA-dependent protein kinase can phosphorylate the terminus of a DSB and initiate the NHEJ (Xu, 2006).

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Finally, a DNA repair ligase, such as ligase D, ligates the broken DNA strands together (Della et al., 2004) (Figure 1.3). To utilize this strategy in chloroplasts, proteins, Cas9, Ku and ligase

D, need to be imported into plastids by fusion with a transit peptide. RNA import is not common in plastids, therefore the challenge is how to import a single guidance RNA (sgRNA) into plastids.

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Figure 1.2 The design of our Cas9-ligation chloroplast genomic DNA engineering process. (A-F) Cas9 first cleaves at a desired site on the plastid genomic DNA to create a DSB under the guidance of a sgRNA. Then a heterodimer factor, Ku, binds to the termini of the DSB, and a DNA-dependent protein kinase phosphorylates the termini of the DSB and initiates the NHEJ. Finally, a DNA repair ligase, ligase D, ligates the broken DNA strands together.

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A Cas9 sgRNA B

B Cas9 sgRNA

C Ku

D DNA-PKcs Ku

E DNA-PKcs Ku

Ligase D F

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1.3 RNA Translocation into Plastids

RNA translocation into subcellular organelles has not been studied very much. The first

case report on the uneven distribution of RNAs in cells was reported in 1983 (Jeffery et al.,

1983). Few cases of intercellular RNA translocation from the nucleus to plastids have been

reported.

RNAs are typically classified into messenger RNAs (mRNAs) that can be translated to

proteins, and noncoding RNAs (ncRNAs) that are not translated. The translocation of mRNA

from the nucleus into plastids has not been reported. While some reports indicate that there are

specific ncRNAs encoded by nuclear genomes that have been found in chloroplasts, evidence

of importation into chloroplasts is still lacking (Lung et al., 2006).

Viroids are single-stranded, self-complementary, circular ncRNAs, functioning as

infectious agents that can move into organelles and induce pathogenesis in the host plant cells

(Wang & Ding, 2010). So far, 42 viroids plus 6 related candidate viroids have been accepted

by the International Committee on Taxonomy of Viruses (Daròs, 2016; Owens et al., 2012).

Among these 48 viroid species, 5 are from the Avsunviroidae family, while the remaining 43

are from the Pospiviroidae family (Owens et al., 2012). Viroids from the Avsunviroidae family

share the same molecular mechanisms of folding into a quasi-rod-like secondary structure with two bifurcations at both ends, replicating within the chloroplasts following a symmetric pathway and possessing intrinsic ribozyme activities (Daròs, 2016; Wang & Ding, 2010).

Whereas, viroids from the Pospiviroidae family share the same features that folding into a rod-

like secondary structure and replicating within the nucleus but follow an asymmetric pathway

(Figure 1.4).

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Figure 1.3 The structural models and replication pathways of the Pospiviroidae family and the Avsunviroidae family (after Gago-Zachert, 2016). Pospiviroidae viroids fold into a rod-like secondary structure and replicates in the nucleus following an asymmetric pathway (left). Avsunviroidae viroids fold into a quasi-rod-like secondary structure and replicates in the chloroplasts following a symmetric pathway (right). The key difference in the “life-cycle” of Pospiviroidae and Avsunviroidae viroids is that replication of Pospiviroidae viroids occurs in the nucleus while Avsunviroidae viroids replicate in chloroplasts. This replication in the chloroplast requires an import mechanisms for the RNA

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viroid across the outer and inner chloroplast envelope membranes (Wang & Ding, 2010). One

significant case was reported by Goméz and Pállas in 2010 where a sequence of the Eggplant

latent viroid (ELVd), a member from the Avsunviroidae family, can mediate the translocation of mRNAs from the cytosol into chloroplasts (Gómez & Pallás, 2010b).

ELVd was first found by Fagoaga. (1994) in Solanum melongena L. (eggplant). Elvd consists of 332 to 335 nucleotides (nt) that can cause symptomless infections and is transmitted between plants though mechanical contact and in seeds (Daròs, 2016). ELVd possesses an intercellular traffic ability that first translocates into the nucleus and then translocates into the chloroplasts to replicate (Gómez & Pallás, 2012b). Although the mechanism of how the ELVd infects plant cells and its ability to integrate into the nucleus and translocates it’s RNA into the chloroplasts remains unclear, there is some evidence that a 168-nt long fragment in the left terminal region of the molecule between positions 15 and 181 is sufficient to mediate the mRNA translocation into the nucleus (Gómez & Pallás, 2012a). Similarly, a 110 nt long fragment in the left terminal region between positions 52 and 150 is sufficient to mediate the mRNA translocation into chloroplasts (Gómez & Pallás, 2010a). Deletions in this 110 nt region will disrupt the ability of ELVd to infect eggplant and translocate into chloroplasts (Martínez et al., 2009). All these lines of evidence suggest that the left terminal region controls ELVd localization.

Utilization of the mechanism that enables ELVd to effectively mediate mRNAs translocation into chloroplasts consists of three basic processes (Gómez & Pallás, 2010b). First, a chimeric sequence is constructed by inserting a piece of complementary DNA (cDNA) of the derived ELVd sequence into the 5’ untranslated region (5’ UTR) of the gene of interest.

Second, this chimeric sequence is transformed into the nucleus in plant cells using

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Agrobacterium. Finally, the transcript product of this chimeric sequence, an mRNA of the gene

of interest fused with a ncRNA of ELVd, will translocate into chloroplasts.

Another relevant study, was reported by Nicolaï et al. (2007), showing that the mRNA

of eucaryotic translation initiation factor 4E (eIF4E) was originally transcribed in the nucleus

and imported into chloroplats. However, using bombardment of GFP fused with eIF4E target

sequence failed to show GFP expression inside chloroplasts (Nicolaï et al., 2007).

1.4 Mitochondrial Genome Editing

For mitochondrial genome editing, the situation is even more complicated as there is

only one case reported in yeast that accomplished mitochondrial genome editing using

bombardment (Johnston et al., 1988), but not a single successful case of genome engineering

of mitochondria in plants has been reported. The translocating function of ELVd is restricted

to chloroplasts and does not work in other subcellular organelles, such as mitochondria

(Ahmad & AbouHaidar, 2016).

Nonetheless, there is one report that used a fusion construct of target-specific sgRNA with a mitochondrial targeting sequence (MTS) from subunit VIII of human cytochrome c oxidase to import RNA into mitochondria (Jo et al., 2015). They attempted to cleave

mitochondrial DNA in human embryonic kidney cells (HEK-293T) cells. However, Jo et al.

failed to prove directly that mitochondrial DNA did get cleaved directly, which can easily be done using reverse transcription polymerase chain reaction (RT-PCR). On the other hand, they

proved that the downstream protein was expressed. Secondly, they failed to prove that the

sgRNA did get imported into the mitochondria.

Another possible option is described in a patent by Belmonte et al. (2016). They

proposed to edit the mitochondrial genome by injecting restriction endonucleases, such as

24

XmaI and other enzymes such as transcription activator-like effector nucleases (TALENs) and zinc finger nucleases (ZFNs), in a form of either mRNA or protein into a female gametocyte using transformation. It is a very practical idea since importing TALENs or ZFNs as proteins into mitochondria can be easily accomplished by fusing a β-signal to the desired protein. In plants, fusion genes of these enzymes could be transformed into the nucleus for expression of the mitochondrial target protein fusions. However, maintaining and producing a desired

TALEN or ZFN are always challenges.

