The Role of Gibberellic Acid in Aphid-Plant-Arbuscular Mycorrhizal Fungus Interactions

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Master's Theses Student Research

5-8-2020

THE ROLE OF GIBBERELLIC ACID IN APHID-PLANT-ARBUSCULAR MYCORRHIZAL FUNGUS INTERACTIONS

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Recommended Citation Olson, Drew, "THE ROLE OF GIBBERELLIC ACID IN APHID-PLANT-ARBUSCULAR MYCORRHIZAL FUNGUS INTERACTIONS" (2020). Master's Theses. 164. https://digscholarship.unco.edu/theses/164

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UNIVERSITY OF NORTHERN COLORADO

Greeley, Colorado

The Graduate School

THE ROLE OF GIBBERELLIC ACID IN APHID- PLANT-ARBUSCULAR MYCORRHIZAL FUNGUS INTERACTIONS

A Thesis Submitted in Partial Fulfillment of the Requirements for the Degree of Master of Science

Drew Olson

College of Natural and Health Sciences School of Biological Sciences

May 2020

This Thesis by Drew Olson

Entitled: The role of gibberellic acid in aphid-plant-arbuscular mycorrhizal fungus interactions has been approved as meeting the requirement for the Degree of Master of Science in College of Natural and Health Sciences in School of Biological Sciences, Program of Biological Sciences

Accepted by the Thesis Committee:

Susana Karen Gomez, Ph.D., Chair

Mitchell McGlaughlin, Ph.D., Committee Member

Hua Zhao, Ph.D., Committee Member

Accepted by the Graduate School

Cindy Wesley, Ph.D. Interim Associate Provost and Dean Graduate School and International Admissions

ABSTRACT

Olson, Drew. The role of gibberellic acid in aphid-plant-arbuscular mycorrhizal fungus interactions. Unpublished Master of Science thesis, University of Northern Colorado, 2020.

Arbuscular mycorrhizal (AM) fungi form beneficial associations with the roots of most terrestrial plants, which is mostly characterized by increased plant nutrient acquisition (phosphate and nitrogen) by the fungi in exchange for carbon and lipids from the plants. These associations can also enhance plant coping mechanisms to abiotic and biotic stresses. However, insect herbivores, such as aphids, can benefit from AM fungi- plant symbioses by altering plant defenses that are driven mainly by the phytohormones jasmonic acid (JA) and salicylic acid (SA). Nevertheless, other phytohormones can modulate plant defenses by altering the balance of the JA and SA signaling pathways, such as the growth promoting phytohormone gibberellic acid (GA). Although GA signaling plays an important role in modulating plant defenses during plant-pathogen interactions, its role in insect-plant and insect-plant-beneficial microbe interactions remains largely unknown. Therefore, the current study set out to investigate the following two objectives: 1) examine the role of exogenous GA application in regulating plant defenses during aphid-plant-AM fungus interactions, and 2) evaluate the role of GA signaling in modulating plant defenses during aphid-plant interactions. In objective 1, we found that exogenous GA negatively affected aphid fitness on GA-treated mycorrhizal plants 7-days post pea aphid feeding. Exogenous GA had a synergistic effect on JA- related plant gene expression in shoots, and increased JA and SA levels in leaves and

iii petioles after 36 hours of aphid feeding. Pea aphid feeding for 36 hours induced SA- related defense gene expression and increased SA levels in leaves and petioles. In addition, root colonization by the AM fungus resulted in induced JA- and SA-related plant defense gene expression in shoots and roots, but decreased SA levels in leaves and petioles after 36 hours of aphid feeding. In objective 2, we found that GA signaling had an antagonistic effect on JA-related defense signaling after 36 hours of aphid feeding. In summary, this study serves as a foundation for future studies examining the role of GA in regulating plant defenses during insect-plant and insect-plant-AM fungus interactions.

ACKNOWLEDGEMENTS

I first and foremost want to thank my advisor, Dr. Karen Gomez for her amazing professional and personal support and commitment to graduate student success. Karen may not be the easiest on her students, but she expects the best from us, therefore bringing out the most of us. I have grown immensely over the past two years and could not have become who I am today without Karen pushing me to learn and grow. Although there were many tough times and long weeks, she was always there to lend out a helping hand. I have faced many challenges during my time at the University of Northern

Colorado, however I was able to persevere through these because of Karen’s guidance and support. With that being said, thank you for everything!

I would like to thank my committee members, Dr. Mitchell McGlaughlin and Dr.

Hua Zhao for their patience, commitment, and guidance through this process. In addition,

I could not have succeeded without the help of Dr. Cris Argueso and her Ph.D. candidate,

Hannah Berry. Thank you, Cris for allowing me to use lab space and help fund the phytohormone analyses for my research. Thank you, Hannah Berry for taking the time to help me every step of the way through the phytohormone extraction and analyses. I cannot thank either of you two enough for being critical members of my success and I hope that I will be able to return the favor later down the road! I also could not have made it this far without the help of Dr. Jamie Riggs for helping me figure out which statistical analyses to use. I would have lost much more sleep over statistical analyses had she not provided me with her valuable assistance!

I want to thank one of my best friends, soon-to-be Dr. Tyler Sherman. Tyler has helped me through everything since the beginning of my time here at the University of

Northern Colorado. Without him, I would never have had the chance to travel home or elsewhere with my family. Moreover, he has provided me with support throughout the entirety of this process whether it be personal and emotional support or in the lab. I know that I would not be the person that I am proud to be today without sharing friendship with this amazing individual. I also want to thank Christine Williams for helping me through a dark time early on in my graduate school career as well as being a great friend. I am not sure where I would be at had you not allowed me to open up to you as my first friend.

You were the first person that I felt that I could trust after making the long and lonely journey here to Colorado. In addition, I want to thank my most recent of close friends

Christopher Gregory and Lewis Eaton. These two remind me that there is more to life than being a scientist and spending all of my time towards my graduate school success. I am not sure that I would have made it through the past 8 months or so without them pulling me out of the dens of the Gomez lab, the teaching assistant office, or my apartment. In summary, my friends make up more than a large part of my success, but more importantly my happiness during these strenuous times. I hope that I have in some way, shape, or form have provided you all something of personal value as you all have provided to me.

I want to thank my parents, Joel and Kelly Olson, my brother, Jaron Olson, and my grandparents, Don and Karen Olson, for their amazing support through thick and thin.

Even when I began to shut myself out to the world, you all made sure to reach out to me and remind me that all things work out in the end.

I want to thank the entire faculty, staff, graduate students, and undergraduate students at the University of Northern Colorado that have helped me succeed as well as provide an amazing place to grow and thrive. In, addition I want to thank the College of

Natural and Health Sciences and Graduate Student Association for their financial support.

I want to thank David Shimokawa for his undying commitment to graduate student success and making sure that students get the funding they need for research and for travel. I know that my time here would have been a much greater challenge without the funding. David also commits himself to graduate student happiness and health by organizing many events on and off campus to get people together and connected, and for that I am forever grateful. Finally, I want to thank The Plant Cell journal for providing me permission for the non-profit use of the figure provided in Fig. 1.4.

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TABLE OF CONTENTS

CHAPTER I: INTRODUCTION AND REVIEW OF LITERATURE ...... 1 Aims ...... 1 Plant Interactions with Soil Beneficial Microbes: Arbuscular Mycorrhizal Symbioses ...... 3 Establishment of the Arbuscular Mycorrhizal Symbiosis ...... 4 Plant Interactions with Phloem-Feeding Insects ...... 8 The Role of Phytohormones in Plant Biotic Interactions ...... 10 Jasmonic Acid Signaling and Plant Defenses ...... 10 Salicylic Acid Signaling and Plant Defenses ...... 13 Gibberellic Acid Biosynthesis, Metabolism, and Plant Defenses ...... 15 Gibberellic Acid Signaling and DELLA Proteins ...... 18 Roles of Gibberellic Acid in Plant-Microbe and Plant-Insect Interactions ...... 19 Significance...... 21 CHAPTER II: EFFECT OF EXOGENOUS GIBBERELLIC ACID IN REGULATING PLANT DEFENSE RESPONSES DURING PLANT INTERACTIONS WITH APHIDS AND ARBUSUCLAR MYCORRHIZAL FUNGI ...... 24 Abstract ...... 24 Introduction ...... 25 Methods and Materials ...... 30 Results ...... 39 Discussion ...... 61 CHAPTER III: ROLE OF GIBBERELLIC ACID SIGNALING IN MODULATING PLANT DEFENSE RESPONSES DURING PLANT-APHID INTERACTIONS ...... 70 Abstract ...... 70 Introduction ...... 71 Materials and Methods ...... 77

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Results ...... 83 Discussion ...... 92 CHAPTER IV: CONCLUSIONS AND FUTURE DIRECTIONS ...... 97 LITERATURE CITED ...... 106

LIST OF TABLES Table

2.1 Primers used for real-time quantitative PCR of plant and fungal target genes ...... 37 3.1 Primers used for real-time quantitative PCR of plant defense target genes...... 81

LIST OF FIGURES

Figure

1.1 Jasmonic acid structure…………………………………………………... 11 1.2 Salicylic acid structure…………………………………………………… 14 1.3 Gibberellic Acid 3 structure……………………………………………… 16 1.4 Jasmonic acid-gibberellic acid antagonism………………………………. 20 2.1 Impact of exogenous GA, pea aphid (Acyrthosiphon pisum) feeding, and AM fungus (Rhizophagus intraradices) root colonization on Medicago truncatula shoots at 36 hours post-aphid feeding………………………... 40

2.2 Impact of exogenous GA, pea aphid (Acyrthosiphon pisum) feeding, and AM fungus (Rhizophagus intraradices) root colonization on Medicago truncatula roots at 36 hours post-aphid feeding………………………….. 41 2.3 Impact of exogenous GA and AM fungus (Rhizophagus intraradices) root colonization on mean pea aphid (Acyrthosiphon pisum) count per colony (a) and mean colony weight (b) after 36 hours of aphid feeding on Medicago truncatula plants………………………………………………. 42 2.4 Impact of exogenous GA, pea aphid (Acyrthosiphon pisum) feeding, and AM fungus (Rhizophagus intraradices) root colonization on mean gibberellic acid 3 (GA3) concentration in leaf and petiole tissue…………. 43 2.5 Impact of exogenous GA, pea aphid (Acyrthosiphon pisum) feeding, and AM fungus (Rhizophagus intraradices) root colonization on mean jasmonic acid (JA) concentration in leaf and petiole tissue………………. 44 2.6 Impact of exogenous GA, pea aphid (Acyrthosiphon pisum) feeding, and AM fungus (Rhizophagus intraradices) root colonization on mean salicylic acid (SA) concentration in leaf and petiole tissue……………….. 45 2.7 Impact of exogenous GA, pea aphid (Acyrthosiphon pisum) feeding, and AM fungus (Rhizophagus intraradices) root colonization on MtMYC2 relative gene expression in Medicago truncatula shoots…………………. 46 2.8 Impact of exogenous GA, pea aphid (Acyrthosiphon pisum) feeding, and AM fungus (Rhizophagus intraradices) root colonization on MtVSP relative gene expression in Medicago truncatula shoots………………….. 48 2.9 Impact of exogenous GA, pea aphid (Acyrthosiphon pisum) feeding, and AM fungus (Rhizophagus intraradices) root colonization on MtPR1 relative gene expression in Medicago truncatula shoots………………….. 50 2.10 Impact of exogenous GA, pea aphid (Acyrthosiphon pisum) feeding, and AM fungus (Rhizophagus intraradices) root colonization on MtMYC2 relative gene expression in Medicago truncatula root tissue……………… 52

2.11 Impact of exogenous GA, pea aphid (Acyrthosiphon pisum) feeding, and AM fungus (Rhizophagus intraradices) root colonization on MtVSP relative gene expression in Medicago truncatula root tissue……………… 53 2.12 Impact of exogenous GA, pea aphid (Acyrthosiphon pisum) feeding, and AM fungus (Rhizophagus intraradices) root colonization on MtPR1 relative gene expression in Medicago truncatula root tissue…………….. 54 2.13 Impact of exogenous GA, pea aphid (Acyrthosiphon pisum) feeding, and AM fungus (Rhizophagus intraradices) root colonization on RiEF, MtLec5, and MtPT4 relative gene expression on mycorrhizal Medicago truncatula roots………………………………………………………….... 55 2.14 Impact of exogenous GA, pea aphid (Acyrthosiphon pisum) feeding, and AM fungus (Rhizophagus intraradices) root colonization on Medicago truncatula shoot fresh weight at 7 days post-aphid feeding………………. 57 2.15 Impact of exogenous GA, pea aphid (Acyrthosiphon pisum) feeding, and AM fungus (Rhizophagus intraradices) root colonization on Medicago truncatula root fresh weight for 7 days post-aphid feeding…...... 59 2.16 Impact of exogenous GA and AM fungus (Rhizophagus intraradices) root colonization on mean pea aphid (Acyrthosiphon pisum) count per colony and mean colony weights 7 days post-aphid feeding on Medicago truncatula plants…………………………………………………………... 61 3.1 Impact of pea aphid (Acyrthosiphon pisum) feeding for 36 hours on Medicago truncatula shoot fresh weight of wild type (WT) and Mtdella1/Mtdella2 plants………………………………………………… 84 3.2 Impact of pea aphid (Acyrthosiphon pisum) feeding for 36 hours on Medicago truncatula root fresh weight of wild type (WT) and Mtdella1/Mtdella2 plants………………………………………………… 85 3.3 Mean pea aphid (Acyrthosiphon pisum) count per colony (a), and mean pea aphid colony weight (b) after feeding for 36 hours on Medicago truncatula wild type (WT) and Mtdella1/Mtdella2 plants……………….. 86 3.4 MtMYC2 relative gene expression in Medicago truncatula shoots of wild type (WT) and Mtdella1/Mtdella2 plants after 36 hours of pea aphid (Acyrthosiphon pisum) feeding…………………………………………... 87 3.5 MtVSP relative gene expression in Medicago truncatula shoots of wild type (WT) and Mtdella1/Mtdella2 plants after 36 hours of pea aphid (Acyrthosiphon pisum) feeding…………………………………………... 88 3.6 MtPR1 relative gene expression in Medicago truncatula shoots of wild type (WT) and Mtdella1/Mtdella2 plants after 36 hours of pea aphid (Acyrthosiphon pisum) feeding…………………………………………... 89 3.7 Impact of pea aphid (Acyrthosiphon pisum) feeding for 7 days on Medicago truncatula shoot dry weight of wild type (WT) and Mtdella1/Mtdella2 plants………………………………………………… 90 3.8 Impact of pea aphid (Acyrthosiphon pisum) feeding for 7 days on Medicago truncatula root dry weight of wild type (WT) and Mtdella1/Mtdella2 plants………………………………………………… 91

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3.9 Mean pea aphid (Acyrthosiphon pisum) count per colony (a), and mean pea aphid colony weight (b) after feeding for 7 days on Medicago truncatula wild type (WT) and Mtdella1/Mtdella2 plants……………….. 92

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CHAPTER I

INTRODUCTION AND REVIEW OF LITERATURE

Aims

Arbuscular mycorrhizal (AM) fungi associate with plant roots and can indirectly interact with insect herbivores aboveground and belowground through alterations in plant defense responses. However, there are insects such as aphids that can evade typical plant defense responses against insect herbivory. Aphids are known to take advantage of AM fungi-plant associations by altering the balance between the phytohormones jasmonic acid (JA) and salicylic acid (SA). Nevertheless, the phytohormone gibberellic acid (GA) has gained attention in recent studies that focus on plant-pathogen interactions by altering the activity of the JA and SA pathways through interactions with DELLA proteins. GA is well known to play important roles in plant growth and development. GA levels are regulated by DELLA proteins, which are transcriptional repressors. Gibberellins, however, interact with and cause the degradation of DELLA proteins, which can affect other plant signaling pathways. To date, little is known about how GA affects defense responses in plant-aphid and AM fungi-plant-aphid interactions. In the present study, two biological systems were used to further investigate the role of GA in regulating plant defenses. The first system involved pea aphids (Acyrthosiphon pisum), barrel medic plants (Medicago truncatula), and the AM fungus Rhizophagus intraradices. The second system involved pea aphids and two plant genotypes (segregant wild type (WT) and

Mtdella1/Mtdella2 mutant). These two systems were used to evaluate the following objectives and hypotheses:

Objective 1 Examine the role of exogenous GA application in regulating plant defenses during AM fungus-plant-aphid interactions.

H1 Aphid fitness will be positively impacted by exogenous GA application of plants.

H2 Exogenous GA application will result in reduced arbuscule formation.

H3 JA levels and JA-regulated defense gene expression will be downregulated in response to exogenous GA application.

H4 SA levels and SA-regulated defense gene expression will be upregulated in response to exogenous GA application.

Objective 2 Evaluate the role of GA signaling in modulating plant defenses during plant-aphid interactions.

H1 Aphid fitness will be improved after feeding on Mtdella1/Mtdella2 mutants compared to wild type plants.

H2 JA biosynthesis genes and JA-regulated defense genes will be downregulated in Mtdella1/Mtdella2 mutant.

To assess objective one, a three-way interaction experiment was designed composed of eight treatments: 1) -AM fungus/ -Pea aphid / -GA ; 2) -AM fungus/ +Pea aphid / -GA; 3) -AM fungus/ -Pea aphid/ +GA ; 4) -AM fungus/ +Pea aphid/ +GA; 5)

+AM fungus / -Pea aphid/ -GA; 6) +AM fungus/ +Pea aphid/ -GA; 7) +AM fungus/ -Pea aphid/ +GA; and 8) +AM fungus/ +Pea aphid/ +GA. Pea aphids were added only after plants receiving AM fungal inoculum reached at least 50% root length colonization.

Based on previous studies, a 36-hour post aphid feeding time-point was used to analyze changes in plant defense gene expression in shoots (four genes) and roots (six genes) and phytohormone concentration (three phytohormones) in plant shoots following aphid

3 feeding. A 7-day post aphid feeding time-point was used to assess aphid fitness that was measured as abundance and colony weight. To assess objective two, a two-way interaction experiment was designed including four treatments: 1) Segregant wild type

(WT) / -Pea aphid; 2) WT / +Pea aphid; 3) Mtdella1/Mtdella2 mutant / -Pea aphid; and 4)

Mtdella1/Mtdella2/ +Pea aphid. The same two time-points used in objective one were also used to assess objective two. By assessing these two objectives, we hope to better understand the role of GA in modulating plant defenses during aphid-plant and aphid- plant-AM fungus interactions.

Plant Interactions with Soil Beneficial Microbes: Arbuscular Mycorrhizal Symbioses

Plants are primary producers and sessile organisms that are under constant threat from a myriad of heterotrophic organisms such as bacteria, fungi, oomycetes, and insect herbivores above- and below-ground. In response to these threats, plants have developed intricate immune systems that can quickly detect conserved molecules associated with pathogens and insects (2-5). For instance, plants can recognize conserved microbial molecular structures known as pathogen- or microbe-associated molecular patterns

(PAMPs or MAMPs), such as bacterial flagella, chitin, or β-glucans, via cell-surface pattern recognition receptors (PRRs) (5). Perception of these molecular patterns subsequently elicits the plant innate immune system known as PAMP-triggered immunity

(PTI) (5-7). Insect pests can also trigger plant innate immunity through conserved herbivore-associated molecular patterns (HAMPs), such as insect saliva and oviposition chemicals, as well as through mechanical damage known as damage-associated molecular patterns (DAMPs) (6). Nevertheless, microbes and insects can release

4 molecules or proteins that interfere with PRR recognition and PTI signaling, known as effectors, to increase host susceptibility (5). Consequently, plants can maintain resistance to pathogens and insects through cytosolic resistance (R) proteins (5, 8). R proteins are nucleotide-binding leucine rich repeat (NB-LRR) receptor proteins that sense specific effectors and enhance host resistance by activating a robust hypersensitive immune response known as effector-triggered immunity (ETI) (5, 8). Although effectors are important components of host susceptibility by dampening plant innate immune responses, they may also be important for plant recognition of beneficial microbes, such as arbuscular mycorrhizal (AM) symbioses (9).

