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Theses and Dissertations--Plant and Soil Sciences Plant and Soil Sciences

2018

CHEMICAL TOPPING BURLEY TOBACCO CHEMICAL TOPPING BURLEY TOBACCO

Mitchell D. Richmond University of Kentucky, mitchell.richmond@utk.edu Author ORCID Identifier:

https://orcid.org/0000-0002-9492-1315 Digital Object Identifier: https://doi.org/10.13023/ETD.2018.079

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REVIEW, APPROVAL AND ACCEPTANCE REVIEW, APPROVAL AND ACCEPTANCE

The document mentioned above has been reviewed and accepted by the student’s advisor, on

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Mitchell D. Richmond, Student

Dr. William A. Bailey, Major Professor

Dr. Mark S. Coyne, Director of Graduate Studies

CHEMICAL TOPPING BURLEY TOBACCO

DISSERTATION

A dissertation submitted in partial fulfillment of the

requirements for the degree of Doctor of Philosophy in the

College of Agriculture, Food and Environment

at the University of Kentucky

By

Mitchell Dale Richmond

Lexington, Kentucky

Co-Directors: Dr. William A. Bailey, Extension Professor / Tobacco Specialist

Dr. Robert C. Pearce, Extension Professor / Tobacco Specialist

Lexington, KY

2018

Copyright © Mitchell Dale Richmond 2018

ABSTRACT OF DISSERATION

CHEMICAL TOPPING BURLEY TOBACCO

The act of topping tobacco (Nicotiana tabacum L.) involves the removal of the terminal

bud or inflorescence of the tobacco plant. This practice ordinarily is accomplished by

manually removing the top of each tobacco plant in an entire field which is labor

intensive and costly. Chemical topping utilizes sucker control products to inhibit the

terminal bud and axillary bud growth without manually removing the top of the tobacco

plant. There were several research objectives in order to determine the utility of a

chemical topping system: 1) determine if burley tobacco could be chemically topped with

currently registered suckercide products while maintaining control of subsequent sucker

growth; 2) compare chemical topping to manual topping for yield and leaf quality; 3)

identify burley tobacco varieties that are better suited for chemical topping systems; 4)

determine the optimum plant growth stage at which chemical topping treatments should

be applied; and 5) identify genes that are differentially expressed following suckercide

applications. To pursue our objectives, studies were initiated investigating the optimum

timing of application, ideal variety maturity, and efficacy of suckercide applications

using combinations of maleic hydrazide (MH), butralin, and fatty alcohols (FA). The

terminal bud was not well controlled with FA or butralin alone nor was acceptable sucker

control or total yield achieved. Our data suggest that chemically topping burley tobacco

with a tank mixture of MH and a local systemic may be a suitable alternative to manual

topping, as total yield and leaf quality grade index were not significantly different and

total TSNA and MH residues were not significantly higher compared to manual topping.

The 10% button and 50% button application timings were best suited for chemical

topping practices. Treatments that targeted the 10% bloom stage did not completely halt

flower development, but all application timings resulted in excellent sucker control.

Medium and late maturity burley varieties were found to be suitable for chemical topping

methods; however, timing the suckercide application may be less difficult in later

maturing varieties. Chemically topping burley tobacco at 10 to 50% button stages with a

tank mixture of MH and a local systemic suckercide was found to be a suitable

alternative to manual topping, and would potentially result in labor savings for burley

tobacco growers. Expression of genes related to phytohormones, meristem

development, cell division, DNA repair and recombination were affected following

MH treatment, which likely leads to the inhibition of apical and axillary meristem

development.

KEYWORDS: Chemical Topping, Tobacco, Maleic Hydrazide,

Mitchell Dale Richmond

April 10, 2018

CHEMICAL TOPPING BURLEY TOBACCO

By

Mitchell Dale Richmond

Co-Directors: Dr. William A. Bailey

Extension Professor / Tobacco Specialist

Dr. Robert C. Pearce

Extension Professor / Tobacco Specialist

Dr. Mark S. Coyne

Director of Graduate Studies

April 10, 2018

This work is dedicated to the memory of my mother and father.

-Mitchell Dale Richmond

April 10, 2018

iii

ACKNOWLEDGEMENTS

I am very thankful for the opportunity that my advisors, Dr. Andy Bailey and Dr.

Robert Pearce, have provided me at the University of Kentucky. It is hard to believe the

transformation that I have made under their guidance over the last several years. I would

also like to show appreciation for the members of my committee; Dr. Erin Haramoto, Dr.

Tyler Mark, and Dr. Ling Yuan, who have also helped guide me along the way. Bobby

Hill, Sitakanta Pattanaik, Chris Rodgers, Sanjay Singh, and Jack Zeleznik have been a

tremendous help for my research and for everything else that I could ask for.