1.5 RNA Translocation into Mitochondria

Still, if CRISPR/Cas 9 use is desired, it is important to find a strategy to import sgRNA into mitochondria.

Unlike plastids, RNA import into mitochondria is a more common phenomenon. The first case of tRNA being transported into mitochondria was in a ciliate, Tetrahymena pyriformis (Suyama, 1967). Since then, transport of many other types of RNA into

mitochondria have been observed, including tRNAs, 5S ribosomal RNA (rRNA), RNase

mitochondrial RNA processing (MRP) RNA, RNase P RNA, and microRNAs (mitomiRs)

(Bandiera et al., 2011; Holzmann et al., 2008; Reviewed by: Kim et al., 2017; Magalhães et

al., 1998; Sripada et al., 2012; Yoshionari et al., 1994; Chang & Clayton, 1987, 1989; Puranam

& Attardi, 2001). However, most of these RNA transport were demonstrated in mammals. In

plants, the most common type of RNA transported into mitochondria is tRNA. Mitochondria

in some other species (not plants) can synthesize the entire set of tRNAs may also import tRNAs. For example, Saccharomyces cerevisiae, will import tRNAs into mitochondria,

especially under stress, such as temperature (Kamenski et al., 2007; R. Martin et al., 1979).

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The pathway of tRNAs transport into mitochondria is not known, but there is some evidence for a protein import apparatus that also imports tRNA (Murcha et al., 2016). The mechanism of tRNA transport in the yeast Saccharomyces cerevisiae and in the protozoans

Trypanosoma or Leishmania are better understood, as more studies have been done in these species. The tRNA type (neutral or charged), and some other factors, including the import apparatus varies. The process can be either ATP-dependent or -independent (Entelis et al.,

2006; Mukherjee et al., 1999; Rinehart et al., 2005; Sherrer et al., 2003). Nonetheless, tRNA transport in plants requires elements similar to those required for protein transport. Most studies on tRNA transportation are based on in vitro assays (Sieber et al., 2011).

One work indicates that the tRNA translocation into mitochondria, is an ATP-dependent process associated with outer membrane protein receptor(s) (Delage et al., 2003). This process can occur in vitro without a cytosolic protein extract. TRNA translocation into inner mitochondrial membranes requires an electrochemical membrane potential resulted by pumping out protons, (Delage et al., 2003; Bhattacharyya & Adhya, 2004). This might explain the movement of negatively charged tRNAs towards the negative matrix crossing the hydrophobic environment between the two mitochondrial membranes. Delage et al. (2003) also noticed that the efficiency of tRNAs uptake is much higher in vitro, suggesting that the inner membrane of mitochondria is either a strong or a specific barrier for tRNA internalization into the mitochondrial matrix. They observed that a certain type of tRNA out competed the other isoacceptors, although no single “import signature” of the tRNAs has been identified.

They proposed that aminoacyl-tRNA synthetases (aaRSs) may play a role in the recognition of the types of tRNA on the mitochondrial membrane.

26

Other research indicates that plant mitochondrial voltage-dependent anion channel

(VDAC) protein is the major component for tRNA uptakes (Salinas et al., 2006). VDAC is a channel that transports many kinds of metabolites, including ATP, ADP, succinate, malate, and pyruvate. Other results indicate that DNA can be translocated into mitochondria through

VDAC in both mammalians and plant cells (Koulintchenko et al., 2003; Szabò et al., 1998).

Salinas et al. (2006) confirmed that tRNA can be translocated into mitochondria through

VDAC as well. The exclusion limit of VDAC is only around 3 kDa, so how VDAC modifies to let different molecules pass, especially tRNA which are larger because of their L-shaped structure, remains unclear. Additionally, Salinas et al. (2006) verified that two major components from the TOMs, TOM20 and TOM40 function as outer membrane protein receptors responsible for tRNA recognition and binding on the mitochondrial outer membrane surface. Antithetically, the TIMs show little or no contributions to the tRNA import pathway.

Furthermore, Salinas et al. (2006) also noted that both the tRNA binding process with TOM20 and TOM40 as well as the importing process through VDAC consume ATP.

Another set of outer membrane protein receptors are Tric1 and Tric2 from the Tailless

Complex Polypeptide 1 Ring Complex (TRiC) (Murcha et al., 2016). These components originate from the preprotein and amino acid transporters family (PRAT), which is distinguished by a sterile-a-motif (SAM) domain at the C-terminus and are predicted to bind tRNAs.

On the other hand, the sequences as well as the secondary structures of tRNAs determine whether or not the tRNA can be imported, although there is no single “import signature” that is shared by all types of tRNAs. In terms of the tRNA sequences, in higher plants, such as tobacco, neither the D-loop, TΨC loop nor the anticodon loop of a tRNA can be modified for

27

a successful import. In other words, the combination of the D-loop, TΨC loop and the

anticodon loop determines the recognition of tRNAs (Salinas et al., 2005). In other species,

such as Phaseolus vulgaris and the lower plant Marchantia polymorpha, the tRNA sequences

for recognition are more variable and are more specific for each tRNAs (Akashi et al., 1997;

Ramamonjisoa et al., 1998). Only mature tRNAs can be imported into mitochondria, suggesting that the secondary structure plays a role in recognition by the translocator (Delage et al., 2003).

In spite of all the elements discovered so far relating to tRNA transport into mitochondria, there are many questions about the mechanism still need to be answered. Does the tRNA unfold to fit in the size of VDAC? If so, will only mature tRNAs can be recognized? What are the components of the inner membrane of mitochondria that recognize and transport tRNAs?

When these questions are answered, a better strategy for manipulating RNA translocation into mitochondria be established.

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CHAPTER 2

RNA Transport into Chloroplasts Mediated by ELVd

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Introduction

Plastids are one of the two types of semi-autonomous organelles in plant cells. As semi- autonomous organelles, plastids contain their own genomic DNA called the plastome

(Olejniczak et al., 2016). The plastome encodes transfer RNAs (tRNAs) and ribosomal RNAs

(rRNAs), and therefore, they can translate messenger RNA (mRNA) which are essential for plastid development and metabolism (Olejniczak et al., 2016). Plastids are like to have once been free-living organisms, according to the endosymbiotic theory of plastid origins (Martin et al., 2015).

The endosymbiotic origin theory of plastids developed over time because of infoemation from cell and molecule biologyover time with the development of the technologies. A German pathologist, Richard Altmann, first put forward the original idea of coexistence of bioblasts in eukaryotic cells in 1890, although it is under debate whether Altmann discovered mitochondria specifically (Altmann, 1894; Kutschera & Niklas, 2005). Fifteen years later (1905), a Russian botanist, Constantin Mereschkowsky first proposed a fully elaborated version of the endosymbiotic origin theory. He claimed that “chromatophores,” another name for chloroplasts at that time, “are foreign organisms that invaded the colourless plasm of the cell and entered into a symbiotic coexistence with it” (see the translated version in English: Martin

& Kowallik, 1999). Limited by the technology available at that time, the four pieces of evidence presented in C. Mereschkowsky’ paper were not convicing now. He argued that first, chromatophores did not arise de novo but pre-existed in egg cells and were passed down to progeny from there. Second, chromatophores can live, and even divide, when the nucleus is absent. Third, chromatophores were similarly to Zoochlorellae, a respective structure in animals, suggesting they might both be independent organisms that entered colorless cells

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forming symbiotic coexistence later on; and fourth, chromatophores are very similar to free-

living, Cyanophyceae (cyanobacteria). Mereschkowsky’s proposal laid the foundation of the

endosymbiotic origin theory of plastid evolution.