Establishment of the Arbuscular Mycorrhizal Symbiosis

The establishment of arbuscular mycorrhizal (AM) symbioses are facilitated through signals exchanged between fungi and plants (9, 10). AM fungi are obligate biotrophic soil microbes from the phylum Glomeromycota that form symbioses with

~80% of terrestrial plants (11). AM fungi-plant interactions date back ~450 million years and there is surmounting evidence suggesting that AM fungi aided in early land plant colonization during the Ordovician period and that plant algal ancestors already possessed the genetic toolkit for symbioses with AM fungi (12-16). Traditionally, this association is mostly known for acquisition of limiting soil nutrients (predominantly phosphorus) via the fungus, in exchange for photosynthates and lipids provided by the plant (10, 17-20). Phosphorus is an important plant nutrient for many plant functions that can be associated with increased plant biomass as well as crop yield (21-25), however it is only obtainable as inorganic phosphates that are largely unavailable due to immobility

5 within soil substrate (26). Although modern agricultural and horticultural practices still heavily rely on chemical fertilizers, there is evidence pointing towards AM fungi as potential bio-fertilizers for future sustainable practices (19, 27, 28). For instance, greenhouse and field experiments have demonstrated the benefits of AM fungal-mediated plant uptake of phosphate, accumulation in plant shoots, and subsequent increase in crop yields (29-33). Additionally, AM fungi are known to play important roles in water absorption, increased photosynthesis, plant-to-plant communication, tolerance to abiotic stress, and enhanced disease resistance (19, 34, 35).

AM fungi-plant interactions are characterized by the formation of fungal tree-like structures within plant root cortical cells known as arbuscules, which are also the sites of nutrient exchange (10). In addition, AM fungi extend their hyphae within plant roots

(intraradical) and into the immediate rhizosphere (extraradical) for nutrient transport, formation of new arbuscules, and increased surface area in the rhizosphere to obtain more soil nutrients (10). Initial contact between the symbionts begins prior to physical contact through chemicals released within the rhizosphere, which activates the plant symbiosis signaling pathway (SSP). This cross-kingdom communication is initiated when plant roots release strigolactones (SLs), plant hormones, into the rhizosphere, which are subsequently sensed by AM fungal spores (36, 37). Upon SL perception, AM fungi will undergo rapid transcriptional and metabolomic changes allowing for germination and hyphal growth toward potential plant hosts (36, 38). Additionally, the role of SLs in early plant communication belowground has been further demonstrated by parasitic plants. For example, a fava bean (Vicia faba) plant parasite, Orobanche crenata, requires belowground SL for seed germination (37).

As AM fungi reach plant roots, they release chitooligosaccharides or lipochitoologosaccharides known as ‘Myc’ factors (39), which are sensed by plant cell- surface lysine motif (LysM) receptor-like kinases and co-stimulated by other receptor- like kinases to initiate internal signaling and transcription (9). Plant perception of ‘Myc’ factors prepares root epidermal cells for physical contact with AM fungi. Chemical and physical signals sensed by plants triggers extensive cytoskeletal remodeling and organelle repositioning to form prepenetration apparatuses, which allows for AM fungal hyphopodia formation and subsequent hyphal growth into plant epidermal cells (40).

Upon hyphal entry, fungal colonization proceed intra- and inter-cellularly to eventually reach the root cortex where the plant-derived peri-arbuscular membrane forms within root cortical cells, which defines the formation of arbuscule development (41, 42). To achieve this extensive morphological change, plants must regulate appropriate signaling and transcriptional responses (9). For instance, hyphal entry and growth into root cortical cells trigger the generation of nuclear-associated Ca2+ oscillations by activating K+ and nucleotide-gated channels on the nuclear membrane (9). Ca2+ oscillations within nuclei are decoded by calcium and calmodulin-dependent kinases to recruit and phosphorylate another nuclear protein known as CYCLOPS (9). Phosphorylated CYCLOPS proteins recruit yet another class of proteins known as DELLA proteins. DELLAs can act as symbiosis transcription factors to upregulate gene expression of symbiosis genes that play critical roles in arbuscule development, branching, and maintenance (9, 43-46).

Apart from avoiding plant defenses, AM fungi can enhance plant resistance to other heterotrophic organisms above- and belowground through a phenomenon known as priming (6, 7, 34, 47, 48). Plant priming can be described as increased preparedness to

7 abiotic and biotic stress due to previous damage or infection. There are two main priming mechanisms, being systemic acquired resistance (SAR) and induced systemic acquired resistance (ISR). SAR is stimulated through pathogen infection, which confers resistance against a broad spectrum of pathogens in non-exposed distal tissues of the plant through long-distance signaling and the accumulation of PATHOGENESIS RELATED (PR) proteins (49). ISR, on the other hand, induces stronger and more effective defenses locally and systemically against necrotrophic pathogens and insect herbivores upon infection or infestation through production of direct and indirect defenses, such as secondary metabolites and physical barriers, and can be enhanced by beneficial soil microbes (6, 34, 47).

Specifically, AM fungi may confer resistance to above- and below-ground insect herbivores through a mechanism known as mycorrhiza-induced resistance (MIR) (34,

48). For instance, tomato (Solanum persicum) and potato (Solanum tuberosum) plants inoculated with AM fungi exhibited increased anti-herbivore related biosynthesis and defense gene expression, such as proteinase inhibitor-I and proteinase inhibitor-II, in response to chewing insects (50, 51). MIR may not always be limited to individual plants and can induce defenses to nearby plants upon insect herbivory. For instance, tomato and

Nicotiana attenuata plants colonized with AM fungi can prime nearby plants in response to insect herbivory by sending antiherbivore signals through belowground common mycorrhizal networks (CMNs) (52, 53). AM fungi may provide other direct and indirect defenses against insect herbivores through changes in leaf composition or changes in volatile profiles that may recruit natural enemies of insect herbivores (54-56). A meta- analysis revealed that AM fungi can negatively impact generalist chewing insects,

8 meanwhile phloem-feeding and specialist chewer insects are generally positively impacted (57). For instance, grain aphids (Sitobion avenae) can hijack nitrogen from barley (Hordeum vulgare L.) that is provided directly from AM fungi (58). Additionally, it has been demonstrated that pea aphids (A. pisum) prefer plants that are colonized with

AM fungi (59, 60), that aphid feeding can cause changes in plant metabolism belowground (61), and can negatively affect AM fungal colonization (54).

Plant Interactions with Phloem- Feeding Insects

Aphids are phloem-feeders of the superfamily Aphidoidea and make up some of the most important agricultural crop pests in temperate regions across the globe (4, 62-

65). Aphids use their needle-like mouthparts, known as stylets, to feed on plant sieve elements of the phloem by moving intracellularly, which also causes minimal mechanical damage to plant hosts. Aphids can devastate crops quickly through their life history traits, act as pathogen and viral vectors, and overcome plant defenses (4, 63-66). For instance, aphids have short generations, and can undergo an asexual, parthenogenetic mode of viviparous reproduction that enables populations to reach high densities quickly (64).

Upon high population densities and declination of host nutritional quality, aphids can produce winged morphs, which allow them to spread to new hosts and establish new colonies (64). Aphids can feed on a variety of plant hosts; some are specialists that feed on only a select group or family of plant hosts, while others can feed on a broad range of hosts from multiple plant families (63). Among the 275 known insect-born plant viruses, aphids have been shown to transmit about half of those viruses including some of the most agriculturally important plant viruses (62). Furthermore, aphids can support growth

9 of saprophytic fungi by exuding honeydew (a sugar-rich liquid substance), which can further interfere with photosynthesis (62).

Plants can typically defend against aphids through antixenosis (feeding behavior) or antibiosis (detrimental chemical defenses) (65). Antixenosis defenses typically affect aphid behavior through changes in plant morphology, release of volatile organic chemicals (VOCs), or accumulation of phloem proteins or callose (65). For example, surface waxes and trichome presence can act as insect barriers thereby reducing aphid settling and increasing aphid mortality. In addition plants can release VOCs, such as (E)-

β-farnesene and methyl salicylate, that can trigger changes in aphid behavior by acting as deterrents or by promoting dispersal behavior (67-69). In response to aphid feeding, plants can maintain resistance by phloem protein plugging or callose deposition at sites of aphid penetration (65). Comparatively, direct defenses are centered around secondary metabolites or proteinase inhibitors that influence aphid mortality, fecundity, or growth

(65). For example, many plants produce steroidal compounds known as cardenolides, which are well-established inhibitors of animal K+/Na+-ATPase (70). Other secondary metabolites include alkaloids, glucosinolates, and benzoxazinoids, which all maintain biochemical inhibitory activities in insect herbivores (70). However, aphids can avoid these defenses by sequestering and exuding these toxins or avoid toxin activation by minimizing mechanical damage (70). In addition, aphids may employ effectors to avoid typical anti-herbivore defense responses. For example, aphid saliva contains and releases several proteins with biochemical functions, including pectinases, cellulases, polyphenoloxidases, peroxidases, and lipases as well as several other proteins that may influence plant susceptibility to aphid feeding (4, 65). Moreover, aphid feeding is well

10 known to influence plant defenses through modulating the strength of the JA- and SA phytohormone-mediated signaling pathways (4, 70, 71).

The Role of Phytohormones in Plant Biotic Interactions

Phytohormones, are naturally occurring regulatory molecules that cause physiological changes in low concentrations (72). It is well-established that the JA- and

SA signaling pathways are necessary to induce defenses against a wide array of plant pathogens and pests (73, 74). SA is generally associated with defenses against biotrophic pathogens through the production of PR proteins, meanwhile JA is generally associated with defenses against necrotrophic pathogens and pests through production of plant defensins, proteinase inhibitors, and secondary metabolites (73, 74). Other phytohormones, such as such as auxin, gibberellic acid (GA), cytokinin (CK), abscisic acid (ABA), ethylene (ET), and brassinosteroid (BR) can influence plant defenses by modulating the strength of the JA- and SA signaling pathways to orchestrate appropriate defenses and to prioritize between growth and defense (73, 74). For instance, ET and

ABA can enhance activation of JA signaling through two different branches; the JA-ERF

(ETHYLENE RESPONSE FACTOR) and the JA-MYC branches in response to necrotrophic pathogens or pests (75, 76). Meanwhile, other phytohormones can modulate defenses by acting antagonistically with JA signaling (73, 77).

Jasmonic Acid Signaling and Plant Defenses

JA is an oxylipin derived phytohormone (Fig. 1.1) that is not only involved in plant growth and development but is critical for adaptation to abiotic stress and defenses against necrotrophic pathogens and insect pests (78, 79). Anti-herbivory JA signaling is

11 mediated by the MYC branch, which can lead to upregulation of defense genes involved in direct and indirect defenses (76, 79). Under low stress conditions, early JA-regulated gene expression is inhibited by JASMONATE ZIM-domain (JAZ) proteins. JAZs act as

JA transcriptional repressors by binding with MYC2, basic helix-loop-helix leucine zipper transcription factors (TFs) (80). MYC2 TFs are crucial for JA signaling and JA- regulated gene expression. JAZs are also known to recruit corepressors, such as

TOPLESS (TPL) and NOVEL INTERACTOR OF JAZ (NINJA) to further prevent JA- signaling (80).

Figure 1.1: Jasmonic acid structure. Retrieved from: https://pubchem.ncbi.nlm.nih.gov/compound/Jasmonic_acid#section=Structures

Upon elicitation, such as aphid stylet penetration, bioactive JA is rapidly synthesized from α-linolenic acid by JASMONATE RESISTANT1 (79). In response to wounding regardless of feeding type, rapid JA biosynthesis is likely propagated by cytosolic Ca2+ influxes generated by 3 GLUTAMATE RECEPTOR-LIKE proteins (79).

α-LeA is released from chloroplast membranes, which is converted into cis-(+)12-oxo- phytodienoic acid (OPDA) through a series of oxygenation, epoxidation, and cyclization reactions that are catalyzed by the enzymes 13-LIPOOXYGENASE, ALLENE OXIDE

SYNTHASE, and ALLENE OXIDE CYCLASE, respectively (79). OPDA is subsequently transported to and oxidized in peroxisomes to produce the bioactive JA, jasmonyl-L-isoleucine (JA-Ile). Once synthesized, JA-Ile is sensed by CORANANTINE

INSENSITIVE1 (COI1), a component of the E3 ubiquitin ligase complex SCFCOI1 (80).

JA-Ile sensing recruits the mediator coactivating complex MED25, which directly promotes COI1-JAZ interactions on MYC2 target gene promoters (81). This consequently inhibits or leads to polyubiquitination and subsequent degradation of JAZ repressors through the 26S proteasome pathway, therefore relieving MYC2 inhibition

(80, 81).

Although MYC2 is important for early JA-regulated gene expression, it can confer direct and indirect resistance to insects by initiating terpene synthesis through gene expression of TERPENE SYNTHASE (TPS) 10, TPS11, TPS21, and production of proteinase inhibitors and VEGATATIVE STORAGE PROTEINS (VSPs) (78, 79).

Degradation of JAZ repressors also relieve other MYC TFs, such as MYC3, MYC4,

MYC5, which act in concert to initiate gene expression necessary to produce secondary metabolites, such as glucosinolates (79). Additionally, the production of defenses against necrotrophic pathogens follows a separate JA-mediated signaling pathway downstream of

COI1 perception of JA-Ile, which relies on JA/ET synergism and is typically repressed by

MYC2. The JA/ET synergistic branch relies on the perception of ET and subsequent activation of APETALA2/ETHYLENE RESPONSE FACTOR (AP2/ERF) TF family, such as ETHYLENE RESPONSE FACTOR (ERF) 1 and OCTADECANOID-

RESPONSIVE ARABIDOPSIS (ORA) 59, which leads to the upregulation of defense

13 genes including PLANT DEFENSIN 1.2 (PDF1.2) (76, 78). However, MYC2 will repress transcriptional activity of AP2/ERF TFs (76). This separation is likely due to the energy costs associated with induced defenses and to promote appropriate defense responses.

However, the SA signaling pathway can also interact with JA signaling to further orchestrate appropriate defense responses.

Salicylic Acid Signaling and Plant Defenses

SA is a key phytohormone (Fig. 1.2) involved in PTI and ETI as well as directing

SAR in response to biotrophic pathogen infection (49). After SA accumulation transcription of plant genes encoding PR proteins, chitinases, β-1,3-glucanases, and other anti-microbial proteins inhibit biotrophic pathogen success (49). Two pathways are involved in SA biosynthesis, the phenylalanine and the isochorismate pathways, use chorismate form the Shikimate pathway to synthesize SA (82). The phenylalanine pathway converts chorismate into trans-cinnamic acid and benzoic acid, this reaction is catalyzed by the enzymes PHENYLALANINE AMMONIA LYASE and an enzyme that largely remains elusive to date (49, 78). Early evidence shows that benzoic acid may be subsequently converted into SA in a reaction that is catalyzed by benzoic acid 2- hydroxylase (49, 82). However, the isochorismate pathway predominates much in the pathogen-induced SA accumulation (78). The isochorismate pathway first converts chorismate into isochorismate through the enzyme ISOCHORISMATE SYNTHASE 1, which is subsequently converted into SA through the enzyme isochorismate pyruvate lyase (49).

Figure 1.2: Salicylic acid structure. Retrieved from: https://pubchem.ncbi.nlm.nih.gov/compound/salicylic_acid#section=2D-Structure

SA signaling relies heavily on the activation of the regulatory receptor known as

NPR1 (NON-EXPRESSOR OF PATHOGENESIS-RELATED 1) (83). NPR1 is an essential regulatory receptor that senses and binds to SA to initiate SA signaling (83).

Uninfected plants maintain inactive NPR1 in the cytosol as oligomers or inactive monomers in the nucleus by the transcriptional repressors NPR3 and NPR4 (49). Upon

SA accumulation through pathogen infection, cytosolic NPR1 will monomerize and enter the nucleus where nuclear NPR1 will also be relieved by NPR3 and NPR4 repression

(49). Following NPR1 activation and nuclear localization, NPR1 acts as a key co- activator for SA-responsive gene expression by interacting with the transcriptional activators of the TGACG-binding factor (TGA) transcription factor family (49). Once activated by NPR1, TGAs will subsequently bind to promoter regions and activate SA- responsive gene expression, including PR1, PR4, PR5, BGL and BG3, and several chitinase genes (49).

Additionally, JA- and SA signaling are mutually antagonistic pathways. NPR1 monomerization is associated with antagonizing MYC2 activity, however this mechanism

15 is not well-characterized (78). SA biosynthesis and signaling can also antagonize MYC2 activity. For instance, two inducers of reactive oxygen species-mediated SA signaling proteins, enhanced disease susceptibility 1 (EDS1) and phytoalexin deficient 4 (PAD4), are important for upstream positive feedback regulation of SA accumulation, but are also involved in MYC2 inhibition (49, 84). Interestingly, PAD4 is known to be involved in plant defenses against the green peach aphid (Myzus persicae) through inducing cell death, callose build-up, leaf senescence and subsequently affecting aphid feeding (65).

On the other hand, MYC2 can suppress SA accumulation and promote SA regulatory genes. For example, MYC2 can bind directly to the promoters of genes encoding for

NAC TFs, which negatively affect SA accumulation through inhibition of ICS1 gene expression as well as activating downstream SA regulatory genes that encode methyltransferases (76).

It is also well-established that JA- and SA signaling involve multiple other phytohormones to ‘fine tune’ appropriate defenses. ABA is a phytohormone involved in abiotic stress responses that can act synergistically with the MYC branch of JA signaling

(73). CK and BR have characterized roles in pant growth and development but are also well known to play key signaling roles in SA-mediated defense responses (73).

Moreover, another phytohormone, GA, has gained much attention over the past decade due to its putative role in modulation of JA- and SA signaling (73, 85, 86).

Gibberellic Acid Biosynthesis, Metabolism, and Plant Defenses

GA signaling is mediated by tetracyclic diterpenoid carboxylic acids known as gibberellins (GAs) (Fig. 1.3) (87). GAs were first identified as a secondary metabolite

16 from the necrotrophic fungal plant pathogen Gibberella fujikuroi (88), but are present across all taxa of higher plants (87, 89). Currently, ~136 GAs have been identified across plants, bacteria, and fungi, but only four are known to maintain bioactive characteristics in plants (GA1, GA3, GA4, and GA7) (89). Bioactive GAs share a C19 skeleton, meanwhile inactive GAs share a C20 skeleton (87). GAs are crucial for a variety of plant developmental processes, such as cell division and elongation, seed germination, stem elongation, pollen development and flowering (87, 89).

Figure 1.3: Gibberellic Acid 3 structure. Retrieved from: https://pubchem.ncbi.nlm.nih.gov/compound/Gibberellin_A3#section=Structures

GA biosynthesis has largely been studied in Arabidopsis thaliana where trans- geranylgeranyl diphosphate (GGPP) is converted into GA12 (89). Within the plastid,

GGPP is first converted into ent-kaurene via the terpene synthases, ent-COPALYL

DIPHOSPHATE SYNTHASE and ent-KAURENE SYNTHASE (89). This intermediate is then passed through the plastid membrane and endomembrane system where it is converted into GA12 through a series of reactions involving the membrane-associated cytochrome P450 monooxygenases (P450s), ent-KAURENE OXIDASE and ent-

KAURENOIC ACID OXIDASE (89). However, it is still largely unknown as to how

17 these intermediates are transported, or at which point, they are catalyzed throughout the plastid membrane. Upon GA12 synthesis, it is further processed into bioactive GAs within the cytosol through 2-oxoglutarate-dependent dioxygenases (2ODDs), GA 20-oxidase

(GA20ox) and GA 3-oxidase (GA3ox) respectively (89). However, there are two pathways that GA12 can take. GA12 can either undergo direct processing through GA20ox and GA3ox to synthesize GA4 and GA7 or GA12 can first be converted into GA53 and subsequently converted into GA1 and GA3 (89).

