The Pennsylvania State University
The Graduate School
Department of Crop and Soil Sciences
PLANT-INSECT (LEPIDOPTERA) INTERACTION IN MAIZE
A Dissertation in
Agronomy
by
Wen-Po Chuang
2012 Wen-Po Chuang
Submitted in Partial Fulfillment of the Requirements for the Degree of
Doctor of Philosophy
May 2012
ii
The dissertation of Wen-Po Chuang was reviewed and approved* by the following:
Dawn S Luthe Professor of Crop and Soil Sciences Dissertation Advisor Chair of Committee
Surinder Chopra Associate professor of Crop and Soil Sciences
Kathleen Brown Professor of Horticulture
John E. Carlson Professor of Forest Resources
Yinong Yang Associate professor of Plant Pathology
Jack Watson Professor of Crop and Soil Sciences Interim Head of the Department of Crop and Soil Sciences
*Signatures are on file in the Graduate School
iii ABSTRACT
Plants are frequently challenged by insect herbivores and have developed sophisticated defense mechanisms to counter their attack. Which insect elicitors can be recognized by plants and which plant proteins are involved in plant defense against insects are two big questions for plant scientists. In this dissertation, I would like to address these two questions by using maize
(Zea mays) and fall armyworm (Spodoptera frugiperda) as a model system. Although chewing insects cause extensive mechanical damage to plants, this does not account for the entire effect of herbivory. Caterpillar oral secretions, including saliva and regurgitant, trigger herbivore defense responses in plants. Several herbivore-associated molecular patterns (HAMPs) have been found in insect regurgitant. However, there is an argument whether caterpillar larvae actually regurgitate on maize leaves during feeding on maize. Thus, I examined fall armyworm larval saliva to determine if it is an important elicitor of herbivore defenses in maize. Analyses indicated that very little regurgitant was deposited on the maize leaf during caterpillar feeding. Furthermore, caterpillar regurgitant failed to trigger plant defense-related genes in maize. Whereas, leaf tissue immunoblots indicated that glucose oxidase, an abundant saliva protein, was deposited on the leaf when caterpillars fed there. The effect of caterpillar saliva on maize defense gene expression was determined by allowing ablated (no saliva) and non-ablated (saliva present) caterpillars to feed in the maize whorl. The results showed that feeding by unablated caterpillars significantly increased the expression of plant genes involved in jasmonate biosynthesis pathway and direct defenses.
Furthermore, saliva-induced bioassay showed that compared to plants fed on unablated larvae, plants fed by ablated larvae did not induce sufficient direct defense to significantly retard larval growth. Furthermore, the saliva-responsive maize protein profile has been identified from the proteomics analysis. The results of this study show that caterpillar saliva is an important elicitor for triggering herbivore defenses in maize. Another research project was to discover a new way to find plant proteins that defend against insect herbivores. Some important plant defense proteins
iv have been found that can resist digestive proteases in the insect gut and are eliminated in the frass. We used mass spectrometry to identify several maize proteins in fall armyworm frass that may play a role in herbivore defense, including ribosome-inactivating protein 2 (RIP2).
Ribosome-inactivating proteins (RIPs) act by depurinating residues on ribosomal RNA and thereby inhibit translation. Since RIP2 was found in frass, we proposed that it might be involved in defending maize from herbivore attack. Immunoblot analysis indicated that RIP2 is initially synthesized as an inactive proenzyme that can be cleaved to the active form by larval gut extracts.
Also, results indicated that the expression of RIP2 was induced by FAW larval feeding, but not mechanical wounding. The proenzyme form of RIP2 was detected in 13 maize inbred lines and two teosinte subspecies. This data indicates that RIP2 expression in response to insect feeding is a wide spread phenomenon in maize. Quantitative-RT-PCR and immunoblot analysis indicated that
RIP2 is rapidly induced (1 hour) following caterpillar attack and remains at the wound sites for four days after caterpillar removal. Phytohormone application assays determined that RIP2 expression was regulated by several different phytohormones, including ethylene, JA and ABA. It appears that there is no consistent pattern of hormonal regulation of RIP-like protein expression.
The expression profile of RIP2 in maize vegetative stage was examined. The data showed that
RIP2 is expressed locally and during maize vegetative development. Furthermore, when purified recombinant RIP2 was directly tested against fall armyworm larvae in bioassays, the data indicated that the amount of RIP2 typically found in the leaf after caterpillar attack could significantly retard caterpillar growth. We concluded that RIP2 plays an important role in protecting maize against insect herbivores. With these two studies, I was able to take a further step toward understanding the interaction of plant-insect relationship.
Key words: plant-insect interaction, maize (Zea mays), fall armyworm (Spodoptera frugiperda), saliva, ribosome-inactivating protein 2 (RIP2)
v TABLE OF CONTENTS
LIST OF FIGURES ...... viii LIST OF TABLES ...... x ACKNOWLEDGEMENTS ...... xi
Chapter 1 Introduction ...... 1
Chapter 2 Literature review ...... 4
Plant defense ...... 4 Constitutive and induced defenses ...... 5 Direct and indirect defense ...... 5 Mechanical wounding versus caterpillar feeding ...... 7 Insect elicitors ...... 9 HAMPs ...... 9 Saliva ...... 9 Regurgitant ...... 11 Plant antinutritive proteins ...... 12
Chapter 3 Fall armyworm (Spodoptera frugiperda) saliva is an important elicitor of herbivore defenses in maize ...... 15
Introduction ...... 15 Results ...... 18 Little regurgitant is detected on caterpillar-fed maize leaves ...... 18 Caterpillars salivate on maize leaves...... 19 Caterpillar regurgitant failed to trigger plant defense-related genes in maize ...... 20 Saliva induces plant defense-related genes in maize ...... 21 Performance of larvae on previously fed maize leaves ...... 22 Structure of the spinneret and collection of saliva ...... 23 Discussion ...... 23 Materials and methods ...... 27 Plant materials and insect rearing ...... 27 Fluorescence detection of regurgitant ...... 28 Ablation of spinneret ...... 29 Quantitative RT-PCR ...... 29 Tissue printing ...... 30 Saliva-induced bioassays ...... 30 Feeding area calculation ...... 31 Scanning electron microscopy ...... 31
Chapter 4 Ribosome-inactivating protein 2 protects maize against insect herbivores ...... 43
Introduction ...... 43 Results ...... 46 Identification of maize proteins in insect frass ...... 46 Two maize RIP genes respond to insect herbivore differently...... 48 RIP2 can be found in maize leaf and frass ...... 49
vi Effect of heterologous RIP2 on fall armyworm larval growth ...... 50 Insect herbivore, not mechanical wounding, induces RIP2 induction ...... 51 Characterization of RIP2 profile under insect herbivore attacking ...... 52 RIP2 is expressed locally and during maize vegetative development...... 52 Phytohormones regulating RIP2 expression ...... 53 Discussion ...... 55 Maize defensive protein in insect frass ...... 55 Different expressed pattern of RIP genes ...... 56 The role of RIP2 in maize against insect herbivores ...... 58 Materials and Methods: ...... 63 Plant materials, insect rearing, and frass collection ...... 63 Immunoblot analysis ...... 63 RIP2 bioassays and RIP2 digestion ...... 64 Phytohormone treatments ...... 65 Analysis of developmental and systemic expression ...... 66 Insect feeding ...... 66 RIP2 persistance after larval feeding...... 66 Quantitative RT-PCR (qRT-PCR)...... 67 Multidimensional protein identification technology (MudPIT) and protein identification ...... 68
Chapter 5 Proteomics study on maize proteins that respond to fall armyworm (Spodoptera frugiperda) saliva ...... 95
Introduction ...... 95 Results: ...... 96 Maize protein response to ablated or unablated larvae in the susceptible genotype Tx601 ...... 97 Ablated group versus control group ...... 97 Unablated group versus control group ...... 98 Unblated group versus ablated group ...... 98 Maize protein response to ablated or unablated larvae in the resistant genotype Mp708 ...... 99 Ablated group versus control group ...... 99 Unablated group versus control group ...... 100 Unablated group versus ablated group ...... 101 Discussion: ...... 101 Materials and Methods: ...... 103 Plant materials and insect rearing ...... 103 Ablation of spinneret ...... 103 Sample collection ...... 104 Protein extraction and sample preparation for 2D-DIGE ...... 104 Sample labeling for 2D-DIGE ...... 105 Two-dimensional polyacrylamide gel electrophoresis (2D-PAGE) ...... 106 Gel imaging and data analysis ...... 107 Protein spot picking and MS/MS analysis and Protein identification ...... 107
Chapter 6 Conclusion and future directions ...... 137
vii References: ...... 141
viii LIST OF FIGURES
Figure 3-1. The absence of regurgitant with caterpillar feeding on maize leaves...... 33
Figure 3-2. Immunoblot analysis of GOX in Tx601 leaves...... 35
Figure 3-3. Immunoblot analysis of GOX in Mp708 leaves...... 36
Figure 3-4. Immunoblot analysis of GOX in Mp708 leaves...... 37
Figure 3-5. Expression analysis of maize genes in response to fall armyworm larval regurgitant treatments...... 38
Figure 3-6. Expression analysis of maize genes involving in JA biosynthesis in response to various treatments...... 39
Figure 3-7. Expression analysis of maize genes involving in plant defense in response to various treatments...... 40
Figure 3-8. Bioassay of S. frugiperda larvae fed on maize genotypes subjected to various treatments...... 41
Figure 3-9. Amounts of leaf area (cm2) eaten by ablated versus unablated fall armyworm larvae...... 42
Figure 3-10. Scanning electron microscopy of two Lepidopteran larval spinnerets...... 43
Figure 4-1. Expression analysis of RIP1 and RIP2 transcripts in maize leaves in response to caterpillar feeding...... 71
Figure 4-2. Immunoblot analysis of RIP2 in maize leaf tissues and S. frugiperda frass...... 72
Figure 3-10. Immunoblot analysis of RIP2 protein in several maize inbreds and teosinte...... 73
Figure 4-4. Immunoblot analysis of rRIP2 digested by papain-coated beads and S. frugiperda larvae...... 74
Figure 4-5. Effect of recombinant RIP2 on S. frugiperda larvae growth...... 75
Figure 4-6. Immunoblot analysis of rRIP2 digested by Mir1-CP...... 76
Figure 4-7. Analysis of RIP2 transcripts levels in maize leaves in response to various treatments...... 77
Figure 4-8. Immunoblot analysis of RIP2 in maize leaves in response to various treatments...... 78
Figure 4-9. Time course of RIP2 mRNA expression in response to feeding by S. frugiperda larva (one larva per plant)...... 79
ix Figure 4-10. Immunoblot analysis of RIP2 protein accumulation in maize leaves after feeding S. frugiperda larva (one caterpillar per plant) for various time points...... 80
Figure 4-11. Immunoblot analysis of RIP2 protein in maize leaves after larval removal...... 81
Figure 4-12. Systemic expression of RIP2 protein in maize leaves after larval feeding...... 82
Figure 4-13. Developmental expression of RIP2 protein in maize leaves after larval feeding...... 83
Figure 4-14. Effect of exogenous ethylene on RIP2 protein expression in maize leaf after 24 hr treatment...... 84
Figure 4-15. Effect of exogenous methyl jasmonate (MEJA) and ethylene on RIP2 protein expression in maize leaf after 24 hr of treatment...... 85
Figure 4-16. Effect of exogenous EthylBloc (ethylene perception inhibitor) on RIP2 protein expression in maize leaves...... 86
Figure 4-17. Effect of exogenous methyl jasmonate (MEJA) on RIP2 protein expression in maize leaves after 24 hr treatment...... 87
Figure 4-18. Effect of exogenous abscisic acid (ABA) on RIP2 expression in maize leaf after 24 hours treatment...... 88
Figure 5-1. Experimental design of a six-gel 2D-DIGE experiment for comparative study of insect-induced maize proteins...... 109
x
LIST OF TABLES
Table 3-1. The numbers of fluorescent spots were detected on treated leaves (figure 3-1). .... 34
Table 4-1. Maize proteins identified in frass of S. frugiperda fed by two maize genotypes (Tx601 and Mp704)...... 70
Supplemental 4-1. Maize proteins identified in frass of S. frugiperda fed by two maize genotypes (Tx601 and Mp704)...... 89
Table 5-1. Differentially expressed protein spots with associated annotation and fold changes compared with control group, ablated group, unablated group in susceptible genotype Tx601...... 110
Table 5-2. Differentially expressed protein spots with associated annotation and fold changes compared with control group, ablated group, unablated group in resistant genotype Mp708...... 112
Table 5-3. Numerical representation maize leafy proteins responded to fall armyworm ablated or unabalted larvae...... 118
Table 5-4. Differentially expressed protein spots with associated annotation and fold changes compared with ablated group and control group in susceptible genotype Tx601...... 119
Table 5-5. Differentially expressed protein spots with associated annotation and fold changes compared with unablated group and control group in susceptible genotype Tx601...... 121
Table 5-6. Differentially expressed protein spots with associated annotation and fold changes compared with unablated group and ablated group in susceptible genotype Tx601...... 123
Table 5-7. Differentially expressed protein spots with associated annotation and fold changes compared with ablated group and control group in resistant genotype Mp708...... 124
Table 5-8. Differentially expressed protein spots with associated annotation and fold changes compared with unablated group and control group in resistant genotype Mp708...... 128
Table 5-9. Differentially expressed protein spots with associated annotation and fold changes compared with unablated group and ablated group in resistant genotype Mp708...... 133
xi ACKNOWLEDGEMENTS
My appreciation and gratitude belongs to many people who have great contributed to the work presented in this dissertation.
I am grateful for my advisor and mentor, Dr. Dawn S. Luthe. This work would not be possible without her great support and encouragement. I cannot express how much I want to thank her. All I can say is that I am so lucky to have her as my advisor. I also want to thank Dr.
Gary Felton to give me numerous suggestions and support for my dissertation and research.
I would like to thank my committee members, Dr. Surinder Chopra, Dr. John Carlson,
Dr. Kathleen Brown, and Dr. Yinong Yang, for their comments and guidance to my thesis. And I would like to special thanks to Dr. Paul Williams at USDA-ARS to give me a financial support to start my work. I would also like to gratefully acknowledge the support of my lab members, Ms.
Erin Bassford, Mrs. Renuka Shivaji, Dr. Olga Pechanova, Dr. German Sandoya, Dr. Liz Bosak,
Mr. Robert Guyer, Mr. Swayamjit Ray, Mrs. Han Yang, and Ms. Shan Jin. Also, I would like to special thanks Mrs. Michelle Peiffer for her great help on my research.
Finally, I want to thank my friends, Mr. Pohao Wang, Dr. Fangyi Cheng, Mr. Chia-Chen
Liang, Ms. Yi-Hsiu Huang, and my former advisor, Dr. Liang-Jwu Chen, for their continuing encouragement and warmness. In the end, I would like to thank to my parents. Without you, I cannot have chance to finish this work.
Chapter 1
Introduction
Maize (Zea mays L.), often known as corn, is one of world leading crops for food, feed, oil, and biofuel sources. In 2009, maize was the leading cereal in the world and wheat and rice were the second and third respectively (FAOSTAT, 2011). In the United States, maize was the most productive grain crop with 316 million metric tons produced in 2010 (USDA, 2011).
Besides representing a staple food source, maize also is grown annually for industrial products, including ethanol, sweeteners, and other bioproducts.
Maize is a member of Poaceae family. Its life cycle contains vegetative and reproductive stages. The vegetative stage starts from seed germination, emergence of the coleoptile, and leaf development until silks emerge as reproductive stage begins. The reproductive stage starts from silking to physiological maturity (Ritchie, 1986). Maize is hermaphrodite. The female inflorescences are ears which grow from axillary meristematic buds between shoot and leaf. The male inflorescences are tassels at the top of the stem. Maize is outcrossing crop. However, it can be manipulated to self-pollinate.
Maize is a diploid plants with 10 chromosomes (2n = 20). The genome size is nearly
2,300 Mb with 32,000 predicted genes (Schnable et al., 2009). Approximately 85% of genome is composed of transposable elements (Schnable et al., 2009). Maize originated in the Americas. It was domesticated more than 8700 years ago in the highlands of Mexico (Tenaillon and
Charcosset, 2011). It is widely accepted that maize originated from one of teosintes, Zea mays ssp. Parviglumis, (Matsuoka et al., 2002). Evidence shows that maize was domesticated by human selection (Wang et al., 1999). Two key genes in the domestication process were found: teosinte branched 1 (tb1), a transcription factor which represses the development of axillary meristems and results in one dominant axis (Hubbard et al., 2002); and teosinte glume
2 architecture 1 (tga1), a transcription factor which regulated to the naked grain phenotypes (Wang et al., 2005).
Insect feeding results in a huge loss of the crop yield worldwide. During 2001-2003, the actual losses in corn yield caused by pest damage around the world was 9.6 % (OERKE, 2006).
In the United States, the pesticide expense in 2011 was more than ten billion dollars (USDA,
2011). Besides using pesticides to reduce yield loss due to insect feeding, transgenic crops made by using recombinant DNA technology have been developed. Endotoxic Cry protein from
Bacillus thuringiensis (Bt) was introduced into crops to generate insect resistance. In 2011, maize insect-resistant transgenic varieties comprised 65% of the total maize planted in the United States
(USDA, 2011). However, the insect-resistant transgenic crops not only have made a significant beneficial impact, but also accelerated the evolution of Bt-resistant insects. One of the Bt-targeted pests is western corn rootworm. However, Bt-resistant western corn rootworm recently was found in Iowa fields (Gassmann et al., 2011). Besides, transgenic crops also have a negative impact on non-target insects, including natural enemies of crop pests (Gatehouse et al., 2011). Thus, research focusing on alternative strategies to reduce the yield loss caused by insect feeding has been investigated. One of these is to identify and exploit endogenous insect resistance genes from crops. Another strategy is to understand how plants recognize insect feeding, since plants can distinguish the difference between mechanical wounding and insect feeding.
The main goals of my research is to understand how insect saliva alters plant defense in monocot plants and identify plant defensive proteins from those that accumulate in insect frass.
It has been shown that insect saliva can alter plant defense in dicot plants (Musser et al., 2005;
Consales et al., 2011). However, there is no report that insect saliva can be recognized by monocot plants and further alter plant defense. By using proteomics and functional genomics approaches, we intend to study the effect of caterpillar saliva on maize plants and characterize the maize defensive proteins found in caterpillar frass.
3 The objectives of this research are:
1) To prove that fall armyworm saliva is effectively recognized by maize plants and
further triggers plant defense
2) To identify maize defensive proteins from maize-fed caterpillar frass and characterize
their function
3) To identify maize proteins expressed differentially in respond to fall armyworm saliva
4 Chapter 2
Literature review
Plant defense
As land plants are sessile organisms, they have evolved sophisticated defense mechanisms against various environmental stresses. To survive and reproduce, plants adapt to stresses by changing their physiology and gene expression. However, some environmental stresses often come together, such as drought and heat, which make plants need to recognize stresses more precisely and activate proper responses. Insect herbivores are one of major biotic stresses to plants. As plants are the main nutrient sources for these insects, plants have evolved with a number of defense mechanisms to protect themselves. Plant physical barriers, such as trichomes, waxes, and hairs play the first line of defense against herbivores. The trichomes on the plant surface limit insect movement and they may release substances that repel the insects (Howe and Schaller, 2008). In some plant species, the number of trichomes increases when the plants suffer herbivore feeding (Howe and Schaller, 2008). Furthermore, leaf toughness is another plant physical barrier. Based on the feeding styles, insect herbivores obtain the nutrients by tearing the plant tissues with their mouthparts or penetrating into plant tissues with a piercing-sucking structure. The leaf toughness can also damage the caterpillar mandibles (Schoonhoven et al.,
2005; Howe and Schaller, 2008). For some angiosperm plants, when insect herbivores damage the leaf veins, the latex in the veins exudes from the point of damage. The latex may trap the insects or its toxic contents may poison them (Konno et al., 2006). These physical barriers can deter the insect feeding. However, these traits are often not strong enough to completely resist insect attack. Thus, chemicals and enzymes also are involved in plant defense against herbivores.
5 Constitutive and induced defenses
Plant synthesized chemicals and enzymes help protect plants from insect herbivore attack. The insect resistance traits can be expressed as a constitutive or continual defense.
However, constitutively producing defensive chemicals and enzymes is costly and limits resources that could be used for growth and reproduction. Unlike constitutive defense, induced defense is only costly when plants are under herbivore attack. Many studies show that induced defenses reduce the allocation cost more than constitutive defenses (Herms and Mattson, 1992;
Strauss et al., 2002; Zavala et al., 2004). Types of plant defense depend on the interactions with other species (Strauss et al., 2002). In addition to induced defense, plants can tolerate the herbivore attack by re-allocating sugars from source (leaves) to sink (roots) (Schwachtje et al.,
2006).