Modern technology offers more evidence to support this theory. Phylogenetic studies on

the genomes sequences of plastids indicate that the closest extant relative of the plant plastid

is cyanobacteria (Martin et al., 2002, 2003). As the closest relatives, plastids and cyanobacteria

both express some protein components in common, such as the key enzymes in the plastid

metabolic pathway, photosystem I and II (Ben-Shem et al., 2003). Further support for the

endosymbiotic origin theory is that some proteins originated in a proteobacterial ancestor of

both plastids and mitochondria, and are now encoded in the eukaryotic nucleus. For example,

β-barrel-membrane proteins (bbps) in the outer membranes of plastids and mitochondria are

coded by genes in the nucleus and are synthesized in the cytosol in plant cells. They also exist

in the proteobacterial ancestor of mitochondria (Paschen et al., 2003). This last example reveals

the integration of the protein in the genome of the proteobacterial ancestor of plastids and the

nucleus of the host cell.

The genome of plastids is much smaller than that of its free-living ancestors indicating either gene loss or transpositions from endosymbiont to the nucleus of the host cell during evolution (Weeden, 1981). Arabidopsis thaliana, for example, maintains 17 insertions with a total of 11 kilobase pairs (kb) in the nuclear genome conserved from the cyanobacterial ancestor (The Arabidopsis Genome Initiative, 2000). Comparably, gene transposition from the nuclear genome to the organellar genomes are much rarer. There is only one putative case of a

transfer of a nuclear gene from the MutS gene family in a Coral species, Sarcophyton glaucum

reported that might have been transported into the mitochondrial genome (Pont-Kingdon et al.,

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1998). No nuclear to plastid gene transposition has been reported so far. Plastids still maintain

many genes from the cyanobacterial ancestor. These genes encode proteins function in three

different metabolism systems: the photosynthetic apparatus, the transcription/translation

system and biosynthetic processes (Wakasugi et al., 2001).

Plastids differentiate into three types, chromoplasts, chloroplasts, and leucoplasts. In

studies of plastids, biologists focus more on the chloroplast than the other two as it involves photosynthesis, and consider it to be a model organelle. Genomic studies are essential for the

studies of chloroplasts, while genetic engineering is used to construct transgenic plants for

commercial uses. Compared to nuclear genomic engineering, chloroplast genomic engineering

has many potential advantages. For example, as chloroplasts are only passed down from the

maternal plant to the next generation, stable transgenic traits can be reliably inherited for many generations. For the same reason, there will not be trait segregations in the offspring or contaminations spread by pollen. In addition, chloroplasts sequester proteins so it is possible to accumulate certain types of proteins in chloroplasts without affecting other functions of the plant. There are usually multiple chloroplasts in a single cell and multiple copies of genomes in each chloroplast, therefore high yields of proteins are possible (Alberts et al., 2002).

The only successful method of delivering transgenes into chloroplasts is by using a gene gun for “bombardment”. Bombardment requires shooting DNA covered gold particles at high velocity into tissue or protoplasts. Transgenic cells then need be selectively regenerated to construct homoplasmic lines (Bock, 2015).

Transformation of plastid genomes is needed to insert double-stranded DNA (dsDNA) with at least two plastid DNA fragments as flanking sequences into the plastid, and

Homologous Recombination (HR) can integrate the DNA into the chromosome(s). Biolistic

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delivery of double stranded DNA inserts DNA randomly. Selection and regeneration are complex and often require species-specific protocols to regenerate homoplasmic plants

(Daniell et al., 2002). An alternative mechanism might be to insert a single strand of nucleic acid and then produce the desired double-stranded DNA in the plastid itself. A standard method to produce dsDNA in vitro uses Reverse Transcriptase (RT) and an RNA template (Smith,

2010). RT requires a sequence specific primer to start the single-stranded DNA (ssDNA) synthesis from the 5’ end toward the 3’ end. Then RT catalyzes the degradation of the RNA template and uses the ssDNA as template to synthesize the complementary ssDNA (Daniell et al., 2002). Together, these two ssDNA form a dsDNA that can insert into the chloroplasts genomic DNA through HR.

In order to synthesize double-stranded DNA (dsDNA) from a template RNA in vivo in a plastid, RT and template RNA would need to be transported into the plastid. Mechanisms of importing proteins like the RT into chloroplast are well established (Soll & Schleiff, 2004). To be specific, the gene of RT fused with a transient peptide is transformed into the nucleus. Then the DNA fusion is transcribed in the nucleus and translated in the cytoplasm and the nascent polypeptide can be imported into a plastid mediated by the transit peptide. The transit peptide can be recognized and imported into the chloroplast stroma by the translocon on the outer/inner chloroplast membrane complex (TOC/TIC). After import, the transit peptide are removed by peptidase (Soll & Schleiff, 2004).

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Figure 2.1 Proposed chloroplast genomic DNA engineering through reverse transcription and homologous recombination (image provided by Colin Murphree). RT and the RNA template are synthesized in the cytoplasm and the nucleus, and then translocate into chloroplasts. RT reverse transcribe the RNA template into dsDNA and the dsDNA inserts into the chloroplasts genomic DNA through HR. A novel strategy to import RNA into a plastid was reported by Gómez and Pallás

(2010b). A sequence of noncoding RNA from a viroid can mediate foreign mRNA transport into chloroplasts (Gómez & Pallás, 2010b). Insertinon of a modified complementary DNA

(cDNA) sequence of Eggplant latent viroid (ELVd) (from Avsunviroidae family) at the 5’ end of the cDNA of Green Fluorescence Protein (GFP) as a 5’ untranslated region (5’ UTR), enables the RNA to be translocated into chloroplasts. The plastid localized RNA was translated and GFP accumulated in chloroplasts. How ELVd imports mRNA into chloroplasts remains unclear, Goméz and Pállas further confirmed that the secondary structure of the ELVd sequenceplays an important role in this process (Gómez & Pallás, 2010a).

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ELVd has been reported to have a restricted host range. ELVd cannot infect many other

viroid host species through its original mechanical transmission process, including

Lycopersicon esculentum L. (tomato), Cucumis sativus L. (cucumber) and Chrysanthemum ×

morifolium Ramat (chrysanthemum) and Citrus medica L. (citron) (Fagoaga et al., 1994). With the uncertainty of the mechanism of the interaction of ELVd with chloroplasts, it is important

to show how the features of ELVd mediating the translocation of mRNAs into chloroplasts so

it can be used in species other than tobacco and so this feature can more generally enable

plastid transformation.

Another potential mechanism for importing RNA into chloroplasts was also tested.