To maintain proper growth and development, it is critical for plants to regulate hormones. There are several known mechanisms for GA homeostasis. The most widely studied deactivation mechanism is the C-2 2β-hydroxylation (89). This mechanism requires a class of 2ODDs, GA 2-oxidase (GA2ox) family, to maintain GA homeostasis.

The two general classes of GA2ox that are important for this mechanism are C19-GA2ox and C20-GA2ox. C19-GA2ox catalyzes C-2 2β-hydroxylation on bioactive GAs as well as some of their immediate precursors, while C20-GA2ox can only interact with C20-GA precursors (89). Non-13-OH GAs, such as GA4 and GA12, can also be deactivated through a class of the CYP714 family of P450s (89). GA epoxidation is another important mechanism of GA inactivation. Specifically, the GA 16α,17-epoxidases

ELONGATED UPPERMOST INTERNODE (EUI) in rice and the EUI-like1 and EUI- like2 (ELA1 and ELA2) in Arabidopsis thaliana have been shown to produce 16α,17- epoxidates from GA4 as well as from its precursors, GA9 and GA12 (90, 91). GAs can also be deactivated by methylation through the methyltransferases GAMAT1 and GAMAT2 to produce inactive GA methyl esters (92, 93).

Another aspect of GA regulation is GA transport to local or distal tissues via the phloem and xylem (94). GAs, mostly GA intermediates, are transported through the highly conserved protein families, nitrate transporter 1/peptide transporter and SWEET families (94). GA12 and GA20 have been found to be involved in long-distance transport in A. thaliana and Pisum sativa (95, 96). Specifically, GA12 travels from root-to-shoot through the xylem and shoot-to-root through the phloem (95). In cucumber (Cucumis sativa), it was found that GA9 can be transported from the ovaries to the sepals and petals

(97). Furthermore, this resulted in high levels of GA4, therefore proposing that GA9 may be transported to nearby tissue and subsequently converted into bioactive GA4 (97). GA short- and long-distance transport is important because GA-mediated gene expression is initially repressed (77, 98), thus may rely on outside GA to enter the system (95, 97).

Gibberellic Acid Signaling and DELLA Proteins

The GA signaling pathway can be regulated temporally and spatially through various environmental cues, such as light, temperature, water, nutrients, abiotic and biotic stress (89). In absence of stimuli, GA signaling is repressed by nuclear GRAS domain transcriptional repressor proteins known as DELLAs (77, 98). Like all GRAS proteins,

DELLAs contain the conserved C-terminus leucine heptad repeats (LHRI and LHRII) along with three conserved motifs, HIID, PFYRE, and SAW (77, 98). DELLAs differ from other GRAS domain proteins in that their N-terminus contains two conserved domains, DELLA (Asp-Glu-Leu-Leu-Ala) and TVHNYP (77, 98). DELLAs inhibit early

GA gene expression through interactions with GA-related TFs, such as

PHYTOCHROME INTERACTING FACTOR (PIF)3 and PIF4. Upon stimulation, GA enters the system and is perceived by its receptor, GIBBERELLIN INSENSITIVE

DWARF 1 (GID1), to form a GA-GID1 complex, which interacts with the N-terminus domains of DELLAs (77). The GA-GID1-DELLA interaction causes further conformational changes in the DELLA C-terminus GRAS domain (77). Consequently, this conformational change recruits GIBBERELLIN INSENSITIVE 2 (GID2) and

SLEEPY1 (SLY1) F-box protein components of SKP1, CULLIN, F-BOX (SCF) E3 ubiquitin-ligase complexes (SCF GID2/SLY1) to polyubiquitinate DELLAs and tag for degradation via 26S proteasome pathway (77, 98). Therefore, PIF3 and PIF4 are relieved from DELLA repression and subsequently GA-related gene expression takes place and

GA-mediated plant growth and development responses can take place (e.g. stem elongation, pollen maturation, and cell elongation) (87, 89).

Interestingly, DELLAs differ among plant taxa. For example; five DELLAs exist in Arabidopsis thaliana (GAI, RGA, RGL1, RGL2, and RGL3), three exist in Medicago truncatula (DELLA1, DELLA2, and DELLA3), and only one exists in Oryza sativa

(SLR1) (43, 99). To date, DELLAs are known to maintain redundant, but diverse protein- protein interaction properties in their GRAS domain. Furthermore, DELLAs can act as transactivating proteins in other signaling pathways, such as the JA signaling or symbiosis signaling pathways (9, 77, 80).

Roles of Gibberellic Acid in Plant-Microbe and Plant-Insect Interactions

DELLAs can act as transactivating proteins by outcompeting MYC2 for JAZ interactions (77, 80). By interacting with JAZs, DELLAs can relieve MYC2 from JAZ repressing activity, thereby allowing for JA-regulated gene expression to take place (Fig.

1.4) (1). This has been demonstrated in plant-pathogen studies using della mutants in A.

20 thaliana and studies applying exogenous GA treatment to O. sativa (85, 100-103). With the use of quadruple-della mutants in A. thaliana and exogenous GA treatment in tomato, it was found that plants were less susceptible to biotrophs and hemibiotrophs, but more susceptible to necrotrophs (85, 100-103). Therefore, GA is a key regulatory phytohormone in plant disease resistance (86). Nevertheless, GA plays an important role in other microbe-plant interactions through DELLAs. Specifically, GA acts as a negative regulator of arbuscule development in AM fungi-plant associations through DELLA degradation (43, 104). As mentioned earlier, sensing of nuclear Ca2+ oscillations in root cortical cells promotes CCaMK-CYCLOPS-DELLA interactions, which is critical for arbuscule development through upregulation of the gene RAM1 (9, 10, 45). Although GA plays key regulatory roles in microbe-plant interactions, the role of GA in mediating plant-insect interactions is not as well-defined.

Figure 1.4: Jasmonic acid-gibberellic acid antagonism. In uninfected tissue, JAZs will interact with MYC2 TFs and inhibit JA signaling. Upon JA perception, JAZs are either inhibited or degraded. Moreover, DELLAs can outcompete MYC2 for JAZ binding. However, in the presence of JA and GA, JA signaling is hindered due to DELLA degradation (1). To date, there is not much known about the role of GA in modulation of plant defenses during herbivory. It has been found that exogenous GA increases susceptibility to the root-knot nematode (Meloidgyne graminicola), GA signaling is dominant over JA,

21 and that JA-mediated defenses in rice against M. gramicola are dependent on GA repression by SLR1 (105). Another recent study has reported that brown planthopper

(Niaparvata lugens) feeding on rice overexpressing OsGID1 resulted in decreased SA levels and defense signaling compared to wild type (WT) plants (106). Moreover, it was reported that brown planthopper egg hatching rate was significantly higher on WT plants and that adults preferred to feed and lay eggs on WT plants in laboratory experiments

(106). Additionally, there were also significantly fewer adults and nymphs found on plants overexpressing OsGID1 compared to WT in field experiments (106). Another study using A. thaliana quadruple della mutants reported that beet armyworm suppression of JA levels were alleviated in the quadruple della mutants, indicating that

DELLA proteins are important for beet armyworm suppression of JA-related hormone levels (OPDA, JA, and JA-Ile) (107). In addition, the authors also suggest that glucosinolate levels might be positively regulated by DELLA proteins (107). Although there is evidence suggesting multiple roles of GA in modulation of plant defenses during plant-herbivore interactions, these may be context-dependent on pest or insect feeding guild (108).

Significance

AM fungi-plant-insect interactions is an area of research that has gained attention over the past two decades due to ecological and agricultural relevance (47, 57, 109-111).

As mentioned earlier, AM fungi are important plant-associated beneficial soil microbes that form relationships with many terrestrial plants, including many important crops

(112). These relationships can promote plant growth and development through changes in

22 plant nutrition (19, 112), increased crop yield (29-33, 112) and alleviate stress from abiotic (19) and biotic factors (6, 34, 48, 113, 114). In addition, aphids are some of the most important agricultural pests that cause billions of dollars of agricultural damage in temperate regions world-wide (62-64). In addition, aphids can be positively affected by root colonization with AM fungi (57). There is evidence to suggest that aphids can indirectly interact with AM fungi through changes in plant gene expression belowground.

For example, a study using barrel medic plants, pea aphids, and the AM fungus

Rhizophagus irrregularis found that aphid feeding strongly upregulated of GA2ox in plant roots during the tripartite interaction (59). GA2ox catalyzes bioactive GAs, which can negatively regulate AM fungal symbioses (43, 104) and JA-regulated defense gene expression (80).

In the present study we used two separate biological systems to; 1) examine the role of exogenous GA application in regulating plant defenses during AM fungus-plant- aphid interactions and 2) evaluate the role of GA signaling in modulating plant defenses during plant-aphid interactions. The first system involved pea aphids (Acyrthosiphon pisum), barrel medic plants, and the AM fungus Rhizophagus intraradices. The second system involved pea aphids and two plant genotypes (segregant wild type (WT) and

Mtdella1/Mtdella2 mutant). Pea aphids have become an important model organism since they are agriculturally important pests and viral vectors for over 30 plant viruses, including the agriculturally notorious Pea enation mosaic virus and the Bean leaf roll virus (63), and have a sequenced genome (115). These pests feed mostly on leguminous plants in the family Fabaceae (63), including the barrel medic plant (116). The barrel

23 medic plant is an important model organism for studying functional genomics and plant- beneficial microbe interactions in agriculturally relevant crops (117). For instance, the barrel medic plant is a close relative of alfalfa, which is an important foraging crop in the

US, and shares similarities with other cool season leguminous crops (117).

Aphid infestation alone causes over $1 billion in agricultural and forestry productivity in the US (118). Moreover, aphids exhibit tremendous insecticide resistance, which requires an increased amount of insecticide use to combat aphid infestation (118).

Consequently, this imposes major public health and environmental concerns (118), underscoring the importance of developing more sustainable alternatives to deal with aphids. AM fungi may be an alternative to not only minimize fertilizer and water input, but also to minimize insecticide use in agricultural systems (112). Although, aphid feeding is generally positively impacted by AM fungi (57), understanding the mechanisms of aphid suppression of MIR may provide an avenue of biotechnological use of AM fungi. Data from this study may provide the scientific community with greater insight into the mechanisms dictating aphid suppression of MIR and take steps closer to finding sustainable solutions to aphid old problems.

CHAPTER II

EFFECT OF EXOGENOUS GIBBERELLIC ACID IN REGULATING PLANT DEFENSE RESPONSES DURING PLANT INTERACTIONS WITH APHIDS AND ARBUSCULAR MYCORRHIZAL FUNGUS

Abstract

Arbuscular mycorrhizal (AM) fungi form symbioses with most plants helping them acquire limiting nutrients and cope better with abiotic and biotic stresses. However, insects, such as aphids, can take advantage of this interaction by modifying plant defenses. For instance, the phytohormones jasmonic acid (JA) and salicylic acid (SA) are well known for modulating specific defense responses against pathogens and insects.

Nevertheless, this process is not simple as other phytohormones are also involved such as gibberellic acid (GA) that can interact with the JA and SA pathways through DELLA proteins. To determine the role of GA in modulating AM fungi-plant-aphid interactions, we used a system involving pea aphids (Acyrthosiphon pisum), barrel medic plants

(Medicago truncatula), and the AM fungus Rhizophagus intraradices. Plants were subjected to exogenous GA application, aphid herbivory (36 hours and 7 days), and AM fungus root colonization. Shoot/root fresh and dry weight, aphid fitness, GA3, JA, and SA concentrations in leaves and petioles, and shoot and root gene expression for several plant defense marker and AM specific genes were measured. The results indicate that exogenous GA negatively impacts pea aphid fitness on mycorrhizal plants, meanwhile

25 positively impacts pea aphid fitness on non-mycorrhizal plants. Exogenous GA increased

JA and SA levels in plant leaf and petiole tissue 36 hours post aphid feeding. In addition,

AM fungus root colonization decreased SA levels in leaves and petioles, however,

PATHOGENENESIS RELATED (PR)1 gene expression was upregulated by AM fungus root colonization in both shoots and roots 36 hours post aphid feeding. Moreover, gene expression of the JA-regulated defense gene VEGETATIVE STORAGE PROTEIN (VSP) was upregulated in response to both exogenous GA and aphid herbivory. The results indicate that GA may play a role in modulation of plant defenses during plant-aphid and

AM fungi-plant-aphid interactions.

Introduction

Plants are immobile primary producers that interact with heterotrophic organisms in their shoots and roots simultaneously. A type of beneficial plant interaction with soil organisms involves arbuscular mycorrhizal (AM) fungi (17). These fungi are members of the phylum Glomeromycota that form beneficial associations with roots of ~80% of vascular plants (17). These ubiquitous soil microorganisms assist plants in absorption of soil nutrients, especially phosphorus, in exchange for photosynthates and lipids (17, 20).

The proposed use of AM fungi as biofertilizers has gained attention in recent years (19,

27, 119). It has been shown that mycorrhizal plants exhibit increases in crop yield and quality (29-33, 120-123), in addition, AM fungi can help plants cope with abiotic (124-

128) and biotic stresses (6, 34, 47, 48, 109, 110, 114). For instance, AM fungal colonization of roots can enhance plant inducible defenses to a variety of pathogens and insect pests (6, 34, 47, 48, 114).

It is also possible for above and belowground organisms to interact indirectly with each other through plant physiology changes in the shared host. AM fungal colonization of roots can enhance plant resistance to insects and pathogens by ‘priming’ plants through the phytohormone jasmonic acid (JA) (6, 34, 48, 114) and through extraradical mycorrhizal networks (52, 53, 129). Priming can be described as enhanced plant preparedness for subsequent pathogen infection or insect infestation (6, 7, 113).

Beneficial microbes such as AM fungi can prime plants through a mechanism known as induced systemic resistance (ISR), which is characterized by whole plant preparedness for insect attack and is governed by JA (6, 7, 34, 47, 48). However, there are opportunistic organisms that take advantage of plant-beneficial microbe interactions. For instance, a meta-analysis revealed that certain specialist leaf-chewers, and phloem- feeders such as aphids may benefit from the AM symbioses (57). Aphids are notorious agricultural pests that serve as vectors for as many as 275 debilitating crop viruses of economic importance and contribute to enormous agricultural and forestry losses in the

U.S. (62, 118). Most aphids feed on phloem sap through their mouthparts called stylets, which secrete saliva containing biochemically active proteins (e.g. pectinases, cellulases, oxidases, and lipases) as well as other compounds that may be involved in plant susceptibility to aphid feeding (4, 65, 71, 130). In addition, aphids have been shown to have a top-down effect on AM fungal colonization of roots through changes in plant volatiles (54), which can subsequently transfer signals and alert nearby plants, and further alter aboveground defense responses and volatile release (61, 129).

Plant defense signaling is largely dictated by the antagonistic effect of the phytohormones JA and SA (73). JA is a key signaling component to produce defenses

27 such as secondary metabolites and proteinase inhibitors against insect herbivores and necrotrophic pathogens (131), meanwhile, SA signaling contributes to defenses against biotrophic pathogens through the production of PATHOGENESIS-RELATED (PR) proteins (76). Although it is well documented that JA signaling is important for plant resistance against aphids (132-137), the underlying mechanism used by aphids to manipulate these plant defenses is not well understood. Several lines of evidence indicate that aphids can promote host susceptibility by modulating the balance between JA and

SA (130, 138-142). Although, JA and SA are at the front lines of plant defense responses, there are other phytohormones that can influence the balance between JA and SA to ‘fine tune’ appropriate plant defense and growth responses (73).

The phytohormone gibberellic acid (GA) has gained attention over recent years due to its role in modulation of the strength of the JA- and SA-signaling pathways in plant-pathogen studies (1, 73, 85, 86, 101-103). GA signaling has been primarily studied during plant-pathogen interactions, and it has been found to enhance resistance to biotrophic pathogens and promote susceptibility to necrotrophic pathogens by negatively regulating JA signaling (1, 85, 100-102, 143). The GA signaling pathway is characterized by a class of diterpenoid carboxylic acids known as ‘gibberellins’ that play critical roles in seed germination, fruit development, stem elongation, flowering, and other aspects of plant development (89, 144). GA signaling is initially repressed by GRAS domain transcriptional repressor proteins of GA signaling known as DELLAs, which are characterized by their unique N-terminus DELLA and TVHNYP domains (77, 145). GA signaling requires gibberellin perception by the receptor GIBBERELLIN INSENSITIVE

DWARF 1 (GID1), which forms a complex and is subsequently transported to the nucleus where DELLAs are recruited. GA-GID1-DELLA interactions promote DELLA polyubiquitination and degradation via the 26S proteasome pathway, therefore, relieving

GA transcription factors and activating GA-regulated gene expression (77). Moreover,

DELLAs are known to have diverse protein-protein interaction capabilities that allow them to play key regulatory roles in other pathways, such as the JA signaling pathway

(77, 80).

DELLAs have been found to confer resistance to necrotrophic pathogens by acting as important transactivating proteins in the JA signaling pathway (1, 85, 102).

DELLAs support JA signaling by releasing the master regulatory transcription factor

MYC2 from its transcriptional repressors, JASMONATE ZIM DOMAIN (JAZ) proteins

(80). Induction of GA leads to DELLA degradation, which negatively affects JA signaling, thus conferring resistance to biotrophic pathogens, meanwhile decreasing resistance to necrotrophic pathogens (1, 85, 143). Although GA signaling plays a key role in plant immunity during plant-pathogen interactions, its role in insect-plant and insect- plant-AM fungi interactions remain elusive. It has been reported that the labial saliva produced by beet armyworm (Spodoptera exigua) requires DELLAs to modulate JA phytohormone levels in Arabidopsis thaliana quadruple della mutants (107). GA has also been found to compromise rice (Oryza sativa) resistance against root-knot nematodes

(Meloidogyne graminocola) through DELLA degradation (105). Conversely, another study indicates that overexpression of OsGID1 increases rice resistance to the brown planthopper (Nilaparvata lugens), however, resistance may be attributed to other

29 structural mechanisms, such as increased lignin content (106). It was shown recently that pea aphids (Acyrthosiphon pisum) may manipulate gene expression of GA biosynthesis genes belowground, which could interfere with AM fungus-plant symbioses (59).

GA is also an important regulator in AM fungi-plant interactions. DELLAs are components of transcription complexes that positively regulate arbuscule development in root cortical cells (43, 45). Furthermore, gibberellin perception and biosynthesis negatively regulates arbuscule formation (104). In terms of tripartite interactions, it has been shown that pea aphid feeding upregulates gene expression of GA biosynthesis genes in Medicago truncatula roots that are highly colonized with the AM fungus Rhizophagus irregularis (59). Although pea aphid feeding can negatively affect AM fungal root colonization in fava beans (54), the mechanisms dictating aphid suppression of plant defenses during this tripartite interaction still remains unclear. Advances in this field of research may provide insights that can help refine sustainable agricultural and forestry practices. For instance, aphids quickly develop insecticide resistance, which requires increased amounts of insecticide use to control aphid outbreaks (118). Consequently, this causes major public health and environmental concerns (118), therefore underscoring the importance of developing more sustainable alternatives to deal with aphids. By expanding our understanding of the mechanisms that drive plant defenses in AM fungi- plant-aphid interactions, AM fungi may be used as an alternative not only to minimize fertilizer and water input, but also to minimize insecticide use, especially in organic agricultural systems (112).

In the present study, the role of exogenous GA application in regulating plant defenses during AM fungus-plant-aphid interactions was examined by using the AM

30 fungus Rhizophagus intraradices, the model legume M. truncatula, and pea aphids. The following hypotheses were tested: 1) aphid fitness will be positively impacted by exogenous GA application to plants, 2) exogenous GA application will result in reduced arbuscule formation, 3) JA levels and JA-regulated defense gene expression will be downregulated in response to exogenous GA application, and 4) SA levels and SA- regulated defense gene expression will be upregulated in response to exogenous GA application.

Methods and Materials

Plant Growth Conditions

Medicago truncatula Jemalong A17 (wild type) seed sterilization and germination procedures followed a protocol outlined in Maurya et al. (59). Briefly, seeds were scarified for 10 minutes in concentrated H2SO4, rinsed in sterile water three times, sterilized for 10 minutes using 10% (v/v) household bleach in 0.1% (v/v) Tween 20 solution, and rinsed in sterile water five times (59). Seeds were spread on wet filter paper

(sterile) in petri dishes, dishes were sealed with parafilm, and wrapped in aluminum foil.