Direct and indirect defense
Plant direct defenses rely on the accumulation of secondary metabolites, defensive proteins and other toxic compounds (Howe and Jander, 2008). Plant secondary metabolites, which include glucosinolates, 2,4-dihydroxy-7-methoxy-1,4-benzoxazin-3-one (DIMBOA), alkaloids, phenolics, and other compounds, act as toxins in plant-insect interactions. Plant secondary metabolites are the by-products of primary metabolites and are not the essential for plant growth. However, it is widely accepted that plant secondary metabolites are involved in the resistance to pathogens and herbivores (Bennett and Wallsgrove, 1994; Theis and Lerdau, 2003;
Mao et al., 2007; Chen, 2008). Many plant secondary metabolites are toxic to insects. Even if insects have evolved mechanisms to detoxify the compounds, there is a high cost to overcoming the toxicity. The cost may affect the insect survival. For plant defensive proteins, a major class of
6 defensive proteins includes protease inhibitors (PIs) that were first found to rapidly accumulate in potato and tomato after insect feeding (Green and Ryan, 1972). In the insect gut, the PIs interact with gut proteases and thereby inhibit their digestive function. In maize, the maize proteinase inhibitor (MPI) is induced by wounding and caterpillar feeding (Tamayo et al., 2000). MPI can inhibit elastase and chymotrypsin-like enzymes in the larval midgut of Spodoptera littoralis and plays a plant defense role against insect herbivores (Tamayo et al., 2000). Besides proteinase inhibitors, plant proteases also involved in the plant defense. Mir1-CP (maize insect resistance 1- cysteine protease) is a good example (Jiang et al., 1995). In insect-resistant maize, this protein accumulates around the wounding site in response to caterpillar feeding. It damages the peritrophic matrix of caterpillar (Pechan et al., 2000; Pechan et al., 2002), which will retard the caterpillar’s growth and provide effective plant defense (Pechan et al., 2002).
Plant indirect defenses include production of green leaf volatiles and other volatile organic compounds that attract natural enemies of the attacking insects. More than 1700 volatile compounds have been identified from 90 more plant families (Pichersky and Gershenzon, 2002;
Maffei et al., 2011). Plants emit a blend of volatiles for growth, development, and responses to the environment stresses (abiotic and biotic stresses) (Kant et al., 2009). Plants emit floral volatiles to attract pollinators. Insects can detect volatiles through their antennae system on their head. However, plant volatiles are also involved in plant defense against insect herbivores and pathogen. Plants release volatiles constantly. However, the composition and quantity of plant volatiles varies based on the environment stresses. Each plant species under insect infestation emits different blends of volatiles (De Moraes et al., 1998). The predatory and parasitoid insects can distinguish the difference and find their host (De Moraes et al., 1998). Furthermore, maize terpene synthase (TPS10) is the key enzyme involved in the synthesis of herbivore-induced sesquiterpenes in maize (Schnee et al., 2006). Overexpression TPS10 in Arabidopsis plants emitted more sesquiterpenes after insect herbivory and resulted in attaching more female parasitic
7 wasp Cotesia marginiventris (Schnee et al., 2006). Herbivore-induced plant volatiles not only attract predatory and parasitoid insects but also trigger indirect defenses to the neighboring uninfested plants (Kost and Heil, 2006).
Mechanical wounding versus caterpillar feeding
Upon insect attack, many chemical reactions change in the plant cell, such as ion fluxes, formation of reactive oxygen species (ROS), protein modification, and phytohormone (jasmonic acid, ethylene, and salicylic acid) biosynthesis and signaling (Mithöfer and Boland, 2008;
Thivierge et al., 2010; Hogenhout and Bos, 2011). Although chewing insects cause serious mechanical damage to plant tissues, this does not account for the entire effect of herbivory on plants. Due to the high cost of producing defenses, plants need to distinguish between mechanical wounding and herbivore feeding to allocate resources efficiently. Here, I discuss three aspects
(volatiles and gene regulation) to show the differences between mechanical wounding and herbivore feeding.
To respond to herbivore attack, plants often emit the volatiles to provide cues for carnivorous insects to find their prey (Walling, 2000). Although wounded plants emit volatiles, the intensity and type of volatiles differs depending on the type of wounding. Plant tissues wounded by scratching, crushing, or puncturing cannot induce the same volatile emission as herbivore feeding (Turlings et al., 1990; Reymond et al., 2000; Spiteller et al., 2000; Halitschke et al., 2001; Schmelz et al., 2001; Mithöfer and Boland, 2008). These results argue that a single artificial wounding could not mimic continuous caterpillar wounding (Mithöfer et al., 2005).
Thus, it has been suggested that plants recognize herbivore wounding depending on the amount, duration, and pattern of damage (Baldwin, 1990; Bruinsma and Dicke, 2008). MecWorm, a mechanical caterpillar, was designed to answer this question (Mithöfer et al., 2005). The results
8 showed that when lima beans suffered continuous wounding by MecWorm, the plants emitted similar herbivore-related volatiles (Mithöfer et al., 2005). However, not all herbivore-related volatiles responded to the MecWorm wounding. This suggested that there are additional factors, such as oral secretions and saliva, that could be involved in plant volatile emission (Mithöfer and
Boland, 2008).
Plants respond to insects differently depending on the herbivore feeding style. Piercing- sucking insects (ex: Sternorrhyncha) have specialized stylets/tube-like structures to suck liquid contents from plants. Chewing insects (ex: Coleoptera and Lepidoptera) cause extensive plant tissue damages by chewing, snipping, and tearing. It has been shown that chewing insects cause extensive wound damage to plant tissues, but this mechanical damage does not account for the entire herbivore effect on plants (Howe and Jander, 2008). Large-scale analysis of gene expression analysis has surveyed gene transcript differences between mechanical wounding and insect feeding (Reymond et al., 2000; Major and Constabel, 2006). Farmer and his colleagues showed that many wound induced genes in Arabidopsis may not be induced or are expressed less when plants are under herbivore attack (Reymond et al., 2000). In addition, few genes had a similar expression pattern between water-treated and insect regurgitant-treated tobacco plants
(Halitschke et al., 2003). Furthermore, microarray results show that more than half of the genes that responded to insect regurgitant had similar transcriptional changes as the genes regulated by
M. sexta feeding (Halitschke et al., 2003). Another study reported that applying oral secretions on plant wounded sites caused more jasmonic acid (JA) and ethylene production in tobacco than wounding alone (Bruinsma and Dicke, 2008). Thus, these findings showed that plants have different gene regulation in respond to mechanical wounding and herbivore feeding.
9 Insect elicitors
HAMPs
Plants can distinguish damages between insect herbivory or mechanical wounding by perceiving herbivore-associated molecular patterns (HAMPs) from insects (Felton and
Tumlinson, 2008). HAMPs are the insect equivalent of the microbial-associated molecular patterns (MAMPs) which are well-known molecules/proteins involved in plant-pathogen interactions (Felton and Tumlinson, 2008). In contrast to MAMPs, only a few HAMPs are been identified. Several HAMP proteins have been identified, including GOX from corn earworm, alkaline phosphatase from whitefly, β-glucosidase from white cabbage butterfly, and lipase from grasshopper (Mattiacci et al., 1995; Eichenseer et al., 1999; Funk, 2001; Musser et al., 2005;
Mithöfer and Boland, 2008; Wu and Baldwin, 2010; Schäfer et al., 2011). In addition, low MW
HAMPs include fatty acid-amino acid conjugates (FACs) from multiple insect species, inceptins
(proteolytic fragments of chloroplastic ATP synthase γ-subunit) from fall armyworm fed on cowpea or maize, bruchins from pea weevil and cowpea weevil, and caeliferin from grasshopper
(Alborn et al., 1997; Doss et al., 2000; Schmelz et al., 2006; Alborn et al., 2007; Yoshinaga et al.,
2007; Mithöfer and Boland, 2008). Although these HAMPs have been identified, there are no
HAMPs which can induce JA- and ET-related responses in all plant species (Wu and Baldwin,
2010). Thus, HAMPs seem to be highly specific to plant species.
Saliva
Caterpillar saliva is the secretion released from salivary glands. In Lepidoptera, larvae have two pairs of salivary glands: labial glands and mandibular glands (Bordas, 1903;
Parthasarathy and Gopinathan, 2005; Felton, 2008). Long and tubular labial glands secrete watery
10 fluids from the principal secretory structure called spinneret (Carter and Hargreaves, 1986; Felton and Eichenseer, 1999; Liu et al., 2004). Mandibular glands are also tubular, but their secretions are released from the cutting edge of the mandibles and contain more lipophilic substances, such as sterols and triglycerides than the labial glands (Wroniszewska, 1966; Mossadegh, 1978;
Eichenseer et al., 2002; Howard and Baker, 2004; Felton, 2008). Early studies of caterpillar saliva were focused on silk formation and digestive enzymes, such as carbohydrases, proteinases, and lipases (Mathur, 1966; Chattoraj and Mall, 1969; Saxena, 1972; Verma and Balyan, 1972;
Shimada and Hayashiya, 1975; Agarwal, 1976; Verma et al., 1977; Raghavan et al., 1978; Liu et al., 2004). Recently, the sialome (salivary transcriptome and proteome) and structure of labial salivary glands of Hericoverpa armigera have been elucidated (Sorensen et al., 2006; Celorio-
Mancera et al., 2011). Glucose oxidase (GOX), lysozyme, phenolglucosyl transferase, and ascorbate peroxidase were found in the labial saliva (Ahmad and Hopkins, 1992; Mathews et al.,
1997; Eichenseer et al., 1999; Liu et al., 2004). Several digestive proteins (amylase and maltase), lysozymes, proteases, and proteinase inhibitors have been identified from the salivary transcriptome (Celorio-Mancera et al., 2011). Mandibular gland saliva also contains proteins, but appears to be richer in lipids (Wroniszewska, 1966; Mossadegh, 1978; Eichenseer et al., 2002;
Howard and Baker, 2004; Felton, 2008). GOX is the most abundant protein (> 50%) in the labial saliva of Helicoverpa zea (Eichenseer et al., 1999). The enzyme activity of GOX in the saliva among several different insect species was highly variable (Merkx-Jacques and Bede, 2005). In addition, GOX activity also varies with different larval developmental stages (Eichenseer et al.,
1999). After head capsule slippage (molting), GOX in caterpillar saliva has the highest enzyme activity compared with GOX in other insect organs. Several studies showed that GOX in labial saliva could prevent induction of nicotine when caterpillar fed on tobacco (Nicotiana tabacum)
(Musser et al., 2002; Musser et al., 2005). The enzyme catalyzes the following reaction:
Glucose+O2gluconic acid + H2O2
11 Hydrogen peroxide, one of the products of the GOX catalyzed reaction, has been shown to induce salicylic acid (SA) accumulation in plants and further triggers systemic acquired resistance (SAR) (Durner et al., 1998; Niki et al., 1998; Felton and Eichenseer, 1999). Bede and her colleagues (2008) have shown that saliva of beet armyworm, Spodoptera exigua, activates the
SAR pathway to decrease the JA-related plant defense when this insect feeds on Arabidopsis thaliana (L.) Heynh. However, there is still an argument about the role of hydrogen peroxide in the reaction. Musser et al. (2005) showed that SA is not involved in the suppression of tobacco nicotine by H. zea GOX. Another possible function of GOX is that it could inhibit plant oxidative enzymes, which may reduce the nutritive quality of plant tissues (Felton and Summers, 1995;
Felton and Gatehouse, 1996; Eichenseer et al., 1999). Plant polyphenol oxidase plays a defensive role by producing oxidative stress in the insect gut (Constabel and Barbehenn, 2008). GOX would scavenge O2 to help maintain a relative anaerobic environment (Eichenseer et al., 1999).
Lysozyme is another enzyme found in the H. zea saliva (Liu et al., 2004). Lysozyme is known to have an antimicrobial function and it has been speculated that saliva has an antimicrobial effect on the bacteria on the plant surface (Eichenseer et al., 1999). Depending on host plants, lysozyme transcripts in caterpillar saliva were expressed differently. Liu et al. (2004) suggested that lysozyme in saliva could also suppress plant-insect defense response by eliciting plant-pathogen defense response such as GOX in saliva did.
Regurgitant
HAMPs from insect regurgitant have been identified from maize to tobacco (Alborn et al., 1997; Halitschke et al., 2001; Alborn et al., 2007). It was believed that HAMPs from insect regurgitant play a major role in triggering herbivory-specific responses in plants (Bonaventure et al., 2011). The well-known components of HAMPs from insect regurgitant are fatty acid-amino
12 acid conjugates (FACs), such as volicitin (Alborn et al., 1997; Felton, 2008). FACs not only elicit terpenoid and indole emission but also induce the terpenoid and indole biosynthesis gene expression in maize (Alborn et al., 1997; Frey et al., 2000; Shen et al., 2000; Schnee et al., 2002;
Voelckel and Baldwin, 2004). It is accepted that FACs are not the only elicitors in regurgitant that can trigger plant defenses (Felton and Tumlinson, 2008). Inceptin, a fragment of chloroplastic
ATP synthase γ-subunit protein, is the first peptide elicitor found in fall armyworm regurgitant fed when they fed on cowpea (Schmelz et al., 2006; Felton, 2008). Although several studies showed the substances in regurgitant induced the direct or indirect plant defenses (Schmelz et al.,
2006; 2007), the regurgitant may not always enhance the plant defense (Yan et al., 2005; Felton,
2008). Although a plasma membrane protein in maize leaves has been suggested as the receptor of volicitin (one of HAMPs from insect regurgitant), there is no follow-up study to identify the particular volicitin receptor (Truitt et al., 2004).
Plant antinutritive proteins
More and more plant proteins are believed to have an antinutritional function in plant defense (Felton, 2005). Plant protease inhibitors (PIs) have been widely studied for their defensive functions. Ryan and his colleagues (1972) first found that PIs in potato and tomato rapidly accumulate after insect feeding. In the insect gut, the PIs interact with gut proteases and thereby inhibit their digestion function. Many transgenic plants expressing PI genes have been developed and showed detrimental effects on insect growth (Chen, 2008; Howe and Jander, 2008;
Zhu-Salzman et al., 2008). Furthermore, it is shown that plant oxidative enzymes are also involved in plant defense against insect herbivores (Chen, 2008). Polyphenol oxidases (PPOs), peroxidases, lipoxygenases, and ascorbate oxidases oxidize essential nutrients or form electophilic products, which react with the nucleophilic side chains of amino acids (Felton et al.,
13 1994; Constabel, 1999; Chen, 2008). These enzymes lower the nutritive value of plants to insects.
Besides the above proteins, some proteases, such as maize insect resistance 1-cysteine protease
(Mir1-CP) and leucine aminopeptidase (LAP), have been shown to defend plants against insect herbivory (Chao et al., 1999; Chen, 2008; Zhu-Salzman et al., 2008). Howe and his colleagues
(2007) found that some putative plant defense proteins can resist digestion in the insect gut and retain enzymatic activity. These proteins or their fragments may be found in the frass. Some of the plant defense proteins detected in the frass are TD2, LAP-A, PR7, and etc. (Chen et al., 2007).
Here, I briefly described three of the plant antinutritive proteins found in frass.
TD2, one of threonine deaminase isoforms in tomato, has been detected in the tomato leaf and the gut lumen and frass of phytophagous insects (Chen et al., 2005; Chen et al., 2007).
Furthermore, TD enzyme activity can be detected in frass collected from tomato-reared M. sexta larvae (Chen et al., 2007). This finding indicates that some plant defense proteins not only are resistant to digestive proteases but also have a post-ingestive defense function on insect gut lumen
(Chen et al., 2007). TD2 exists as proenzyme form in the tomato leaf and the regulatory domain that inhibits enzyme activity is removed in the insect gut and the processed enzyme is excreted in the frass (Chen et al., 2007). When insects ingest tomato leaves, TD2 is converted to the processed form in the insect gut where it plays an anti-nutrient role by reducing the levels of threonine, a necessary amino acid for phytophagous insects (Chen et al., 2007). A further study confirmed that a chymotrypsin-like protease in the lepidopteran gut processed TD from the proenzyme to the active form (Gonzales-Vigil et al., 2011).
Leucine aminopeptidase A (LapA) in tomato increases in response to wounding, exogenous methyl jasmonate, pathogen infection, and insect feeding (Chao et al., 1999; Chen et al., 2005; Chen et al., 2007; Zhu-Salzman et al., 2008). LapA protein has been detected in the midgut and frass of M. sexta (Chen et al., 2005; Chen et al., 2007), and its overexpression in tomato delays M. sexta growth and development (Lee et al., 2009).
14 Arginase is another antinutritive enzyme. L-arginine (Arg) is one of essential amino acids for plant-feeding insects (Spencer, 2007). Besides Arg its use in protein synthesis, Arg is also the precursor for the biosynthesis of polyamines, proline, glutamate, and nitric oxide (NO). There are two major pathways of Arg metabolism. Arginase and NO synthase are the enzymes involved in each of these pathways (Chen et al., 2008). Arginase hydrolyzes Arg to ornithine, the precursor of polyamines, and urea. ARG2, one of arginase isoform, in cultivated tomato (Solamum lycopersicum) responds to mechanical wounding and methyl jasmonate (Chen et al., 2004). In the midgut of M. sexta larvae reared on tomato plants, both TD2 and ARG2 have been identified
(Chen et al., 2005). Both enzymes are active in the midgut extract of M. sexta (Chen et al., 2005).
Transgenic tomato plants that overexpressed ARG2 reduced the growth of M. sexta larvae (Chen et al., 2005). Also, the available arginine levels are reduced in the ARG2 oveexpressing transgenic lines (Chen et al., 2005). Therefore, both ARG2 and TD2 have the ability to degrade the essential amino acids in the gut of Lepidoptera (Chen et al., 2008).
15 Chapter 3
Fall armyworm (Spodoptera frugiperda) saliva is an important elicitor of herbivore defenses in maize
Introduction
Plants are frequently challenged by insect herbivores. Therefore, to avoid being overeaten by insects, plants have developed very sophisticated defense mechanisms by recognizing cues from insects that they use to trigger plant defense. Trichomes, waxes, and hairs are physical barriers that defend plants against insect herbivory. In addition to these physical defenses, numerous chemical reactions change in the plant cell upon insect attack. These include ion fluxes, formation of reactive oxygen species (ROS), protein modification, and phytohormone (jasmonic acid, ethylene, and salicylic acid) biosynthesis and signaling (Mithöfer and Boland, 2008;
Thivierge et al., 2010; Hogenhout and Bos, 2011). Based on herbivore feeding styles, plants respond to insects differently. Piercing-sucking insects (ex: Sternorrhyncha) have specialized stylets/ tube-like structures to suck liquid contents from plants. Chewing insects (ex: Coleoptera and Lepidoptera) cause extensive plant tissue damages by chewing, snipping, and tearing. It has been known that chewing insects cause extensive wound damage to plant tissues, but this mechanical damage does not account for the entire herbivore effect on plants (Howe and Jander,
2008). Large-scale analysis of gene expression has surveyed transcriptional differences between mechanical wounding and insect feeding (Reymond et al., 2000; Major and Constabel, 2006).
Farmer and his colleagues showed that many wound- induced genes in Arabidopsis may not be induced or are expressed less when plants are under herbivore attack (Reymond et al., 2000). In addition, a few genes had a similar expression pattern between water and insect regurgitant treated tobacco plants (Halitschke et al., 2003). Furthermore, microarray result shows that more than half of the genes that responded to insect regurgitant had similar transcriptional changes as
16 the genes regulated by M. sexta feeding (Halitschke et al., 2003). Another study reported that regurgitant applied to plant wound sites caused more jasmonic acid (JA) and ethylene production in tobacco than wounding alone (Bruinsma and Dicke, 2008). Recently, it was reported that the expression of two defense genes (ERF/AP2TF and protease inhibitor) in Arabidopsis were suppressed by regurgitant from the caterpillars (Consales et al., 2011). Thus, these findings showed that plants have different gene regulation in response to mechanical wounding and herbivore feeding.
In addition to regurgitant, caterpillar saliva also contains chemical cues that can be recognized by plants and further alter plant defenses. This comprehensive finding was demonstrated by cauterizing (or ablating) the caterpillar spinneret which releases saliva (Musser et al., 2002). When ablated caterpillars fed on tobacco, there was more nicotine production in plants (Musser et al., 2002). This finding suggested that components of saliva suppress the production of this secondary metabolite involved in plant defense. Similar suppression results were obtained when M. sexta regurgitant was applied on tobacco (Kahl et al., 2000).
Where as regurgitant arises from the digestive system, saliva is the secretion released from salivary glands. In Lepidoptera, larvae have two pairs of salivary glands: labial glands and mandibular glands (Bordas, 1903; Parthasarathy and Gopinathan, 2005; Felton, 2008). Long and tubular labial glands secrete watery fluids through the principal secretory structure called spinneret (Carter and Hargreaves, 1986; Felton and Eichenseer, 1999; Liu et al., 2004).