Appreciation is also extended to personnel at the University of Kentucky Research and

Education Center and the Agricultural Experiment Station Spindletop Farm for assistance

throughout the last six years. I would also like to thank my fellow graduate students that

have helped in and out of the field. Last but far from least, I want to thank my family and

friends. Specifically, Jenna Richmond, who I am fortunate to call my wife and who has

supported me throughout my time as a graduate student. I am excited to start writing the

next chapter with you.

iv

TABLE OF CONTENTS

Acknowledgements ............................................................................................................ iii

List of Tables ..................................................................................................................... vi

List of Figures .................................................................................................................. viii

Chapter One: Literature Review ..........................................................................................1

Introduction .....................................................................................................................1

Topping and formation of axillary bud growth...............................................................2

Control of axillary bud growth .......................................................................................6

Chemical topping ..........................................................................................................11

Exploring the mode of action of MH ............................................................................12

Fate of maleic hydrazide in plants ................................................................................13

Conclusions and Dissertation Overview .......................................................................15

Chapter Two: The Effect of Suckercide Product and Rate on Chemical Topping of Burley

Tobacco ..............................................................................................................................17

Abstract .........................................................................................................................17

Introduction ...................................................................................................................18

Materials and Methods ..................................................................................................22

Results and Discussion .................................................................................................24

Sucker control ..........................................................................................................24

Plant height ..............................................................................................................25

Total yield ................................................................................................................26

Quality grade index ..................................................................................................28

Maleic hydrazide residues ........................................................................................28

Tobacco-specific nitrosamines.................................................................................30

Cumulative distribution function for cost saving.....................................................31

Conclusion ....................................................................................................................32

Chapter Three: The Effect of Suckercide Application Timing and Variety Maturity on

Chemical Topping of Burley Tobacco ...............................................................................42

Abstract .........................................................................................................................42

Introduction ...................................................................................................................43

Materials and Methods ..................................................................................................44

Results and Discussion .................................................................................................46

Sucker control ..........................................................................................................46

Plant height ..............................................................................................................47

Leaf dimensions .......................................................................................................49

Total yield ................................................................................................................50

Quality grade index ..................................................................................................52

Conclusion ....................................................................................................................52

v

Chapter Four: Global Transcriptomic Changes in Chemically Topped Burley Tobacco ..61

Abstract .........................................................................................................................61

Introduction ...................................................................................................................62

Materials and Methods ..................................................................................................68

Tissues and RNA sequencing ..................................................................................68

Data processing and gene expression quantification ...............................................69

Functional annotation and gene ontology (GO) analysis .........................................70

Quantitative RT-PCR ...............................................................................................70

Results and Discussion .................................................................................................71

Analysis of RNA-seq libraries of MH-treated apical and axillary buds ..................71

MH-treatment significantly alters gene expression both in apical and axillary

buds ....................................................................................................................................72

Gene ontology (GO) enrichment analysis highlights influence of MH on different

developmental and metabolic pathways in ApB and AxB ................................................72

MH perturbs expression of transcription factor genes involved in meristem

maintenance and development ...........................................................................................73

MH-treatment induces defense and secondary metabolism genes in axillary buds

............................................................................................................................................76

MH-treatment affects expression of phytohormone biosynthesis and signaling

genes in apical and axillary buds .......................................................................................77

MapMan visualization highlights the influence of MH-treatment on different plant

metabolic pathways ............................................................................................................79

Quantitative RT-PCR analysis of selected DEGs validates the RNA-seq data .......81

Conclusion ........................................................................................................................99

Literature Cited ................................................................................................................101

Curriculum Vita ...............................................................................................................124

vi

LIST OF TABLES

Table 2.1. Suckercide application rate and date for manual and chemical topping

treatments .......................................................................................................................... 34

Table 2.2. Total weight of suckers per plant prior to harvest for manual and chemical

topping treatments ..............................................................................................................35

Table 2.3. Plant height following manual topping and chemical topping treatments........36

Table 2.4. Effect of manual or chemical topping treatments on burley tobacco yield ......37

Table 2.5. Effect of manual or chemical topping treatments on quality grade index for

type 31 burley tobacco .......................................................................................................38

Table 2.6. Maleic hydrazide residues as affected by manual or chemical topping, upper

and stalk positions, and precipitation .................................................................................39

Table 2.7. Tobacco-specific nitrosamines and nicotine content from differing manual or

chemical topping treatments ..............................................................................................40

Table 3.1. Suckercide application date for manual and chemical topping application

timings................................................................................................................................54

Table 3.2. Sucker control effectiveness as percent of the control for manual and chemical

topping application timings for medium and later maturing varieties ...............................55

Table 3.3. Plant height following manual topping and chemical topping application

timings for medium and late maturing varieties ................................................................56

Table 3.4. Leaf length for tip stalk position following manual topping and chemical

topping application timings for medium and late maturing varieties ................................57