Dishes were incubated in a 4°C refrigerator for three days (dark), kept at room temperature (24°C) for one day (dark), then the aluminum foil was removed and dishes were placed under indirect light (mean: 184 µmol m-2 s-1) for three days. Seedlings were planted in azalea pots (12 cm W x 8.5 cm H) filled with a sterilized mason sand: topsoil mix (9:1) (Pioneer Sand Company, Windsor, CO) and were grown in a growth chamber under a 16-hour photoperiod with the following conditions: 24°C, 40% relative humidity, and light intensity of 285-290 µmol m-2 s-1 provided by fluorescent and halogen light

31 bulbs. The sand was thoroughly washed with tap water and the topsoil was sieved (sieve no. 8) prior to autoclaving both substrates three times (each cycle: 60 minutes, 121°C, at

15 psi). The sand: topsoil mix was saturated with ½ strength modified Hoagland’s solution (100µM P, 15mM N, pH 6.1) prior to transplant (146). Azalea pots were covered by clear plastic humidity domes (54.6 cm H×28 cm W×17.8 cm D) for one week after transplant. Once domes were removed, plants were watered daily with 50 mL of Milli-

Q® water or received fertilizer treatment twice a week with 50 mL ½ strength modified

Hoagland’s solution (100µM P, 15mM N, pH 6.1) (146).

Rhizophagus intraradices Inoculation

Fifteen days after M. truncatula seedlings were transplanted into azalea pots, seedlings with two or three trifoliolate leaves were transplanted into smaller pots (6.35 cm W x 9 cm H) with either a 1:10 dilution of R. intraradices inoculum or a 1:10 dilution of mock inoculum (root exudates without AM fungi). Mock and R. intraradices (UT118,

IA506, and Co204) inocula were purchased from the International Culture Collection of

(Vesicular) Arbuscular Mycorrhizal Fungi (INVAM) (Morgantown, West Virginia,

USA). The inoculation procedure consisted of a 2.5 cm bottom layer of sterilized sand: topsoil substrate (9:1), followed by a 1.5 cm layer of either a 1:10 R. intraradices inoculum or 1:10 mock inoculum, and a 1.5 cm top layer of reused soil substrate from the seedlings’ azalea pots. Substrate or inoculum layers were moistened with ½ strength modified Hoagland’s solution (100 µM P, 15 mM N, pH 6.1) prior to transplanting (146).

Plants were covered by clear plastic humidity domes (54.6 cm H×28 cm W×17.8 cm D) for one week and were grown in growth chambers for the remainder of the experiment.

Extra plants were inoculated to assess root colonization levels by R. intraradices prior to adding aphids to experimental plants. To assess the level of R. intraradices colonization, roots were cleared with 10% (w/v) KOH (85°C for 4 h), rinsed using deionized water, and stained with 5% (v/v) Sheaffer black ink that was prepared in 5%

(v/v) acetic acid (147). After staining, R. intraradices colonization was quantified using the modified gridline-intersect method with the aid of an Olympus SZX10 stereo microscope (Leica Microsystems, Wetzlar, Germany) (148). Plants receiving mock inoculum did not show staining for fungal structures. Aphids were added to experimental plants when R. intraradices colonization reached at least >50% of root length colonization (59).

Gibberellic Acid 3 Treatment

-1 A 50 mg mL GA3 stock solution (PhytoTechnology Laboratories, Lenexa, KS) was made, which was further diluted into a 10-6 M working solution using 100% ethanol and Milli-Q® water (43). One week after inoculation with AM fungi and mock inocula, plants received 17.5 mL of 1 µM GA3 or control treatments daily (except when fertilized)

(43) and 17.5 mL of ½ strength Hoagland’s solution (100 µM P, 15 mM N, pH 6.1) twice a week (146) for the remainder of the experiment.

Pea Aphid Infestation

Parthenogenetic, wingless female pea aphids were provided by Dr. Kenneth Korth

(University of Arkansas, Fayetteville, AR) and reared on fava bean (V. fava) plants in insect tents in the laboratory under a 16-h photoperiod. One thousand six-day old aphids were synchronized (138) by adding wingless adult aphids to a separate insect tent with non-infested fava beans. The adults were removed the following day leaving only

33 nymphs. Nymphs were raised on fava beans for an additional five days. After R. intraradices colonization reached at least 50%, 15 six-day old aphids were added to each plant receiving aphids. Harvests took place in the middle of the subjective day (10:00 –

16:00 h) when jasmonate levels peak (149). One replicate of every treatment was harvested at the same time to account for changes in gene expression that are regulated by circadian rhythms.

The present study consisted of eight treatments: 1) -AM fungus/ -Pea aphid / -GA

; 2) -AM fungus/ +Pea aphid / -GA; 3) -AM fungus/ -Pea aphid/ +GA ; 4) -AM fungus/

+Pea aphid/ +GA; 5) +AM fungus / -Pea aphid/ -GA; 6) +AM fungus/ +Pea aphid/ -GA;

7) +AM fungus/ -Pea aphid/ +GA; and 8) +AM fungus/ +Pea aphid/ +GA. Immediately after aphid infestation took place, all plants including non-infested plants were covered with organza drawstring gift bags (15.24 x 22.86 cm, SumDirect). Aphids fed continuously on plants for 36 hours (138) and 7 days (59). At the end of each feeding period, aphids were gently collected from each plant using a vacuum device and were immobilized at -20°C prior to counting and weighing them. Non-infested plants were also exposed to the vacuuming effect. The vacuum device consisted of a 4.8 mm diameter

Tygon tubing, connected to a 50 mL conical tube (Fisher Scientific), and a cut 200 µL pipette tip. Plant fresh weights for roots and shoots were measured using an analytical scale prior to freezing the tissue in liquid nitrogen and storage at -80°C.

Phytohormone Quantification

Phytohormone extraction and concentration was determined using ultra- performance liquid chromatography - mass spectrometry (UPLC-MS), and was performed only on plant shoot tissues following the procedure outlined in Sheflin et al.

(150) with some modifications. Plant shoot samples were stored at -80°C post-harvest, lyophilized and subsequently extracted using a monophasic methyl-tert-butyl ether

(MTBE) extraction protocol (150). First, 19.4 mg of lyophilized leaf and petiole tissue was measured using an analytical balance and subsequently treated with 990 µL of HPLC grade MTBE and 10 µL of internal standard mix and vortexed at 4°C for 2 hours.

Samples were centrifuged at 3,500 x g at 4°C for 15 minutes, and 750 µL of supernatant was transferred to new microcentrifuge tubes to incubate overnight at -80°C. Next, samples were centrifuged at 18,000 x g at 4°C for 20 minutes. Samples were extracted under nitrogen conditions at room temperature and resuspended in 100 µL of HPLC grade methanol and stored at -80°C. Samples were sent to the Proteomics and

Metabolomics Facility at Colorado State University, Fort Collins, CO where UPLC-MS was conducted (150).

Ribonucleic Acid Isolation and Complementary Deoxyribonucleic Acid Synthesis

RNA isolation and cDNA synthesis followed the protocol outlined in (151). Three biological replicates were selected randomly within each of the treatments for gene expression analyses. Roots and shoots were ground separately using a mortar and pestle in liquid nitrogen. RNA was extracted using the RNeasy Plant Mini Kit according to the manufacturer’s instructions (Qiagen Inc., Germantown, MD, USA). RNA samples were treated with 87 µL of nuclease-free water, 10 µL of 10X reaction buffer, and 3 µL of

TurboTM DNase (2 units µL-1) for a total volume of 100 µL and incubated at 37°C for 40 minutes. DNAse-treated RNA samples were purified using the RNeasy MinElute

Cleanup kit (Qiagen Inc.). An additional DNAse treatment was performed using the

DNA-freeTM DNA removal kit (Thermo Fisher Scientific) according to the manufacturer’s protocol for RNA samples that showed trace amounts of genomic DNA contamination. For cDNA synthesis, 1 µg of total RNA was mixed with 1 µL dNTPs (10

-1 mM each) and 1 µL anchored oligo dT22 (500 ng µL ) and incubated at 65°C for 5 minutes using a T100™ Thermal Cycler (Bio-Rad Laboratories Inc., Hercules, CA,

USA).. Following this step, 4 µL of SuperScript® IV Buffer, 1.2 µL Nuclease-free water,

1 µL of DTT (100 µM), 0.5 µL of RNaseOUTTM (Thermo Fisher Scientific), and 0.3 µL of SuperScript® IV (Thermo Fisher Scientific) were added to each sample (total volume

20 µL). Samples were incubated at 50°C for 10 minutes and 80°C for 10 minutes using a

T100™ Thermal Cycler (Bio-Rad Laboratories Inc). cDNA quality was assessed via reverse-transcription polymerase chain reaction (RT-PCR) (26 cycles of 95°C for 30 seconds, 59°C for 30 seconds, and 72°C for 30 seconds) using the reference gene

ELONGATION FACTOR 1 α (EF1- α). Products were visualized using a 0.5X TAE 2%

(w/v) agarose gels.

Plant Gene Expression During Arbuscular Mycorrhizal Fungus-Plant-Aphid Interactions

Plant gene expression was assessed to investigate the role of exogenous GA3 in plant defense signaling. Markers for plant defense responses such as the SA-regulated defense gene PATHOGENESIS RELATED 1 (MtPR1) (152), and the JA-regulated defense genes MtMYC2 and MtVSP (153) were chosen. Also, marker genes involved in arbuscule presence such as the plant phosphate transporter gene MtPT4 (146), the AM

36 specific lectin (MtLec5) (154, 155), and the AM fungal gene RiEF (156) were included for root samples.

To assess gene expression, 1 µL of cDNA template (1:5), 5 µL of

PowerSYBR®Green Master Mix (Thermo Fisher Scientific), 2 µL of autoclaved Milli-

Q®water, and 1 µL of 3 µM forward and reverse primers was used. Each of the 384-well plates were run on a C1000®Touch ThermalCycler (Bio-Rad, Hercules, CA, USA) and each run included two technical replicates and 3-4 biological replicates per treatment.

The thermal profile was comprised of an initial incubation at 95°C for 10 minutes, followed by 40 cycles at 95°C for 15 seconds, an annealing/extension at 53.3-63.3°C for

1 minute, and a melt curve analysis ranging from 65-95°C that increases incrementally by

5°C. Oligonucleotide sequences and annealing temperatures used for RT-qPCR are reported in Table 2.1. The 2-ΔCq method was used to calculate relative expression (157) of six gene targets. Each gene was calibrated to the reference gene GLYCERALDEHYDE-3-

PHOSPHATE DEHYDROGENASE (GAPDH) (158, 159).

Table 2.1. Primers used for real-time quantitative PCR of target genes Gene Forward Primer (5’-3’) Reverse Primer (5’-3’) Average Annealing Temperature GAPDH AACATCATTCCCAGCAGCAA AACATCGACGGTAGGCACAC 56.6°C MtMYC2 GTCACAGTTCGTCGCTGGTG CGCCTCTGCTGCTTGATTTC 57.65°C MtVSP GACCTTTGGGTGTTTGACATTGA TCCTTCTGTTTGAGTGGTCTTCCT 58°C MtPR1 ATCCCCCAGAACATTGCTCG CCATCCAACACCACTACCCC 57.85°C MtPT4 GACACGAGGCGCTTTCATAGCAGC GTCATCGCAGCTGGAACAGCACCG 60°C MtLec5 TCAAGTTGCTGAAACACATGAT GAGCAGAACCATTGCAACAA 58.6°C RiEF GCTATTTTGATCATTGCCGCC TCATTAAAACGTTCTTCCGACC 60°C

Statistical Analyses

All statistical analyses were carried-out using SAS 9.4 for Windows (SAS

Institute Inc, Cary, NC, USA). Normal distribution of raw data was determined using

Shapiro-Wilk and Anderson-Darling tests (P>0.05). Three-factor analyses of variance

(ANOVA) were used to determine the interaction effect of GA3 application, aphid herbivory, and AM fungus root colonization on plant growth parameters and plant defense gene expression. Data on plant growth and gene expression that were not normally distributed were subsequently analyzed by three-factor analyses of maximum likelihood using log-linked gamma distributions. Three-factor analyses of maximum likelihood using log-linked gamma distributions were also used to determine the interaction effect of GA3 application, aphid herbivory, and AM fungus root colonization on phytohormone concentration for the phytohormones GA3, JA, and SA. Two-factor

ANOVAs were used to determine the effect of GA3 application and AM fungus root colonization on mean aphid colony weights. Aphid colony weight data that did not follow the assumptions of normal distribution were subsequently analyzed using two-factor analyses of maximum likelihood using log-linked gamma distributions. Mean aphid count per colony data were analyzed using two-factor analyses of maximum likelihood using Poisson or negative binomial distributions. In addition, two-factor ANOVAs were used to determine the effect of GA3 application and aphid feeding on AM fungal-related gene expression.

Results

Effect of Exogenous Gibberellic Acid 3 Application on Plant Shoot and Root Fresh Weight After 36 Hours of Aphid Feeding

Overall, the interaction among exogenous GA, aphid herbivory, and AM fungus root colonization (GA*PA*AMF) did not have a significant effect on mean shoot fresh weight at 36 hours post aphid feeding (Fig. 2.1a; P=0.1660). At 36 hours post feeding, there were no significant effects observed from the interaction between exogenous GA and aphid herbivory (GA*PA) regardless of AM fungus root colonization status (+/-

AMF) (Fig. 2.1a; P=0.4832) or the interaction between aphid herbivory and AM fungus root colonization (PA*AMF) of exogenous GA (+/-GA) (Fig. 2.1a; P=0.2085). However, the interaction between exogenous GA and AM fungus (GA*AMF) regardless of aphid herbivory (+/-PA) had a significant effect on mean plant shoot fresh weight (Fig. 2.1b; p=0.0179). The post-hoc analysis revealed that plants with AM fungus colonized roots only and plants receiving exogenous GA with AM fungus root colonization regardless of aphid herbivory (+/-PA) had the highest mean fresh shoot weight (Fig. 2.1b).

Additionally, the interaction among exogenous GA, aphid herbivory, and AM fungus root colonization (GA*PA*AMF) did not have a significant effect on mean root fresh weight

(Fig. 2.2; P=0.1589).

Figure 2.1: Impact of exogenous GA, pea aphid (Acyrthosiphon pisum) feeding, and AM fungus (Rhizophagus intraradices) root colonization on Medicago truncatula shoots at 36 hours post-aphid feeding. The interaction among exogenous GA, pea aphid feeding, and AM fungus (GA*PA*AMF) did not have statistically significant effects on shoot fresh weight according to the three-factor analysis of maximum likelihood (P=0.1660) (a), but the interaction between exogenous and AM fungus was statistically significant according to the two-factor analysis of maximum likelihood using a log-linked gamma distribution (P=0.0179) (b). Values represent the mean shoot fresh weight ± SEM (n= 7 or 8) per treatment. Different letters represent significant differences among groups using Tukey-Kramer post-hoc tests (P<0.05).

There were no other significant effects reported for mean root fresh weight (Fig.

2.2), indicating that mean root fresh weight was not significantly affected by exogenous

GA, pea aphid feeding, or AM fungus root colonization at 36 hours post aphid feeding.

Figure 2.2: Impact of exogenous GA, pea aphid (Acyrthosiphon pisum) feeding, and AM fungus (Rhizophagus intraradices) root colonization on Medicago truncatula roots at 36 hours post-aphid feeding. The interaction among exogenous GA, pea aphid feeding, and AM fungus (GA*PA*AMF) did not have statistically significant effects on shoot fresh weight according to the three-factor analysis of variance (P=0.1589). Values represent the mean shoot fresh weight ± SEM (n= 7 or 8) per treatment.

Effect of Exogenous Gibberellic Acid 3 Application on Pea Aphid Count and Colony Weight at 36 Hours Post Feeding

The interaction between exogenous GA and AM fungus root colonization

(GA*AMF) did not have a significant effect on mean aphid count per colony at 36 hours post aphid feeding (Fig. 2.3a; P=0.6474). However, the interaction between exogenous

GA and AM fungus root colonization (GA*AMF) did have a significant effect on mean aphid colony at 36 hours post aphid feeding (Fig. 2.3b; P=0.0410). Nevertheless, a post- hoc analysis revealed that there were no differences among the groups (Fig. 2.3b).

Figure 2.3: Impact of exogenous GA and AM fungus (Rhizophagus intraradices) root colonization on mean pea aphid (Acyrthosiphon pisum) count per colony (a) and mean colony weight (b) after 36 hours of aphid feeding on Medicago truncatula plants. The interaction between exogenous GA and AM fungus root colonization (GA*AMF) had no statistically significant effect on mean pea aphid count per colony according to the two-factor analysis of maximum likelihood using Poisson distributions with scaled Pearson χ2 (P=0.6474) (a). The interaction between exogenous GA and AM fungus root colonization (GA*AMF) had a significant effect on mean pea aphid colony weight according to the two-factor analysis of variance (P=0.0410), however there were no differences among the treatment groups according to the Tukey-Kramer post-hoc test (P>0.05) (b). Values represent the mean shoot fresh weight ± SEM (n= 7 or 8) per treatment.

Effect of Exogenous Gibberellic Acid 3 Application and Arbuscular Mycorrhizal Fungus Root Colonization on Leaf and Petiole Levels of JA, SA, and GA3 at 36 Hours Post Aphid Feeding

The overall interaction among exogenous GA, aphid herbivory, and AM fungus root colonization (GA*PA*AMF) did not have a significant effect on mean GA3 concentration in leaf and petiole tissue at 36 hours post aphid feeding (Fig. 2.4a;

P=0.1223). However, the interaction between exogenous GA and AM fungus root colonization (GA*AMF) regardless of aphid feeding (+/- PA) had a significant effect on mean GA3 concentration (Fig. 2.4b; P=0.0282).

Figure 2.4: Impact of exogenous GA, pea aphid (Acyrthosiphon pisum) feeding, and AM fungus (Rhizophagus intraradices) root colonization on mean gibberellic acid 3 (GA3) concentration in leaf and petiole tissue. The interaction among exogenous GA, pea aphid feeding, and AM fungus root colonization (GA*PA*AMF) had no statistically significant effect on mean GA3 concentration according to a three-factor analysis of maximum likelihood using a log-linked scaled Pearson χ2 gamma distribution (P=0.1223) (a), but the interaction between exogenous GA and AM fungus root colonization (GA*AMF) did have a significant effect on mean GA3 according to a two-factor analysis of maximum likelihood (P=0.0282) (b). Values represent the mean ± SEM (n = 4). Different letters represent significant differences among groups using Tukey-Kramer tests (P<0.05).

The interaction among exogenous GA, aphid herbivory, and AM fungus root colonization (GA*PA*AMF) also did not have a significant effect on mean JA concentration in shoot leaf and petiole tissue (Fig. 2.5a; P=0.9135). Meanwhile, the interaction between exogenous GA and AM fungus root colonization (GA*AMF) regardless of aphid herbivory (+-PA) did have a significant effect on mean JA concentration (Fig. 2.5b; P=0.0389). Moreover, the post-hoc analysis indicated that plants receiving exogenous GA only regardless of aphid herbivory (+-PA) exhibited the highest mean JA concentration (Fig. 2.5b).

Figure 2.5: Impact of exogenous GA, pea aphid (Acyrthosiphon pisum) feeding, and AM fungus (Rhizophagus intraradices) root colonization on mean jasmonic acid (JA) concentration in leaf and petiole tissue. The interaction among exogenous GA, pea aphid feeding, and AM fungus root colonization (GA*PA*AMF) had no statistically significant effect on mean JA concentration according to a three-factor analysis of maximum likelihood using a log-linked gamma distribution with a scaled Pearson χ2 (P=0.9135) (a), but the interaction between exogenous GA and AM fungus root colonization (GA*AMF) did have a significant effect on mean JA concentration according to a two-factor analysis of maximum likelihood (P=0.0389) (b). Values represent the mean ± SEM (n = 4). Different letters represent significant differences among groups using Tukey-Kramer tests (P<0.05).