Mandibular glands are also tubular, but their secretions are released from the cutting edge of the mandibles and contains more lipophilic substances, such as sterols and triglycerides than the labial glands (Wroniszewska, 1966; Mossadegh, 1978; Eichenseer et al., 2002; Howard and
Baker, 2004; Felton, 2008). Early studies of caterpillar saliva were focused on silk formation and digestive enzymes, such as carbohydrases, proteinases, and lipases (Mathur, 1966; Chattoraj and
Mall, 1969; Saxena, 1972; Verma and Balyan, 1972; Shimada and Hayashiya, 1975; Agarwal,
17 1976; Verma et al., 1977; Raghavan et al., 1978; Liu et al., 2004). Recently, glucose oxidase
(GOX), lysozyme, phenolglucosyl transferase, and ascorbate peroxidase were found in the labial saliva (Ahmad and Hopkins, 1992; Mathews et al., 1997; Eichenseer et al., 1999; Liu et al.,
2004). Mandibular gland saliva also contains proteins, but appears to be richer in lipids
(Wroniszewska, 1966; Mossadegh, 1978; Eichenseer et al., 2002; Howard and Baker, 2004;
Felton, 2008). GOX is the most abundant protein (> 50%) in the labial saliva of Helicoverpa zea
(Eichenseer et al., 1999). The enzymatic activity of GOX in the saliva among several different insect species was highly variable (Merkx-Jacques and Bede, 2005). In addition, GOX activity also varies with different larval developmental stages (Eichenseer et al., 1999). After head capsule slippage (molting), when feeding begins GOX in caterpillar saliva was the highest.
Several studies showed that GOX in labial saliva could prevent induction of nicotine when caterpillar fed on tobacco (Nicotiana tabacum) (Musser et al., 2002; Musser et al., 2005).
Lysozyme is another enzyme found in H. zea saliva (Liu et al., 2004). Lysozyme is known to have an antimicrobial function and it has been speculated that saliva has antimicrobial effects on the bacteria present on the plant surface (Eichenseer et al., 1999). Depending on host plants, lysozyme transcripts in caterpillar saliva were expressed differently. Liu et al. (2004) suggested that lysozyme in saliva could also suppress plant-insect defense response by eliciting plant- pathogen defense response as GOX in saliva did.
While it is known that insect saliva can alter plant defenses in dicot species, there is no report that insect saliva can be recognized by monocot plants and further alter plant defenses. In this paper, we studied the effect of caterpillar saliva on defense expression in maize plants.
18 Results
Little regurgitant is detected on caterpillar-fed maize leaves
Plants can recognize elicitors from insect regurgitant and further trigger plant defense against insect herbivores (Howe and Schaller, 2008). However, there is a dispute on whether caterpillars actually regurgitate during most feeding bouts (Peiffer and Felton, 2009). Thus, to test whether fall armyworm larvae regurgitate on maize, we first fed fall armyworm (Spodoptera frugiperda) larvae with artificial diet containing a fluorescence dye. Then larvae were allowed to feed on maize leaf tissues of two maize genotypes (Tx601 and Mp708). Tx601 is susceptible to fall armyworm feeding (Williams et al., 1989), whereas Mp708 is highly resistant to a number of lepidopteran pests including S. frugiperda (Williams et al., 1990). The leaf tissues where larvae fed were collected and examined for the deposition of regurgitant using confocal fluorescence microscopy. Only a few, faintly fluorescent spots were detected on the leaves of insect- susceptible and insect-resistant genotypes, whereas larval regurgitant directly collected from larvae had very strong fluorescence intensity (Figure 3-1). The number of fluorescence spots found on each leaf also was determined (Table 3-1). These results indicated that only miniscule amounts of regurgitant were deposited on approximately one-third of the leaf samples. The estimated amount of regurgitant which S. frugiperda fed on another maize genotype (B73) was
1.20 nl (Peiffer and Felton, 2009). This implies that fall armyworm larvae does not significantly regurgitate on maize plants during their eating period. This result was similar to the findings for several other caterpillar species (Peiffer and Felton, 2009).
19 Caterpillars salivate on maize leaves
Caterpillar saliva may also contain elicitors that can trigger plant defense (Felton, 2008).
Glucose oxidase (GOX) is one of major proteins in the saliva of noctuid caterpillars (Felton,
2008). A large scale survey of GOX activity across two superfamilies of Lepidoptera indicated that GOX could be detected in the labial glands of eighty-three out of eighty-eight species including the fall armyworm (Eichenseer et al., 2010). In our study, we determined if fall armyworm larvae salivate on maize leaves by detecting GOX protein deposited on the leaf surface. Proteins on leaf surface were transferred to nitrocellulose membrane using a leaf printing method (Peiffer and Felton, 2005) and GOX distribution on leaf surface was determined by immunoblot analysis. We also cauterized the larval spinneret that releases saliva to determine whether GOX comes from labial gland saliva (Musser et al., 2006). In Tx601 (Figure 3-2), abundant levels of GOX were detected around the caterpillar feeding sites on maize leaves, but little GOX was detected on the non-fed leaves or leaves that were fed on by larvae with their spinneret ablated. We also examined insect resistant maize inbred (Mp708) (Figures 3-3 and 3-
4). Again, GOX was detected around the caterpillar feeding sites, but not on the non-fed leaves
(Figure 3-3). In addition, more GOX was detected on the plants fed on by unablated than ablated larvae (Figure 3-4). For both maize genotypes, small amounts of GOX were detected maize leaves that were fed on by ablated larvae (Figure 3-2 and 3-4). It is possible that this GOX might have been secreted from the mandibular glands (Eichenseer et al., 1999). Taken together these data indicate that that fall armyworm larvae routinely deposit their GOX-containing saliva on maize leaves during feeding.
20 Caterpillar regurgitant failed to trigger plant defense-related genes in maize
Although little regurgitant was detected on maize leaves, we determined if several plant herbivore defense genes responded to the application of regurgitant. Maize plants were mechanically wounded (see description of wounding tool in Materials and Methods) and caterpillar regurgitant was applied to the wound sites. Control plants were infested with unablated fall armyworm larvae for 24 hrs. The caterpillar regurgitant was collected from fall armyworm larvae that were fed with artificial diet or maize leaves before the experiment. Maize defense gene expression was analyzed by quantitative real-time polymerase chain reaction (qRT-
PCR). Lipoxygenase (LOX) and allene oxide synthase (AOS) catalyze steps in the JA- biosynthetic pathway. LOX functions in the initial step of JA biosynthesis pathway by catalyzing the oxidation of α-linolenic acid to (9S)- or (13S)-hydroperoxy-octadecadi(tri)enoic acid acid (9-
HPOT, 13-HPOT) depending on the specific type of LOX. Then, AOS converts 9-HPOT to 9,
10(S)-epoxy-octadecatrienoic acid (9, 10-EOT) and 13-HPOT to 12, 13-EOT. AOS is the first committed step in the pathway (Schaller and Stintzi, 2009). Other maize defense genes have been tested including maize proteinase inhibitor (MPI) that is induced by wounding and caterpillar feeding (Tamayo et al., 2000). MPI can inhibit elastase and chymotrypsin-like enzymes in the larval midgut of Spodoptera littoralis and plays a plant defense role against insect herbivores
(Tamayo et al., 2000). Maize peroxidase (POX) is another gene associated with environmental stresses (Dowd and Johnson, 2005). The expression data showed that the response of several maize defense-related genes to caterpillar regurgitant and wounding treatment were similar
(Figure 3-5). Furthermore, the results showed that the source of caterpillar regurgitant (diet fed or plant fed) did not affect the response of the maize plants. Although regurgitant from caterpillars fed on maize leaves induced MPI gene expression as the same level as unablated caterpillars, regurgitant failed to induce the three other genes tested. Thus, this result indicated that in addition
21 to caterpillar regurgitant, there should be another insect elicitor that can be recognized by maize plants.
Saliva induces plant defense-related genes in maize
To determine if caterpillar saliva can affect the expression of maize defenses, we tested several genes involved with JA-biosynthesis and direct defenses in maize. Plants were infested with ablated or unablated fall armyworm larvae for 24 hrs. In addition to the genes tested in regurgitant experiment (Figure 3-5), we also tested phospholipase D (PLD) and12- oxophytodienoic acid-10,11-reductases (OPR) that function in the JA-biosynthetic pathway. PLD first hydrolyzes phospholipids to release linolenic acid from the chloroplast membrane (Wang et al., 2000; Shivaji et al., 2010) while OPR catalyzes the conversion of 12-oxo-phytodienoic acid
(OPDA) to 3-oxo-2-(2-(Z)-pentenyl)-cyclopentane-1 octanoic acid (OPC-8:0). OPC is further processed to become jasmonic acid (JA). Another maize defense gene, would-induced protein
(WIP1) a Bowman-Birk type proteinase inhibitor that responded to wounding in maize coleoptiles, was tested (Rohrmeier and Lehle, 1993). We also tested one of the maize aquaporins, which are plant plasma membrane intrinsic proteins (PIPs), as a negative control. In maize roots,
PIP transcripts responded to day-night cycles (Lopez et al., 2003). The gene expression results indicated that unablated caterpillars, which have ability to release saliva, triggered the highest expression of JA-biosynthetic pathway genes and maize herbivore defense genes compared with other treatments (Figures 3-6 and 3-7). Although ablated caterpillars triggered defense gene expression, it was not significantly different from that in control or wounded plants. Thus, these data indicated that caterpillar saliva can be recognized by maize and elicit herbivore defense gene expression.
22 Performance of larvae on previously fed maize leaves
Expressions of defense-related genes in maize leaves were induced in response to caterpillar saliva. However, transcription of these genes does not necessarily mean that they are translated into functional proteins that mediate insect defense. Plants were treated with ablated or unablated larvae for 24 hr as previously described and these plants were used in larval feeding bioassays to determine if they exhibited a resistance phenotype. These plants were fed to naïve fall armyworm larvae that had no prior feeding experience on maize. Both the insect susceptible
(Tx601) and resistant (Mp708) genotypes were tested. In Tx601, naïve larvae that fed on plants treated with unablated larvae (unablated group) had the lowest larval weight (Figure 3-8). Naïve larvae that fed on plants treated with ablated larvae (ablated group) had lower larval weights than larvae reared on control plants, but the weights were not statistically different. Naïve larvae that fed on Mp708, showed the same trends as those fed on Tx601 (Figure 3-8). Both data showed that larvae fed with fully induced plants have statistically significant lower larval weight compared to those that fed on plants that were treated with ablated larvae or on uninfested control plants. Furthermore, leaves from both genotypes treated with unablated larvae retarded caterpillar growth by approximately 25% when compared to control plants (Figure 3-8). These results indicated that there is no difference in the response between insect-susceptible and insect-resistant inbreds. To ensure that there was no difference in the amount of feeding damage between ablated and unablated larvae, we measured the amount of leaf area consumed both maize genotypes
(Figure 3-9). The leaf feeding areas were not statistically different between the two treatments within the same maize genotype. Thus, this result excludes the possibility that ablated larvae eat less due to damage to the spinneret and attenuate the defense response. These data indicate that maize plants can recognize an elicitor(s) in caterpillar saliva and further trigger defenses against insect herbivory.
23 Structure of the spinneret and collection of saliva
Attempts to directly collect saliva from the fall armyworm spinnerets as previously described for corn earworm (H. zea) were not successful (Musser et al., 2002). It was impossible to see droplets of saliva on the spinneret of fall armyworm larvae. We then examined the spinneret structure of these two insects by scanning electron microscopy. The results showed that spinneret structures between corn earworm and fall armyworm larvae were quite different (Figure
3-10). The corn earworm spinneret is a tubular style or spigot-like (Figure 3-10, left). However, the fall armyworm spinneret is a scroll-like structure with a connection on the tip (Figure 3-10, middle and right). This unusual structure might explain why drops of saliva could not be collected from fall armyworm spinneret. Although salivary gland homogenates have been used previously to test the effect of saliva on plant defenses, we did not use this approach in our experiments. The salivary gland homogenates contains hundreds of non-secreted proteins and thus may confound the interpretation of results. We believe the experiments using ablation of the spinneret are the least ambiguous and most definitive approach for testing the effects of saliva.
Discussion
Plants can distinguish damage between insect herbivory or mechanical wounding by perceiving herbivore-associated molecular patterns (HAMPs) from insects (Felton and
Tumlinson, 2008). HAMPs are the herbivore equivalent of the microbial-associated molecular patterns (MAMPs) which are well-known molecules/proteins involved in plant-pathogen interactions (Felton and Tumlinson, 2008). In contrast to MAMPs, only a few HAMPs have been identified. HAMP proteins include GOX from corn earworm, alkaline phosphatase from a whitefly, β-glucosidase from white cabbage butterfly, and lipase from a grasshopper (Mattiacci et
24 al., 1995; Eichenseer et al., 1999; Funk, 2001; Musser et al., 2005; Mithöfer and Boland, 2008;
Wu and Baldwin, 2010; Schäfer et al., 2011). In addition, low MW HAMPs include fatty acid- amino acid conjugates (FACs) from multiple insect species, inceptins (proteolytic fragments of chloroplastic ATP synthase γ-subunit) from fall armyworm fed on cowpea or maize, bruchins from pea weevil and cowpea weevil, and caeliferin from grasshopper (Alborn et al., 1997; Doss et al., 2000; Schmelz et al., 2006; Alborn et al., 2007; Yoshinaga et al., 2007; Mithöfer and Boland,
2008). Although these HAMPs are been identified, there are no HAMPs which can induce JA- and ET-related responses in all plant species (Wu and Baldwin, 2010). Thus, HAMPs seems to be very specific recoginition by several plant species.
Since some of the low MW HAMPs have been identified from lepidopteran regurgitant, it has been proposed that when caterpillars feed on plants, they not only create wounding sites but also release low MW HAMPs by regurgitation. Thus, plants activate their defense system by recognition of these HAMPs. However, Felton and his colleagues showed that some species of caterpillars do not regurgitate during most feeding bouts (Peiffer and Felton, 2009). We used the same fluorescent labeling method to determine if fall armyworm larvae regurgitated on maize and found that very little regurgitant was deposited on the leaves of two inbreds differing in susceptibility to fall armyworm feeding (Table 3-1). This result was similar to the previous study using another maize inbred, B73 (Peiffer and Felton, 2009). These data indicate that fall armyworm larvae do not routinely regurgitate during feeding on maize.
Another potential source of HAMPs in fall armyworm is in saliva. Caterpillar saliva has lubrication, predigestion, and antimicrobial functions (Riberiro, 1995). However, HAMPs in saliva also may be recognized by plants and further trigger plant defenses. Immunoblot results showed that fall armyworm larvae consistently deposited the secreted salivary enzyme GOX through the spinneret onto wounds on the maize leaf (Figure 3-2~3-4). This was not unexpected since GOX activity was found in the labial glands of eighty-eight insect species including fall
25 armyworm (Eichenseer et al., 2010). However, there is a report showing that a small portion of total GOX activity can be detected in the mandibular gland and residual, ingested GOX in the caterpillar gut (Eichenseer et al., 1999). This might explain why we detected small amounts of
GOX on the leaves when ablated larvae with intact mandibular glands feed on the plants.
Alternatively, ablation of the spinneret in the fall armyworm may not be 100% successful in preventing salivation.
In this study, data from gene expression analyses and bioassays clearly indicated that fall armyworm saliva plays an important role in eliciting herbivore defenses in maize. Genes involving in JA biosynthetic pathway were up-regulated by fall armyworm saliva (Figure 3-6) indicating that it could trigger plant defense in maize by elevating JA production. Furthermore, maize genes encoding proteins involved in direct defenses were also induced by fall armyworm saliva (Figure 3-7). Although ablated larvae also induced JA-biosynthetic pathway and direct defense genes, the induction was not statistically different than that of unwounded or mechanical damaged plants. These results demonstrated that ablated larvae that were unaffected in their ability to regurgitate were not able to trigger direct defenses in maize. This was supported by the bioassay data that feeding by ablated larvae did not induce sufficient direct defenses to significantly retard larval growth (Figure 3-8). Although it could be questioned that ablated larvae ate less leaf tissue due to spinneret damage, there was no significant difference between amounts of leaves consumed by ablated larvae and unablated larvae (Figure 3-9). In addition, in the gene expression and bioassay experiments, we placed three larvae on each plant to induce plant defenses and observed that both ablated and unablated larvae seriously damaged the plants.
Despite the similar levels of damage, plants responded to the ablated and unablated larvae very differently. These results indicate that saliva from fall armyworm larvae has the capacity to elicit direct defense proteins in maize, but the specific eliciting component(s) in the saliva has yet to be
26 identified. Experiments with purified GOX, suggest that it does not elicit a defense response in maize (unpublished data).
HAMPs from insect regurgitant have been identified in plants ranging from maize to tobacco (Alborn et al., 1997; Halitschke et al., 2001; Alborn et al., 2007). It was believed that
HAMPs from insect regurgitant played a major role in triggering herbivore-specific responses in plants (Bonaventure et al., 2011). However, our study showed that the ablated fall armyworm larvae with unimpaired ability to regurgitate failed to activate defenses in maize. Furthermore, it is questionable if caterpillars regularly regurgitate during eating (Peiffer and Felton, 2009).
Labial salivary glands in the silk moth Bombyx mori are referred to as silk glands that produce silk proteins in the last stage of larval development. These silk proteins are the most well-known labial saliva proteins (Mondal et al., 2007; Celorio-Mancera et al., 2011). However, the components of labial salivary glands in other Lepidoptera have not been completely identified. Recently, the sialome (salivary transcriptome and proteome) and structure of labial salivary glands of Hericoverpa armigera have been reported (Sorensen et al., 2006; Celorio-
Mancera et al., 2011). Several digestive proteins (amylase and maltase), lysozymes, proteases, and proteinase inhibitors have been identified from the salivary transcriptome (Celorio-Mancera et al., 2011). This comprehensive study suggests that labial salivary glands in Lepidoptera not only functions as silk glands but also secrete proteins involved in pre-digestion, insect immunity, detoxification, and other unknown functions. Our saliva study provides evidence that saliva from labial salivary glands of fall armyworm also is an important mediator of defenses in maize. The identity of the salivary HAMP(s) in the fall armyworm awaits identification.
We attempted to collect saliva directly from the spinneret in order to directly collect saliva but were unable to do so. The spinneret structure of larval Bombyx mori that is used to produce silk fibroin fibers has been extensively studied (Asakura et al., 2006). However, there have been few reports of spinneret structures for other Lepidoptera species (Sorensen et al.,
27 2006). The spinnerets of H. zea and H. armigera are tubular, spigot-liket structures (Sorensen et al., 2006; Felton, 2008). The spinneret base is mushroom-like and the tip region forms a circular structure in H. armigera larvae (Sorensen et al., 2006). Compared with these two insect species, the spinneret structure of fall armyworm larvae is quite different (Figure 3-10). The tip of spinneret is a fixed connection structure between two tubular or scroll-like structures. The saliva might be released from the inner region of two tubular structures or the inner region of connection structure on the top. This might explain why we were unable to directly collect the saliva from the spinneret of fall armyworm larvae. Alternative approaches are therefore necessary to characterize the HAMP(s) components of fall armyworm saliva. In summary, the source of HAMPs or elicitors in the oral secretions of the fall armyworm arise from saliva and not from regurgitant.
The majority of HAMPS have been identified from the regurgitant of caterpillars. However, in the case of the fall armyworm, larvae deposit saliva on maize leaves and as yet unknown elicitor(s) in their saliva is recognized by maize plants which then triggers plant defenses.
Materials and methods
Plant materials and insect rearing
Maize (Zea mays) genotypes that are resistant (Mp708) and susceptible (Tx601) to fall armyworm feeding were obtained from Dr. W. Paul Williams (USDA-ARS, Mississippi State
University). Seeds were sown in pots filled with topsoil (Hagerstown loam) in the Crop and Soil
Sciences greenhouse at The Pennsylvania State University, University Park, PA. Well-watered maize plants were grown until they were at the V8 stage (eight fully emerged leaves) (Ritchie,
1986). The wounding tool used in regurgitant application experiment was described by Bosak
(2011).
28 Plant tissues surrounding the wounded area near the whorl were collected after caterpillar feeding or wounding by a paper punch (6mm diameter, ten times per plant). The tissue samples were stored at -80°C after immediately freezing in liquid nitrogen.
Fall armyworm (Spodoptera frugiperda) eggs were obtained from USDA-ARS Corn
Host Plant Resistance Research Laboratory at Mississippi State University. Larvae were reared on an artificial diet (Peiffer and Felton, 2005) in a 27°C incubator with a 16 h photoperiod and newly molted 5th instar larvae were used for all experiments except for bioassays with neonate larvae.
Corn earworm (Helicoverpa zea) eggs were initially purchased from Bioserv
(Frenchtown, NJ), or from colonies were established in the Felton laboratory. Larvae were reared on a casein based artificial diet (Jacob and Chippendale, 1971) in 1 ounce plastic cups (XpedX,
Camp Hill, PA) in a growth chamber maintained at 26° C with a 14:10 light dark cycle. After pupation, pupae were transferred to a mating chamber where they emerged. Moths were fed 10% sucrose in water and remained in the chamber for 5 days, then eggs were collected to continue the colony.
Fluorescence detection of regurgitant
The newly molted 5th instar fall armyworm larvae were fed with fluorescence dye (Alexa
Fluor 488, Invitrogen, Carlsbad, CA) on artificial diet. Then, larvae were placed on small pieces of the insect-susceptible (Tx601) and insect-resistant (Mp708) maize leaves and allowed to feed.
After few bites on the leaves, the leaf material and larval regurgitant were collected. The materials were analyze for a fluorescence signal using an Olympus Fluoview FV1000 (Olympus,
Center Valley, PA). The data was analyzed by the software of Olympus Fluoview FV1000.