Nicolaï et al. (2007). MRNA of the nuclear-encoded eukaryotic translation factor 4E (eIF4E)

is translocated into chloroplasts in four species, Arabidopsis thaliana, Nicotiana tabacum,

Lactuca sativa (lettuce) and Spinacia oleracea (spinach) (Nicolaï et al., 2007). The authors

constructed a chimeric gene cassette of eIF4E fused with a codon modified GFP (mGFP5), and delivered this into A. thaliana using particle bombardment. However, even though they detected the mRNA of this chimeric gene cassette, they failed to detect any proteins translated from either eIF4E or mGFP.

The focus of the research reported here is to identify sequences from ELVd and eIF4E

that confer ability to transport nuclear-encoded RNA into plastids.

Methods and Materials

Plant materials

Seeds of A. thaliana and N. benthamiana were vernalized with 0.1% agarose and stored at 4 ℃ for 48 hours (hr). Then the seeds were planted on the surface of autoclaved soil. Plants

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were maintained at 25 ℃ w i th 16:8 hours (hr) light:dark period. The ecoild type A. thaliana

Columbia (Col-0) was used for transformation.

Plasmid construction

The chimeric DNA segments of enhanced green fluorescent protein (eGFP) fused with

ELVd (Appendix B) was synthesized as gBlocks® Gene Fragments (Integrated DNA

Technologies [IDT], Coralville, Iowa). The DNA segments were ligated into cloning vector pUC 19 using SmaI blunt end restriction enzyme (New England Biolabs Inc. [NEB], Ipswich,

Massachusetts) and T4 DNA ligase (NEB, Ipswich, MA). Plasmids with chimeric DNA segments were transformed into Escherichia coli (E. coli). One milliliter (mL) of the transgenic

E. coli cultures were mixed with one mL of 70% glycerol, incubated at room temperature for one hr, and frozen at -80 ℃ for long term storage. The remainder of the transgenic E. coli cultures were used for plasmid isolation using the QIAprep Spin Miniprep Kit (Qiagen,

Germantown, Maryland), and the plasmids were sent for sequencing (Genewiz, Morrisville,

North Carolina; Eurofins Genomics, Louisville, Kentucky) to verify the sequences.

The transgenic E. coli cultures with confirmed sequence of the chimeric DNA segments were then cultured in a large volume. After plasmid isolation using the ZymoPURE™ Plasmid

Midiprep Kit (Zymo research, Irvine, California), restriction enzymes XbaI and BamHI were used to cut the chimeric DNA segments out of pUN19 and to linearize the expression vector

PC-GW-BAR (KP826773.1) (NEB, Ipswich, MA). Digested chimeric DNA segments and linearized PC-GW-BAR vector DNA were purified using the QIAquick® Gel Extraction Kit

(Qiagen, Germantown, MD) T4 DNA ligase was used to ligate the chimeric DNA segments into the PC-GW-BAR vector (NEB, Ipswich, MA).

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Transformations

Fifty microliters (µL) of E. coli strain DH5α competent cells (Lucigen Corporation,

Middleton, Wisconsin) were chilled on ice for 20 minutes (min) and mixed with 5 μL of ligation product. The cell mixtures were transferred to a water bath at 42 ℃space and incubated for a 90 second (sec) heat shock. Then the cell mixtures were quickly chilled on ice for two

min. Each sample tube was filled with 300 μL LB and incubated in a shaker at 37 ℃ for 1 hr to recover. 100 μL of recovered cells were spread on LB plates appropriate with proper antibiotics using glass beads, and incubated at 37 ℃ overnight (Lucigen Corporation, n.d.).

Purified plasmids were diluted to 15 ng/µL. 50 µL of Agrobacterium tumefaciens strain

GV3101 competent cells were chilled on ice for 20 min and then mixed with 2 μL of the diluted plasmids. Cell mixtures were transferred into precooled 0.1 centimeter (cm) cuvettes and shocked with a preset program for agrobacteria (2400 mV, 200 Ω, 250 µFD) using the Gene

Pulser Xcell™ Electroporation System (BIO-RAD Laboratories, Inc., Hercules, CA). Each tube was filled with 1 mL LB was added to each tube and incubated for 2.5 hr in a shaker at 28 ℃ for cells to recover. 70 µL of recovered cells were spread on LB plates with appropriate antibiotics and incubated at 28 ℃ for two to three days (Weise, 2013).

Transgenic plants selection

T1 seeds were collected from transformed A. thaliana plants and vernalized in the same

way as wildtype. 50 μg/mL BASTA herbicide (Glufosinate-ammonium from Sigma-Aldrich

Corporation, Raleigh, NC) was applied by spraying on T1 plants after they grew two to four

true leaves. Transgenic plants carrying the bar gene are resistant to BASTA while non- transformed or non-transgenic seedlings are sensitive and will die after BASTA application. It

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sometimes took up to 5 sprayings (3 days apart) to achieve reliable selection of the transgenic

plants.

Total DNA isolation

100 mg of leaf tissue (ca. one leaf) from each A. thaliana T1 plant was collected, frozen

in liquid nitrogen and ground well using a bead beater for 45 sec. Cetyltrimethylammonium bromide (CTAB) extraction buffer (Appendix A) was premixed with β-mercaptoethanol at a

ratio of 600:1 (v/v) and the mixture was preheated to 65 ℃. Each tissue sample was mixed with

600 μL CTAB and β-mercaptoethanol mixture using beads beater and incubated at 65 ℃ for

15 min. A 600-μL aliquot of a chloroform and isoamyl alcohol mixture (ratio of 24:1 [v/v])

was added to each sample mixed and then centrifuged at 13,000 g for 10 min at room

temperature. The aqueous phases were then transferred to new tubes and vortexed with an

equal volume of isopropanol and then held at -20 ℃ overnight. Samples were thawed at room

temperature and centrifuged at 13,000 g for 10 min, and the supernatant discarded. The pellets

were washed with 1mL of 70% ethanol and 0.5 mL of 100% ethanol in turn and then left to

dry in air. DNA samples were resuspended in 100-200 µL of TE buffer (Appendix A) and

stored at -20 ℃.

Total RNA isolation and RT-PCR analysis

100 mg of leaf tissue (ca. one leaf) from A. thaliana T1 plants were collected, frozen in

liquid nitrogen and ground well using a bead beater for 45 sec. An RNeasy Plant Mini Kit

(Qiagen, Germantown, MD) was used to extract total RNA. DNA was removed using the

TURBO DNA-free™ Kit TURBO™ DNase Treatment and Removal Reagents (Invitrogen by

Thermo Fisher Scientific, Grand Island, NY). Isolated RNA was used as templates to

synthesize cDNAs using RNA to cDNA EcoDry™ Premix (Random Hexamers) (Clonetech

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Laboratories, Inc., Mountain View, CA). CDNAs were then used as template in Polymerase

Chain Reaction (PCRs) to amplify target genes. PCR products were sent out to verify

sequences.

Total protein isolation and western blotting

100 mg of leaf tissue from each plant, N. benthamiana, or A. thaliana was collected,

frozen in liquid nitrogen and ground to a fine powder using a bead beater for 45 sec. Each

sample was then vortexed in 200 μL of buffer E (Appendix A), ground in a bead beater and

then centrifuged at 10,000 g for 10 min. Supernatants were transferred to new tubes and

centrifuged again. Supernatants were then transferred to new tubes, mixed with 20 µL buffer

Z (Appendix A) and stored at -20 ℃.