Similarly, the interaction among exogenous GA, aphid herbivory, and AM fungus root colonization (GA*PA*AMF) did not have a significant effect on mean SA concentration in shoot leaf and petiole tissue (Fig. 2.6a; P=0.4278). In addition, none of the two-factor interactions had significant effects on mean SA concentration (Fig. 2.6a).

However, the three main effects: exogenous GA, aphid herbivory, and AM fungus root colonization did have significant effects on mean SA concentration (Fig. 2.6 b, c, d;

P<0.0001, P=0.0378, and P<0.0001).

Figure 2.6: Impact of exogenous GA, pea aphid (Acyrthosiphon pisum) feeding, and AM fungus (Rhizophagus intraradices) root colonization on mean salicylic acid (SA) concentration in leaf and petiole tissue. The interaction among exogenous GA, pea aphid feeding, and AM fungus root colonization (GA*PA*AMF) had no statistically significant effect on mean SA concentration according to a three-factor analysis of maximum likelihood using a log-linked gamma distribution (P=0.4278) (a), but the three main effects: exogenous GA (P<0.0001) (b), pea aphid feeding (P=0.0378) (c), and AM fungus root colonization (P<0.0001) (d) were statistically significant according to Wald’s χ2 tests. Values represent the mean ± SEM (n = 4).

Effect of Exogenous Gibberellic Acid 3 and Arbuscular Mycorrhizal Fungus Root Colonization in Modulating Defense Gene Expression in Shoots After 36 Hours of Aphid Feeding

The interaction among exogenous GA, aphid herbivory, and AM fungus root colonization (GA*PA*AMF) did not have a significant effect on MtMYC2 relative gene expression in plant shoots at 36 hours post aphid feeding (Fig. 2.7a; P=0.2933).

Similarly, none of the two-factor interactions had a significant effect on MtMYC2 relative gene expression in plant shoots (Fig. 2.7a). However, the main effect AM fungus root colonization did have a significant effect on MtMYC2 relative gene expression in plant shoots (Fig. 2.7; P=0.0001).

Figure 2.7: Impact of exogenous GA, pea aphid (Acyrthosiphon pisum) feeding, and AM fungus (Rhizophagus intraradices) root colonization on MtMYC2 relative gene expression in Medicago truncatula shoots. The interaction among exogenous GA, pea aphid feeding, and AM fungus root colonization (GA*PA*AMF) had no statistically significant effect on MtMYC2 relative gene expression according to a three-factor analysis of maximum likelihood using a log-linked gamma distribution (P=0.2933) (a), but the main effect AM fungus root colonization (AMF) was statistically significant according to a Wald’s χ2 test (P=0.0001) (d). Values represent the mean ± SEM (n = 3 or 4).

In addition, the interaction among exogenous GA, aphid herbivory, and AM fungus root colonization (GA*PA*AMF) did not have a significant effect on MtVSP relative gene expression in plant shoots at 36 hours post aphid feeding (Fig. 2.8;

P=0.6219). Meanwhile, the three main effects: exogenous GA, aphid herbivory, and AM fungus root colonization did have significant effects on MtVSP relative gene expression in M. truncatula shoots (Fig. 2.6 b, c, d; P<0.0001, P=0.0378, and P<0.0001).

Figure 2.8: Impact of exogenous GA, pea aphid (Acyrthosiphon pisum) feeding, and AM fungus (Rhizophagus intraradices) root colonization on MtVSP relative gene expression in Medicago truncatula shoots. The interaction among exogenous GA, pea aphid feeding, and AM fungus root colonization (GA*PA*AMF) had no statistically significant effect on MtVSP relative gene expression according to a three-factor analysis of maximum likelihood using a log-linked gamma distribution (P=0.2933) (a), but the three main effects: exogenous GA (P=0.0010) (b), pea aphid feeding (P<0.0001) (c), and AM fungus root colonization (P<0.0001) (d) were statistically significant according to Wald’s χ2 tests. Values represent the mean ± SEM (n = 3 or 4).

The interaction among exogenous GA, aphid herbivory, and AM fungus root colonization (GA*PA*AMF) did not have a significant effect on MtPR1 relative gene expression in plant shoots at 36 hours post aphid feeding (Fig. 2.9a; P=0.3434). However, the interaction between exogenous GA and aphid herbivory (GA*PA) regardless of AM

49 status (+/-AMF) did have a significant effect on MtPR1 relative gene expression in plant shoots (Fig. 2.9b; P=0.0357). A post-hoc analysis revealed that plants that received exogenous GA and aphid herbivory regardless of AM status (+/-AMF) differed among the rest of the groups and exhibited the lowest MtPR1 relative gene expression in plant shoots (Fig. 2.9b). Additionally, the interaction between exogenous GA and AM fungus root colonization (GA*AMF) regardless of aphid herbivory (+/-PA) had a significant effect on MtPR1 relative gene expression in plant shoots (Fig. 2.9c; P=0.0449). A post- hoc analysis revealed that MtPR1 relative gene expression was highest in plants with AM fungus root colonization regardless of aphid herbivory (+/-PA) compared to the other treatment groups (Fig. 2.9c). Although the interaction between aphid herbivory and AM fungus root colonization (PA*AMF) regardless of exogenous GA (+/-GA) did not have a significant effect on MtPR1 relative gene expression in plant shoots (Fig. 2.9a;

P=0.3713), the main effects: aphid herbivory and AM fungus root colonization did have significant effects on MtPR1 relative gene expression in M. truncatula shoots (Fig. 2.9 d and e; P=0.0234 and P<0.0001).

Figure 2.9: Impact of exogenous GA, pea aphid (Acyrthosiphon pisum) feeding, and AM fungus (Rhizophagus intraradices) root colonization on MtPR1 relative gene expression in Medicago truncatula shoots. The interaction among exogenous GA, pea aphid feeding, and AM fungus root colonization (GA*PA*AMF) had no statistically significant effect on MtPR1 relative gene expression according to a three-factor anlysis of variance (P=0.2933) (a). The interaction effects of exogenous GA and aphid herbivory (GA*PA) (P=0.0357) (b), and exogenous GA and AM fungus root colonization (GA*AMF) (P=0.0449) (c) were statistically significant according to the two-factor analyses of variance. In addition, two main effects: aphid herbivory (P<0.0001) (d) and

AM fungus root colonization (P<0.0001) (e) were statistically significant according to the Student’s t tests. Values represent the mean ± SEM (n = 3 or 4). Different letters represent significant differences among groups using Tukey-Kramer pot-hoc tests (P<0.05).

Effect of Exogenous Gibberellic Acid 3 and Arbuscular Mycorrhizal Fungus Root Colonization on Defense Gene Expression in Roots After 36 Hours of Aphid Feeding

The overall interaction among exogenous GA, aphid herbivory, and AM fungus root colonization (GA*PA*AMF) did not have a significant effect on MtMYC2 relative gene expression in plant roots at 36 hours post aphid feeding (Fig. 2.10a; P=0.2539).

Likewise, none of the two-factor interactions had significant effects on MtMYC2 relative gene expression in plant roots (Fig. 2.10a). However, two main effects: aphid herbivory and AM fungus root colonization did have significant effects on MtMYC2 relative gene expression in plant roots (Fig. 2.10 b and c; P=0.0002 and P=0.0087).

Figure 2.10: Impact of exogenous GA, pea aphid (Acyrthosiphon pisum) feeding, and AM fungus (Rhizophagus intraradices) root colonization on MtMYC2 relative gene expression in Medicago truncatula root tissue. The interaction among exogenous GA, pea aphid feeding, and AM fungus root colonization (GA*PA*AMF) had no statistically significant effect on MtMYC2 relative gene expression according to a three-factor analysis of maximum likelihood using a log-linked gamma distribution (P=0.2539) (a), but two main effects: aphid herbivory (P=0.002) (b) and AM fungus root colonization (P=0.0087) (c) were statistically significant according to Wald’s χ2 tests. Values represent the mean ± SEM (n = 3 or 4).

The interaction among exogenous GA, aphid herbivory, and AM fungus root colonization (GA*PA*AMF) did not have a significant effect on MtVSP relative gene expression in plant roots at 36 hours post aphid feeding (Fig. 2.11a; P=0.3298). Similarly, none of the two-factor interactions had significant effects on MtVSP relative gene expression in plant roots (Fig. 2.11a). However, the main effect AM fungus root

53 colonization main did have a significant effect on MtVSP relative gene expression in plant roots (Fig. 2.11b; P<0.0001).

Figure 2.11: Impact of exogenous GA, pea aphid (Acyrthosiphon pisum) feeding, and AM fungus (Rhizophagus intraradices) root colonization on MtVSP relative gene expression in Medicago truncatula root tissue. The interaction among exogenous GA, pea aphid feeding, and AM fungus root colonization (GA*PA*AMF) had no statistically significant effect on MtVSP relative gene expression according to a three-factor analysis of maximum likelihood using a log-linked gamma distribution (P=0.3298) (a), but the main effect AM fungus root colonization (P<0.0001) (b) was statistically significant according to a Wald’s χ2 test. Values represent the mean ± SEM (n = 3 or 4).

There was no significant effect of the interaction among exogenous GA, aphid herbivory, and AM fungus root colonization (GA*PA*AMF) on MtPR1 relative gene expression observed in plant roots at 36 hours post aphid feeding (Fig. 2.12a; P=0.7807).

Additionally, none of the two-factor interactions had significant effects on MtPR1 relative gene expression in plant roots (Fig. 2.12a). However, the main effect AM fungus root colonization did have a significant effect on MtPR1 relative gene expression in plant roots (Fig. 2.12b; P<0.0001).

Figure 2.12: Impact of exogenous GA, pea aphid (Acyrthosiphon pisum) feeding, and AM fungus (Rhizophagus intraradices) root colonization on MtPR1 relative gene expression in Medicago truncatula root tissue. The interaction among exogenous GA, pea aphid feeding, and AM fungus root colonization (GA*PA*AMF) had no statistically significant effect on MtPR1 relative gene expression according to a three-factor analysis of variance (P=0.7807) (a), but the main effect AM fungus root colonization was statistically significant according to a Student’s t test (P<0.0001) (b). Values represent the mean ± SEM (n = 3 or 4).

Effect of Exogenous Gibberellic Acid 3 and Aphid Herbivory on Fungus Root Colonization Markers After 36 Hours of Aphid Feeding

The interaction between exogenous GA and aphid herbivory (GA*PA) had no significant effect on RiEF relative gene expression in plant roots 36 hours after aphid feeding (Fig. 2.13a; P=0.0998). The two main effects: exogenous GA and aphid herbivory had no significant effects on RiEF relative gene expression in plant roots (Fig.

2.13a; P=0.5835 and P=0.4338). The interaction between exogenous GA and aphid herbivory (GA*PA) also had no significant effect on MtLec5 relative gene expression in plant roots (Fig. 2.13b; P=0.2712). Likewise, the two main effects: exogenous GA and

55 aphid herbivory had no significant effects on MtLec5 relative gene expression in plant roots (Fig. 2.13b; P=0.03424 and P=0.9817). The interaction between exogenous GA and aphid herbivory (GA*PA) had a significant effect on MtPT4 relative gene expression in plant roots (Fig. 2.13c; P=0.0335), however a post-hoc analysis revealed that there were no differences among the treatment groups (Fig. 2.13c).

Figure 2.13: Impact of exogenous GA, pea aphid (Acyrthosiphon pisum) feeding, and AM fungus (Rhizophagus intraradices) root colonization on RiEF, MtLec5, and MtPT4 relative gene expression on mycorrhizal Medicago truncatula roots. The interaction between exogenous GA, and pea aphid feeding (GA*PA) had no statistically significant effect on RiEF (a) or MtLec5 (b) relative gene expression (P=0.2933) (a), but this interaction did have a statistically significant effect on MtPT4 relative gene expression (c) according to the two-factor analyses of variance (P=0.0335) (c). No differences in MtPT4 relative gene expression were observed among treatment groups

56 according to Tukey-Kramer post-hoc tests (P>0.05). Values represent the mean ± SEM (n = 3 or 4).

Effect of Exogenous Gibberellic Acid 3 Application and Arbuscular Mycorrhizal Fungus Root Colonization on Plant Shoot and Root Fresh Weight After 7 Days of Aphid Feeding

Overall, the interaction among exogenous GA, aphid herbivory, and AM fungus root colonization (GA*PA*AMF) did not have a significant effect on M. truncatula shoot fresh weight 7 days post aphid feeding (Fig. 2.14a; P=0.3086). The interaction between exogenous GA and aphid herbivory (GA*PA) regardless of AM status (+/-AMF) did have a significant effect on plant shoot fresh weight (Fig. 2.14b; P=0.0111). Also, the interaction between exogenous GA and AM fungus root colonization (GA*AMF) regardless of aphid herbivory (+/-PA) did have a significant effect on plant shoot fresh weight (Fig. 2.14c; P<0.0001). The interaction between aphid herbivory and AM fungus root colonization (PA*AMF) regardless of exogenous GA (+/-GA) did not have a significant effect (Fig. 2.14a; P=0.0894), however, the two main effects: aphid herbivory and AM fungus root colonization did have significant effects on plant shoot fresh weight

(Fig. 2.14 d and e; P<0.0001 and P<0.0001).

Figure 2.14: Impact of exogenous GA, pea aphid (Acyrthosiphon pisum) feeding, and AM fungus (Rhizophagus intraradices) root colonization on Medicago truncatula shoot fresh weight at 7 days post-aphid feeding. The interaction among exogenous GA, pea aphid feeding, and AM fungus (GA*PA*AMF) did not have statistically significant effects on shoot fresh weight according to the three-factor analysis of maximum likelihood (P=0.3086) (a), but the interactions of exogenous GA and aphid herbivory

(GA*PA) (P=0.0111) (b) and exogenous GA and AM fungus (GA*AMF) (P<0.0001) (c) were statistically significant according to two-factor analyses of maximum likelihood using log-linked gamma distributions. In addition, two main effects: aphid herbivory (P<0.0001) (d) and AM fungus colonization (P<0.0001) (e) were statistically significant according to Wald’s χ2 tests. Values represent the mean shoot fresh weight ± SEM (n= 6 to 8) per treatment. Different letters represent significant differences among treatment groups using Tukey-Kramer post-hoc tests (P<0.05).

The interaction among exogenous GA, aphid herbivory, and AM fungus root colonization (GA*PA*AMF) did not have a significant effect on plant root fresh weight 7 days post aphid feeding (Fig. 2.15a; P=8616). The interactions of exogenous GA and aphid herbivory (GA*PA) regardless of AM root colonization (+/-AMF), and aphid herbivory and AM fungus root colonization (PA*AMF) regardless of exogenous GA (+/-

GA) did have significant effects on plant root fresh weight 7 days post aphid feeding

(Fig. 2.15 b and c; P=0.0488 and P=0.0440). However, a post-hoc analysis revealed that there were no differences among the groups in the interaction between aphid herbivory and AM fungus root colonization (PA*AMF) regardless of exogenous GA (+/-GA) (Fig.

2.15c).

Figure 2.15: Impact of exogenous GA, pea aphid (Acyrthosiphon pisum) feeding, and AM fungus (Rhizophagus intraradices) root colonization on Medicago truncatula root fresh weight for 7 days post-aphid feeding. The interaction among exogenous GA, pea aphid feeding, and AM fungus (GA*PA*AMF) did not have statistically significant effects on root fresh weight according to the three-factor analysis of variance (P=0.8616) (a), but the interactions of exogenous GA and aphid herbivory (GA*PA) (P<0.0488) (b) and aphid herbivory and AM fungus root colonization (PA*AMF) (P=0.0440) (c) were statistically significant according to two-factor analyses of maximum likelihood using a log-linked gamma distributions. Values represent the mean shoot fresh weight ± SEM (n= 6 to 8) per treatment. Different letters represent significant differences among treatment groups using Tukey-Kramer pot-hoc tests (P<0.05).

Effect of Exogenous Gibberellic Acid 3 on Pea Aphid Count Per Colony and Colony Total Weight 7 Days After Feeding

The interaction between exogenous GA and AM fungus root colonization

(GA*AMF) did have a significant effect on both mean aphid count per colony and mean aphid colony weight 7 days post aphid feeding (Fig 2.16 a and b; P=0.0010 and

P=0.0003). Mean aphid colony count and mean aphid colony weight were higher for aphids that fed continuously for 7 days on plants that were colonized by AM fungi without receiving GA compared to the other treatment groups, whereas these two fitness parameters had the lowest values for aphids that feed on non-mycorrhizal plants without receiving GA (Fig. 2.16a and b). However, mean aphid count per colony and mean aphid colony weights for aphids that fed continuously for 7 days on non-mycorrhizal plants did not differ from aphids that fed on non-mycorrhizal plants or mycorrhizal plants that received GA. In addition, aphid mean count per colony and mean aphid colony weight for aphids that fed continuously for 7 days on mycorrhizal plants that received GA did differ from aphids that fed on non-mycorrhizal only and mycorrhizal only plants.

Figure 2.16: Impact of exogenous GA and AM fungus (Rhizophagus intraradices) root colonization on mean pea aphid (Acyrthosiphon pisum) count per colony and mean colony weights 7 days post-aphid feeding on Medicago truncatula plants. The interaction between exogenous GA and AM fungus (GA*AMF) had a statistically significant effect on mean pea aphid count per colony according to the two-factor analysis of maximum likelihood using negative binomial distributions with scaled Pearson χ2 (P=0.0010) (a). The interaction between exogenous GA and AM fungus (GA*AMF) had a significant effect on mean pea aphid colony weights (P=0.0003) according to the two-factor analysis of maximum likelihood using a gamma distribution with scaled Pearson χ2 (P=0.0003) (b). Values represent the mean aphid count per colony or mean aphid colony weight ± SEM (n= 6 to 8) per treatment. Different letters represent significant differences among treatment groups using Tukey-Kramer post-hoc tests (P<0.05).

Discussion

In the present study, we investigated the role of exogenous GA in regulating plant defenses during AM fungi-plant-aphid interactions and tested the following hypotheses:

1) aphid fitness would be positively impacted by exogenous GA application of plants, 2) exogenous GA application would result in reduced arbuscule formation, 3) JA levels and

JA-regulated defense gene expression would be downregulated in response to exogenous

GA application, and 4) SA levels and SA-regulated defense gene expression would be upregulated in response to exogenous GA application.

As expected, aphid fitness parameters (abundance and aphid colony weight) were not impacted by the interaction between exogenous GA and AM fungus root colonization after 36 hours of feeding, although a trend showing a slightly higher mean colony weight was observed on non-mycorrhizal plants that received exogenous GA (Fig. 2.3).

Nevertheless, the interaction between exogenous GA and AM fungus root colonization had a significant impact on both aphid fitness parameters measured after 7 days of continuous feeding (Fig. 2.16). The data agree with previous studies showing that pea aphids benefit from feeding on plants that are highly colonized by AM fungi compared to non-mycorrhizal control plants (54, 59). This study revealed that aphids are affected differently when they feed on non-mycorrhizal and mycorrhizal plants that are treated with GA, indicating that GA signaling may be involved in regulating plant defenses during AM fungi-plant-aphid interactions. Aphid abundance and colony weight after feeding on mycorrhizal plants receiving GA were not as high as on mycorrhizal plants without GA. Even though the data did not support the hypothesis that aphid fitness would be positively impacted by exogenous GA application to plants, the results indicate that

GA regulates AM fungus-plant-aphid interactions. In addition, there is evidence suggesting that exogenous GA application can be cytotoxic in high concentrations against the melon fruit fly (Bactrocera cucurbitae (Coquillett), Egyptian cotton leafworm larvae

(Spodoptera littoralis) and migratory locust (Locusta migratoria migratoria) and negatively affect fall armyworm (Spdoptera frugiperda) food consumption and female oviposition (160-163). We did not find clear evidence of cytotoxicity against aphids due to exogenous GA when comparing non-mycorrhizal plants only and GA treated non-

63 mycorrhizal plants. However, aphids that fed on mycorrhizal plants not treated with GA did better than those on mycorrhizal plants treated with GA.