29 Ablation of spinneret
The late 4th instar fall armyworm larvae were placed on ice until flaccid. The larvae were immobilized with a hair clip and the spinnerets cauterized with a heat pen (Electron Microscopy
Sciences, Hatfield, PA). This method has been described by Peiffer and Felton (2005). The cauterized larvae were placed on artificial diet to recover and after molting.
Quantitative RT-PCR
Maize plants were fed by ablated larvae and unablated larvae for 24 hr. Three larvae were placed in one plant. A paper punch was used to mimic mechanical wounding (multiple wounding sites per plant). Undamaged plants were controls. Each treatment had four biological replicates.
Total RNA from leaf tissue was isolated using the TRIzol Reagent (Invitogen) and DNase
(Promega) following the manufacturer’s instructions. The first-strand cDNA was synthesized with High Capacity cDNA Reverse Transcription Kit (ABI, Foster City, CA) with Oligo (dT)20 primers following the manufacturer’s instructions. qRT-PCR was carried out in an ABI 7500 Fast
Real Time PCR System. The primers were designed by Primer Express software for real-time
PCR (Version 3.0) (ABI, Foster City, CA). The PCR was conducted by following the default conditions: Step 1: 50°C for 2 min and 95°C for 10 min, Step 2: 95°C for 15 sec and 60°C for 1 min repeated 40 cycles, Step 3: 72°C for 10 min, Step 4: dissociation stage. The relative quantification of gene expression was analyzed by ABI 7500 Fast SDS Software (Version 1.4)
(ABI, Foster City, CA). The data set was normalized using actin as a control. Gene specific forward (F) and reverse (R) primers used to generate data present in this study were: ACTIN-F
(accession number: U60511.1) 5’- GGA GCT CGA GAA TGC CAA GAG CAG-3’, ACTIN-R
5’- GAC CTC AGG GCA TCT GAA CCT CTC-3’, AOS-F (NM_001111774) 5’-CAA ACC
30 GAC GAA TTT GAG CAA-3’, AOS-R 5’- GGA GGC TCG CAA CAA GTT G-3’, LOX1-F
(AF271894) 5’-CGT TCC GTG AAG TGT GGT TCT-3’, LOX1-R 5’- CTG TAA GGA GTA
CTT GGC ATA TTT GC-3’, MPI-F (X78988) 5’-GCG GAT TAT CGC CCT AAC C-3’, MPI-R
5’-CGT CTG GGC GAC GAT GTC-3’, OPR2-F (NM_001112435.1) 5’-CAA AGC GAG CAA
CAG AGC AG-3’, OPR2-R 5’- CTA TTT GCT CTC GGT CGG TCA-3’, PIP1-1-F (X82633)
5’-GCG CCG CCG TAA TTT ACA-3’, PIP1-1-R 5’-GCC GAC CCA GAA GA TCC A-3’,
PLD-F (NM_001112216.1) 5’-CGG GAA CAA GTC GGA TTA CCT-3’, PLD-R 5’-GCC TGC
AGT GTG CAC TCT ATG-3’, POX-F (TA112628_4577) 5’-CGA GGC CAC GTG GAA GAC-
3’, POX-R 5’-TCG ATC TGC CCC ATC TTG A-3’, WIP1-F (X71396) 5’-AGC TCA AGT
GCT GCA CCA ACT-3’, WIP1-R 5’-GAC GTC GTC GCA GGT GTA GA-3’, The results were analyzed by statistic software (SAS, Cary, NC).
Tissue printing
Maize plants were fed by ablated and unablated caterpillar for few hours until several feeding sites were evident. Fed leaf tissues were immediately blotted onto the nitrocellulose membrane using the Panther Semi-Dry Electroblotter (Thermo Scientific Owl, MA). Undamaged plants were used as controls. The detection of GOX on the membrane was conducted by immunoblot analysis as previously described (Peiffer and Felton, 2005).
Saliva-induced bioassays
Maize plants were fed by ablated larvae and unablated larvae for 24 hr. Three larvae were placed in each plant and 18 to 30 total plants were infested. The fed leaf tissues were collected, cut into approximately 1 cm pieces and placed in diet cups. Paper punch was used to mimic
31 mechanical wounding (multiple wounding sites per plant). Undamaged plants were controls.
Naïve neonate larvae (5 to 7.5 mg) were placed into diet cups with treated leaf tissues. Each treatment had 30 larvae. Bioassays using the inbred Tx601 were done one time and those with
Mp708 were repeated three times. The plates were incubated at 26ºC for 4 days in 16 hr photoperiod before the final larval weights were determined. The results were analyzed by statistical software (SAS, Cary, NC).
Feeding area calculation
Maize leaves (4 cm by 5 cm) were detached from the plant, scanned, and placed in 1% agar petri dish (VWR international, Radnor, PA). Fifth instar larvae were allowed to feed for 24 hours (one larva per leaf) and then leaves were scanned. Both scanned files of leaf tissues before caterpillar feeding and after caterpillar feeding were analyzed by SigmaScan Pro 5 (SPSS
Science, Chicago, IL). The feeding area results were analyzed by statistic software (SAS, Cary,
NC).
Scanning electron microscopy
Larvae were cleaned before fixation by sonicating in 1% Triton X-100 for 30 seconds.
After 3 rinses in dH20, larvae were patted dry and prepared as described by Hayat
(1989). Briefly, the larvae heads were removed and fixed overnight in 2.5% glutaraldehyde,
1.5% formaldehyde (Electron Microscopy Sciences, Fort Washington, PA) in 0.1 M sodium cacodylate, pH 7.4. Larvae were then rinsed in 0.1 M sodium cacodylate and treated with 2% aqueous osmium tetroxide (EMS) for two hours. Following water rinses the larvae were dehydrated in graded ethanol solutions (70% to 100%), then dried in a Baltec SCD030 (Techno
32 Trade, Manchester, NH) critical point dryer. Dried samples were mounted and coated with gold/palladium to increase conductivity (Baltec SCD050, Techno Trade, Manchester,
NH). Samples were imaged at 20 KV in a JSM 5400 scanning electron microscope (JEOL,
Peabody MA), and images were captured using IMIX-PC v.10 software (Princeton Gamma Tech,
Princton NJ).
33
(A) (B)
Leaf 1 Leaf 1
Leaf 2 Leaf 2
Regurgitant Regurgitant
Light microscopy Fluorescence microscopy Light microscopy Fluorescence microscopy
Figure 3-1. The absence of regurgitant with caterpillar feeding on maize leaves. (A) Tx601 (B) Mp708. The caterpillar fed Tx601 and Mp708 leaves were examined under
fluorescence and light microscopy. Leaf 1 was the sample that had no fluorescence signal
(Upper). Leaf 2 was the sample that had one green fluorescence spot (Middle). Regurgitant is the
caterpillar regurgitant, which was the positive control (Lower). The red arrows indicate the
fluorescence spots.
34 Table 3-1. The numbers of fluorescent spots were detected on treated leaves (figure 3-1). The number in the table represents total number of fluorescence spots detected in all leaf pieces examined.
Tx601 Mp708
Fall armyworm fed leaves 5/15 7/15
Control leaves 0/2 0/2
35
(A) (B) (C)
Figure 3-2. Immunoblot analysis of GOX in Tx601 leaves. (A) maize plants without infestation by fall armyworm (B) maize plants were fed on by cauterized fall armyworm for 2 hrs (B) maize plants were fed on by unablated fall armyworm for
1 hrs. The leaf tissues were transferred to nitrocellulose membrane. The membrane was hybridized with anti-GOX antibody. The black dots (GOX) were abundant near the wounding sites.
36
(A) Uninfested plants (B) Infested plants
Figure 3-3. Immunoblot analysis of GOX in Mp708 leaves. (A) uninfested maize leaves and (B) maize leaves that were fed on fall armyworm for 24 hrs. The leaf tissues were transferred to nitrocellulose membrane. The membrane was hybridized with anti-GOX antibody. The black dots (GOX) were abundant near the wounding sites.
37
(A) Ablated plants (B) Unablated plants
Figure 3-4. Immunoblot analysis of GOX in Mp708 leaves. (A) maize leaves were fed on by cauterized fall armyworm for 24 hrs (B) maize leaves were fed on by unablated fall armyworm for 24 hrs. The leaf tissues were transferred to nitrocellulose membrane and the membrane was hybridized with anti-GOX antibody. The black dots (GOX) were abundant near the wounding sites.
38
Figure 3-5. Expression analysis of maize genes in response to fall armyworm larval regurgitant treatments. Maize plants were wounded by wounding tool only (wounded), wounded by wounding tool and treated with regurgitant of S. frugiperda larvae which fed with artificial diet (diet) or maize leaves
(maize), or by unablated S. frugiperda larvae (unablated) for 24 hrs. Undamaged plants were used as a control. Total RNA was isolated from wounding site of maize leaves. The relative expression levels were determined in each biological sample and normalized to actin gene expression.
Letters indicate significant differences by LSD test (p<0.05) (n=3~4, error bar indicates SE).
39
Figure 3-6. Expression analysis of maize genes involving in JA biosynthesis in response to various treatments. Maize plants were wounding by paper punch (wounded), fed by ablated S. frugiperda larvae
(Ablated) or by unablated S. frugiperda larvae (unablated) for 24 hrs. Undamaged plants were used as a control. Total RNA was isolated from wounding site of maize leaves. The relative expression levels were determined in each biological sample and normalized to actin gene expression. Letters indicate significant differences by LSD test (p<0.05) (n=4, error bar indicates
SE).
40
Figure 3-7. Expression analysis of maize genes involving in plant defense in response to various treatments. Maize plants were wounding by paper punch (wounded), fed by ablated S. frugiperda larvae
(Ablated) or by unablated S. frugiperda larvae (unablated) for 24 hrs. Undamaged plants were used as a control. Total RNA was isolated from wounding site of maize leaves. The relative expression levels were determined in each biological sample and normalized to actin gene expression. Letters indicate significant differences by LSD test (p<0.05) (n=4, error bar indicates
SE).
41
(A) (B)
Figure 3-8. Bioassay of S. frugiperda larvae fed on maize genotypes subjected to various treatments. (A) Tx601 (B) Mp708 Maize plants were treated as described in Figures 3 and 4. The leaf
tissues were fed by naïve larvae (5.0-7.5mg) for 4 days. The final weight of larvae was recorded.
Letters indicate significant differences by LSD test (p<0.05) n=16-28 (Tx601); n=98-
115(Mp708), error bar indicates SE.
42
Figure 3-9. Amounts of leaf area (cm2) consumed by ablated versus unablated fall armyworm larvae. Detached leaf tissues were fed by fall armyworm larvae (4th instar) for 24 hours. The leaf areas of detached leaves before and after caterpillar feeding were scanned by SigmaScan Pro 5. The differences between original leaf area and the remaining leaf area after caterpillar feeding represented area eaten by fall armyworm larvae. Letters indicate significant differences by LSD test (p<0.05) n=10-11 (Tx601); n=9-10(Mp708), error bar indicates SE.
43
Figure 3-10. Scanning electron microscopy of two Lepidopteran larval spinnerets. Corn earworm (left); fall armyworm (middle and right). Scale bar = 10 µm. The red arrow indicates the larval spinneret.
Chapter 4
Ribosome-inactivating protein 2 protects maize against insect herbivores
Introduction
Plants are frequently attacked by voracious insect herbivores, and consequently they have evolved a broad range of defense mechanisms to combat these pests. The most effective plant defense mechanisms combine physical traits such as trichomes, thorns, and cuticles (Levin,
1973), and chemical substances to deter feeding and “poison” the insects. Inducible plant defense are classified into two types. Plant indirect defenses include production of green leaf volatiles and other volatile organic compounds that attract natural enemies of the attacking insects. Plant direct defenses rely on the accumulation of secondary metabolites, defensive proteins, and other toxic compounds (Howe and Jander, 2008). Plant secondary metabolites, which include glucosinolates,
44 2,4-dihydroxy-7-methoxy-1,4-benzoxazin-3-one (DIMBOA), alkaloids, and phenolics and other compounds, act as toxins in plant-insect interactions. A major class of defensive proteins includes protease inhibitors (PIs) that were first found to rapidly accumulate in potato and tomato after insect feeding (Green and Ryan, 1972). In the insect gut, the PIs interact with gut proteases and thereby inhibit their digestion function. Many transgenic plants expressing PI genes have been developed and showed detrimental effects on insect growth (Chen, 2008; Howe and Jander, 2008;
Zhu-Salzman et al., 2008). Furthermore, it is shown that plant oxidative enzymes are also involved in plant defense against insect herbivores (Chen, 2008). Polyphenol oxidases (PPOs), peroxidases, lipoxygenases, and ascorbate oxidases oxidize essential nutrients or form electophilic products, which react with the nucleophilic side chains of amino acids (Felton et al.,
1994; Constabel, 1999; Chen, 2008). These enzymes lower the nutritive value of plants to insects.
Besides the above proteins, some proteases, such as maize insect resistance 1-cysteine protease
(Mir1-CP) and leucine aminopeptidase (LAP), have been shown to defend plants against insect herbivory (Chao et al., 1999; Chen, 2008; Zhu-Salzman et al., 2008).
Herbivore or jasmonic acid inducible plant proteins have been found by microarray and proteomics analyses of insect infested plants (Reymond et al., 2000; Collins et al., 2010) . Most of these inducible proteins are found in plant tissues after insect feeding but before insect ingestion. However, recently it has been shown that several plant defense proteins can successfully survive digestion in the insect gut and are eliminated in the frass. In fact, it has been demonstrated that several jasmonate-inducible proteins (JIPs) have enzymatic activity in the insect midgut (Chen et al., 2005). Therefore, proteomic analysis of frass is a new approach that has been used to identify plant proteins that could play a role in defense against insect herbivory
(Chen et al., 2007). Using this technique, several JIPs were found in frass of Manduca sexta that reduce the ability of the insect to obtain essential nutrients from the plant (Chen et al., 2007).
One of these proteins is threonine deaminase isoform 2 (TD2) that was found in the frass of M.
45 sexta reared on tomato (Chen et al., 2007). TD2 is synthesized as proenzyme form in the tomato.
When insects ingest tomato leaves, TD2 is converted to the processed form in the insect gut where it plays an anti-nutrient role by reducing the levels of threonine, a necessary amino acid for phytophagous insects (Chen et al., 2007). In this study, we have used proteomic analysis of fall armyworm (Spodoptera frugiperda) frass to determine if there are similar herbivore-induced defensive proteins in maize. One of predominant proteins found in this analysis was identified as ribosome-inactivating protein 2 (RIP2).
Ribosome-inactivating proteins (RIPs) are enzymes that have site-specific RNA N- glycosidase activity that arrest translation (Bass et al., 2004). RIPs block translational elongation by depurinating residues on the large ribosomal RNA component of the ribosome (Endo et al.,
1987; Endo and Tsurugi, 1987; Nielsen and Boston, 2001). In 1925, it was first reported that pokeweed RIPs inhibit viral infection (Irvin, 1983; Nielsen et al., 2001). Since then it has been shown that plant RIPs play a role in defense against viruses, pathogens, and insects (Nielsen et al., 2001; Peumans et al., 2001; Bertholdo-Vargas et al., 2009). Based on protein structure, plant
RIPs are classified into three types (Nielsen and Boston, 2001). Type 1 RIPs consist of a single polypeptide chain with an approximate molecular mass of 30 KDa (Nielsen and Boston, 2001).
Pokeweed antiviral protein, soapwort saporin, and barley translation inhibitor are grouped as
Type 1 RIPs (Nielsen and Boston, 2001). Type 2 RIPs have two polypeptide subunits, which have an enzymatic domain and galactose binding domain linked by disulfide bonds (Nielsen and
Boston, 2001). Ricin and abrin are well-known, highly toxic type 2 RIPs (Nielsen and Boston,
2001). Type 3 RIPs are synthesized as inactive precursors (proRIPs) that require proteolytic processing to form the processed RIP (Nielsen and Boston, 2001). JIP60 in barley and RIP in maize are classified as type 3. Two RIP isoforms (RIP1 and RIP2) have been identified in maize
(Bass et al., 2004). Maize RIP1 is expressed in kernel, where it is believed to protect the seed from pathogen infection (Nielsen et al., 2001). Over-expression of maize RIP1 in transgenic
46 tobacco and rice enhanced their resistance to insects and pathogens (Kim et al., 2003; Dowd et al., 2006). Unlike RIP1, which is found only in the kernel, RIP2 is expressed throughout the plant from the leaves to the tassel, but it is not present in the kernel. RIP2 can be induced by water stress (Bass et al., 2004). The physical locus of RIP2 gene on the maize genome is chromosome
7, bin 7.04, where there is a strong quantitative trait loci for caterpillar resistance (Bass et al.,
1995; Bass et al., 2004; Brooks et al., 2005). These studies suggest that maize RIP2 may play an important defensive role against insects in the plant’s vegetative tissues, as RIP1 does in kernel.
This study was undertaken to determine if putative plant herbivore defense proteins could be found in the frass of fall armyworm caterpillars feeding on maize. We also determined if one of these proteins, RIP2 was toxic to fall armyworm larvae and investigated the factors regulating its accumulation in maize.
Results
Identification of maize proteins in insect frass
Since the previous analysis of M. sexta frass proteome focused on identification of herbivore defense proteins on dicot, tomato (Chen et al., 2007), we were interested in finding potential defensive proteins in frass collected from caterpillars that fed on the monocot, maize.
Plants from two maize genotypes (Tx601 and Mp704) were used to feed fall armyworm larvae for 24 hrs to trigger plant defenses. Tx601 is susceptible to fall armyworm feeding (Williams et al., 1989), whereas Mp704 is highly resistant to a number of lepidopteran pests (Williams and
Davis, 1982). Frass was collected from the insects and its proteome was determined by mass spectrometry. The proteome was analyzed by Dr. Gregg Howe at Michigan State University.
47 Table 4-1 shows the most abundant maize proteins are both found in frass from insects reared on either insect-susceptible (Tx601) or insect-resistant (Mp704) maize genotypes. The complete list of maize proteins identified in the frass is showed in Supplemental 4-1. The number of peptide counts in Table 4-1 and Supplemental 4-1 represents the number of peptides for a particular protein in the frass sample that were identified by mass spectrometry. These results supported the findings of Chen et al., (2007) demonstrating that potential plant defensive proteins can resist digestive proteases in the insect gut and are eliminated in the frass. Beta-D-glucosidase in maize has been shown to activate DIMBOA, a secondary metabolite that is toxic to the
European corn borer (Ostrinia nubilalis) against insects (Yu et al., 2009). Lipoxygenase (LOX) and allene oxide synthase (AOS) catalyze steps in the jasmonic acid (JA) biosynthetic pathway and it is the production of JA in response to insect feeding that triggers many plant defenses against herbivory (Howe and Jander, 2008). Overexpressing different potato AOS (StAOS2) alleles in Arabidopsis resulted in varying levels of pathogen resistance to Erwinia carotovora
(Pajerowska-Mukhtar et al., 2008). Furthermore, 9-LOX (ZmLOX3) in maize plays a defense role against nematodes (Meloidogyne incognita) and fungi (Aspergillus flavus and Aspergillus nidulan) (Gao et al., 2008; Gao et al., 2009). Maize RIP1 has a toxic effect on fungi and insects
(Dowd et al., 1998; Nielsen et al., 2001). A putative fruit protein with unknown function that contains an oxidoreductase domain also was identified. A pathogen-responsive oxidoreductase
(drd-1) in potato is induced in response to E. carotovora (Montesano et al., 2003). Endo-1, 3- glucanase (PR-2) in tomato accumulates in response to virus infection (citrus exocortis viroid)
(Domingo et al., 1994). In rice, the expression of one endo-beta-glucanase isoform responded to wounding, methyl jasmonate, and ethephon (Akiyama et al., 2009). In maize, one of peroxidase isoforms (px5) is associated with plant disease resistance (Dowd and Johnson, 2005). The growth rate of two major maize pests (Helicoverpa zea and Lasioderma serricorne ) was slowed when they were fed another maize peroxidase (px11) (Chen et al., 2008). Another frass protein was
48 similar to a GDSL-like lipase (GLIP1) found in Arabidopsis that is involved in plant defense against the necrotrophic fungus Alternaria brassicicola (Oh et al., 2005). GLIP1, which is regulated by ethylene, could trigger systemic resistance signaling in plants after fungal infection
(Oh et al., 2005). GDSL-like lipase (GLIP2) in Arabidopsis also plays a role in plant defense against pathogens (Lee et al., 2009). Chitinases, which also were found in frass, have been shown to be defensive proteins in plants. For example, the over-expression poplar chitinase (WIN6) in tomato retarded the development of Colorado potato beetle (Lawrence and Novak, 2006). Ectopic expression of a rice chitinase in peanuts enhanced fungus resistance to Cercospora arachidicola
(Iqbal et al., 2011). In addition, transcripts of the maize chitinase gene Prm3 increase in response to fall armyworm feeding in maize (Shivaji et al., 2010). We also identified several peptides of leucine aminopeptidase from the frass (Table 4-1). Leucine aminopeptidase A (LapA) in tomato increases in response to wounding, exogenous methyl jasmonate, pathogen infection, and insect feeding (Chao et al., 1999; Chen et al., 2005; Chen et al., 2007; Zhu-Salzman et al., 2008). LapA protein has been detected in the midgut and frass of M.sexta (Chen et al., 2005; Chen et al.,
2007), and its overexpression in tomato delays M. sexta growth and development (Lee et al.,
2009). These proteomic studies indicated that putative maize herbivore defense proteins are present in fall armyworm frass and there was little difference in the proteins found in the frass from the two inbreds differing in resistance to fall armyworm.