40 μL of each protein sample was mixed with 10 μL buffer Z, and 5 μL BLUeye

Prestained Protein Ladder (Gel Company, San Francisco, CA) was vortexed briefly with 45

μL buffer Z. Samples were incubated at 85 ℃ for 5 min and then loaded onto NuPAGE™ 4-

12% Bis-Tris Protein Gels (1.5 mm, 10 well, Invitrogen by Thermo Fisher Scientific, Grand

Island, NY). Gels were run at 200 V until chlorophyll ran off the gel. Proteins were transferred

onto PVDF membranes using electrode cassettes submerged in freshy made transfer buffer

(Appendix A) and run at 50 V at 4 ℃ overnight. Blots were then blocked with 3% BSA/TBST

for at least one hr. Blots were probed with eGFP Tag Monoclonal Antibody (Invitrogen by

Thermo Fisher Scientific, Grand Island, NY) at a dilution of 1: 1,000 (v/v) in BSA/TBST at

room temperature overnight. Blots were then incubated with Goat anti-Mouse IgG (H+L)

Secondary Antibody, HRP (horseradish peroxidase) (Invitrogen by Thermo Fisher Scientific,

Grand Island, NY) at a dilution of 1: 10,000 (v/v) in BSA/TBST at room temperature for least

one hr. Blots were incubated with SuperSignal™ West Pico PLUS Chemiluminescent

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Substrate (Invitrogen by Thermo Fisher Scientific, Grand Island, NY) to detect eGFP. Blots

were detected with CL-XPosure™ Film (Thermo SCITIFIC by Thermo Fisher Scientific,

Rockford, IL) and developed with SRX-101A Tabletop Processor (Konica Minolta Healthcare

Americas, Inc., Wayne, NJ).

Confocal microscopy and lambda scan

For N. benthamiana, leaf tissues were freshly cut from transient transgenic plants and,

for A. thaliana, leaf tissues were freshly cut off from basta-resistant T1 plants, 25 days old.

Tissues were placed on a standard glass slides with the abaxial surface up. Slides were then

inverted and placed on the platform of a confocal microscope with Airyscan, ZEISS LSM 710

for N. benthamiana and ZEISS LSM 880 for A. thaliana (ZEISS, the U.S.). Laser excitation

wavelength was set at 488 nm for the fluorescence signal with an emission window of 503 nm

to 619 nm and a 9-nm gap in between for lambda scan. ImageJ was used to edit images of

confocal microscopy and lambda scans.

Results

The primary purpose of this research project was to test the ability of the plastid ELVd

sequence and eIF4E to transport nuclear-encoded RNA into the plastids of A. thaliana plants

as an example of a non-host plant for the viroid (Fadda et al., 2003). N. benthamiana was used

as a control for transformation and DNA transfer, because it had been successful for Gómez and Pallás (2010b).

Plasmid Constructions

The ELVd cDNA sequence was obtained from Gómez and Pallás (2010b), while eGFP cDNA sequence was obtained from the PC-GW-EGFP vector (KP826772.1) (Dalal et al., 2015;

Gómez & Pallás, 2010b) (Appendix B). The two restriction sites, XbaI and BamHI, were

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selected for double restriction site cloning to excise the ccdB gene. A chimeric cDNA sequence was constructed starting with the XbaI restriction site, followed by ELVd and eGFP, and ending with a BamHI restriction site. The chimeric sequence was sent for synthesis as gBlocks® DNA fragments (IDT, Coralville, Iowa). As only 80% of synthesized gBlock®

DNA fragments are error free, the DNA fragments were first cloned into the cloning vector

pUC19 to confirm the sequence. Clone with the correct sequence were selected for further use.

The chimeric cDNA fragment was excised from pUC19 and ligated into BASTA resistant

expression vector PC-GW-BAR (KP826773.1) at the same restriction sites as above. The

ligated expression vector was verified again by sequencing. The expression vector was finally

transformed into Agrobacterium tumefaciens strain GV3101, which is rifampicin and

gentamicin resistant. This expression cassette uses the Cauliflower Mosaic Virus (CaMV) 35S

promoter (GenBank: AB626665) and CaMV 35S terminator (GenBank: HM750245.1).

The transit peptide fused with the eGFP chimeric cDNA and eIF4E fused with eGFP

chimeric cDNA were cloned into the same expression vector using the methods as described

above. For a positive control, the eGFP DNA fragment was PCR amplified from the PC-GW-

EGFP vector and cloned into the expression vector PC-GW-BAR. The empty PC-GW-BAR

vector was transformed into Agrobacterium and into plants as a negative control.

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Figure 2.2 Vector map. The PC-GW-BAR possesses a BASTA resistant gene (BAR) and a ccdB gene, which can function as a selective marker for E. coli. The ELVd-GFP DNA fragment was ligated into PC-GW-BAR using restriction-ligation cloning. The expression cassette uses CaMV 35S promoter and terminator. Tobacco Transformation

Agrobacterium was used to transfer DNA into N. benthamiana leaves. After a 2-day

dark treatment, the leaf tissues of transgenic N. benthamiana were sent for confocal

microscopy (Cellular and Molecular Imaging Facility, NCSU) (Figure 2.3). The ELVd fused

with eGFP (ELVd::eGFP) transgenic lines showed the same expression pattern as the

Chloroplast Transit Peptide fused with eGFP (CTP::eGFP) transgenic line that the eGFP signal

overlapped the autofluorescence signal of chloroplasts. However, the eIF4E fused with eGFP

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(eIF4E::eGFP) transgenic line did not show a GFP signal in confocal microscopy, suggesting

this candidate could not mediate the RNAs translocation into chloroplasts.

Figure 2.3 Confocal microscopy of N. benthamiana transgenic plant leaf tissue. Laser excitation was set at 488 nm to detect the eGFP signal. Pixel dwell was set at 3.15 microseconds (μs) to capture clear details, and the brightness and construct were adjusted to 400% using ImageJ to show the details. The ELVd::eGFP transgenic lines showed the same expression pattern as the CTP::eGFP transgenic line that the eGFP signal overlapped the autofluorescence signal of chloroplasts. A western blot of leaf tissues from the same transgenic plants shown above was carried

out to verify the expression of eGFP (Figure 2.4). The unmodified eGFP line showed a band

at 33 kDa corresponding to the full size of eGFP (32.7 kDa). No eGFP was detected in the

ELVd::eGFP or in the eIF4E::eGFP transgenic lines. The results with ELVd::eGFP are in

conflict with the confocal microscopy. No GFP protein signal was observed (microscopically)

or detected (western blot) in the transgenic plants carrying the eIF4E-fusion.