The interaction between exogenous GA and AM fungus root colonization had an effect on GA3 levels in leaves and petioles at 36 hours post aphid feeding. GA3 levels were highest in mycorrhizal plants that were treated with GA compared to plants that did not receive GA, whereas the GA3 levels in non-mycorrhizal plants that received GA did not differ from the other treatments. Likewise, the interaction between exogenous GA and AM fungus root colonization had an effect on JA levels in leaves and petioles. JA levels were high in plants that were treated with exogenous GA regardless of aphid herbivory but did not differ from mycorrhizal plants regardless of aphid herbivory (Fig.

2.5b). Although the SA levels were the highest in leaves and petioles compared to the levels of JA and GA3, only the main effects had a significant impact on SA levels. Plants that received GA, plants exposed to aphid feeding, and non-mycorrhizal plants accumulated more SA in leaves and petioles.

Only the main effect AM fungus root colonization had a significant effect on

MtMYC2 relative gene expression in shoots at 36 hours post aphid feeding. Interestingly, the treatments that were colonized by AM fungi had higher levels of MtMYC2 relative gene expression compared to the non-mycorrhizal treatments. In the case of the JA- regulated defense gene MtVSP, the three main effects (exogenous GA, pea aphid feeding, and AM fungus root colonization) had a significant impact on MtVSP relative gene expression. Plants that received GA, plants exposed to aphid feeding, and mycorrhizal plants had higher levels of MtVSP relative gene expression.

Increased JA levels, upregulated MtVSP mean relative gene expression, and lack of MtMYC2 downregulation in shoots by exogenous GA indicates that GA signaling may play a positive role in regulation of plant defenses during plant-insect interactions. This does not agree with previous plant-pathogen studies indicating that GA signaling suppresses JA signaling (1, 85, 86, 100, 164) However, one study indicates that GA and

JA signaling work synergistically to induce TERPENE SYNTHASE genes (TPS11 and

TPS21) in A. thaliana plants (165). Here, DELLAs negatively influenced MYC2 regulation of TPS11 and TPS21, meanwhile repression of DELLAs induced TPS11 and

TPS21 gene expression, therefore indicating that GA signaling can have a positive association with JA signaling (165). In addition, the researchers found that production of the plant volatile (E)-β-caryophyllene required the participation of GA signaling and JA signaling (165). (E)-β-caryophyllene has been found to aid in plant defenses against sucking insects, including aphids directly and indirectly (166-168). However, (E)-β- caryophyllene production has been found to be decreased in mycorrhizal fava beans plants that were fed on by pea aphids (54). Therefore, the GA-mediated increased JA levels and MtVSP induction in conjunction with the lack of MtMYC2 reduction may be due to a synergistic effect of exogenous GA and JA signaling, however further investigation is needed.

Mean SA concentration in shoot tissue was positively impacted by exogenous GA and aphid feeding but was negatively impacted by AM fungus root colonization 36 hours post aphid feeding (Fig. 2.6b, c, and d). Contrastingly, MtPR1 mean relative gene expression in plant shoots was positively impacted by AM fungus root colonization, meanwhile exogenous GA antagonized AM fungus mediated MtPR1 relative gene

65 expression (Fig. 2.9c). Additionally, aphid herbivory positively impacted both mean SA concentration and MtPR1 relative gene expression in shoots 36 hours post aphid feeding

(Fig. 2.6c and Fig. 2.9d). This is consistent with other research that found that pea aphid feeding triggers SA accumulation 48 h after aphid feeding in M. truncatula leaf tissue

(138). However, the present study indicate that AM fungi and aphids may alter SA- regulated defense signaling through separate mechanisms. Future research should focus on SA biosynthesis, upstream SA signaling mechanisms, and other SA-related defense genes.

Belowground defenses were mostly dictated by presence of AM fungus root colonization. Like aboveground defense gene expression, MtMYC2, MtVSP, and MtPR1 mean relative gene expression in root tissue were positively impacted by AM fungus root colonization 36 hours post aphid feeding (Fig. 2.10, Fig 2.11, and Fig. 2.12). MtMYC2 gene expression only differed in that aphid herbivory upregulated belowground MtMYC2 mean relative gene expression (Fig. 2.10c). Interestingly, the mean relative gene expression for arbuscule-specific and AM-specific genes (MtPT4 and MtLec5) were not impacted by exogenous GA or aboveground aphid herbivory (Fig. 2.13), indicating that once arbuscules developed in a root system, they are unaffected by either treatment.

Previous research has found that exogenous GA treatment starting at 6 days post inoculation (dpi) prevents arbuscule development in plant roots, and that pea aphid feeding can negatively impact AM fungus colonization of roots (43, 54, 59). In addition, we used the RiEF gene to quantify and compare fungal biomass across treatments.

Although fungal biomass was not significantly impacted by exogenous GA in this study, this may be due to hyphal growth throughout the root length. Previous research has

66 shown that the AM fungal phenotype in plants that received exogenous GA treatment at 6 dpi is the same as in Mtdella1/Mtdella2 mutant plants. In both cases, arbuscule development is severely decreased, meanwhile there were no differences between wild type and Mtdella1/Mtdella2 in root length colonization (43). This may be due to increased hyphal branching, which was found to be increased by exogenous GA treatment, meanwhile arbuscules were nearly undetectable (169). In the present study,

GA treatments were started at 7 dpi, root length colonization was higher, AM fungal species, the substrate and Hoagland’s fertilizer were different which could account for the differences in the results. In addition, we inoculated plants that were 22 days old, meanwhile previous research inoculated 2 day old plants, which may have impacted our results (43). Our results may also vary compared to Floss et al. in that we fertilized plants twice a week, whereas the researchers fertilized once a week (43). In contrast to the present study, exogenous GA was also applied in one-tenth-strength Hoagland’s solution

(169). This extra day of fertilizer instead of GA treatment in the present study may have allowed for arbuscules to develop. Although GA has typically been characterized as a negative regulator of arbuscule development, it can positively influence AM fungal development. For instance, it was found in another model legume Lotus japonicus that inhibition of GA biosynthesis negatively affects AM fungal development (169). It has also been recently found that exogenous GA may affect different AM types differently

(170). For example, the AM fungus Rhizophagus irregularis forms Arum-type with model legumes but can exist as Paris-type in slow growing plants (171). Arum-type mycorrhizas are characterized by intercellular hyphal growth and arbuscules, meanwhile

Paris-type are characterized by intracellular hyphal growth coils, and arbuscules (171).

Arum-type are commonly found in association with crop plants. Exogenous GA positively impacts AM fungal establishment of R. irregularis in the flowering plant

Eustoma grandiflorum by promoting hyphopodia and subsequent arbuscule formation

(170). In order to determine the impact of exogenous GA on arbuscule development during tripartite interactions, future studies should consider coupling molecular markers with measurements of arbuscule density via microscopy (43).

The interaction between exogenous GA and AM fungus root colonization did have a significant effect on plant shoot fresh weight 36 hours post aphid feeding (Fig.

2.1). Mycorrhizal plants and mycorrhizal plants that received GA had the highest shoot fresh weight means and did not differ from one another, meanwhile non-mycorrhizal plants that received exogenous GA had higher shoot fresh weight than non-mycorrhizal plants that did not receive exogenous GA (Fig. 2.1b). Plant root fresh weight 36 hours post aphid feeding was unaffected by all treatments (Fig 2.2). The interaction between exogenous GA and aphid herbivory regardless of AM fungus colonization did have a significant impact on mean plant shoot fresh weight 7 days post aphid feeding. Shoot fresh weight of plants that received exogenous GA and aphid herbivory regardless of mycorrhizal status was lower than the weight in other groups (Fig. 2.14b). In addition, the interaction between exogenous GA and AM fungus colonization regardless of aphid herbivory had an impact on plant shoot fresh weight 7 days post aphid feeding (Fig.

2.14c). Mycorrhizal only plants exhibited the highest mean shoot fresh weight.

Mycorrhizal plants that received exogenous GA had lower mean shoot fresh weight than plants colonized with AM fungi only but were still higher than plants receiving exogenous GA only and plants that did not receive either treatment (Fig. 2.14c). This

68 may indicate that the AM fungus-plant symbiosis was interrupted regardless of aphid feeding. Mean shoot fresh weight was reduced by aphid herbivory, meanwhile mean shoot fresh weight was increased by AM fungus root colonization. Similar to previous research, our results indicate that AM fungi increase plant shoot biomass (19) (Fig. 2.14d and e). Plant root fresh weight 36 hours post aphid feeding was unaffected by all treatments. The interaction between exogenous GA and aphid herbivory regardless of

AM fungus root colonization significantly impacted mean root fresh weight 7 days post aphid feeding. All groups were the same except for roots from plants that received both exogenous GA and aphid herbivory, which were lower than the rest. In addition, the interaction of aphid herbivory and AM fungus root colonization regardless of exogenous

GA did impact mean root fresh weight. However, there were no differences among the groups.

The present study revealed that GA may play a role in the regulation of plant defenses during AM fungi-plant-aphid interactions. The data supported previous research indicating that MIR can promote aboveground and belowground defense signaling through JA- and SA-related defense gene expression, meanwhile AM fungus root colonization negatively impacted shoot SA levels and had a negligible impact on JA levels. The data did not support the hypotheses that exogenous GA would negatively impact JA levels as well as downregulate JA-related defense gene expression.

Surprisingly, the opposite was observed, that exogenous GA positively impacted shoot

JA levels and may upregulate JA-related defense gene expression, such as MtVSP.

Although shoot SA levels were increased by exogenous GA, exogenous GA seemed to have an antagonizing effect on MtPR1 regulation. In addition, the data did not support the

69 hypothesis that aphid fitness improves when feeding on plants treated with exogenous

GA. Finally, the data showed that exogenous GA treatment did not affect arbuscule presence or fungal biomass based on plant and fungal marker genes. This study serves as a focal point for future research seeking to elucidate the complex role of GA in regulating plant defenses in plant-aphid and AM fungus-plant-aphid interactions.

CHAPTER III

ROLE OF GIBBERELLIC ACID SIGNALING IN MODULATING PLANT DEFENSE RESPONSES DURING PLANT- APHID INTERACTIONS

Abstract

Since plants cannot evade immediate abiotic and biotic stresses, they rely on intricate regulatory mechanisms driven by phytohormones to prioritize growth or defense. Specifically, the phytohormones jasmonic acid (JA) and salicylic acid (SA) initiate appropriate defense signaling against pathogens and insect pests, however, aphids, can modify plant defenses in their favor. Nevertheless, another phytohormone, gibberellic acid (GA) can modulate plant defenses through interactions with DELLA proteins. DELLAs are GA transcriptional repressors with diverse protein-protein interaction properties that are degraded upon GA perception, therefore modulating other signaling pathways. The present study used two genotypes of Medicago truncatula plants, segregant wild type (WT) and Mtdella1/Mtdella2 mutant; and the pea aphid

(Acyrthosiphon pisum) to evaluate the role of GA signaling in mediating plant defense responses after 36 hours and 7 days of aphid feeding. The results indicate that 36 hours of pea aphid feeding resulted in downregulation of gene expression of the JA-regulated transcription factor MYC2 in Mtdella1/Mtdella2 shoots compared to aphid-infested WT shoots. Overall, gene expression of the JA-regulated defense gene VEGETATIVE

STORAGE PROTEIN (VSP) was reduced in Mtdella1/Mtdella2 shoots compared to WT

71 shoots. Gene expression of the SA-regulated gene PATHOGENESIS RELATED (PR) 1 did not differ between plant genotypes whether they were aphid-infested or not. The aphid fitness parameters measured in this study were not altered by feeding for 36 hours and 7 days in WT and Mtdella1/Mtdella2 shoots. In summary, we provide intriguing evidence showing that GA signaling may be involved in JA-regulated defense gene expression during M. truncatula - pea aphid interactions.

Introduction

Arthropod herbivores such as aphids can contribute up to 20% of agricultural losses worldwide (172, 173). Aphids are a diverse group (over 4000 species) of phloem- feeders that can decrease crop yields and exhibit insecticide resistance rather quickly (62,

64, 118). In addition, aphids serve as vectors of approximately 275 plant viruses of economic importance because these viral diseases lead to devastating yield losses worldwide (62, 66). Aphids can be specialists, such as the Russian wheat aphid

(Diuraphis noxia), that feed only on few select plant species or a single plant family, meanwhile generalists, such as the cotton or melon aphid (Aphis gossypii) feed on broad groups of plant families (63, 64). Most aphids feed on phloem sap through their modified mouthparts called stylets, which pierce into plant tissue and travel intercellularly to feed on plant sieve elements to obtain plant photoassimilates (65). Aphid feeding relies on two separate types of salivation, a gelling saliva, and a watery saliva to influence host susceptibility (4, 65, 174). First, the gelling saliva provides a lubricating sheath to allow movement to the plant sieve elements through the plant apoplast, meanwhile minimizing mechanical plant damage and avoiding activating plant wound responses (65).

Meanwhile, the watery saliva serves as a secondary measure to modify plant defense responses and contains biochemically active proteins (e.g. pectinases, cellulases, oxidases, and lipases) as well as other compounds that may influence plant susceptibility to aphid infestation, which is produced after the stylet has reached the sieve elements (4,

65, 130, 174, 175).

Plants can defend against aphid feeding through a mechanisms known as sieve element occlusion (SEO), which can plug and seal areas of sieve elements where aphid stylets have penetrated (65). Initially, plants rely on a quick plugging mechanism by phloem (P) proteins in Brassicaceae plants or the structurally similar proteins known as forisomes in leguminous plants, which are encoded by the SEO gene family (176). In leguminous plants, such as fava beans (Vicia faba), forisome activation can be triggered by aphid feeding and can negatively affect feeding behavior of the potato aphid (Myzus persicae) and the vetch aphid (Megoura viciae), meanwhile pea aphid (Acyrthosiphon pisum) feeding can avoid forisome activation (174, 177, 178). In addition, it was found that vetch aphids can reverse forisome activation by injecting watery saliva into the sieve element, therefore preventing SEO (174). Another aspect of SEO is callose deposition.

Callose is a β-1-3 glucan that is synthesized at the plasma membrane and contributes to plant resistance to several aphid species by acting as a sealing mechanism to inhibit aphid feeding (65). However, aphid salivation can also desist callose deposition and may not be a suitable defense mechanism against growing numbers of aphid invaders (65). Although aphids can avoid SEO through compounds in their watery saliva, there are other modes of plant defenses that aphids must evade as well (4).

The front-line of plant innate immunity is triggered by the plant perception of conserved molecular patterns known as pathogen- or microbe-associated molecular patterns (PAMPs or MAMPs) (3, 5, 179). For instance, plants can sense bacterial flagellin or fungal chitin through cell surface receptors known as pattern recognition receptors (PRRs), which trigger signaling cascades that promote PAMP-triggered immunity (PTI) (3, 5, 179). Plants can also recognize and activate resistance to insect herbivores through herbivore-associated molecular patterns (HAMPs) found in insect oral secretions or oviposition fluids (74, 179). In addition, chewing and piercing from insect herbivory causes mechanical damage that can activate plant innate immunity through plant perception of damage-associated molecular patterns (DAMPs) (74, 79, 179).

Nevertheless, pathogenic microbes and insects can evade PTI by releasing molecules or proteins known as effectors that can attenuate appropriate defense responses and promote host susceptibility (3, 5, 179). Consequently, plants can maintain resistance to pathogens and insects through resistance (R) genes. A common feature of R genes is that most encode nucleotide-binding leucine rich repeat (NB-LRR) receptor proteins that sense specific effectors and enhance host resistance (3, 5, 179). Detection of effectors triggers a robust and longer-lasting hypersensitive immune response known as effector-triggered immunity (ETI) (3, 5, 179). For the most part, R genes have been associated with plant innate immunity against pathogens, however there are some that confer resistance against aphid herbivory (180-184). For example, it has been demonstrated that Medicago truncatula resistance to the bluegreen aphid (Acyrthosiphon kondoi) and its close relative the pea aphid (Australian biotype) is conferred by the R genes Acyrthosiphon kondoi resistance (AKR) and Acyrthosiphon pisum resistance (APR) (182). Additionally, both R

74 genes are located in chromosomal regions rich in NB-LRRs (183, 185). In addition to

AKR and APR, two other R genes, Mi-1.2 and virus aphid transmission (Vat), confer resistance to potato aphids and melon aphids respectively (180, 181, 184, 186, 187).

Together these indicate that NB-LRRs may play an important role in aphid resistance, however, specific elicitors have yet to be identified, and these NB-LRRs may be aphid species or biotype specific (65).

Plant immune responses are predominately regulated by the phytohormones jasmonic acid (JA) and salicylic acid (SA) (49, 73, 79, 188). It is known that the JA and

SA pathways interact, sometimes resulting in antagonism between these pathways. SA signaling has been shown to support host resistance to biotrophic pathogens (49, 179), meanwhile, JA signaling is critical for defense responses against necrotrophic pathogens and a wide-range of insect herbivores, including chewing and phloem-feeding insects

(73, 74, 78, 79, 188, 189). Although the mechanisms driving aphid suppression of plant defenses is still poorly understood, it has been demonstrated that JA signaling induces direct and indirect plant defenses against aphids and that suppression of JA signaling can promote aphid herbivory (130, 132-137, 179). Although SA has been linked with aphid suppression of plant immunity, there is evidence suggesting that SA signaling can promote basal plant immunity against aphid herbivory (70, 180, 181, 190-194). For instance, it has been found in tomato that SA signaling regulates Mi-1.2 gene mediated resistance to potato aphids (180, 181). In addition, SA signaling can also confer resistance to aphids through regulation of PHYTOALEXIN DEFICIENT4 (PAD4) (192,

195, 196). PAD4 encodes a nucleocytoplasmic protein that not only is involved with pathogen resistance, but also plays an important role in basal resistance to the green

75 peach aphid (Myzus persicae (Sülzer)) through multiple mechanisms, including callose deposition (65, 192, 195, 196). In addition PAD4 expression is a key component of a positive regulatory SA feedback loop that negatively regulates MYC2 expression, indicating that basal resistance to aphids may be regulated by SA (84, 196). Although JA and SA are key phytohormones involved in modulating plant defenses, other phytohormones may also regulate plant-insect interactions.

In recent years, the phytohormone gibberellic acid (GA) has gained attention due to its new role in altering the activity of the JA and SA signaling pathways, specifically during plant-pathogen interactions (1, 73, 85, 86, 101-103). In summary, GA signaling has been found to enhance plant resistance to biotrophic pathogens, and promote susceptibility to necrotrophic pathogens by negatively regulating JA signaling (1, 77, 85,

98, 100-102, 143-145). GA signaling is characterized by a class of diterpenoid carboxylic acids known as gibberellins and plays important roles in many aspects of plant growth and development (98, 144, 145). Initially, GA signaling is repressed by GRAS domain transcriptional repressor proteins known as DELLAs, which are characterized by their unique N-terminus DELLA and TVHNYP domains (77, 145). GA signaling first requires gibberellin perception by the receptor GIBBERELLIN INSENSITIVE DWARF (GID) 1

(98). This interaction forms a complex and is subsequently transported to the nucleus where DELLAs are recruited. gibberellin-GID1-DELLA interactions promote DELLA polyubiquitination and degradation via the 26S proteasome pathway, therefore, relieving

GA transcription factors and activating GA-regulated gene expression (77). Moreover,

DELLAs are known to have diverse protein-protein interaction capabilities allowing them to regulate other pathways such as the JA signaling pathway (77, 80).

DELLAs have been found to confer plant resistance to necrotrophic pathogens by acting as important transactivating proteins in the JA signaling pathway (1, 85, 102).