Two maize RIP genes respond to insect herbivore differently
Two RIP genes (RIP1 and RIP2) that share 70% amino acid sequence identity have been identified in maize (Bass et al., 2004). A previous study showed that RIP2 transcripts could be detected in leaf tissues but not in the kernel where the RIP1 gene was specifically expressed (Bass et al., 2004). Furthermore, it has been shown that the RIP2, and not RIP1 protein, was expressed
49 in vegetative tissues (Bass et al., 2004). These finding suggested that that RIP2 was probably the isoform found in caterpillar frass. However, due to high similarity of RIP1 and RIP2 sequences, it was difficult to determine if RIP1, or RIP2, was found in the frass sample. Thus, we used quantitative real-time-PCR (qRT-PCR) to detect RIP1 and RIP2 gene expression in maize plants infested with fall armyworm larvae. RIP2 transcripts were highly expressed in caterpillar fed leaves and those for RIP1 were undetectable in both control and 24h fed plants (Fig. 4-1). Thus, it is most likely that RIP2, not RIP1, is the isoform present in the frass of caterpillars reared on maize leaves (Table 4-1).
RIP2 can be found in maize leaf and frass
Because RIP2 is synthesized as an inactive proenzyme in maize (Bass et al., 2004), the form of RIP2 present in fall armyworm-fed maize leaves and caterpillar frass was determined using immunoblot analysis. The proenzyme form of RIP2 with a predicted size of 30 kD size was detected in the leaf tissues (Fig. 4-2). However, there was a smaller polypeptide in the frass sample that could be the processed form of RIP2 (pRIP2) (Fig. 4-2). Combined with the frass proteomics data, it appears that the processed RIP2 (pRIP2), not the proenzyme form, was detected by mass spectrometry in the frass. Due to the relatively high number of peptide counts for RIP2 in the frass data (Table 4-1), it appears that RIP2 may be abundant in the frass of caterpillars reared on maize. To determine if proenzyme form of RIP2 is commonly induced in insect-fed maize, immunoblot analysis was used to detect RIP2 in 13 maize inbred lines including
Mo17, B73, and W64A, and two teosinte subspecies. In the remaining articles in this dissertation, if there is no specific description in the experiment, the immunoblot analysis result was only detected proenzyme form of RIP2. The proenzyme form of RIP2 was detected in all of the tested maize genotypes (Fig. 4-3). This result shows that proenzyme form of RIP2 expression in
50 response to insect feeding is a wide spread phenomenon in maize. However, we failed to detect proenzyme form of RIP2 in sorghum (data not shown).
Effect of heterologous RIP2 on fall armyworm larval growth
The RIP1 protein has been shown to reduce the growth of several insect pests and fungal pathogens, including Aspergillus flavus (Dowd et al., 1998; Nielsen et al., 2001). To investigate whether RIP2 protein also is toxic to lepidopteran larvae, we over-expressed and purified recombinant RIP2 (rRIP2) from E. coli. Since a previous study indicated that that rRIP2 could be processed by papain (Bass et al., 2004), purified rRIP2 was treated with papain-coated beads to generate the processed form rRIP2 (prRIP2). To investigate that whether rRIP2 can be processed by caterpillars, the frass from FAW larvae reared on artificial diet containing rRIP2 was collected. Fig. 4-4 shows maize RIP2 in FAW-induced maize leaf, processed maize RIP2 in FAW frass, purified rRIP2, prRIP2 which rRIP2 treated with papain-coated beads, and prRIP2 collected from frass which FAW larvae fed with purified rRIP2. The polypeptides of purified rRIP2 containing 45 histidine resuidues fused with maize RIP2 coding sequence were bigger than maize
RIP2 in leaf (Bass et al., 2004). Maize RIP1 can be processed to become two polypeptides of processed RIP1 (Dowd et al., 1998). The processed prRIP2 treated with papain shows the predicted two prRIP2 fragments. Fig. 4-4 (lane 5) shows that rRIP2 can be processed by the caterpillar to become prRIP2. However, most of larger prRIP2 polypeptides were digested by
FAW larvae. This result could explain why we can only detected one processed RIP2 polypeptides in FAW frass.
Prior to caterpillar feeding bioassays, the amount of RIP2 expressed in plants exposed to larval feeding was determined to be approximately 0.8 µg/100 mg fresh weight. Thus, this concentration of either rRIP2 or processed prRIP2 was incorporated into artificial diet and used
51 for bioassays. After 5 days of incubation, both rRIP2 and prRIP2 fed larvae had lower larval weights than those fed either bovine serum albumin (BSA) or buffer (Fig. 4-5). A comparison of
RIP2 fed larvae with controls indicated that their weight was suppressed by 26%. These results indicate that both forms of RIP2 are toxic to fall armyworm larvae at a concentration typically found in caterpillar fed whorls. The data in Fig. 4-4 indicated that both rRIP2 and prRIP2 have the same inhibitory effect on insect growth. We have shown that caterpillar digestive enzymes can process RIP2 from proenzyme to processed form (Fig. 4-4). In addition to caterpillar enzymes, we also wanted to know whether plant proteases are able to process proRIP2. Since
Mir1-CP is an insecticidal cysteine protease in maize (Pechan et al., 2002), rRIP2 was incubated with this enzyme and the results (Fig. 4-6) indicated that Mir1-CP can process rRIP2. This shows that in addition to disrupting the fall armyworm peritrophic matrix (PM) (Pechan et al., 2002;
Mohan et al., 2006), Mir1-CP can also process the precursor of RIP2 and could enhance its insecticidal effects.
Insect herbivore, not mechanical wounding, induces RIP2 induction
A previous study reported that RIP2 transcripts were induced by water stress (Bass et al.,
2004). Since we have detected RIP2 in the caterpillar frass and determined that RIP2 transcripts are highly expressed in the maize leaf under herbivore attack, we propose that feeding by fall armyworm larvae elicits RIP2 expression in maize. To test this, maize plants were fed with fall armyworm larvae for 24 hrs or wounded by paper punch to mimic mechanical wounding. Figure
4-7 shows that RIP2 transcripts increased approximately 100-fold in abundance after herbivore attack. However, plants without insect feeding (control group) and those that were mechanical wounded did not accumulate RIP2 transcripts. Immunoblot analysis further showed that RIP2 protein abundantly accumulated in response to herbivory, but not other treatments (Fig. 4-8).
52 Both RIP2 transcripts and protein expression data show that caterpillar feeding is needed to trigger RIP2 expression but mechanical wounding is not.
Characterization of RIP2 profile under insect herbivore attacking
Since fall armyworm feeding triggered the accumulation of both RIP2 transcripts and protein, we wanted to determine the rapidity of this response. RIP2 transcript levels increased slightly 30 min after insect feeding, but were not significantly higher than the control. However, there was a significant increase in transcript levels between 1 and 4h and the levels remained high at 24 h (Fig. 4-9). The immunoblot analysis indicates that RIP2 was present in low levels prior to fall armyworm feeding and increased in abundance up to 24h (Fig. 4-10). These data indicated that continuous caterpillar feeding will induce the accumulation of RIP2 transcripts and protein.
The presence of a RIP2 polypeptide in caterpillar frass (Table 4-1) suggests that RIP2 protein may be very stable in the leaf. To test this,, maize plants were first fed by fall armyworm larvae (one larva per plant) for 24 hours to induce RIP2 protein expression. Afterward, the larva was removed and leaf samples were collected at subsequent time points. After 24h of feeding
RIP2 protein was abundant in the leaf up to 4 days after caterpillar removal and dramatically decreased in abundance at 5 days (Fig.4-11). These results indicate that the RIP2 protein is very stable in the plant even in the absence of fall armyworm feeding.
RIP2 is expressed locally and during maize vegetative development.
In dicot plants, such as tomato, there often is a systemic induction of defense proteins such as PIN1 (Montesano et al., 2003). To determine if fall armyworm feeding could systemically induce RIP2 accumulation, larvae were placed in “cages” on the maize leaves to limit their
53 movement. Several leaf samples were collected distally from the feeding site and non-fed leaves.
Immunoblot analysis of these samples showed that RIP2 accumulation was induced only near the feeding site (Fig. 4-12), hence it appears that there is no widespread systemic induction of RIP2.
The vegetative portion of the maize lifecycle can be roughly divided into two stages, juvenile and adult. The juvenile stage persists from germination until the plants have five to six leaves and then there is a transition to the adult stage (Freeling, 1992). The time of transition is positively correlated with the survival and growth rates of insect herbivores, i.e. plants that undergo this transition earlier in their life cycle are more resistant to caterpillar feeding (Williams et al., 1998; Brooks et al., 2007). To determine if there were temporal differences in RIP2 expression during maize development, samples were collected from the whorls of plants at several vegetative stages and analyzed for RIP2 protein expression in control and caterpillar- infested plants. Figure 4-13 shows that RIP2 accumulation was induced by caterpillar feeding in the juvenile stages (V2 and V4), during the transition from juvenile to adult (V6), and the adult
(V8, V10) vegetative stages. These results indicate that RIP2 can protect maize against insect herbivory during all of the tested developmental stages.
Phytohormones regulating RIP2 expression
Several phytohormones salicylic acid (SA), jasmonic acid (JA), and ethylene (ET), are known to be involved in regulating plant responses to biotic stresses (Park et al., 2002). Since caterpillar feeding induces RIP2 expression, it is likely that one or more of these phytohormones is involved in regulating its expression. When unwounded plants were treated with MeSA,
MeJA, and ethephon (to generate ET), there was no induction of RIP2 expression (data not shown). Since caterpillars damage plant tissues during feeding, we mechanically wounded the plants immediately prior to hormone treatment. Plants that were both wounded and treated with
54 ethephon accumulated RIP2 (Fig. 4-14). JA is usually involved in regulating plant defenses against chewing insect herbivores. MeJA treatment after wounding failed to trigger RIP2 expression (Fig. 4-15), but treatment with a combination of wounding, MeJA and ethephon resulted in induction. This result suggests that ET, not MeJA, regulates RIP2 expression. The combination of these two phytohormones did not induce RIP2 expression in the absence of mechanical wounding. These data indicate that wounding alone is not sufficient to trigger RIP2 expression and that it is dependent on ET. To determine if this was the case, EthylBloc (1- methyl-cyclopropene) that irreversibly binds to ET receptors, was used to test RIP2 expression when ET perception was blocked. Fig. 4-16 shows that there was a 2 hr delay in RIP2 accumulation in response to caterpillar feeding when the plants were treated with EthylBloc. This analysis combined with the treatment with ethephon suggests that ET is needed for RIP2 expression. Analysis of the RIP2 promoter sequence from the maize genome database
(www.maizesequence.org) (Schnable et al., 2009) using PLANTCARE (Lescot et al., 2002), predicted that it has several cis-acting regulatory elements that should respond to MeJA. In a previous study, it was shown that the same MeJA concentration (0.01%) that was used in this experiment induced the expression of another maize insect defense protein, Mir1-CP in the genotype Mp708 (Ankala et al., 2009). However, Mp708 has constitutively elevated levels of JA and hence application of a lower exogenous MeJA concentration could have been sufficient to stimulate defense protein accumulation in this inbred (Shivaji et al., 2010). Therefore, a range of
MeJA concentrations (0.01% to 0.1%) was tested to determine if MeJA was involved in regulating RIP2 expression. In the presence of wounding, 0.03% MeJA induced RIP2 expression.
However, only plants treated with the highest concentration of MeJA (0.1%) induced RIP2 expression in the absence of wounding (Fig. 4-17). These data suggest that maize plants might need to be treated with relatively high MeJA concentrations to trigger RIP2 accumulation in the absence of wounding. MeSA did not induce RIP2 expression, even in combination with
55 mechanical wounding (data not shown). Since a previous study reported that RIP2 could be induced by water stress (Bass et al., 2004), plants were treated with abscisic acid (ABA), the major plant hormone involved in the drought response (Jeffers et al., 2005). ABA and wounding treatment induced RIP2 expression (Fig. 4-18) and suggesting that water stress plays a role in its expression (Bass et al., 2004).
Discussion
Maize defensive protein in insect frass
Plants have evolved with a number of defense mechanisms to protect themselves against insect herbivory. Because insect herbivores consume foliage and use it for their growth and development, plants can counteract herbivory by synthesizing a number of antinutritional or toxic substances (Berenbaum, 1995; Felton and Gatehouse, 1996; Felton, 2005; Zhu-Salzman et al., 2008). When insects ingest this cocktail of antinutritional proteins, it causes “indigestion” and severely limits their ability to fully utilize plant nutrients and impairs their growth (Felton, 2005).
Several studies showed that some ingested plant proteins can remain intact in the insect gut either further move across the gut into hemolymph or be eliminated in the frass (Chen et al., 2005;
Jeffers et al., 2005; Chen et al., 2007; Zhu-Salzman et al., 2008). In this study, we identified several maize proteins in fall armyworm frass that could cause this “indigestion.” Lipoxygenase, peroxidase, and the putative fruit protein containing oxidoreductase may play an antinutritive role on plant defense (Zhu-Salzman et al., 2008). Lipoxygenase may not only be involved in the JA biosynthetic pathway, but could also produce substances that cause the loss of essential amino acids needed by the insects (Felton, 2005). It has been shown that lipoxygenase inhibits larval growth when corn earworm (Helicoverpa zea) feed on diets containing this enzyme (Felton et al.,
56 1994). Peroxidases produce quinones that are electrophiles and interact with the nucleophilic side chains of both free and protein amino acids (Felton, 2005). Chinitase and leucine aminopeptidase may display toxicity by attacking the insect digestive system (Zhu-Salzman et al., 2008). Insect chinitases have been shown to the damage the insect peritrophic matrix (PM) both in vitro and in vivo (Kramer and Muthukrishnan, 1998; Lawrence and Novak, 2006). Although there is no direct evidence showing that plant chitinases damage the insect PM, over-expression of the poplar chinitase, WIN6, in tomato retards insect development (Lawrence and Novak, 2006). Leucine aminopeptidase activity is detectable in the frass of tomato reared M. sexta (Chen et al., 2007).
With the proteomics data from M. sexta midgut and frass, leucine aminopepetidase A may function in the insect midgut to release Arg from the N terminus of peptides to reduce the food quality or damage insect gut (Chen et al., 2005; Chen et al., 2007; Zhu-Salzman et al., 2008).
Another protein found in the frass, beta-D-glucosidase may not involve in the activation
DIMBOA but could also be an elicitor to attract host parasitoids (Mattiacci et al., 1995). In this study, we have shown that RIP2 is induced by herbivory in maize and that it is toxic to fall armyworm larvae at a physiologically relevant concentration. Although the mode of action of the ribosome-inactivating proteins in the insect gut is unknown, it appears that gut enzymes are capable of processing proRIP2 to the processed form. RIP2 could harm the nutrient uptake by arresting the protein synthesis in the insect midgut cells.
Different expressed pattern of RIP genes
There have been many reports demonstrating that RIPs are developmentally regulated in plants. The highly toxic protein, ricin is only expressed in the seeds of castor bean (Ricinus communis). There are two ricin isoforms, D and E that are differentially expressed in seeds of different sizes (Despeyroux et al., 2000; Sehgal et al., 2011). Saporin in soapwort has several
57 isoforms that are differentially expressed in various organs (leaf, root, and seed) (Ferreras et al.,
1993). Saporin-L and saporin-S have different expression patterns during leaf development.
(Tartarini et al., 2010). Single-chain, ribosome-inactivating protein (SCRIP) in bitter melon is highly expressed in seeds and flowers and only moderately expressed in roots and stems (Xu et al., 2007). Two beetin proteins in sugar beet are detected in the adult plants but not in seeds or young plants (Iglesias et al., 2008). PAP-1 and PAP-2, two RIP paralogs in pokeweed, are expressd in spring and summer leaves, but PAP-S is expressed in seeds (Irvin, 1983; Irvin and
Uckun, 1992; Kawade et al., 2008). Cinnamomin in camphor is stored in cotyledons and degraded after germination (Liu et al., 2002; Kawade et al., 2008). Two RIP genes (SoRIP1 and
SoRIP2) are involved in the somatic embryogenesis in spinach (Kawade et al., 2008). In addition to these proteins, it appears that maize RIP1 and RIP2 expression is differentially regulated in seed and vegetative tissues, respectively.
Several different phytohormones are involved in the regulation of RIP expression. For example, jasmonate-induced protein (JIP60), a RIP gene in barley, is regulated by jasmonate
(Reinbothe et al., 1994), whereas SCRIP in bitter melon is up-regulated by abscisic acid (ABA)
(Xu et al., 2007). In the rice genome, 31 RIP isoforms been identified (Jiang et al., 2008), but of these, only 20 transcripts are detected through plant development and only three respond to ABA
(Jiang et al., 2008). SoRIP1 and SoRIP2 in spinach are up-regulated by gibberellic acid (GA) and down-regulated by ABA (Kawade et al., 2008). Saporin-L and saporin-S in soapwort are expressed in fully expanded leaf tissues, but also differentially respond to mechanical injury and
ABA treatment (Tartarini et al., 2010). Beetins from leaves of sugar beet are up-regulated by salicylic acid and hydrogen peroxide (Iglesias et al., 2008). SCRIP in bitter melon and PAP-H in pokeweed are induced by ethylene (Park et al., 2002; Xu et al., 2007). In this study, we determined that RIP2 expression also was regulated by several different phytohormones,
58 ethylene, JA and ABA. Thus, it appears that there is no consistent pattern of hormonal regulation of RIP-like protein expression.
The role of RIP2 in maize against insect herbivores
Our results show that RIP2 accumulates in maize leaves as proenzyme form that appears to be proteolytically cleaved to the processed form that is found in frass (Fig. 4-2). This processing is similar to that of the tomato herbivore defense protein, threonine deaminase 2
(TD2) (Chen et al., 2007). The processed form of TD2 has been detected in tomato and the gut lumen and frass of phytophagous insects (Chen et al., 2005; Chen et al., 2007). Furthermore, TD2 enzyme activity can be detected in frass collected from tomato-reared M. sexta larvae (Chen et al., 2007). This finding indicates that some plant defense proteins not only are resistant to digestive proteases, but also have post-ingestive defense function on insect gut lumen (Chen et al., 2007). TD2 exists as proenzyme form in the tomato leaf and the regulatory domain that inhibits enzyme activity is removed in the insect gut and the processed enzyme is excreted in the frass (Chen et al., 2007). A further study confirmed that a chymotrypsin-like protease in the lepidopteran gut processed TD from the proenzyme to the active form (Gonzales-Vigil et al.,
2011). Our results suggest that insect gut proteases also process proRIP2 to the active form as is the case with TD2.
Proenzymes, the inactive form of enzymes, have suppressed activity until they are proteolytically processed to the active form (Neurath and Walsh, 1976; Neurath, 1989; Balconi et al., 2010). The internal segment of maize RIP1 is domain thought to suppress its enzymatic activity until proteolytic cleavage has occurred (Hey et al., 1995). Due to the highly similarity of protein sequence between RIP1 and RIP2, RIP2 might be regulate its enzyme activity in a similar manner to RIP1. The enzymatic activity of plant RIPs have been broadly studied since several
59 RIPs are used in medical research (Barbieri et al., 1979; Gasperi-Campani et al., 1985; Stirpe et al., 1992; Nielsen and Boston, 2001). Animal and plant ribosomes are susceptible to plant RIPs, such as PAP (Hartley and Lord, 1993; Hartley et al., 1996; Madin et al., 2000; Nielsen and
Boston, 2001). However, most cereal RIPs exhibit low activity with plant ribosomes. For example, maize ribosomes are resistant to processed RIP1 (Bass et al., 1992; Hey et al., 1995;
Krawetz and Boston, 2000; Bass et al., 2004). Thus, the idea that RIP2 synthesized as proenzyme form in order to protect maize ribosomes themselves may not the true. Due to long half-life of
RIP2, it suggests that RIP2 is synthesized as proenzyme form to be more stable in the leaf tissues until proteolytic cleavage occurs.