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Figure 2.4 Western blot for full-length eGFP of N. benthamiana. The unmodified eGFP line showed a band at 33. The ELVd::eGFP or in the eIF4E::eGFP transgenic lines did show bands. EGFP Was Not Detected in Arabidopsis

The results of N. benthamiana transient expression showed that ELVd may mediate the translocation of RNA while eIF4E may not. To verify the ability of translocating of these two molecule candidates, the same empty PC-GW-BAR vector, and expression vectors of

ELVd::eGFP, eIF4E::eGFP and CTP::eGFP, were transformed into A. thaliana using floral dip. Each construct was transformed into three plants and all T1 seeds were collected. A

homozygous eGFP transgenic line constructed in previous project was used as a positive

control. All of the T1 seeds of the ELVd::eGFP transgenic line were screened with BASTA

and 11 plants survived. No GFP signal was observed in any of the 11 T1 plants under confocal

micrscopy. Half of the T1 seeds of the eIF4E::eGFP transgenic line were screened with BASTA

but no transgenic plant survived. Half of the T1 seeds of the CTP::eGFP transgenic line were

screened with BASTA and six plants survived. A GFP signal was observed in all six T1 plants

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under confocal microscopy. One-fourth of the T1 seeds of the empty vector transgenic line were screened with BASTA and fiver plants survived, and were used as negative control in confocal microscopy. However, the emission of the ELVd::eGFP transgenic line is brighter

than the empty vector negative control but darker than the CTP::eGFP positive control (Figure

2.5). Therefore, a lambda scan was used on the same leaf tissue of the ELVd::eGFP transgenic

plant but in a different area to avoid the bleaching (Figure 2.6). A lambda scan under confocal

microscopy captures a stack of images at a serial emission wavelength under the same laser

excitation wavelength. In this lambda scan, the region of emission wavelength was set from

503 nm to 619 nm with a 9-nm gap between each scan.

Figure 2.5 Confocal microscopy of A. thaliana T1 transgenic plant leaf tissues. Laser excitation was set at 488 nm to detect eGFP signal. Pixel dwell was set at 2.45 μs for empty vector, transit peptide, and ELVd::eGFP transgenic lines; and 1.97 μs for unmodified eGFP to capture the best brightness. The emission of the ELVd::eGFP transgenic line is brighter than the empty vector negative control but darker than the CTP::eGFP positive control.

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Chloroplasts reveal a broad emission range of the autofluorescence with two domain emission wavelength regions in the yellow wavelength region (520-561 nm) and the red wavelength region (648-709 nm) under the excitation wavelength of 488 nm (Kodama, 2016).

The emission wavelength region of eGFP is 475 nm to 575 nm under the excitation wavelength of 488 nm (Song, 2007). Therefore, if it were an eGFP signal, it would be lost if the wavelength went beyond 575 nm in a lambda scan. However, in the region of wavelength from 503 nm to

619 nm, the signal did not disappear at any point. On the contrary, the signal became stronger at the beginning, indicating that this signal was chloroplast autofluorescence.

Figure 2.6 Lambda scan of the same leaf tissue of ELVd::eGFP transgenic line from figure 2.5. The region of wavelength from 503 nm to 619 nm with a 9-nm gap in between. The signal did not disappear at any point. To verify GFP expression, RT-PCR and western blots were used to detect GFP mRNA or protein. Three plants from each transgenic line were randomly selected, including the plants used for confocal microscopy above. Total DNA was isolated using a CTAB extraction from each sample BAR including the empty vector control line. Two samples of the CTP::eGFP

57 transgenic line, three samples of the homozygous eGFP transgenic line and the ELVd::eGFP transgenic line were randomly selected for PCR amplification. ELVd::eGFP expression vector, and empty PC-GW-BAR vector were positive controls. The forward primer: 5’-

GGATTGATGTGATATCTCCAC-3’, which paired with the PC-GW-BAR backbone, and the reverse primer: 5’-CCTTACTTGTACAGCTCGTCC-3’, which paired with the eGFP cDNA

(Figure 2.7A) were used to amplify the entire ELVd::eGFP DNA fragment. The size of the amplified ELVd::eGFP fragment was 1182 bp. Although this first pair of primers were not specific enough that all templates showed a band, faint or bright, it clearly amplified a sequence of about 1200 bp from two out of the three samples of the ELVd::eGFP transgenic lines. The gel-purified PCR products of these two samples were sent for sequencing. ELVd::eGFP 2 showed a continuous sequence of around 400 bp, while ELVd::eGFP 3 showed a continuous sequence of around 600 bp that corresponded to the full-length eGFP sequence. The forward primer: CAAATACAAATACATACTAAGGGTTTC and the reverse primer:

CAAGACCTTCCTCTATATAAGG, which paired with BAR DNA (Figure 2.7B) were used to amplify the BAR sequence. The size of the BAR sequence was 765 bp. This pair of primers amplified a sequence of around 750 bp in each sample, except the three samples of the homozygous eGFP line, which were negative control.

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Figure 2.7 PCR amplification gel electrophoresis. (A) Primers amplified the entire chimeric cDNA of ELVd::eGFP. Two out of the three samples of the ELVd::eGFP transgenic lines amplified a clear band; (B) Primers amplified BAR. Only the three negative control samples of the homozygous eGFP line did not amplify any bands.

As demonstrated by PCR, two out of the three samples of the ELVd::eGFP transgenic

line carried the targeted gene and were selected for RT-PCR. Total RNA was isolated from

leaf tissues of these two plants and reverse transcribed into cDNA. Then a PCR was performed

with two pairs of primers. First was the forward primer: 5’-GTTGGCGAAACCCCATTTC-

3’, which paired with ELVd, combined with the reverse primer: 5’-

CACCATGGGAACAAATCCTGG-3’, which paired with eGFP. This pair of primers

amplified the entire ELVd::eGFP construct of 1057 bp. Second was the forward primer: 5’-

CCTTACTTGTACAGCTCGTCC-3’, which pairs with eGFP, combined with the reverse

primer: 5’-CACCATGGGAACAAATCCTGG-3’, which paired with eGFP. This pair of

primers amplified eGFP (720 bp). Both pairs of primers amplified a clear band for each sample

(Figure 2.8). The gel-purified PCR products of the entire ELVd::eGFP chimeric construct were

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sequenced and each sample showed a continuous sequence of around 180 bp corresponding to

the full-length eGFP.

Figure 2.8 RT-PCR for two ELVd::eGFP transgenic lines. The two sample of the ELVd::eGFP transgenic line from Figure 2.7 amplified both the entire ELVd::eGFP construct (left) and the eGFP sequence (right). To further verify eGFP expression, a western blot was done with leaf tissues collected

from the same nine transgenic plants used for DNA amplification. The three homozygous

eGFP transgenic samples showed a clear single band with a size of around 33 kDa (Figure 2.9),

while the other samples did not show bands under this exposure. An overexposure of the same

blot showed that the band pattern of the samples of the ELVd::eGFP transgenic plants were

the same as the band patterns of the empty PC-GW- BAR transgenic plants. The band pattern

of homozygous eGFP transgenic samples differed from the band pattern of both of the samples

of the empty PC-GW-BAR vector transgenic plants and the ELVd::eGFP transgenic plants.

However, there was no band at a size of around 33 kDa in the samples of CTP::eGFP transgenic

line (data not shown).

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Figure 2. 9 Western blot for full-length eGFP. The three homozygous eGFP transgenic samples showed a clear single band with a size of around 33 kDa, while the other samples did not show bands under this exposure.

Discussion

ELVd is the only species of the monotypic genus Elaviroid of Avsunviroidae family.