DELLAs regulate JA signaling by releasing the master regulatory transcription factor

MYC2 from its transcriptional repressors, JASMONATE ZIM DOMAIN (JAZ) proteins

(80). Induction of GA leads to DELLA degradation, which negatively affects JA signaling, thus conferring resistance to biotrophic pathogens, while decreasing resistance to necrotrophic pathogens (1, 85, 143). Although GA plays a key role in modulating plant defense signaling during plant-pathogen interactions, its role in plant-insect interactions remains largely unknown. It has been reported that beet armyworm labial saliva can only modulate JA levels in Arabidopsis thaliana wild type plants compared to quadruple della mutants, indicating that DELLAs may act as a central regulatory signaling node in this interaction (107). It has also been found that GA decreases rice (Oryza sativa) resistance against root-knot nematodes (Meloidogyne graminocola) through DELLA degradation

(105). On the other hand, another study found that GA hypersensitive rice plants overexpressing OsGID1 increased rice resistance to the brown planthopper (Nilaparvata lugens) (106). Although GA signaling has been reported to regulate plant-insect interactions, to our knowledge, there are no studies that investigated the role of GA signaling in plant-aphid interactions.

In the present study, the role of GA signaling in regulating plant defenses during plant-aphid interactions was evaluated by using the model legume Medicago truncatula and pea aphids (Acyrthosiphon pisum). The following hypotheses were tested: 1) aphid fitness will be improved after feeding on Mtdella1/Mtdella2 mutants compared to wild type plants, and 2) JA-regulated defense gene expression will be downregulated in

Mtdella1/Mtdella2 mutants. Pea aphids are notorious agricultural pests that feed on many leguminous plants in the Fabaceae family and spread important plant viruses, such as the

Pea enation virus and Bean leafroll virus (63). In addition, pea aphids have been reported to promote SA levels in M. truncatula (116, 197), indicating that pea aphids may modulate JA and SA to promote host susceptibility.

Materials and Methods

Plant Growth Conditions

Medicago truncatula segregant wild type (WT) and Mtdella1/Mtdella2 seeds were kindly provided by Dr. Maria Harrison (Boyce Thompson Institute for Plant

Research, Ithaca, NY). The seed sterilization and germination procedures followed a protocol outlined in Maurya et al. (59). Seeds were scarified for 10 minutes in concentrated H2SO4, rinsed in sterile water, sterilized for 10 minutes using 10% (v/v) household bleach in 0.1% (v/v) Tween 20 solution, and rinsed in sterile water (59). Seeds were spread on wet filter paper (sterile) in petri dishes, dishes were sealed with parafilm, and wrapped in aluminum foil. Dishes were incubated at 4°C for three days (dark), set in room temperature (24°C) for one day (dark), aluminum foil was removed and dishes were then set under indirect light (mean: 184 µmol m-2 s-1) for three days. Seedlings were then planted in azalea pots (12 cm W x 8.5 cm H) with a sterilized sand: topsoil mix (9:1)

(Pioneer Sand Company, Windsor, CO). All plants used in the experiment were grown in a growth chamber under the following conditions: 16 h photoperiod, 24°C, 40% relative humidity, and light intensity of 285-290 µmol m-2 s-1 provided by fluorescent and halogen bulbs. The sand was thoroughly washed with tap water, and topsoil was sieved

78 prior to autoclaving three times each substrate (each cycle: 60 minutes, 121°C, and

15psi). The sterile sand: topsoil mixture was saturated with ½ strength modified

Hoagland’s solution (500 µM P, 15 mM N, pH 6.1) prior to transplant (146). Azalea pots were covered with clear plastic humidity domes (54.6 cm H×28 cm W×17.8 cm D) for one week after transplant. Once domes were removed, plants in each pot were watered with 50 mL of Milli-Q® water five days a week and fertilized twice a week with 50 mL

½ strength modified Hoagland’s solution (500µM P, 15mM N, pH 6.1) (146). After 15 days, seedlings with two or three trifoliolate leaves were transplanted into smaller pots

(one seedling per pot) (6.35 cm W x 9 cm H) for the remainder of the experiment. Pots were covered with clear plastic humidity domes (54.6 cm H×28 cm W×17.8 cm D) for one week after transplant. Every pot was watered daily with 17.5mL Milli-Q® water or received fertilizer twice a week with 50 mL ½ strength modified Hoagland’s solution

(500µM P, 15mM N, pH 6.1) (146).

Pea Aphid Infestation

Parthenogenetic, wingless female pea aphids that were kindly provided by Dr.

Kenneth Korth (University of Arkansas, Fayetteville, AR) were reared on fava beans (V. fava) plants in insect tents under laboratory conditions under a 16-h photoperiod. One- thousand six-day old aphids were synchronized (197) by adding wingless adult aphids to a separate insect tent with non-infested fava beans. The adults were removed the following day leaving only nymphs that were reared on fava beans for additional five days. Twenty eight days after plants were transplanted into smaller pots (6.35 cm W x 9 cm H pots), 15 six-day old aphids were added to experimental plants (197). Harvests took place in the middle of the subjective day (10:00 h – 16:00 h) when jasmonate levels peak

(149). One replicate of every treatment was harvested at the same time to account for changes in gene expression that are controlled by circadian rhythms.

In the present study, there were four treatments: 1) segregant WT /-Pea aphid; 2) segregant WT / +Pea aphid; 3) Mtdella1/Mtdella2 mutant / -Pea aphid; and 4)

Mtdella1/Mtdella2/ +Pea aphid. After receiving aphids, every plant including plants without aphids were caged with white organza drawstring gift bags (15.24 x 22.86 cm,

SumDirect). Aphids fed continuously on plants for 36 hours (197) and 7 days (59). At the end of each feeding period, aphids were collected from each plant using a mild vacuum device and were immobilized at -20°C prior to counting and weighing them. Non- infested plants were also exposed to the vacuuming effect. The vacuum device consisted of a 4.8 mm diameter Tygon tubing, connected to a 50 mL conical tube (Fisher

Scientific), and a cut 200 µL pipette tip. Plant fresh weights for roots and shoots were measured prior to freezing the tissue in liquid nitrogen and storage at -80°C. Plant dry weights were recorded for plants seven days post aphid feeding. Four randomly selected plants from each treatment from the seven-day time-point were measured for shoot and root dry weights. Selected samples were incubated at 60°C for three days prior to measuring. Dry weights were then compared with their respective fresh weights to calculate average percent weight change for each treatment. Average percent weight change was then used to calculate dry weights for all other shoot and root samples.

Ribonucleic Acid Isolation and Synthesis of Complementary Deoxyribonucleic Acid

RNA isolation and cDNA synthesis followed the protocols described previously

(151). Three biological replicates were selected randomly within each of the treatments

80 for gene expression analyses. Shoot tissues were ground using a mortar and pestle in liquid nitrogen, and RNA was extracted using the RNeasy Plant Mini Kit according to the manufacturer’s instructions (Qiagen Inc., Germantown, MD, USA). RNA samples were

DNAse-treated by adding 87µL of nuclease-free water, 10 µL of 10X reaction buffer, and

3 µL of TurboTM DNase (2 units µL-1) and were incubated at 37°C for 40 minutes.

DNAse-treated RNA samples were purified using the RNeasy MinElute Cleanup kit

(Qiagen Inc.). Samples were further treated for genomic DNA contamination using a

DNA-freeTM kit (Thermo Fisher Scientific). For cDNA synthesis, 1 µg of total RNA was

-1 mixed with 1 µL dNTPs (10 mM each) and 1 µL anchored oligo dT22 (500 ng µL ) and incubated at 65°C for 5 minutes using a T100™ Thermal Cycler (Bio-Rad Laboratories

Inc, Hercules, CA). Following this step, 4 µL of SuperScript® IV Buffer, 1.2 µL

Nuclease-free water, 1 µL of DTT (100 M), 0.5 µL of RNaseOUTTM (Thermo Fisher

Scientific), and 0.3 µL of SuperScript® IV (Thermo Fisher Scientific) were added to each sample (total volume 20 µL). Samples were incubated at 50°C for 10 minutes and

80°C for 10 minutes using a T100™ Thermal Cycler. cDNA quality was assessed via reverse-transcription polymerase chain reaction (RT-PCR) (26 cycles of 95°C for 30 seconds, 59°C for 30 seconds, and 72°C for 30 seconds) using the reference gene

ELONGATION FACTOR 1 α (EF1- α). Products were visualized using a 0.5X TAE 2%

(w/v) agarose gels.

Plant Gene Expression During Plant-Aphid Interactions

Plant gene expression measurements were taken to evaluate the role GA signaling in aboveground plant defense signaling. Genes involved in plant defenses such as the SA

81 defense signaling gene [PATHOGENESIS RELATED 1 (MtPR1)] (152), and the JA- related defense genes MtMYC2 and MtVSP (153) were selected.

To assess gene expression, 1 µL of cDNA template (1:5), 5 µL of

PowerSYBR®Green Master Mix (Thermo Fisher Scientific), 2 µL of autoclaved Milli-

Q®water, and 1 µL of 3 µM forward and reverse primers were used. Each of the 384- well plates were run on a C1000®Touch ThermalCycler (Bio-Rad Laboratories Inc) and each run included two technical replicates and 3-4 biological replicates per treatment.

The thermal profile was comprised of an initial incubation at 95°C for 10 minutes, followed by 40 cycles at 95°C for 15 seconds, an annealing/extension at 56.6-58°C for 1 minute, and a melt curve analysis ranging from 65-95°C that increases incrementally by

5°C. Oligonucleotide sequences and annealing temperatures used for real-time quantitative PCR (RT-qPCR) are reported in Table 3.1. The 2-ΔCq method was used to calculate relative expression (157), and each target gene was calibrated to the reference gene GLYCERALDEHYDE-3-PHOSPHATE DEHYDROGENASE (GAPDH) (158, 159).

Table 3.1. Primers used for real-time quantitative PCR of target genes Gene Forward Primer (5’-3’) Reverse Primer (5’-3’) Average Annealing Temperature GAPDH AACATCATTCCCAGCAGCAA AACATCGACGGTAGGCACAC 56.6°C MtMYC2 GTCACAGTTCGTCGCTGGTG CGCCTCTGCTGCTTGATTTC 57.65°C MtVSP GACCTTTGGGTGTTTGACATTGA TCCTTCTGTTTGAGTGGTCTTCCT 58°C MtPR1 ATCCCCCAGAACATTGCTCG CCATCCAACACCACTACCCC 57.85°C

Statistical Analyses

All statistical analyses were carried out using SAS 9.4 for Windows (SAS

Institute Inc, Cary, NC, USA). Normal distribution of raw data was determined using

Shapiro-Wilk and Anderson-Darling tests. Two-factor analyses of variance (ANOVA) were used to evaluate the role of GA signaling and pea aphid feeding (Genotype*PA) on shoot and root dry and fresh weights. Interaction effects were further analyzed by pairwise comparisons using the Tukey-Kramer post-hoc tests. If the interaction effect was not significant, then Student’s t tests were used to evaluate the main effects (Genotype;

PA). Plant root dry weight after seven days of aphid feeding did not meet the assumptions of normality under the Shapiro-Wilk and Anderson-Darling tests (P < 0.05), therefore, the data were analyzed using log-linked Gamma distribution. Two-factor ANOVAs were also used to evaluate the role of GA signaling in modulation of plant defense gene expression during aphid feeding. Interaction effects were further investigated using

Tukey-Kramer post-hoc tests. If the interaction effect was not significant, then Student’s t tests were used to analyze the main effects. Aphid colony weight data were analyzed using Student’s t tests. Aphid total colony number data were analyzed using Poisson or negative binomial distributions.

Results

Effect of Gibberellic Acid Signaling and Pea Aphid Feeding on Shoot and Root Fresh Weight After 36 Hours of Aphid Feeding

The interaction between plant genotype and pea aphid feeding (Genotype*PA) did not have a significant effect on plant shoot fresh weight after aphid feeding for 36 hours

(Fig. 3.1 a). However, the main effect plant genotype did have a significant effect on plant shoot fresh weight 36 hours after aphid feeding (Fig. 3.1 b). The mean shoot fresh weight of wild type plants was higher compared to the mean shoot fresh weight of

Mtdella1/Mtdella2 plants regardless of aphid feeding status. The main effect pea aphid feeding did not have a significant effect on shoot fresh weight after 36 hours of aphid feeding (Fig. 3.1 a). The interaction between plant genotype and pea aphid feeding

(Genotype*PA) did not have a significant effect on root fresh weight 36 hours after aphid feeding, however, the main effect plant genotype did have an effect on root fresh weight

36 hours after aphid feeding (Fig. 3.2 a and b). Like the results for shoots, the mean root fresh weight of wild type plants was higher compared to the mean shoot fresh weight of

Mtdella1/Mtdella2 plants regardless of aphid feeding status. The main effect pea aphid feeding did not have an effect on root fresh weight at 36 hours post aphid feeding.

Figure 3.1: Impact of pea aphid (Acyrthosiphon pisum) feeding for 36 hours on Medicago truncatula shoot fresh weight of wild type (WT) and Mtdella1/Mtdella2 plants. The interaction between plant genotype and pea aphid feeding (Genotype*PA) did not have a statistically significant impact on shoot fresh weight according to the two- factor analysis of variance (P=0.5092) (a), but the main effect plant genotype was

85 statistically significant according to the Student’s t test (P<0.0001). Values represent the mean shoot fresh weight ± SEM (n = 7 or 8) per plant genotype.

Figure 3.2: Impact of pea aphid (Acyrthosiphon pisum) feeding for 36 hours on Medicago truncatula root fresh weight of wild type (WT) and Mtdella1/Mtdella2 plants. The interaction between plant genotype and pea aphid feeding (Genotype*PA) did not have a statistically significant impact on root fresh weight according to the two- factor analysis of variance (P=0.2360) (a), but the main effect plant genotype was statistically significant according to the Student’s t test (P<0.0001). Values represent the mean shoot fresh weight ± SEM (n = 7 or 8) per plant genotype.

Effect of Gibberellic Acid Signaling on Pea Aphid Fitness After 36 Hours of Aphid Feeding

Plant genotype did not have a significant effect on the aphid fitness parameters that were measured in the present study such as mean aphid count per colony and mean aphid colony weight after 36 hours of feeding (Fig. 3.3 a, b).

Figure 3.3: Mean pea aphid (Acyrthosiphon pisum) count per colony (a), and mean pea aphid colony weight (b) after feeding for 36 hours on Medicago truncatula wild type (WT) and Mtdella1/Mtdella2 plants. Plant genotype had no statistically significant effect on mean aphid count per colony (a) or mean aphid colony weight based on Student’s t tests (P>0.05). Values represent the mean ± SEM (n = 8) per plant genotype.

Role of Gibberellic Acid Signaling in Plant Defense Gene Expression 36 Hours After Aphid Feeding

The interaction between plant genotype and pea aphid feeding (Genotype*PA) did have a significant effect on MYC2 relative gene expression in shoots 36 hours after aphid feeding (Fig. 3.4). Shoots of aphid-infested wild type plants had increased MYC2 gene expression compared to aphid-infested Mtdella1/Mtdella2 plants, while MYC2 gene expression in non-infested wild type and Mtdella1/Mtdella2 plants did not differ from any of the treatments.

Figure 3.4: MtMYC2 relative gene expression in Medicago truncatula shoots of wild type (WT) and Mtdella1/Mtdella2 plants after 36 hours of pea aphid (Acyrthosiphon pisum) feeding. The interaction between plant genotype and pea aphid feeding (Genotype*PA) did have a significant effect on MtMYC2 relative gene expression in shoot based on a two-factor analysis of variance (P=0.00140). Values represent the mean ± SEM (n = 3 or 4) per plant genotype.

The interaction between plant genotype and pea aphid feeding (Genotype*PA) did not have a significant effect on VSP relative gene expression in shoots at 36 hours post aphid feeding (Fig. 3.5 a). However, the main effects plant genotype and pea aphid feeding did have a significant effect on VSP relative gene expression at 36 hours post aphid feeding (Fig. 3.5 b and c). The mean VSP relative gene expression in shoots of

Mtdella1/Mtdella2 plants was downregulated compared to the mean gene expression in wild type plants regardless of pea aphid feeding status (Fig. 3.5 b). Meanwhile, pea aphid feeding regardless of plant genotype downregulated VSP relative gene expression in shoots (Fig. 3.5 c).

Figure 3.5: VSP relative gene expression in Medicago truncatula shoots of wild type (WT) and Mtdella1/Mtdella2 plants after 36 hours of pea aphid (Acyrthosiphon pisum) feeding. The interaction between plant genotype and pea aphid feeding (Genotype*PA) did not have a significant effect on VSP relative gene expression in shoot based on a two-factor analysis of variance (P=0.3462) (a), however, the main effects (plant genotype; aphid feeding) did have a significant effect on VSP relative gene expression based on Student’s t tests (b and c). Values represent the mean ± SEM (n = 3 or 4) per plant genotype.

In contrast, the interaction between plant genotype and pea aphid feeding

(Genotype*PA), and the main effects (Genotype; PA) did not alter PR1 gene expression in shoots 36 hours after aphid feeding (Fig. 3.6).

Figure 3.6: MtPR1 relative gene expression in Medicago truncatula shoots of wild type (WT) and Mtdella1/Mtdella2 plants after 36 hours of pea aphid (Acyrthosiphon pisum) feeding. The interaction between plant genotype and pea aphid feeding (Genotype*PA) did not have a significant effect on MtPR1 relative gene expression in shoot based on a two-factor analysis of variance (P=0.7987). Values represent the mean ± SEM (n = 3 or 4) per plant genotype.

Effect of Gibberellic Acid Signaling and Pea Aphid Feeding on Plant Shoot and Root Dry Weight After 7 Days of Aphid Feeding

The interaction between plant genotype and pea aphid feeding (Genotype*PA) did not have a significant effect on plant shoot dry weights 7 days after aphid feeding (Fig.

3.7 a). However, the main effect plant genotype did have a significant effect on plant shoot dry weights 7 days after aphid feeding (Fig. 3.7 b). The mean shoot dry weight of wild type plants was higher compared to the mean shoot fresh weight of

Mtdella1/Mtdella2 plants regardless of aphid feeding status. The main effect pea aphid feeding did not have a significant effect on shoot dry weight after 7 days of aphid feeding.

Figure 3.7: Impact of pea aphid (Acyrthosiphon pisum) feeding for 7 days on Medicago truncatula shoot dry weight of wild type (WT) and Mtdella1/Mtdella2 plants. The interaction between plant genotype and pea aphid feeding (Genotype*PA) did not have a statistically significant impact on shoot dry weight according to the two- factor analysis of variance (P=0.4165) (a), but the main effect plant genotype was statistically significant according to the Student’s t test (P<0.0001). Values represent the mean shoot dry weight ± SEM (n = 6-8) per plant genotype.

The interaction between plant genotype and pea aphid feeding (Genotype*PA) did have a significant effect on root dry weights 7 days after aphid feeding (Fig. 3.8). Non- infested wild type plants had higher root dry weight compared to both non-infested and aphid-infested Mtdella1/Mtdella2 plants. However, root dry weight of aphid-infested wild type plants did not differ from any of the treatments (Fig. 3.8).

Figure 3.8: Impact of pea aphid (Acyrthosiphon pisum) feeding for 7 days on Medicago truncatula root dry weight of wild type (WT) and Mtdella1/Mtdella2 plants. The interaction between plant genotype and pea aphid feeding (Genotype*PA) did have a statistically significant impact on root dry weight according to the two-factor analysis of variance (P=0.0139). Values represent the mean shoot fresh weight ± SEM (n = 6-8) per plant genotype. Different letters represent significant differences among groups using Tukey-Kramer post-hoc tests (P<0.05).

Effect of Gibberellic Acid Signaling on Pea Aphid Fitness After 7 Days of Aphid Feeding

Plant genotype did not have a significant effect on the aphid fitness parameters measure in this study such as mean aphid count per colony and mean aphid colony weight after 7 days of aphid feeding (Fig. 3.9 a and b).

Figure 3.9: Mean pea aphid (Acyrthosiphon pisum) count per colony (a), and mean pea aphid colony weight (b) after feeding for 7 days on Medicago truncatula wild type (WT) and Mtdella1/Mtdella2 plants. Plant genotype had no statistically significant effect on mean aphid count per colony (a) or mean aphid colony weight (b) based on Student’s t tests (P>0.05). Values represent the mean ± SEM (n = 6-8) per plant genotype.