Due to the rapid induction of RIP2 in response to caterpillar feeding, it might play a role in the early defense against insect herbivory (Fig. 4-9~4-10). Several well-known maize defense proteins are involved in the rapid response to insect herbivore attack. Maize insect resistance 1- cysteine protease (Mir1-CP) accumulated near the wounding site as early as 1 hour after insect infestation (Pechan et al., 2000) and is known to be involved in maize defense against insect herbivory (Pechan et al., 2000). With the finding that Mir1-CP can process recombinant RIP2 to processed RIP2 and that both Mir1 and RIP2 are expressed near the wounding site as early as one hour, these two proteins may synergize the maize herbivore defense pathway. Other genes that are rapidly induced in response to wounding in maize include wound-induced protein (WIP1) transcripts in maize coleoptile that are induced as early as 30 minutes after wounding (Rohrmeier and Lehle, 1993) and maize proteinase inhibitor (MPI) transcripts that increase as early as 20 minutes after wounding (Tamayo et al., 2000). Therefore it is likely that RIP2 and Mir1-CP,
WIP1, and MPI are involved in the early stages of maize defense against herbivory
In maize leaves, RIP2 may play a specialized role. This idea is supported by the fact that induced RIP2 protein is very stable in the leaf tissue up to four days (Fig. 4-11). One explanation for this long half-life of RIP2 protein is that it responds to the insect feeding behavior. Mir1-CP
60 has been shown to persist up to seven days (Pechan et al., 2002). When insect herbivores feed on plants, their eating pattern is not continuous with no pause. Their feeding period often is separated by multiple interfeeding gaps (Reynolds et al., 1986). During these gaps, they may need time to digest leaf tissue or prepare to molt without ingesting food. Because it usually takes one or two days to molt in the field, plants need to trigger and maintain their defense system for the next insect attack. RIP2 would be one of the defense proteins that plants employ in this type of defense. Another explanation could be that when insect herbivores feed on maize, they create many open wound sites that could be the site for fungal infection. Because kernel RIP1 can inhibit fungal growth, it is possible that RIP2 in the leaf might function in a similar manner
(Nielsen et al., 2001).
We have observed that the size of the RIP2 protein varies among several maize inbreds
(Fig. 4-3). We have checked RIP2 transcript sequences in two maize genotypes which have different polypeptides sizes (Fig. 4-3 lane 8 and 11) and found that they are the same. Therefore post-translational protein modification could cause the different protein sizes. Protein glycosylation has been shown to occur in RIPs. The difference between the two RIP proteins
(BE27 and BE29) in the sugar beet leaf is the level of glycosylation on the same polypeptide chain (Iglesias et al., 2005). Furthermore, the level of glycosylation of ricin from castor bean is correlated with its toxicity (Sehgal et al., 2011). With above studies, the effect of RIP2 glycosylation might affect its half-life or enzymatic activity. Further studies will address this question.
The physical locus of maize RIP2 is in bin 7.04 on chromosome 7 of the maize genome, unlke RIP1 that is located in bin 8.05 on chromosome 8 (Bass et al., 1995; Bass et al., 2004).
Maize chromosomes 1, 5, 7, and 9 contain major loci for insect resistance to fall armyworm and southwestern corn borer (Diatraea grandiosella) (Brooks et al., 2007). The RIP2 locus is found in a strong quantitative trait loci for insect resistance in maize (Brooks et al., 2005; Brooks et al.,
61 2007). Due to the 26% reduction rate at RIP2 bioassay (Fig. 4-5) and its physical locus, RIP2 could be an important caterpillar resistance gene in maize.
More than 130 RIP genes have been identified in a number of different plant species
(Girbes et al., 2004; Jiang et al., 2008). However, there is no RIP ortholog in A. thaliana
(Peumans et al., 2001; Kawade et al., 2008). It has been shown that there are at least 31 RIPs in the rice genome (Jiang et al., 2006). Some of these rice genes are regulated by abiotic and biotic stresses (Jiang et al., 2006). This suggests that each RIP gene could respond to a specific environmental stress or combination of stresses. RIPs have been reported to be regulated by abiotic stresses, such as drought, salt, H2O2, and heat (Stirpe et al., 1996; Rippmann et al., 1997;
Bass et al., 2004; Iglesias et al., 2005; Jiang et al., 2008). RIP2 transcripts in maize are up- regulated after water deficit and returned to basal level after re-watering (Bass et al., 2004). This pattern is similar to rab17 that is an ABA-regulated gene in maize (Vilardell et al., 1990; Bass et al., 2004). RIPs also respond to biotic stresses. For example, SCRIP in bitter melon is expressed in very low amount in leaves during development but is up-regulated by fungal infection and wounding (Xu et al., 2007). Six of the thirty-one RIP genes in rice responded to the bacterium
Xanthomonas oryzae pv oryzae (Jiang et al., 2008). Beetins from sugar beet leaves are induced by the artichoke mottled crinkle virus (AMCV) (Iglesias et al., 2008). Pokeweed root exudates containing PAP-H, a type I ribosome-inactivating protein, inhibited the growth of soil-borne fungi (Park et al., 2002). Maize RIP1 in kernel reduced the survival of multiple insects and reduced their growth rate (Gatehouse et al., 1990; Dowd et al., 1998). Furthermore, Maize RIP1 also reduces Aspergillus nidulans and A. flavus infestation (Nielsen et al., 2001). In our study, maize RIP2 expression responded to insect feeding and treatment with ABA. Our results suggest that maize RIP2 can be regulated by abiotic and biotic stresses as other plant RIPs.
Our data showed that recombinant RIP2 has toxic effects on fall armyworm (Fig. 4-5).
RIP2 retards the larval growth by 26%. Several other RIP genes have been transformed into
62 several crops to study their ability to counteract biological stresses. Coexpression of tobacco peroxidase and maize RIP1 in transgenic tobacco enhanced resistance to Helicoverpa zea and
Lasioderma serricorne (Dowd et al., 2006). Furthermore, transgenic rice that co-expressed a modified maize RIP1 and rice chitinase gene improved fungal resistance to sheath blight (Kim et al., 2003). When pokeweed PAP was transformed into tobacco and potato, these plants had increased resistance to several different viruses (Lodge et al., 1993). Ectopic expression of pokeweed RIP in tobacco also increased fungus resistance (Corrado et al., 2005). It also has been reported that expression of an elderberry RIP gene in tobacco enhanced insect resistance
(Shahidi-Noghabi et al., 2009). Taken together, these observations demonstrate that plant RIPs are toxic to a variety of plant viruses, pathogens, and insects herbivores.
Further studies reported that several plant type 1 RIPs have been evaluated for their entomotoxic effect on Anticarsia gemmatalis and S. frugiperda (Bertholdo-Vargas et al., 2009).
Two RIPs, ricin and saporin have been shown to suppress the development of Lepidoptera and
Coleoptera (Gatehouse et al., 1990). In addition to inhibiting the growth of A. flavus (Krawetz and Boston, 2000), maize RIP1 also inhibited the growth of multiple Lepidoptera and Coleoptera species (Dowd et al., 1998). A few studies have investigated the mechanism of RIP toxicity to insects. SNA-I from elderberry caused cell apoptosis in the gut tissues of Acyrthosiphon pisum and S. exigua (Shahidi-Noghabi et al., 2010). SNA-I and SNA-II also induced caspase-3 like activity in midgut cell line of Lepidoptera (Shahidi-Noghabi et al., 2010). These findings suggest that RIPs are toxic to insect herbivores because they trigger apoptosis in the midgut. These studies might explain the toxic effect of RIP2 on S. frugiperda.
63 Materials and Methods:
Plant materials, insect rearing, and frass collection
Maize (Zea mays) genotypes that are resistant (Mp708) and susceptible (Tx601) to fall armyworm feeding were obtained from Dr. W. Paul Williams (USDA-ARS Corn Host Plant
Resistance Research Laboratory, Mississippi State University). Both genotypes were fed to caterpillars for the frass proteomic analysis. Unless otherwise noted, Tx601 was used for the experiments reported here. Seeds were sown in pots filled with topsoil (Hagerstown loam) in the
Crop and Soil Sciences greenhouse at The Pennsylvania State University, University Park, PA.
Well-watered maize plants were grown until they were at the V8 stage (eight fully emerged leaves) (Ritchie, 1986). Plant tissues surrounding the wounded area near the whorl were collected after caterpillar feeding or wounding with a paper punch (6mm diameter, ten times per plant). The tissue samples were stored at -80°C after immediately freezing in liquid nitrogen.
Fall armyworm (Spodoptera frugiperda) eggs were obtained from USDA-ARS Corn
Host Plant Resistance Research Laboratory, Mississippi State University. Larvae were reared on an artificial diet (Peiffer and Felton, 2005) in a 27°C incubator with a 16 h photoperiod until they molted to the 5th instar before using. For the collection of S. frugiperda frass, maize plants were fed by 5th instar S. frugiperda larvae for 24 h. The leaves around the feeding sites were collected, placed in diet cups and fed to previous (5th instar) S. frugiperda larvae. Frass was collected within 24 hours after feeding and stored at -80°C after immediately freezing in liquid nitrogen.
Immunoblot analysis
The leaf tissue and caterpillar frass were frozen with liquid nitrogen and homogenized using the Geno/Grinder 2000 (SPEX CertiPrep, Metuchen, NJ). These samples were then
64 extracted with SDS-PAGE sample buffer (Eichenseer et al., 2010). The protein concentration in the extraction was determined by RC-DC Protein assay (Bio-Rad. Hercules, CA). Equal amount of protein were loaded on each lane of a gel and separated by SDS-PAGE (ref). The SDS-PAGE gel was blotted onto the nitrocellulose membrane by Panther Semi-Dry Electroblotter (Thermo
Scientific Owl, MA). Polyclonal antibody to RIP was obtained from Dr. Rebecca Boston (North
Carolina State University, Raleigh). This anti-RIP antibody cross reacted with both RIP1 and
RIP2 (Bass et al., 2004). Western blots were carried out using 1: 10,000 diluted anti-RIP antibody and 1:10,000 diluted anti-rabbit secondary antibody with HRP conjugated (Thermo Fisher
Scientific, Rockford, IL). Immunoreacting proteins were detected by chemiluminescence (West
Femto Maximum Sensitivity Substrate, Thermo Scientific, MA).
RIP2 bioassays and RIP2 digestion
Recombinant RIP2 (rRIP2) was produced and purified as described by Bass et al.(2004).
Processed RIP2 (prRIP2) was made by treating the recombinant protein with immobilized papain
(Thermo Fisher Scientific, Rockford, IL). The papain-beads-prRIP2 mixture was centrifuged and the supernatant containing the processed RIP2 (prRIP2) was collected. For toxicity bioassays, the appropriate amount of bovine serum albumin (BSA), rRIP2, or prRIP2 (0.8 µg protein/100mg artificial diet) was placed into 24-well Multiwell plate (BD Falcon, Franklin Lakes, NJ) containing artificial diet (Peiffer and Felton, 2005). An equal volume of phosphate buffered saline (PBS) buffer served as the control. The plates were air-dried and a single larva (one day after hatching) was placed in each well. The plates were covered with Breath EasyTM (USA
Scientific Inc, FL). Each treatment had 24 larvae. The plates were incubated at 26ºC incubator for
5 days with a 16 hr photoperiod before the final larval weight was determined. The results were analyzed by SAS statistical software (SAS Institute, Cary, NC).
65 Recombinant Mir1-CP (10µg/ml) from a previous study (Mohan et al., 2008) was mixed with recombinant RIP2 (1mg/ml) at 37°C incubator for 24 h. The cleavage of rRIP2 was examined by immunoblot analysis.
Phytohormone treatments
The effect of exogenous ethylene application was tested by spraying maize plants with 3 mM 2-chloroethylphosphonic acid (Ethephon, Sigma-Aldrich, St. Louis, MO). The ethephon solution was sprayed on the plants. Methyl jasmonate (MEJA, Bedoukian Research Inc.,
Danbury, CT) was first diluted 10-fold with ethanol to become 10% MEJA solution (vol/vol).
The fresh working solution of 0.01% MEJA was prepared with sterile water. The 0.01% MEJA and 1 mM methyl salicylate (MESA, MP Biomedicals Inc., Solon, OH) was sprayed on the plants. The ethephon, MEJA, and MESA concentrations used were similar to those used on maize in a previous study (Ankala et al., 2009). Abscisic acid (ABA, Sigma-Aldrich, St. Louis, MO) was first dissolved as 50 mM ABA in ethanol. The fresh working solution of 300 µM ABA was sprayed on plants. Each plant hormone was sprayed on two individual plants for both wounded and unwounded treatments. Plants were wounded with a paper punch to mimic mechanical wounding. For controls the same amount of 0.1 % ethanol in sterile water was sprayed on wounded and unwounded plants. Leaf tissue was collected after a 24 h treatment.
Ethylene perception was blocked by treating plants with 1-methylcyclopropene (1-MCP)
(EthylBloc, Floralife, Walterboro, SC). Plants were placed in plastic chambers (1.2m x 0.395m x
0.395m). The EthylBloc powder (950 mg or 0.14% 1-MCP) was dissolved in 19 ml EthylBloc mixing solution in a 50ml centrifuge tube inside the chambers. The 1-MCP concentration was similar to those used for maize in a previous study (Ankala et al., 2009).
66 Analysis of developmental and systemic expression
Maize plants of different vegetative stages (V2, V4, V6, V8, and V10) (Ritchie, 1986) were infested with two or three of 3th or 5th instar larvae per plant. In each stage, 2 to 6 plants were harvested for control and feeding treatments. Leaf tissue adjacent to the feeding sites was collected and immediately frozen in liquid nitrogen.
For systemic analysis of RIP2 expression five S. frugiperda larvae were placed in the center of unexpanded eleventh leaf in the whorl of V8 stage plants for 24 hrs. A small bag that limited their movement was placed around the larvae. Leaf samples from the feeding site (0 cm) and distal to the site (3 and 5 cm) were collected. Samples also were taken from other leaves
(unexpanded 9th and 10th leaf) of the same plant. Tissues from four to five different plants were harvested for control and fed plants.
Insect feeding
Maize plants were infested with newly molted 5th instar larvae for various time points.
Unwounded plants were as controls. A paper punch was used to mimic mechanical wounding to plants (multiple wounding sites per plants). The leaf tissue around the wounding sites was collected and immediately frozen in liquid nitrogen and stored at -80°C.
RIP2 persistance after larval feeding
Maize plants were infested with newly molted 5th instar larvae (one larva per plant) for
24 h. Unwounded plants were as controls. The larvae were removed after 24 h and this was designated as 0 day. Samples were collected at several time points (1 to 6 days) after larval
67 removal. Leaf tissue around the feeding site was collected and immediately frozen in liquid nitrogen and stored at -80°C.
Quantitative RT-PCR (qRT-PCR)
Total RNA from leaf tissues was isolated using the TRIzol Reagent (Invitrogen) and
DNase (Promega) following the manufacturer’s instructions. The first-strand cDNA was synthesized with High Capacity cDNA Reverse Transcription Kit (ABI, Foster City, CA) with oligo (dT)20 primers following the manufacturer’s instructions. qRT-PCR was carried out in an
ABI 7500 Fast Real Time PCR System. The primers were designed by Primer Express software for real-time PCR (version 3.0) (ABI, Foster City, CA). The PCR conditions were as follows:
Step 1: 50°C for 2 min and 95°C for 10 min, Step 2: 95°C for 15 sec and 60°C for 1 min repeated
40 cycles, Step 3: 72°C for 10 min, Step 4: dissociation stage. The relative quantification of gene expression was analyzed by ABI 7500 Fast SDS Software (version 1.4) (ABI, Foster City, CA).
The data set was normalized using actin as a control. Gene specific forward (F) and reverse(R) primers used to generate data present in this study were: ACTIN-F (accession number: U60511.1)
5’- GGA GCT CGA GAA TGC CAA GAG CAG-3’, ACTIN-R 5’- GAC CTC AGG GCA TCT
GAA CCT CTC-3’, RIP2-F (L26305) 5’-GAG ATC CCC GAC ATG AAG GA-3’, RIP2-R 5’-
CTG CGC TGC TGC GTT TT-3’, RIP1-F(M83926) 5’-TGT GAT CCC CGA CAT GCA-3’,
RIP1-R 5’-CGA TCC TCG CTG CTT CGT-3’. The result was analyzed by SAS statistical software (SAS, Cary, NC).
68 Multidimensional protein identification technology (MudPIT) and protein identification
Proteins were extracted using a modified phenol-based extraction method described elsewhere (Chen et al., 2007) and quantified with a Bradford assay. The proteome was analyzed by Dr. Gregg Howe at Michigan State University. For mass spectrometry analysis, 100 µg of total protein were run 1 cm into a 10% denaturing polyacrylamide gel. The gel was stained with
Coomassie Brilliant Blue and destained overnight. The piece of gel containing the proteins was excised and subjected to in-gel trypsin digestion according to Shevchenko’s method with modifications (Mattiacci et al., 1995). Briefly, gel bands were dehydrated using 100% acetonitrile and incubated with 10mM dithiothreitol in 100mM ammonium bicarbonate, pH8, at 56°C for 45 min, dehydrated again and incubated in the dark with 50mM iodoacetamide in 100mM ammonium bicarbonate for 20min. Gel bands were then washed with ammonium bicarbonate and dehydrated again. Sequencing grade modified trypsin was prepared to 0.01ug/uL in 50mM ammonium bicarbonate and ~50uL of this was added to each gel band so that the gel was completely submerged. Bands were then incubated at 37C° overnight. The extracted peptides were re-suspended in a solution of 2% Acetonitrile/0.1% Trifluoroacetic Acid to 20 uL. From this
10uL were automatically injected by a Waters nanoAcquity Sample Manager (www.waters.com) and loaded for 5 minutes onto a Waters Symmetry C18 peptide trap (5um, 180um x 20mm) at
4uL/min in 2%ACN/0.1%Formic Acid. The bound peptides were then eluted using a Waters nanoAcquity UPLC (Buffer A = 99.9% Water/0.1% Formic Acid, Buffer B = 99.9%
Acetonitrile/0.1% Formic Acid) onto a Michrom MAGIC C18AQ column (3u, 200A, 100U x
150mm, www.michrom.com) and eluted over 240 min with a gradient of 5% B to 30% B in
210min at a flow rate of 1ul/min. Eluted peptides were sprayed into a ThermoFisher LTQ-FT
Ultra mass spectrometer (www.thermo.com) using a Michrom ADVANCE nanospray source.
Survey scans were taken in the FT (25000 resolution determined at m/z 400) and the top ten ions
69 in each survey scan are then subjected to automatic low energy collision induced dissociation
(CID) in the LTQ. The resulting MS/MS spectra are converted to peak lists using BioWorks
Browser v3.3.1 (ThermoFisher) using the default parameters and searched against an appropriate protein database using the Mascot searching algorithm, v 2.2 or 2.2.03 with a fragment ion mass tolerance of 0.80 Da and a parent ion tolerance of +/- 10.0 ppm allowing for up to 2 missed tryptic sites. Oxidation of cysteine (carbamidomethyl cysteine) as well as variable modification of methionine oxidation were specified in Mascot (www.matrixscience.com). The Mascot output was then analyzed using Scaffold (version Scaffold-01_07_00, Proteome Software Inc., Portland,
OR) (www.proteomesoftware.com) to probabilistically validate protein identifications using the
ProteinProphet computer algorithm (Lawrence and Novak, 2006). At the time of analysis corn had no comprehensive proteome information available therefore putative peptides were generated by six frame translation of the TIGR plant transcript assembly sequences (Kramer and
Muthukrishnan, 1998) (Database issue:D846-51. http://plantta.tigr.org/).Protein identifications were accepted if they could be established at greater than 95.0% probability and contained at least
2 identified peptides.
70 Table 4-1. Maize proteins identified in frass of S. frugiperda fed by two maize genotypes (Tx601 and Mp704).
71
) 60 B actin 50
40
30
20
10 A N.D. N.D. 0
Relative expression (target gene/ (target expression Relative RIP1 Control RIP1 Fed RIP2 Control RIP2 Fed
Figure 4-1. Expression analysis of RIP1 and RIP2 transcripts in maize leaves in response to caterpillar feeding. Maize plants were fed by S. frugiperda larvae for 24 hrs. Undamaged plants were used as the control. Total RNA was isolated from feeding site on maize leaves. The relative expression levels were determined using qRT-and gene expression was normalized to that of actin. (n=3~4, error bar indicates SE). Letters indicate significant differences by LSD test (p<0.05).
72
L F
30- RIP2
15-
pRIP2
Figure 4-2. Immunoblot analysis of RIP2 in maize leaf tissues and S. frugiperda frass. Maize plants were fed by S. frugiperda larvae for 24 hrs. Leaf tissue at the feeding site (L) was collected. The insect fed leaf tissue was used to feed S. frugiperda larvae in diet cups. Then, frass pellets of S. frugiperda (F) were collected. Equal amounts of protein (5 µg) were loaded in each well. The proteins were separating by SDS-PAGE and analyzed on immunoblots using anti-RIP antibody. The polypeptide labeled RIP2 corresponds to 30 kD proenzyme form of RIP2. The polypeptide labeled RIP2 corresponds to 15kD processed form of RIP2.
73
1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 35 - 35 -
25 -
Figure 3-10. Immunoblot analysis of RIP2 protein in several maize inbreds and teosinte. Plants were fed by S. frugiperda larvae for 24 hrs. Leaf tissues at the feeding sites (L) were
collected. Equal amounts of proteins (50µg) were loading in each well. The resulting protein was
separating by SDS-PAGE and analyzed by immunoblot using anti-RIP antibody. The polypeptide
labeled RIP2 corresponds to 30 kD proenzyme form of RIP2. Lane 1 to 15 are the following. 1:
Mp704, 2: Mp496; 3: CML131; 4: OH43; 5: Mo17; 6: Ab24E; 7: CML139; 8: Mp708; 9: KI3;
10: CML67; 11: Tx601; 12: B73; 13: W64A; 14: Zea mays parviglumis; 15: Zea mays mexicana.