ELVd is diverse in that the nine variants fall into four groups based on sequence similarities

(Daròs, 2016). ELVd is a single-stranded circular noncoding RNA that folds into a quasi-rod-

like secondary structure (Daròs, 2016). As a single-stranded RNA, the strands of

complementary polarity both accumulate in the host cell during replication. Therefore, the

strand that accumulates at a higher concentration is assigned arbitrarily as the plus (+) polarity

(Daròs, 2016; Fadda et al., 2003), while the complementary strand is assigned as the minus (-

) polarity. Strands of both polarities of ELVd contain conserved nucleotides in double-stranded

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elements that form two hammerhead ribozymes that induce self-cleavage (Fadda et al., 2003).

According to the direction of replication, the upstream hammerhead ribozyme is assigned as

plus (+) polarity, and the downstream hammerhead ribozyme is assigned as minus (-) polarity.

A classic ELVd (AJ536613) sequence folds into its secondary structure (Daròs, 2016) (Figure

2.10). Based on the replication mechanism in other members of the family Avsunviroidae, the replication process of ELVd can be described in three steps. First, ELVd self-cleaves into linear monomers. Second, both of the linear monomers get transcribed by a nuclear-encoded RNA polymerase (NEP). Finally, the two linear monomeric sequences get circularized by a host

RNA ligase (B. Navarro et al., 2012; Flores et al., 2000; J. Navarro et al., 2000).

Figure 2.10 The secondary structure of ELVd (AJ536613) predicted by Mfold (Daròs, 2016). Yellow indicates the domain of + hammerhead ribozyme and orange indicates the domain of – hammerhead ribozymes. Arrows indicate the ribozyme cleavage sites. The ELVd sequence provided by Goméz and Pállas (2010b) was constructed with a fragment of the minus strand of ELVd (AJ536613, position 54–267) followed by a fragment of the plus strand of ELVd (AJ536613, position 54–261). This construct excluded any of the ribozymes. Only the fragment of the + polarity between the positions 52-150 was essential for translocation (Gómez & Pallás, 2012b). ELVd possesses a nuclear localization step before it

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translocates to the chloroplasts for replication. The fragment of the + polarity between the

positions 15-181 was responsible for the nuclear localization (Gómez & Pallás, 2012a, 2012b).

A full-length ELVd of + polarity (AJ536613, position 1–333) could also mediate the

translocation (Gómez & Pallás, 2012a, 2012b). The influence of the minus strand in the

construct ELVd is not clear, nor is the influence of its absence on self-cleavage. The ELVd sequence used in this project was derived from the minus strand of ELVd followed by a fragment of the plus strand. Therefore, it was possible that the sequence elements that carried out the RNA translocation into A. thaliana chloroplasts caused the failure of the eGFP expression.

The most likely RNA ligase in A. thaliana chloroplasts is the tRNA ligase (Martínez et al., 2009). Although the A. thaliana tRNA ligase circularized monomeric RNAs, these RNAs opened at different positions relative to the self-cleavage sites (Englert et al., 2007; Nohales et al., 2012). Therefore, the stroma environment in A. thaliana chloroplasts could cause some other self-cleavages of the chimeric ELVd::eGFP transcript or distributed the function of

ELVd as 5’-UTR, and caused the failure of the eGFP expression.

The same GFP pattern was observed in N. benthamiana (Gómez & Pallás, 2010b)

(Figure 2.3). This result was a strong encouragement to apply this strategy to other species to

translocate RNAs into chloroplasts under the mediation of ELVd. However, in our experiments

an eGFP signal was not detected in A. thaliana. It is possible that ELVd cannot mediate the

translocation of RNA in A. thaliana, or it is also possible that the RNA chimera of ELVd and

eGFP did get imported into chloroplasts, but the yield of translation products was too low to

be detected in our tests. The western blot, failed to detect eGFP protein from the CTP::eGFP

transgenic plants, which showed a strong eGFP signal in confocal microscopy. Possibly the

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failure of the western blot could be due to the SuperSignal™ West Pico PLUS

Chemiluminescent Substrate used in this project to detect eGFP protein might not be sensitive enough.. A potential improvement for the western blot is to use SuperSignal™ West Femto

Maximum Sensitivity Substrate instead. Besides, the RNA to cDNA EcoDry™ Premix

(Random Hexamers) (Clonetech, Mountain View, CA) kit was used to reverse transcribe RNA

into cDNA, which can reverse transcribe all RNAs with or without poly-A tails. Considering that a poly-A tail is essential for translation, it is possible that these transcripts did not have poly-A tails and therefore failed to be translated. A potential method to refute this possibility is to use the RNA to cDNA EcoDry™ Premix (Oligo dT) (Clonetech, Mountain View, CA) kit, which can only reverse transcribe RNAs with poly-A tails.

In all the three possible situations, a chloroplast isolation combined with RT-PCR or an in situ hybridization could be sufficient to verify each of the possibilities.

In order to capture distinct contrast of the autofluorescence and the eGFP signal in N. benthamiana leaves, the pixel dwell was set at 3.15 microns for ZEISS LSM 710. Pixel dwell can be understood as exposure time. However, the brightness of the images was too low to be clearly visualized. Therefore, the brightness, as well as the contrast, of these N. benthamiana confocal images were adjusted to 400%. The confocal microscopy performed with the ZEISS

LSM 880 showed strong autofluorescence that was almost as strong as the eGFP fluorescence.

One of the reasons was that the image editing software, ImageJ, caused a color anamorphosis when all the parameters from the ZEISS LSM settings were maintained. Another reason is that the excitation wavelength of eGFP fluorescence, 488 nm, can also cause high autofluorescence

(Kodama, 2016). It is worth noting that the excitation wavelength region of CFP (cyan

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fluorescence protein) fluorescence is 380-480 nm, which would be a better choice as a fluorescence emitter (Song, 2007).

Another interesting phenomenon we observed was that none of the transgenic A. thaliana plants were able to survive through the late stage of flowering except the homozygous eGFP transgenic plants. All these plants died eventually and even when they set some siliques, these siliques were seedless. None of the transgenic A. thaliana plants of the eIF4E::eGFP line survived during the BASTA screening. Both of these phenomena suggested a possibility that the accumulation of the chimeric transcripts or the transgenesis itself might cause lethality. To solve this problem, other promoters and terminators besides the CaMV 35S promoter and the

CaMV 35S terminator can be used to drive an inducible- or tissue- specific expression to prevent the death caused by the constitutive promoter.

There are many unresolved questions remaining. Failing to detect the expression of eGFP does not mean that this strategy does not work because there are many possibilities that still need to be tested. Even if this strategy does not work in A. thaliana, it worth testing in other species such as in Solanum melongena (eggplant), which is a natural host of ELVd, which therefore contains all the components required by ELVd. Eggplant is an important commercial crop itself.