Discussion

In the present study, we investigated the role of GA signaling in regulating plant defenses during plant-aphid interactions and tested the following hypotheses: 1) aphid fitness will be improved after feeding on Mtdella1/Mtdella2 mutants compared to wild type plants and 2) JA-regulated defense genes will be downregulated in

Mtdella1/Mtdella2 mutants.

The two parameters of aphid fitness (abundance and insect colony weight) measured in the present study were not affected by plant genotype at the two time points that were chosen, indicating that GA signaling might not be involved in modulating plant defenses against aphids (Fig. 3.3 and 3.9). Future studies may benefit from extending feeding time or adding other aphid fitness parameters such as relative growth rate, aphid survivorship, or reproduction as they may be altered by changes in JA or SA (138, 142).

Interestingly, JA-regulated gene expression in shoots was altered by GA signaling at 36

93 hours post aphid feeding. MtMYC2 relative gene expression in shoots was impacted by the interaction between plant genotype and pea aphid feeding, showing that aphid- infested Mtdella1/Mtdella2 shoots had reduced levels of MtMYC2 transcripts compared to the levels in aphid-infested wild type plants (Fig. 3.5). Previously, MYC2 was found to be positively regulated by DELLAs through JAZ inhibition, therefore releasing MYC2 and subsequently allowing JA-related defense gene expression (80). Here, we demonstrate that DELLAs are important for MtMYC2 expression in early plant responses against pea aphid herbivory. MtVSP relative gene expression in shoots was not impacted by the interaction between plant genotype and pea aphid feeding (Fig. 3.4 a), however the main effects plant genotype and pea aphid feeding did have significant effects on the mean VSP relative expression in shoots (Fig. 3.4 b and c). Overall, the mean MtVSP relative gene expression in shoots was repressed in Mtdella1/Mtdella2 mutants compared to wild type plants, as well as in treatments that received aphids. This indicates that

DELLAs may be important for regulating MtVSP gene expression and that pea aphid feeding negatively impacts MtVSP gene expression in Mtdella1/Mtdella2 mutants. In addition, the interaction and main effects did not alter MtPR1 relative gene expression in shoots at 36 hours post aphid feeding, indicating that MtPR1 gene expression may not be regulated by GA signaling or pea aphid feeding (Fig. 3.6). Similarly, it was found that

MtPR1 gene expression in M. truncatula plants during pathogen and rhizobia infection was relatively stable and no significant changes were reported (152). However, another

SA defense marker gene, MtPR5, and the SA biosynthesis gene Phenylalanine ammonia- lyase (MtPAL) were significantly affected in early stages of rhizobial nodulation or pathogen infection (152). Therefore, in order to rule out the involvement of GA signaling

94 in regulating the SA pathway, future research designed to assess gene expression of SA biosynthesis genes, SA levels, and other SA-related defense genes should be considered.

In addition, assessment of gene expression focusing on the JA/ethylene pathway, which is important for JA-regulated defenses against necrotrophic pathogens and is mostly antagonistic with the MYC branch of JA signaling should be taken into consideration (76,

78).

Previously, it has been found that JA-regulated defense gene expression is important for M. truncatula resistance to the bluegreen aphid (135). However, it was also reported that pea aphid susceptible M. truncatula plants exhibit increased gene expression of the SA-regulated gene β-1-3 GLUCANASE (BGL) 36 hours post aphid feeding, meanwhile resistant plants expressed BGL transcripts as early as 24 hours post aphid feeding, and MtPR5 gene expression was constant throughout the time-points (116). Both resistant and susceptible plants lacked upregulation of gene expression of the JA-related genes MtVSP and proteinase inhibitor (116). Although, we found that MtMYC2 gene expression was upregulated by pea aphid feeding in the WT plants, MtVSP gene expression was downregulated by aphid feeding 36 hours post aphid feeding regardless of plant genotype. In addition, changes in SA-related defenses were not found in MtPR1 gene expression. It was shown previously that JA levels were severely decreased between

24- and 48-hours post pea aphid feeding in M. truncatula plants (138). In addition, pea aphid susceptible M. truncatula plants that were fed on by pea aphids continuously for 7 days exhibited higher fold changes in gene expression of BGL and MtPR5, indicating that these defenses may not be suitable for pea aphid resistance (59).

The interaction between plant genotype and pea aphid feeding did not have a significant effect on plant shoot or root fresh weight 36 hours post aphid feeding (Fig. 3.1 a and Fig. 3.2 a), but the main effect of plant genotype did have a significant effect on shoot and root fresh weight (Fig. 3.1 b and Fig. 3.2 b). Overall, wild type plants exhibited higher plant shoot fresh and dry weights compared to Mtdella1/Mtdella2 plants regardless of aphid feeding status 36 hours post aphid feeding (Fig. 3.1 b). Root fresh weight was also significantly higher in wild type plants compared to Mtdella1/Mtdella2 plants regardless of aphid feeding 36 hours post aphid feeding (Fig. 3.2 b). Similar results were observed in plant shoot dry weight 7 days post aphid feeding (Fig. 3.7). However, the interaction between plant genotype and pea aphid feeding did have a significant effect on plant root dry weight 7 days post aphid feeding (Fig. 3.8). Here, it was found that non- infested wild type plants had higher root dry weight compared to both non-infested and aphid-infested Mtdella1/Mtdella2 plants. However, root dry weight of aphid-infested wild type plants did not differ from any of the treatments (Fig. 3.8). These results agree with previous data showing that root fresh weight in Mtdella1/Mtdella2 mutants is reduced compared to wild type (43). Although, shoot length was not measured in the present study, Mtdella1/Mtdella2 plants are taller compared to wild type plants (43).

The present study revealed that GA signaling may play a role in modulating JA- regulated defense gene expression (MtMYC2 and MtVSP) but had no impact in modulating gene expression of the SA-regulated defense gene MtPR1. The data did not support the hypothesis that JA-regulated defense genes will be downregulated in

Mtdella1/Mtdella2 mutants. Even though, MtMYC2 and MtVSP gene expression was

96 repressed in Mtdella1/Mtdella2 shoots, the aphid fitness parameters measured in the present study did not correlate with the gene expression data showing that aphid fitness will be improved after feeding on Mtdella1/Mtdella2 mutants compared to wild type plants. Future studies evaluating the role of GA signaling in mediating plant-aphid interactions may benefit by including other aphid fitness parameters such as relative growth rate, aphid survival, and reproduction, which have been shown to be altered by M. truncatula and other legume plant defenses (138, 142). This study serves as foundation for future endeavors aiming to discern the role of GA signaling in mediating plant-aphid interactions.

CHAPTER IV

CONCLUSIONS AND FUTURE DIRECTIONS

The present research focused on two objectives; 1) examine the role of exogenous

GA in regulating plant defenses during AM fungus-plant-aphid interactions, and 2) evaluate the role of GA signaling in modulating plant defenses during plant-aphid interactions. To achieve our objectives, we used two study systems. The first system involved pea aphids (Acyrthosiphon pisum), barrel medic plants (Medicago truncatula), and the AM fungus Rhizophagus intraradices. The second system involved pea aphids and two plant genotypes (segregant wild type (WT) and Mtdella1/Mtdella2 mutant).

The barrel medic plant has been a keystone model legume organism for over three decades due to its importance in comparative and functional genomics, which greatly aids agricultural and ecological advancements (198, 199). The barrel medic plant is a fairly used foraging crop that is closely related to alfalfa (Medicago sativa), which is the most widely used foraging crop in the United States (11 million hectares) and is an important crop globally (199). In addition, the barrel medic serves as a foundational model organism for studying plant-beneficial microbe interactions, such as nitrogen fixing bacteria (rhizobia) or AM fungi (199). Advancements in understanding these mechanisms not only affect agricultural output through increased plant nutrition but allow for more comprehensive plant-microbe studies that map out signaling events that allow for plant- beneficial microbe associations as well as how pathogens or parasitic plants may hijack plant immunity (199-201). These advancements benefit other leguminous crops, such as

98 peas, beans, and lentils, which are important crops in the United States (202).

Additionally, barrel medic plants can share similar insect pests with other leguminous plants, such as the pea aphid, therefore allowing for more wide-ranging pest management research (64).

Pea aphids are agriculturally important crop pests and legume specialists that not only maintain short generation times, exhibit incredible dispersal behavior, and devastate crops quickly, and they are viral vectors for over 30 agriculturally important plant viruses, such as the Pea enation mosaic virus and the Bean leafroll virus (62-64).

However, pea aphids have become an important model organism not only for studying plant-aphid interactions but also for beneficial microbe-plant-aphid interactions (115).

Although rhizobia are limited to leguminous plants, AM fungi are known to form beneficial associations with most terrestrial plants including important agricultural and horticultural crops (17, 29). AM fungi-plant-insect interactions have been a growing field due to tremendous agricultural and ecological relevance (47, 109, 110, 203). Although

AM fungi exist belowground, they can play an important role in aboveground plant- insect interactions and vice versa through changes in plant physiology (54).

Plants maintain strict regulatory systems relying on potent phytohormones to prioritize growth over defense as well as maintain or restrict AM fungal growth in plant roots (73, 74, 204, 205). AM fungi have been reported to ‘prime’ plant defenses aboveground for insect attack, however this mechanism may be dependent upon insect feeding type as well as insect specialism (6, 7, 34, 48, 57, 113). In addition, there is relatively little consensus as to how phloem-feeding insects, such as aphids, can manipulate plant defense signaling as well as take advantage of AM fungal-plant

99 symbioses. It has been found that pea aphid feeding aboveground can negatively impact

AM fungal colonization belowground. For instance, pea aphid feeding can induce GA biosynthesis belowground during AM fungi-plant-aphid interactions (59). GA has been typically known to negatively regulate AM fungal-plant symbioses (114). In addition,

GA has been found to negatively regulate JA-related defenses in plant-pathogen studies

(1, 73, 85, 86, 100-102). This indicates that pea aphid herbivory may undermine antiherbivore defenses and indirectly interact with belowground AM fungi through changes in plant metabolites. However, there are few studies to date that have tested the role of

GA signaling in plant-insect interactions. To our knowledge this is the first study that has attempted to evaluate the role of GA in AM fungi-plant-aphid interactions or GA signaling in plant-aphid interactions.

The assessment of our first objective (described in Chapter II) agreed with previous research that AM fungal root colonization positively impacts plant shoot biomass (19). In the 7 days post aphid feeding time point, we found that shoot fresh weight for mycorrhizal plants receiving GA regardless of aphid feeding did not differ from non-mycorrhizal plants receiving GA, but were lower than mycorrhizal only plants, indicating that GA may antagonize the symbiosis to an extent. This may be due to the fact that plant shoot weights are not heavily impacted by 36 hours of pea aphid feeding.

As expected, pea aphid herbivory negatively impacted shoot fresh weight, meanwhile

AM fungus root colonization positively impacted shoot fresh weight. Fresh root weight was unaffected 36 hours post-aphid feeding and negligibly affected 7 days post aphid feeding. Despite there being obvious differences in plant shoot and root weights, the use of plant fresh weight is not an accurate measure of plant biomass. Traditionally, plant dry

100 weight is the more accurate approach, however the use of frozen tissue for gene expression ruled out the possibility of dry weight measurements.

As expected, there were no differences in aphid fitness parameters after 36 hours of feeding. Interestingly, aphid fitness parameters were differentially affected after continuous feeding for 7 days. Here, the aphid fitness parameters for both aphid count per colony (abundance) and aphid colony weights followed a similar pattern. Aphids that fed on plants only colonized by the AM fungus exhibited highest abundance and aphid colony weight, which confirms previous research (59). In contrast with our hypothesis, aphid fitness parameters for aphids feeding on plants receiving GA were not higher than treatments not receiving GA. Although GA treatment did improve aphid fitness parameters in comparison to non-mycorrhizal plants, aphid fitness parameters for aphids feeding on mycorrhizal plants receiving GA were lower than that for aphids that fed on mycorrhizal only plants and did not differ from aphids that fed on non-mycorrhizal plants receiving GA. It is possible that this study could have benefitted from other aphid parameters, such as relative growth rate, aphid survival, and aphid reproduction, which have been shown to be altered by JA and SA changes (138, 142). In addition, exogenous

GA in corn (Zea mays) has been shown to interrupt host selection, food consumption, and female reproduction in the fall armyworm (Spodptera frugiperda), indicating that other antixenotic measures may be important to incorporate in future studies (163). Our results indicate that aphid fitness on mycorrhizal plants is negatively affected by exogenous GA.

We also analyzed phytohormone concentrations in leaves and petioles 36 hours post aphid feeding. Here we found that the interaction between GA and AM fungus root

101

colonization regardless of aphid herbivory impacted GA3 levels in leaves and petioles.

Interestingly, mycorrhizal plants receiving GA had the highest levels of GA3 levels in leaves and petioles. JA levels were similarly affected by the interaction between exogenous GA and AM fungus root colonization. Interestingly, JA levels were high in non-mycorrhizal plants receiving GA. GA and JA typically undergo antagonistic crosstalk, however the data here indicate that exogenous GA positively affected JA levels in leaves and petioles. Although most studies have found antagonistic crosstalk between the two phytohormones, it has been reported that GA and JA signaling can work synergistically to initiate sesquiterpene production (1, 85, 86, 101, 102, 165). SA levels were positively impacted by exogenous GA and pea aphid herbivory, meanwhile AM fungus root colonization negatively impacted SA levels. It was expected that exogenous

GA would increase SA levels due to suppression of JA signaling. Interestingly, our data show that exogenous GA increased both JA and SA levels in leaves and petioles. Our data also agree with prior research that has shown that pea aphid feeding can increase SA levels in leaves between 24 h and 48 h post aphid feeding (138). SA is known to negatively affect AM fungal penetration in plant roots and AM fungi are known to induce

JA signaling, it was expected that AM fungus root colonization would negatively impact

SA levels in roots (34, 114). Our results indicate that plant phytohormone crosstalk may not be as straight forward as the literature may suggest, especially during tripartite interactions.

We also assessed plant defense gene expression in shoots and roots. Here we found that JA-regulated gene expression for MtMYC2 and MtVSP was induced by AM fungus root colonization in both shoot and root tissues. MtMYC2 gene expression only

102 differed in belowground expression where MtMYC2 gene expression was upregulated due to pea aphid feeding aboveground. However, MtVSP gene expression in shoots was also impacted by exogenous GA and pea aphid herbivory. In accordance with increased JA levels in leaves and petioles, MtVSP gene expression was increased by exogenous GA.

This further indicates a possible synergistic effect in GA-JA crosstalk. Surprisingly, pea aphid herbivory increased MtVSP gene expression. Our results show that plant responses to aphid herbivory and AM fungi when receiving exogenous GA contradict previous plant-pathogen studies indicating that GA signaling suppresses JA signaling (1, 85, 86,

100, 164).

We also assessed SA-regulated defense gene expression using the marker gene

MtPR1 (152). In shoot tissue, MtPR1 expression was positively impacted by AM fungus root colonization and pea aphid herbivory. Similarly, with increased SA levels in leaves and petioles, MtPR1 expression was higher in plants that were fed on by pea aphids. This is consistent with studies that have shown that SA levels and SA-related gene expression increase in M. truncatula leaves and petioles after pea aphid infestation or the closely related bluegreen aphid (Acyrthosiphon kondoi) infestation (116, 135, 138). In addition,

AM fungus root colonization upregulated MtPR1 gene expression in root tissue, which may be indicative of mycorrhiza-induced resistance (MIR) against root pathogens (206).

Interestingly, the present study indicate that AM fungi and aphids may alter SA-regulated defense signaling through separate mechanisms.

Finally, we assessed the role of exogenous GA and aphid herbivory on arbuscule development and AM fungal biomass using the arbuscule-specific plant genes MtPT4 and

MtLec5 and the fungal gene RiEF (43, 155, 156). Although MtPT4 gene expression was

103 significantly impacted by the interaction between exogenous GA and pea aphid herbivory, there were no differences among the means. In addition, MtLec5 was unaffected by either treatment. Future research would require quantifying arbuscule density microscopically to better observe the effects of GA or pea aphid herbivory on

AM fungus-plant symbioses (43, 169). Fungal biomass was also unaffected using RiEF as a marker. However, this agrees with previous research showing that GA can stimulate hyphal branching, therefore allowing for similar fungal biomass among treatments (43,

169).

The assessment of our second objective focused on the role of GA signaling in modulating plant defenses during plant-aphid interactions. Here we found that plant shoot fresh and dry weight was lower in Mtdella1/Mtdella2 mutants than in segregant wild type

(WT) plants 36 hours and 7 days post aphid feeding. Plant root fresh weight 36 hours post aphid feeding also followed the trend showing that Mtdella1/Mtdella2 mutants had lower root fresh weigh than WT plants. However, plant root dry weight 7 days post aphid feeding was impacted by the interaction between plant genotype and pea aphid feeding.

Non-infested WT plants had the highest root dry weights, meanwhile non-infested and aphid infested Mtdella1/Mtdella2 plants had the lowest root dry weights. Aphid-infested

WT plants did not differ from any other groups. Our results agree with previous research showing that root fresh weight in Mtdella1/Mtdella2 was lower than in WT plants (43).

Unexpectedly, aphid fitness parameters were unaffected by plant genotype 36 hours and 7 days post aphid feeding. However, this may be due to the fitness parameters that were used. As mentioned earlier, relative growth rate, aphid survival, and aphid reproduction have been shown to be altered by JA and SA changes and may have

104 benefitted this study (138, 142). Perhaps an extending feeding time-point would have benefitted the study as well. Regardless, the current data show no clear evidence of GA signaling altering aphid fitness.

We also assessed plant defense gene expression for the JA-regulated defense marker genes MtMYC2 and MtVSP and SA-regulated defense marker gene MtPR1.

MtMYC2 gene expression was affected by plant genotype and pea aphid herbivory. Here it was found that DELLAs are important for MtMYC2 gene expression during pea aphid herbivory. These data were further supported by MtVSP gene expression. Here we found that MtVSP gene expression was lower in Mtdella1/Mtdella2 mutants compared to WT plants, indicating that DELLAs are important for downstream JA-regulated defenses. In addition, MtVSP was downregulated by pea aphid feeding regardless of plant genotype, which is consistent with previous studies indicating that JA-regulated defense gene expression in aphid susceptible plants is not upregulated after pea aphid or bluegreen aphid feeding (116, 135). In addition, MtPR1 gene expression was unaffected, indicating that SA signaling is not impacted by GA signaling or pea aphid herbivory. However, involvement of SA signaling cannot be ruled out. Previous studies have shown that other

SA biosynthesis or SA-regulated defense markers related to pathogen or rhizobia infection may be involved in M. truncatula susceptibility and resistance to aphid herbivory (116, 135).

In summary, data from our first experiment found that GA may alter plant defenses leading to increased pea aphid resistance in mycorrhizal plants through GA-JA synergism. In all, our results from this experiment provide evidence supporting mycorrhiza-induced resistance (MIR) (34, 48, 113). In addition, we propose that AM

105 fungi may alter SA-related defense gene expression through a separate mechanism. Data from our second experiment showed that GA signaling is involved in JA-regulated defense gene expression in plant-aphid interactions. However, SA signaling does not seem to be involved.

To conclude, this study provides novel information on the role of the phytohormone GA in tripartite interactions as well as the role of GA signaling in plant- aphid interactions. Future studies should continue to study the role of GA-JA synergism in tripartite interactions. It may be beneficial to include studies involving volatile compounds as well as aphid behavior and other aphid fitness parameters to indicate antibiotic or antixenotic means of GA-JA synergism, since it has been shown that this positive crosstalk can affect volatile production (165). In addition, future research should focus on other JA-regulated genes that may be upregulated in this GA-JA synergism as well as further investigate how AM fungi may regulate SA-related defense expression.

Other future studies in pairwise plant-aphid interactions should include other JA- and SA- related biosynthesis and defense genes as well as metabolites to further elucidate the role of GA signaling in this interaction. Finally, further research in this field may provide greater outlets for more sustainable agricultural practices in handling important pests, such as aphids, that are more economically and environmentally stable.

106

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