74
1 2 3 4 5 55 - 40 - 35 - 25 -
15 -
10 -
Figure 4-4. Immunoblot analysis of rRIP2 digested by papain-coated beads and S. frugiperda larvae. (1) maize Tx601 leaf tissues fed by S. frugiperda larvae for 24 hr (11 µg), (2) S. frugiperda frass which larvae fed with maize Tx601 leaf tissues (38µg), (3) recombinant RIP2 (rRIP2)(0.25 µg),
(4) recombinant RIP2 (rRIP2) incubated with papain-coated beads (0.15 µg), (5) S. frugiperda frass which larvae fed with insect artificial diet containing rRIP2 (1.9 µg). The resulting protein was separating by SDS-PAGE and analyzed on an immunoblot using anti-RIP antibody. The 35
KD polypeptide corresponds to rRIP2 (lane 3). The two small polypeptides correspond to processed form of rRIP2.
75
35 A 30 A
25 B B 20
15
10
5 (mg) weight Larvae 0
BSA PBS rRIP2 prRIP2 Figure 4-5. Effect of recombinant RIP2 on S. frugiperda larvae growth. Recombinant RIP2 (rRIP2) and RIP2 processed with papain (prRIP2, see Materials and Methods) were placed into artificial diet at a ratio of 0.8µg protein/100mg diet. One day after hatching, neonate larvae were fed with protein/diet for 5 days and the final weight of larvae was determined . Letters indicate significant differences by LSD test (p<0.05) n=21-24, error bar indicates SE.
76
rRIP2 + + + + - - + Mir1-CP + + + + + + -
Incubation times 0 1 3 24 0 24 24 prRIP2
40 -
35 -
25 -
15 -
Figure 4-6. Immunoblot analysis of rRIP2 digested by Mir1-CP. Recombinant RIP2 (rRIP2) was incubated with Mir1-CP for several hr. Mir1-CP and rRIP2
mixture has equal loaded amount (3.45 µg). Mir1-CP only had loaded with 2µg purified protein.
rRIP2 only had loaded with 4.9µg purified protein. prRIP2 only had loaded with 3 µg purified
protein. The resulting protein was separating by SDS-PAGE and analyzed on an immunoblot
using anti-RIP antibody. The last lane was loaded with small polypeptides of rRIP2 processed by
papain-coated beads
77
300 B
250
200
150
100
Relative expressionRelative 50
A A 0 Control Mechanical wounding Larval feeding Figure 4-7. Analysis of RIP2 transcripts levels in maize leaves in response to various treatments. Maize plants were wounding by paper punch (mechanical wounding) or fed by S. frugiperda larvae for 24 hrs. Undamaged plants were used as a control. Total RNA was isolated from wounding site of maize leaves. The relative expression levels were determined using qRT-PCR and normalized to the expression of the reference gene, actin. (n=4, error bar indicates SE) Letters indicate significant differences by LSD test (p<0.05).
78
C W F K
Figure 4-8. Immunoblot analysis of RIP2 in maize leaves in response to various treatments. Maize plants were wounded with a paper punch (W) or fed by S. frugiperda larvae (F) for 24 hrs.
Undamaged plants were collected as a control (C). Maize kernel (K) showed RIP1 in kernel.
Equal amounts of protein (50µg) were loaded in each well. The resulting protein was separating by SDS-PAGE. The gel was hybridized with anti-RIP antibody.
79
45 ) B
40 actin 35 B 30
25
20
15 B 10 B 5 A A Relative expression (RIP2/ expression Relative 0 0 hr 0.5 hr 1 hr 2 hr 4 hr 24 hr
Figure 4-9. Time course of RIP2 mRNA expression in response to feeding by S. frugiperda larva (one larva per plant). Relative transcript levels were determined by q RT-PCR. Total RNA was isolated from feeding
sites and unwounded plants (0 hr control). Gene expression was normalized to that of actin. (n=4,
error bar indicates SE) Letters indicate significant differences by LSD test (p<0.05).
80
Hrs 0 1 2 4 6 12 24
Figure 4-10. Immunoblot analysis of RIP2 protein accumulation in maize leaves after feeding S. frugiperda larva (one caterpillar per plant) for various time points. Proteins were extracted from leaf tissues at the larval feeding sites. Uninfested plants were collected as a control (0 hr). Each time point represents four biological samples. Equal amounts of protein (50µg) were loaded in each well.
81
Days C0 0 1 2 3 4 5 6 C6
Figure 4-11. Immunoblot analysis of RIP2 protein in maize leaves after larval removal. Maize plants were first fed by S. frugiperda larva (one caterpillar per plant) for 24 hrs. The larvae were removed and this was designated as 0 day. Samples from other time points (to 6 days) were collected. Proteins extracted from leaf tissues at the feeding sites. Uninfested plants were collected as controls. The samples of control (C0 and C6) were collected at the same time on 0 and 6 day. Each time point represents two biological samples. Equal amounts of protein (50 µg) were loaded in each well.
82
Control Caterpillar fed
1’ 2’ 3’ 4’ 5’ 1 2 3 4 5
Figure 4-12. Systemic expression of RIP2 protein in maize leaves after larval feeding. Maize V8 stage plants were fed by S. frugiperda larvae for 24 hours. The larvae were placed on eleventh leaf in a small bag that limited their movement. Leaf samples from the feeding site (0 cm; lane 1) and distal to the site (3 and 5 cm; lane 2 and 3) were collected. Samples were taken from other leaves (unexpanded ninth and tenth leaf; lane 4 and 5) of the same plant. Undamaged plants were as control groups. Lane 1’ to 5’ represent the various tissue sections of undamaged plants. Each treatment represents four to five biological samples. Equal amounts of protein
(50µg) were loaded in each well.
83
Stages V2 V4 V6 V8 V10
C F C F C F C F C F
Figure 4-13. Developmental expression of RIP2 protein in maize leaves after larval feeding. Maize plants at various vegetative stages (V2, V4, V6, V8, and V10) were fed with larvae for 24 hr (larval feeding=F). Undamaged plants were as control groups (control=C). In each stage, 2 to 6 plants were harvested for control and feeding treatments. Equal amounts of protein (50 µg) were loaded in each well.
84 C W WE
Figure 4-14. Effect of exogenous ethylene on RIP2 protein expression in maize leaf after 24 hr treatment. Maize plants were damaged by a paper punch (wounded only=W). Wounded plants were spraying with 3mM 2-chloroethylphosphonic acid (ethephon) (wounded+ethephon=WE).
Undamaged plants without ethephon treatment were the control (control=C). Leaf samples were collected after 24hr of treatment. Each treatment represents two biological samples. Equal amounts of protein (50µg) were loaded in each well.
85
C W WJ WJE JE
Figure 4-15. Effect of exogenous methyl jasmonate (MEJA) and ethylene on RIP2 protein expression in maize leaf after 24 hr of treatment. Maize plants were damaged by paper punch (wounded only=W). Wounded plants were spraying with 0.01% MEJA and 3mM 2-chloroethylphosphonic acid (ethephon) (wounded+MEJA=WJ; wounded+MEJA+ethephon=WJE). Undamaged plants were also treated with 0.01% MEJA and
3mM 2-chloroethylphosphonic acid (ethephon) (MEJA+ethephon=JE). Undamaged plants without treatment were controls (control=C). Each treatment represents two biological samples.
Equal amounts of protein (50µg) were loaded in each well.
86
EC EF BF EF BF EF Hrs 1 1 1 2 2 4
Figure 4-16. Effect of exogenous EthylBloc (ethylene perception inhibitor) on RIP2 protein expression in maize leaves. Maize plants were first treated with EthylBloc (E) or buffer only (B) for 14 hr. Plants were then infested with S. frugiperda larva (one per plant) for several hours (one to four hours). Leaves from fed plants (F) or control plants (C) were collected. Each time point represents two biological samples. Equal amounts of protein (50µg) were loaded in each well.
87
MEJA 0.01% 0.03% 0.10%
C W J WJ C W J WJ C W J WJ
1 2 3 4 5 6 7 8 9 10 11 12 Figure 4-17. Effect of exogenous methyl jasmonate (MEJA) on RIP2 protein expression in maize leaves after 24 hr treatment. Maize plants were damaged by paper punch (wounded only=W). Wounded plants were sprayed
with 0.01% to 0.1% MEJA (wounded+MEJA=MJ). Undamaged plants were also treated with
0.01% to 0.1% MEJA (MEJA=J). Undamaged plants without treatment were as control groups
(control=C). Leaf samples were collected after 24 hr treatment. Each treatment represents two
biological samples. Equal amounts of protein (50µg) were loaded in each well.
88
C A W WA
Figure 4-18. Effect of exogenous abscisic acid (ABA) on RIP2 expression in maize leaf after 24 hours treatment. Maize plants were damaged by paper punch (wounded only=W). Wounded plants were sprayed with 300µM ABA (wounded+ABA=WA). Undamaged plants were also treated with 300µM
ABA (ABA=A). Undamaged plants without treatment were as control groups (control=C). Leaf samples were collected after 24 hours treatment. Each treatment represents two biological samples. Equal amounts of protein (50µg) were loaded in each well.
89 Supplemental 4-1. Maize proteins identified in frass of S. frugiperda fed by two maize genotypes (Tx601 and Mp704).
90
91
92
93
94
Chapter 5
Proteomics study on maize proteins that respond to fall armyworm (Spodoptera frugiperda) saliva
Introduction
Plants are frequently challenged by insect herbivores and have developed sophisticated defense mechanisms to counter their attack. Although chewing insects cause extensive mechanical damage to plants, this does not account for the entire effect of herbivory. Caterpillar oral secretions, including saliva and regurgitant, trigger herbivore defense responses in plants.
The effect of fall armyworm oral secretions on the maize defense response was examined in the chapter 3. In this chapter, maize proteins changed in abundance in respone to ablated or unablated fall armyworm larvae were examined by proteomic analysis.
Proteomics is a powerful tool in the post-genomic era because it generates information about integrated protein expression profiles. It provides information about protein identity, abundance, interactions, and modifications. Thus, proteomics provides a comprehensive approach to understanding the entire dynamic protein network in the cell. In addition, proteomics also provides information that transcriptome data cannot. Although evidence often shows positive correlations between transcript and protein abundance in the cell, there are also several studies to show that this is not always the case (Baginsky, 2009).
The classic proteomics approach for separating proteins from protein mixture is two- dimensional polyacrylamide gel electrophoresis (2D-PAGE). 2D-PAGE separates proteins by isoelectric point and mass. However, 2D-PAGE has the limitations of being time-consuming, labor intensive, and variable from gel to gel. Gel to gel inconsistency makes it difficult to
96 quantify protein expression levels within gels. Thus, a method named “difference in gel electrophoresis (DIGE),” in which two samples stained with different fluorescence dyes and run in the same 2D gel, was developed to decrease the gel to gel variation (Ü nlü et al., 1997).
Proteins labeled with different fluorescence dyes can then be quantified. DIGE technology decreases gel variation, enhances the sensitivity, and increases the reliability. 2D-DIGE has been used to study the abitoic and biotic stresses in maize (Casati et al., 2005; Pechanova et al., 2011).
In this study, we used 2D-DIGE technology to identify the maize proteins responded to fall armyworm saliva by comparing maize whorls infested with ablated or unablated larvae. The main goal of this study was to identify saliva-responsive proteins from susceptible and resistant maize genotypes. The results would give us a better understanding of how maize plants respond to insect elicitors.
Results:
The purpose of this study was to examine protein expression changes in maize leaves after challenging with ablated or unablated fall armyworm larvae and identify saliva-responsive proteins. The insect-susceptible genotype (Tx601) and insect-resistant genotype (Mp708) at V8 stage were infested with 5th instar ablated or unablated larvae for 24 hr (Ritchie, 1986). Leaf tissues near the wounding area were collected. Undamaged plants were used as controls. To compare the differential protein expression among three treatments (control, ablated, unablated),
2D-DIGE technology was used. The experiment design was described in the section of Materials and Methods. Protein spots expressed differently (at p-value <0.05) as determined by software analysis were subjected to MALDI MS/MS for protein identification. The identified protein spots with information regarding fold-change, protein characteristics, and biological process as
97 determined by Gene Ontology are listed in Table 5.1 (Tx601) and Table 5.2 (Mp708). Numbers of differentially expressed protein spots among the three treatments are summarized in Table 5.3.
Maize protein response to ablated or unablated larvae in the susceptible genotype Tx601
When the protein expression among the control, ablated and unablated groups were compared, sixty-six protein spots were differentially expressed in susceptible genotype Tx601
(Table 5.1). Thirty-three protein spots were up-regulated and thirty-three protein spots were down-regulated among three treatment comparisons. Each treatment comparison is discussed in detail on the following paragraphs.
Ablated group versus control group
The differentially expressed protein profile between the ablated and control groups is listed in Table 5.4. Forty-five protein spots were ablated larvae responsive proteins. Seventeen protein spots were up-regulated and twenty-eight protein spots were down-regulated. From all 17 up-regulated spots, only four spots exhibited a greater increase than 2-fold, including
Os10g0431900 (spot 1), lovastatin insensitive 1 (spot 2), chorismate synthase 2 (spot 6), and ribosome-inactivating protein (spot 5). Examples of other up-regulated proteins, which include superoxide dismutase (spot 74), proteasome subunit alpha type 2 (spot 490), and isoflavone reductase IRL (spot 110), exhibited an increase less than 2-fold. For these down-regulated proteins, two proteins were grouped as ATP synthase (spots 138 and 144) and three protein spots were RUBISCO related-proteins (spots 34, 167, and 452). Other down-regulated proteins included protochlorophyllide reductase B (spot 44), geranylgeranyl hydrogenase (spot 43), and
NADP-malic enzyme (spot 112).
98 Unablated group versus control group
The differentially expressed protein profile between the unablated and control group is listed in Table 5.5. Forty-four protein spots were unablated larvae responsive proteins. Twenty- four protein spots were up-regulated and twenty protein spots were down-regulated. From all 24 up-regulated spots, only five spots exhibited an increase greater than 2-fold, including
Os10g0431900 (spot 1), lovastatin insensitive 1 (spot 2), chorismate synthase 2 (spot 6), ribosome-inactivating protein (spot 5), and GDP-mannose 3, 5-epimerase 1 (spot 15). Examples of other up-regulated proteins include superoxide dismutase (spot 74), S-adenosylmethionine synthetase (spot 31), peroxidase 39 (spot 51), isoflavone reductase IRL (spot 110), and glutathione S-transferase (spot 692). From all down-regulated proteins, two proteins were grouped as ATP synthase (spot 138 and 144) and three protein spots were grouped as RUBISCO related-proteins (spot 22, 34, and 167). Other down-regulated proteins include protochlorophyllide reductase B (spot 44), geranylgeranyl hydrogenase (spot 43), and NADPH- protochlorophyllide oxidoreductase (spot 72).
Unblated group versus ablated group
The differentially expressed protein profile between the unablated and ablated group is listed in Table 5.6. Eight protein spots were saliva-responsive proteins. Four protein spots were up-regulated and four protein spots were down-regulated. Examples of up-regulated proteins include cytosolic glyceroldehyde-3-phosphate dehydrogenase GAPC2 (spot 78), atp1 (spot 541), and two unknown proteins (spot 273 and 149). From all four down-regulated proteins, two protein spots were grouped as ATP synthase (spot 144) and three protein spots were grouped as
99 RUBISCO related-proteins (spot 22). Other down-regulated proteins include NADPH- protochlorophyllide oxidoreductase (spot 72) and fructose-bisphosphate aldolase (spot 240).
Maize protein response to ablated or unablated larvae in the resistant genotype Mp708
When the protein expression among the control, ablated and unablated groups were compared, one hundred and eighty-two protein spots were differentially expressed in resistant genotype Mp708 (Table 5.2). Ninety-nine protein spots were up-regulated and eighty-three protein spots were down-regulated among three treatment comparisons. Eight protein spots were down-regulated in the ablated and unablated group when compared to the control, but protein expression in the unablated group decreased less than in the ablated group. One protein spot was up-regulated in the ablated but down-regulated in the unablated group when compared to the control. Each treatment comparison is discussed in detail in the following paragraphs.
Ablated group versus control group
The differentially expressed protein profile between the ablated and control group is listed in Table 5.7. Eighty protein spots were ablated larvae responsive proteins. Thirty-five protein spots were up-regulated and forty-five protein spots were down-regulated. From all 35 up- regulated spots, nine spots exhibited an increase greater than 2-fold, including lipoxygenase (spot
12, 367, and 381), RUBISCO related genes (spots 18 and 381), patatin T5 (spot 5), and ribosome- inactivating protein (spot 8). Examples of other up-regulated proteins include dehydrin (spot 45), aspartate aminotransferase (spot 138 and 278), aminomethyltransferase (spot 848), and UDP- glucose 6-dehydrogenase (spots 221 and 246). For the down-regulated proteins, three proteins were grouped as ATP synthase (spots 220, 390, and 1230) and two spots as RUBISCO related-
100 proteins (spots 238 and 711). Other down-regulated proteins include protochlorophyllide reductase (spots 3, 29, and 141), geranylgeranyl hydrogenase (spot 50), and abscisic stress- ripening protein (spot 70).
Unablated group versus control group
The differentially expressed protein profile between the unablated and control group is listed in Table 5.8. One hundred and thirteen protein spots were unablated larvae responsive proteins. Fifty-five protein spots were up-regulated and fifty-eight protein spots were down- regulated. From all 55 up-regulated spots, eight spots exhibited an increase greater than2-fold, including lipoytgenase (spot 12 and 367), RUBISCO related gene (spot 18), patatin T5 (spot 5 and 13), ribosome-inactivating protein (spot 8), O-succinylhomoserine sulfhydrylase (spot 10), and NADH dehydrogenase subunit J (spot 17). Examples of other up-regulated proteins include dehydrin (spot 45), aspartate aminotransferase (spot 138 and 278), acetolactate synthase (spot 259 and 298), 12-oxo-phytodienoic acid reductase (spot 262), and UDP-glucose 6-dehydrogenase
(spot 221 and 246). From all down-regulated proteins, six protein spots were grouped as ATP synthase (spots 60, 69, 220, 238, 273, and 1230) and seven protein spots were grouped as
RUBISCO related-proteins (spots 59, 70, 76, 256, 283, 365, and 374). Other down-regulated proteins include protochlorophyllide reductase (spot 3, 29, and 141), geranylgeranyl hydrogenase
(spot 50), s-adenosylmethionine synthetase (spot 204 and 923), and abscisic stress-ripening protein (spot 70).
101 Unablated group versus ablated group
The differentially expressed protein profile between the unablated and ablated group is listed in Table 5.9. Eighty-one protein spots were saliva-responsive proteins. Fourty-nine protein spots were up-regulated in unablated plants and thirty-two protein spots were down-regulated.
Examples of up-regulated proteins include glutathione S-transferase (spot 327, 624 and 852), acetolactate synthase (spot 259), aspartate aminotransferase (spot 1017), and ascorbate peroxidase
(spot 1047 and 1025). From all down-regulated proteins, four protein spots weregrouped as ATP synthase (spot 60, 65, 263, and 309) and six proteins spots were grouped as RUBISCO related- proteins (spot 69, 76, 365, 374, 378, and 378). Other down-regulated proteins included NADP- malic enzyme (spot 431, 521, and 654) and fructose-bisphosphate aldolase (spot 449).
Discussion:
In this study, insect-susceptible and insect-resistant maize genotypes were used to study the saliva-responsive proteins in maize. We found 81 protein spots that were saliva-responsive proteins in Mp708 genotype. However, in Tx601 genotype, only eight protein spots were identified that were expressed differently in respond to fall armyworm saliva. There could be several possibilities to explain this result. One of possible reasons could be that Mp708 has more insect-responsive genes than Tx601. Another possibility could be that the 2D-DIGE was not optimal for the Tx601 sample. Due to lack of repeat, we cannot rule of this possibility.
In this proteomics study, there are very few protein spots that were differentially expressed more than 2-fold. This was lower than the expectation. There are several reasons that could explain the lower expression levels and the numbers of protein spots that were identified.
102 1. In this study, we did not remove the Rubisco, which is highly abundant in green tissues, from the samples. Rubisco would contribute up 30% total soluble protein content in C4 leaf tissues (Sugiyama et al., 1984). Several studies showed that highly abundant proteins in 2D-
PAGE would affect the gel performance due to co-migratory and overloading effects (Corthals et al., 2000; Krishnan and Natarajan, 2009). Furthermore, low abundance proteins are hard to identify due to the presence of highly abundant proteins in the samples. Highly abundance proteins also affected the quantification process. Thus, several methods to remove or reduce portion of Rubisco have been developed and should have been employed in this study (Xi et al.,
2006; Cellar et al., 2008; Cho et al., 2008; Krishnan and Natarajan, 2009; Widjaja et al., 2009).
2. When started this study, maize sequence genome project was not yet completed. Thus, the database was built from NCBI (non redundant selection of the viridiplant taxa level).