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APPENDICES

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Appendix A: Recipes and Protocols

1. Antibiotics and herbicide

Component Stock concentration (mg/mL) Working concentration (μg/mL) Kanamycin 50 50 Carbenicillin 100 100 Rifampicin 25 15 Gentamicin 100 50 BASTA 10 50

2. CTAB DNA extraction buffer

Component Amount Note 0.5 M Ethylenediaminetetraacetic acid (EDTA) 4.4 mL pH=8 Cetyl trimethylammonium bromide (CTAB) 0.8 g N-Laurosylsacosine (Sarkosyl) 1 g 1 M Tris base 22 mL pH=8 D Sorbitol 2.55 g NaCl 4.68 g Deionized water (diH2O) Bring to 100 mL

3. TE buffer

Component Amount Note 1 M Tris base 1 mL pH=8 0.5 M EDTA 0.2 mL pH=8 diH2O Bring to 100 mL

4. Buffer E

Component Amount Note Tris-HCl 3 g Sodium dodecyl sulfate (SDS) 2 g Glycerol 20 mL Sodium metabisulfite 19 g diH2O Bring to 200 mL pH=8.8

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5. Buffer Z

Component Amount Note Tris-HCl 3 g Sodium dodecyl sulfate (SDS) 24 g 2-Mercaptoethanol 44 mL Bromophenol blue 0.002 g diH2O Bring to 200 mL pH=6.8

6. Lysogeny broth (LB) liquid medium

For E. coli LB liquid medium, 12.5 gram (g) of LB broth (Fisher Bioreagents, Waltham,

MA) was dissolved in 500 mL distilled water and autoclaved for 20 minutes (min). Media were stored at room temperature.

For agrobacterium LB liquid medium, 5 g of LB broth (Fisher Bioreagents, Waltham,

MA) was dissolved in 200 mL distilled water and autoclaved for 20 min. Agrobacterium LB liquid media were always made freshly. Media were stored at room temperature.

7. LB-agar plates

For E. coli LB plates, 12.5 g of LB broth (Fisher Bioreagents, Waltham, MA) was dissolved in 500 mL distilled water mixed with 7.5 g agar (Sigma-Aldrich Corporation,

Raleigh, NC) and autoclaved for 20 min. Plates were stored in refrigerator at 4 ℃.

For agrobacterium LB plates, 5 g of LB broth (Fisher Bioreagents, Waltham, MA) was dissolved in 200 mL distilled water mixed with 3 g agar (Sigma-Aldrich Corporation, Raleigh,

NC) and autoclaved for 20 min. Agrobacterium LB plates were always made freshly.

8. Plasmids screening and extraction

Plasmids were transformed into E. coli and selected on LB-agar plates with proper antibiotics, 100 μg/mL carbenicillin for pUC 19 and pUC 57, 50 μg/mL kanamycin for PC-

GWs. Colonies were picked and incubated in LB liquid media with proper antibiotics, 100

μg/mL carbenicillin for pUC 19 and pUC 57, 50 μg/mL kanamycin for PC-GWs at 37 ℃

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overnight. Plasmids were extracted using QIAprep® Spin Miniprep Kit (Qiagen, Germantown,

MD). After purification, plasmids of pUC 19, PC-GW-BAR, PC-GW-mCherry with noncoding RNA were sent for sequencing (Genewiz, Morrisville, North Carolina; Eurofins

Genomics, Louisville, Kentucky) to confirm DNA segments; while

9. Floral dip for A. thaliana

After transformation, agrobacteria were plated on LB plates with kanamycin, carbenicillin, rifampicin and incubated at 28 ℃ for at least two days till single colonies grew up. Single colonies were transferred into 50 mL liquid LB media with kanamycin, carbenicillin, rifampicin and incubated at 28 ℃ for 12 to 24 hr till OD600 reached 0.8.

Agrobacteria were harvested by centrifuging at 5,000 rpm for 5 min and then suspended in 5%

sucrose. Right before dipping, Silwet L-77 were added to a final concentration of 0.03%. Each

plant was dipped for exact 30 sec and then covered with black plastic bag to block light and

keep moisture for 24 hr. A second dip was applied seven days after the first dip and blocked

from light for 24 hr. Plants were washed thoroughly after the second dip.

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Appendix B: DNA Segments Sequences and Primers

1. Primers Name Sequence Tm/℃ Binding position Sq_M_F GCTGACCCTGAAGTTCATC 60 on eGFP Sq_M_R GCTCAGGTAGTGGTTGTC 60 on eGFP Sq_ELVd_F GGATTGATGTGATATCTCCAC 60 on PC-GW-BAR Sq_ELVd_R GCACCCGACATAGATAATG 60 on PC-GW-BAR Sq_ELVd_1_F GTTGGCGAAACCCCATTTC 65 on ELVd Sq_ELVd_1_R CCTTACTTGTACAGCTCGTCC 65 on eGFP Sq_BAR_F CAAATACAAATACATACTAAGGGTTTC 59 on PC-GW-BAR Sq_BAR_R CAAGACCTTCCTCTATATAAGG 59 on PC-GW-BAR EGFP F ATGGTGAGCAAGGGCGAG 65 on eGFP EGFP R CTTGTACAGCTCGTCCATGC 65 on eGFP

2. ELVd::eGFP

AATTAATCTAGAGTTGGCGAAACCCCATTTCGACCTTTCGGTCTCATCAGGGGTGGCACACACCAC CCTATGGGGAGAGGTCGTCCTCTATCTCTCCTGGAAGGCCGGAGCAATCCAAAAGAGGTACACCC ACCCATGGGTCGGGACTTTAAATTCGGAGGATTCGTCCTTTAAACGTTCCTCCAAGAGTCCCTTCC CCAAACCCTTACTTTGTAAGTGTGGTTCGGCGAATGTACCGTTTCGTCCTTTCGGACTCATCAGGG AAAGTACACACTTTCCGACGGTGGGTTCGTCGACACCTCTCCCCCTCCCAGGTACTATCCCCTTTC CAGGATTTGTTCCCATGGTGAGCAAGGGCGAGGAGCTGTTCACCGGGGTGGTGCCCATCCTGGTC GAGCTGGACGGCGACGTAAACGGCCACAAGTTCAGCGTGTCCGGCGAGGGCGAGGGCGATGCCA CCTACGGCAAGCTGACCCTGAAGTTCATCTGCACCACCGGCAAGCTGCCCGTGCCCTGGCCCACCC TCGTGACCACCCTGACCTACGGCGTGCAGTGCTTCAGCCGCTACCCCGACCACATGAAGCAGCAC GACTTCTTCAAGTCCGCCATGCCCGAAGGCTACGTCCAGGAGCGCACCATCTTCTTCAAGGACGAC GGCAACTACAAGACCCGCGCCGAGGTGAAGTTCGAGGGCGACACCCTGGTGAACCGCATCGAGCT GAAGGGCATCGACTTCAAGGAGGACGGCAACATCCTGGGGCACAAGCTGGAGTACAACTACAAC AGCCACAACGTCTATATCATGGCCGACAAGCAGAAGAACGGCATCAAGGTGAACTTCAAGATCCG CCACAACATCGAGGACGGCAGCGTGCAGCTCGCCGACCACTACCAGCAGAACACCCCCATCGGCG ACGGCCCCGTGCTGCTGCCCGACAACCACTACCTGAGCACCCAGTCCGCCCTGAGCAAAGACCCC AACGAGAAGCGCGATCACATGGTCCTGCTGGAGTTCGTGACCGCCGCCGGGATCACTCTCGGCAT GGACGAGCTGTACAAGTAAGGCGCGCCGAGCTCCCTGCAGGGGATCCAATTAA

ELVd eGFP Restriction digestion sites