However, proteomic analysis heavily relies on well annotated databases. Therefore, we were not able to identify several protein spots from the NCBI database. There were 31.9 % (31 of 97) of total protein spots that were not identified in Tx601 study. There were 15.7 % (34 of 216) of the total protein spots that were not identified in Mp708 study. Furthermore, in the Tx601 study, there were 48.5% (32 of 66) of the total identified spots assigned as proteins with known function in maize. In the Mp708 study, there were only 38.4% (70 of 182) of the total identified spots assigned as proteins with known function in maize.
3. In this study, samples were collected after insect infestation for 24 hr. Thus, the proteomics results showed the profile of the plant’s late defense to insect herbivory. It could have missed the information for plant early defense and signaling proteins. These proteins would be up-regulated only at very early time points when plants are under insect attack and then their expression level could be lowered to that of the uninfested phase. Furthermore, we would not get plant defensive proteins which are quickly up-regulated by insect elicitors but are slowly up- regulated by other stresses/elicitors. Ribosome inactivating proteins 2 (RIP2) is one of example.
103 The details of the RIP2 study are in Chapter 4. There was a significant difference in defense gene expression between ablated and unablted samples. However, we failed to detect large changes in the protein expression level.
Materials and Methods:
Plant materials and insect rearing
Maize (Zea mays) genotypes that are resistant (Mp708) and susceptible (Tx601) to fall armyworm feeding were obtained from Dr. W. Paul Williams (USDA-ARS, Mississippi State
University). Seeds were sown in pots filled with topsoil (Hagerstown loam) in the Crop and Soil
Sciences greenhouse at The Pennsylvania State University, University Park, PA. Well-watered maize plants were grown until they were at the V8 stage (eight fully emerged leaves) (Ritchie,
1986).
Fall armyworm (Spodoptera frugiperda) eggs were obtained from USDA-ARS Corn
Host Plant Resistance Research Laboratory at Mississippi State University. Larvae were reared on an artificial diet (Peiffer and Felton, 2005) in a 27°C incubator with a 16 h photoperiod and newly molted 5th instar larvae were used for all experiments except for bioassays with neonate larvae.
Ablation of spinneret
The late 4th instar fall armyworm larvae were placed on ice until flaccid. The larvae were immobilized with a hair clip and the spinnerets cauterized with a heat pen (Electron Microscopy
104 Sciences, Hatfield, PA). This method has been described by Peiffer and Felton (2005). The cauterized larvae were placed on artificial diet to recover after molting.
Sample collection
Maize plants were fed by ablated larvae and unablated larvae for 24 hours. Three larvae were placed in one plant. Plant tissues surrounding the wounded area near the whorl were collected after ablated or unablated caterpillar feeding. The tissue samples were stored at -80°C after immediately freezing in liquid nitrogen. Undamaged plants were as controls. Each treatment had 3 biological samples (6 plants per sample).
Protein extraction and sample preparation for 2D-DIGE
Three grams of frozen leaf samples were ground with liquid nitrogen. The protein was extracted by a phenol-based procedure with modification (Pechanova, 2006). The plant powder was put in 50ml falcon tube on ice. Two and half volume (7.5 ml) of extraction buffer (0.9 M sucrose, 0.5M Tris-base, 0.05M Na2-EDTA, 0.1M KCL, 2% β-mercaptoethanol, pH 8.7) was added and mixed by vortexing for 10 min at 4 °C. Two and half volumes (7.5ml) of Tris- saturated phenol, pH 8.0 was added and mixed by vortexing for 10 min at room temperature. The homogenate was centrifuged at 5,000g for 20 min at 4 °C. The phenol phase (the upper phase) was collected and transferred to another 15ml falcon tube. An equal volume of extraction buffer was added and mixed with phenol phase solution for 10 min at room temperature. The mixture was centrifuge at 5,000g for 20 min at 4 °C. This extraction process was repeated two times.
After the last extraction, protein was precipitated from the phenol phase with five volumes of cold methanol solution containing 0.1M ammonium acetate and 1% β-mercaptoethanol in 50ml falcon
105 tubes. Precipitation was carried out overnight at -70 °C. The precipitation was collected by centrifugation at 5,000g for 10 min at 4 °C and washed three times with five volumes of cold methanol solution. Each wash was followed by centrifugation at 5,000g for 10 min at 4 °C. After last methanol wash, the protein pellet was washed with three times with five volumes of cold
80% acetone. Each wash was followed by centrifugation at 5,000g for 5 min at 4 °C. After last acetone wash, the protein pellet was air-dried and stored at -20 °C.
Sample labeling for 2D-DIGE
For 2D-DIGE, the protein pellet was resuspended in sample/rehydration buffer (6M urea,
2M thiourea, 1% CHAPS, and 1% ASB14). Any un-dissolved material was removed by centrifugation at 14,000rpm for 5 min and supernatant was collected. The protein concentration was determined by RC-DC kit (Bio-Rad, Hercules, CA). The protein solution was mixed with
10% volume of 100mM DTT and 2% volume of pH 3-11NL IPG buffers (GE Healthcare, United
Kingdom). Three biological replicas of each treatment were used for one 2D-DIGE run. The experimental design for one six-gel 2D-DIGE experiments is described in Figure 5.1. Due to the
DIGE comparison, we combined three biological replicas together as the fourth sample. Thus, each treatment had four samples: sample 1, sample 2, sample 3, sample 4(sample 1, 2, and 3 mixture).
Protein samples were labeled with CyDye DIGE Fluor minimal dyes (GE Healthcare,
United Kingdom) as follows: 12.5µg protein sample was labeled with Cy3 or Cy5. The 12.5 µg of internal standard was labeled with Cy2. The internal standard was an equal proportion mixture from three biological replicas of each treatment. The labeling sequence in each gel was listed in
Figure 5.1. The protein sample was incubated with 100 pmol of CyDye dye in dark and at 4 °C for 30 min. Stop solution (1 µl of 1 mM lysine) was added and incubated in dark and at 4 °C for
106 10 min to stop the reaction. For each analytical gel, 12.5 µg of Cy3 labeled sample, 12.5 µg of
Cy5 labeled sample, and 12.5 µg of Cy2 labeled internal standard were pooled with sample/rehydration buffer (6M urea, 2M thiourea, 1% CHAPS, and 1% ASB14) and a sufficient amount of 100mM DTT and pH 3-11NL IPG to obtain a final volume of 450µl. For the preparative gel, 450µg of unlabeled protein from three biological replicas of each treatment (50µg of each) was pooled with 12.5 µg of Cy3 labeled sample, 12.5 µg of Cy5 labeled sample, and
12.5 µg of Cy2 labeled internal standard with sample/rehydration buffer (6M urea, 2M thiourea,
1% CHAPS, and 1% ASB14) and added sufficient amount of 100mM DTT and pH 3-11NL IPG to the final volume of 450µl.
Two-dimensional polyacrylamide gel electrophoresis (2D-PAGE)
Isoelectic focusing (IEF) was carried out in Bio-Rad PROTEIN IEF CELL on 24cm IPG strip with non-linear 3-11 pH range (Bio-Rad, Hercules, CA). The labeled protein mixture was loaded on each strip and passive rehydration was performed at 20 °C for 9 hours. Focusing was performed at 20 °C for a total of 67400 Vh. IPG strips were equilibrated with equilibrium buffer I
(6 M urea, 1.5 M pH 8.8 Tris, 50 % glycerol, 2 % SDS, 1 % DTT) with shaking at room temperature for 15 min. Then the IPG strips were equilibrated with equilibrium buffer II (6 M urea, 1.5 M pH 8.8 Tris, 50 % glycerol, 2 % SDS, 2.5 % DTT, a trace of Bromophenol Blue) with shaking at room temperature in dark for 15 min. For the second dimension, equilibrated IPG strips were place on a 12.5 % (w/v) sodium dodecyl sulfate polyacrylamide gel (SDS-PAGE) with instrument (Ettan DALTsix Electrophoresis) (GE Healthcare, United Kingdom) at 1 watt/gel.
107 Gel imaging and data analysis
Both analytical and preparative gels were scanned on a Typhoon 9400 Imager (GE
Healthcare, United Kingdom) with three different channels for three fluorescent dyes.
Excitation/emission wavelengths were 532nm/580nm for Cy3, 633nm/670nm for Cy5, and
488nm/520nm for Cy2. Images were analyzed by Progenesis Samespots (Nonlinear USA,
Durham, NC) that does the comparative analysis within gels and determines the relative quantification of proteins. Statistical analysis (one way ANOVA) included in the software is set up at p-value less than 0.05 (p < 0.05).
Protein spot picking and MS/MS analysis and Protein identification
Protein spots that were expressed differently as shown by the software analysis at p-value less 0.05 were subjected to an automatic spot picking robotic system Etta Spot Picker (GE
Healthcare, United Kingdom). Selected gel pieces were collected in 96-well plates designed for the Proteineer dp automated digester (Bruker, Bremen, Germany). Briefly: gels pieces were washed with three successive soaking in 100% ammonium hydrogenocarbonate 50mM, and a mix of 50% acetonitrile 50% ammonium hydrogenocarbonate 50mM. Two additional washes were performed with 100% acetonitrile to dehydrate the gel. Three µl of freshly activated trypsin
(Roche, porcine, proteomics grade) 10ng /µl in ammonium hydrogenocarbonate was used to rehydrate the gel pieces at 8°C for 30 minutes. Trypsin digestion was performed for 3h at 30°C.
Peptide extraction was performed with 10µl of 1% formic acid for 30 minutes at 20°C.
Protein digests (3µl) were adsorbed for 3 minutes on prespotted anchorchips (R) using the Proteineer dp automaton. Spots were washed "on-target" using 10mM dihydrogeno-
108 ammonium phosphate in 0.1% TFA-MilliQ water to remove salts. High throughput spectra acquisition was performed using an Ultraflex II MALDI mass spectrometer (Bruker) in positive reflectron mode, with close calibration enabled, Smartbeam laser focus set to medium, and a laser fluency setting of 65 to 72% of the maximum. Delayed extraction is set to 30 ns. Steps of 100 spectra in the range of 860 to 3800 Da are acquired at a 200 Hz LASER shot frequency with automated evaluation of intensity, resolution and mass range. 600 successful spectra per sample are summed, treated and de-isotoped in line with an automated SNAP algorithm using Flex
Analysis 2.4 software (Bruker), and subsequently submitted in the batch mode of the Biotools 3.0 software suite (Bruker) with an in-house hosted Mascot search engine (MatrixScience.com) to the database (NCBI non redundant selecting viridiplant taxa level). A mass tolerance of 80 ppm with close calibration and one missing cleavage site were allowed. Partial oxidation of methionine residues and complete carbamylation of cystein residues are considered. The probability score calculated by the software was used as one criterion for correct identification. Experimental and Mascot results molecular weights and pI were also compared.
109 (A)
(B)
Gel Fluorescence dye Fluorescence dye Internal standard (Cy2) labeling (Cy3) labeling (Cy5) 1 12.5 µg control 1 12.5 µg ablated 1 12.5 µg 9 sample mixture Preparative gel (1.39 µg each sample) (with 450 µg unlabeled proteins) 2 12.5 µg control 2 12.5 µg ablated 2 12.5 µg 9 sample mixture Analytical gel (1.39 µg each sample) 3 12.5 µg ablated 3 12.5 µg unablated 4 12.5 µg 9 sample mixture Analytical gel (1.39 µg each sample) 4 12.5 µg unablated 1 12.5 µg control 3 12.5 µg 9 sample mixture Analytical gel (1.39 µg each sample) 5 12.5 µg unablated 2 12.5 µg control 4 12.5 µg 9 sample mixture Analytical gel (1.39 µg each sample) 6 12.5 µg ablated 4 12.5 µg unablated 3 12.5 µg 9 sample mixture Analytical gel (1.39 µg each sample) Figure 5-1. Experimental design of a six-gel 2D-DIGE experiment for comparative study of insect-induced maize proteins. (A) The six-gel arrangement (B) table showing details of labeling samples and loading amounts
of six-gel 2D-DIGE experiment
110 Table 5-1. Differentially expressed protein spots with associated annotation and fold changes compared with control group, ablated group, unablated group in susceptible genotype Tx601.
111
112 Table 5-2. Differentially expressed protein spots with associated annotation and fold changes compared with control group, ablated group, unablated group in resistant genotype Mp708.
113
114
115
116
117
118 Table 5-3. Numerical representation maize leafy proteins responded to fall armyworm ablated or unabalted larvae. Number refers to differentially expressed proteins that were up-regulated or down-regulated among three treatments (control group, ablated group, and unablated group).
Genotype Expression level Ablated vs Control Unablated vs Control Unablated vs Ablated Up 17 24 4 Tx601 Down 28 20 4 Up 35 55 49 Mp708 Down 45 58 32
119 Table 5-4. Differentially expressed protein spots with associated annotation and fold changes compared with ablated group and control group in susceptible genotype Tx601.
120
121 Table 5-5. Differentially expressed protein spots with associated annotation and fold changes compared with unablated group and control group in susceptible genotype Tx601.
122
123 Table 5-6. Differentially expressed protein spots with associated annotation and fold changes compared with unablated group and ablated group in susceptible genotype Tx601.
124 Table 5-7. Differentially expressed protein spots with associated annotation and fold changes compared with ablated group and control group in resistant genotype Mp708.
125
126
127
128 Table 5-8. Differentially expressed protein spots with associated annotation and fold changes compared with unablated group and control group in resistant genotype Mp708.
129
130
131
132
133 Table 5-9. Differentially expressed protein spots with associated annotation and fold changes compared with unablated group and ablated group in resistant genotype Mp708.
134
135
136
Chapter 6
Conclusion and future directions
Insect feeding which results in a huge loss of the crop yield worldwide is a serious problem to agriculture. Plant scientists have developed several strategies to reduce the yield loss.
Insect-resistant transgenic crops are one of examples. However, the Bt resistant insects have been found in the field. Due to increasing demand of crop production, the insect-resistant transgenic crops would have a bigger negative impact on our ecology. Thus, to identify and exploit endogenous insect resistant genes from crops and understand how plants recognize insect feeding are two big issues to be examined. We have been focused on the effect of caterpillar saliva on maize plants and characterized of the maize defensive proteins found in caterpillar frass. The objectives of this research were:
1) To prove that fall armyworm saliva is effectively recognized by maize plants and
further triggers plant defense (Chapter 3)
2) To identify maize defensive proteins from maize-fed caterpillar frass and characterize
their function (Chapter 4)
3) To identify maize proteins that respond differently to fall armyworm saliva (Chapter 5)
In chapter 3, I have examined the effect of fall armyworm (Spodoptera frugiperda) oral secretions on the maize (Zea mays) defense response. Analyses indicated that very little regurgitant was deposited on the maize leaf during caterpillar feeding. Furthermore, caterpillar regurgitant was failed to trigger plant defense-related genes in maize. Whereas, leaf tissue immunoblots indicated that glucose oxidase, an abundant saliva protein, was deposited when caterpillars fed on the leaf. The effect of caterpillar saliva on maize defense gene expression was determined by allowing ablated (no saliva) and non-ablated (saliva present) caterpillars to feed in
138 the maize whorl. The expression of a group of maize defense genes including those from the jasmonate biosynthesis pathway and those involved in direct defenses were monitored using qRT-
PCR. The results showed that feeding by unablated caterpillars significantly increased the expression of these genes. Furthermore, a saliva-induced bioassay showed that compared to plants fed on unablated larvae, plants fed by ablated larvae did not induce sufficient direct defense to significantly retard larval growth. The results of this study show that caterpillar saliva is an important elicitor for triggering herbivore defenses in maize. We further generated the proteomics data for saliva-responsive proteins in maize (Chapter 5). Eighty-one protein spots in
Mp708 and eight protein spots in Tx601 were saliva-responsive proteins. Although very few protein spots were identified in Tx601 genotype, the identified protein spot list in Mp708 data indicated the saliva-responsive protein profile in maize.
Based on our results, the source of elicitors in the oral secretions of the fall armyworm arises from saliva and not from regurgitant. There are three aspects should be examined in the future. First, the unknown elicitor(s) in caterpillar saliva need to be identified. Furthermore, the plant receptor which can recognize insect elicitor(s) should be identified. Second, the spinneret structure of Lepidoptera larvae should be further studied. Third, early insect-induced defense profile need to be identified. The profile will help us understand the regulation mechanism of how plants respond to insect attack.
In Chapter 4, I have examined the identified plant defensive proteins from insect frass and characterized one of them that plays an important role in protecting maize against insect herbivores. With collaboration with Dr. Gregg Howe at Michigan State University, several maize proteins, including ribosome-inactivating protein 2 (RIP2), in fall armyworm frass were identified by mass spectrometry. Ribosome-inactivating proteins (RIPs) act by depurinating residues on ribosomal RNA and thereby inhibit translation. Since RIP2 was found in frass, we proposed that it might be involved in defending maize from herbivore attack. The data showed that RIP2 is
139 initially synthesized as an inactive proenzyme that can be cleaved to the active form by larval gut extracts. Also, results indicated that the expression of RIP2 was induced by FAW larvae feeding, but not mechanical wounding. The proenzyme form of RIP2 was detected in 13 maize inbred lines and two teosinte subspecies. These data indicate that RIP2 expression in response to insect feeding is a wide spread phenomenon in maize. Quantitative-RT-PCR and immunoblot analysis indicated that RIP2 is rapidly induced (1 hour) following caterpillar attack. Surprisingly, RIP2 remains at the leaf wounding sites for four days after caterpillar removal. Phytohormone testing assays determined that RIP2 expression was regulated by several different phytohormones, including ethylene, JA and ABA. It appears that there is no consistent pattern of hormonal regulation of RIP-like protein expression. The expression profile of RIP2 in the maize vegetative stage was examined. The data showed that RIP2 is expressed locally and during maize vegetative development. Furthermore, when purified recombinant RIP2 was directly tested against FAW larvae in bioassays, the data indicated that amount of RIP2 typically found in the leaf after caterpillar attack significantly retards caterpillar growth. These results indicate that RIP2 plays an important role in protecting maize against insect herbivores.
Based on our results, the frass proteome is a good approach of identifying the plant proteins that could play a role in defense against insects. We have been examined one of plant proteins identified from caterpillar frass that plays a defense function in maize. There are three aspects should be examined in the future. First, the molecular mechanism of RIP2 retardation of caterpillar growth should be examined. The fall armyworm gut enzyme that can process RIP2 to processed RIP2 needs to be identified. Recently, a chymotrypsin-like protease from lepidopteran gut which processes TD2 to processed TD2 was identified (Gonzales-Vigil et al., 2011). Second, several candidate proteins identified from caterpillar frass should be examined in the future.
Several studies have showed that beta-D-glucosidase, allene oxide synthase, lipoxygenase, and etc. are involved in plant-insect/pathogen interaction (Howe and Jander, 2008; Yu et al., 2009).
140 Third, it could be another possibility that insect frass itself can be recognized by plants and further trigger plant defense.
Plant-insect interactions are a very complicated relationship. Plants need insects as pollinators. On the other hand, plants are also the major food source for insects, including belowground and aboveground insects. Furthermore, insect herbivores can overcome plant defense responses in the coevolutionary process. Thus, it is a challenge to understand how plants recognize and further respond to insect herbivores. In this dissertation, we have examined the effect of caterpillar saliva on plant defense in maize and identified plant defensive proteins in caterpillar frass. These findings would help us have a further step to understand the plant-insect interaction.
141 References:
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VITA Wen-Po Chuang
Education 2006– 2012 Ph.D., Agronomy program, Department of Crop and Soil Sciences, The Pennsylvania State University, University Park, PA 1998 – 2002 B.S., Agronomy, Department of Agronomy, National Taiwan University, Taipei, Taiwan Professional Experiences 2006-2012 Graduate Research Assistant/Fellow, Agronomy Program, Department of Crop and Soil Sciences, Penn State University 2009 (Sep-Dec) Visting Student, Unité d'Entomologie fonctionnelle et évolutive, Université de Liège -Gembloux Agro-Bio Tech, Belgium. 2005-2006 Graduate Research Assistant/Fellow, Biochemistry and Molecular Biology, Mississippi State University 2004-2005 Graduate Research Assistant/Fellow, The Institution of Molecular Biology, National Chung Hsing University, Taiwan 2001-2002 Summer Intern, Department of Agronomy, National Taiwan University, Taiwan 2000 Summer Intern, Division of Molecular and Genomic Medicine, National Health Research Institutes, Taiwan 1999 Summer Intern, Institute of Molecular Medicine, National Taiwan University, Taiwan Honors and Awards 2012 Recipient of American Society of Plant Biologists travel grant 2011 Recipient of College of Agricultural Sciences Travel Grant Award, Penn State University 2010 Invited student speaker of Penn State University & Max Planck Institute for Chemical Ecology minisymposium, Jena, Germany 2006 Recipient of Maize genetics conference travel grant Teaching assistant Agro 028: Principles of Crop Management, Penn State University, Fall, 2007 Agro 410W: Physiology of Crop Plants, Penn State University, Spring, 2008 PLBIO 512: Plant Resource Acquisition and Utilization, Penn State University, Fall, 2011 (invited speaker)