Table 3.5. Leaf width for tip stalk position following manual topping and chemical

topping application timings for medium and late maturing varieties ................................58

Table 3.6. Total yield following manual topping and chemical topping application

timings for medium and late maturing varieties ................................................................59

Table 3.7. Quality grade index following manual topping and chemical topping

application timings for medium and late maturing varieties .............................................60

Table 4.1. Summary of sequencing and read mapping in different RNA-seq libraries .....82

Table 4.2. Expression of different phytohormone biosynthesis and signaling genes in

MH-treated apical buds of tobacco ....................................................................................83

Table 4.3. Expression of different phytohormone biosynthesis and signaling genes in

MH-treated axillary buds of tobacco ........................................................................... 84-85

vii

Table 4.4. Primers used to conduct RT-qPCR analysis. ....................................................86

viii

LIST OF FIGURES

Figure 2.1. Cumulative distribution function for manual and chemical topping burley

tobacco ...............................................................................................................................41

Figure 4.1. Schematic diagram of the experimental design for chemical topping and gene

expression analysis.............................................................................................................87

Figure 4.2. Overview of RNA sequencing analysis ...........................................................88

Figure 4.3. Differentially expressed genes in maleic hydrazide (MH)-treated apical and

axillary buds .......................................................................................................................89

Figure 4.4. Gene ontology (GO) analyses of DEGs in maleic hydrazide (MH)-treated

apical and axillary buds ............................................................................................... 90-91

Figure 4.5. Effect of maleic hydrazide on expression of key transcription factors involved

in apical and axillary bud development .............................................................................92

Figure 4.6. Phylogenetic and gene expression analysis of KNOX gene family from

tobacco ...............................................................................................................................93

Figure 4.7. Differentially expressed transcription factor (TF) genes in chemically topped

axillary and apical buds .....................................................................................................94

Figure 4.8. MapMan visualization of differential gene expression in chemically topped

apical bud compared with control ......................................................................................95

Figure 4.9. MapMan visualization of differential gene expression in chemically topped

axillary bud compared with control ...................................................................................96

Figure 4.10. Validation of RNA-seq results using quantitative real-time PCR (qRT-

PCR) ...................................................................................................................................97

Figure 4.11. A model depicting the effects of maleic hydrazide on different

developmental and metabolic processes in apical (ApB) and axillary (AxB) buds of

tobacco ...............................................................................................................................98

1

Chapter One: Literature Review

Introduction

Kentucky is the leading state for production of type 31 light air-cured burley

tobacco (Nicotiana tabacum L.), accounting for over 70% of the estimated 150 million

pounds produced in the United States (USDA NASS, 2017). The estimated average yield

of burley tobacco produced in Kentucky from 2013 to 2017 was 2,180 kg ha-1 (USDA

NASS, 2017). Burley tobacco is predominately used as a component in the

manufacturing of blended cigarettes (Palmer and Pearce, 1999).

The intensive labor requirement for producing tobacco coupled with fluctuating

market prices and increasing costs of inputs leads to uncertainty in profitability. Studies

dealing with tobacco production have suggested that it takes 371-494 hours of labor to

grow one hectare of burley tobacco, even with increased labor efficiency from changes

such as float systems and baling of cured leaves (Snell and Powers, 2013; Duncan and

Wilhoit, 2014). Current challenges within the tobacco industry include delivering

increasingly regulated and potentially reduced-risk tobacco products to a decreasing

number of consumers (Snell, 2017). Maximizing yields and reducing input costs will be

vital in maintaining a profitable tobacco operation in the face of a rapidly changing

marketplace. Therefore, research to improve the efficiency of production is worth

investigating since burley tobacco is still important to Kentucky’s economy.

Removal of the terminal bud or inflorescence of the tobacco plant, commonly

known as topping, is usually accomplished by manually removing the top of each tobacco

plant in an entire field, which is labor intensive and costly. Removal of the terminal bud

or inflorescence prevents reproductive development (i.e. seed head) and results in energy

transfer to increased leaf size, weight, nicotine content, and other chemical constituents

2

(Tso, 1990). Topping results in a loss of apical dominance and the stimulation of axillary

bud growth, known as suckers (Decker and Seltmann, 1971). Controlling sucker growth

is positively correlated with yield, where greater sucker control is associated with higher

yielding tobacco (Collins and Hawks, 1993).

Topping and stimulation of axillary bud growth

Unlike most other row crops, tobacco is valued primarily for the vegetative

growth, so the terminal bud or inflorescence is removed from the plant. This operation is

known as topping and results in the loss of apical dominance. The plants start to produce

axillary buds in the leaf axil region known as suckers (Tso, 1990; Decker and Seltmann,

1971). There are other crops that also benefit from topping. It has been shown in cotton

(Gossypium hirsutum L.) that a higher number of bolls per plant were retained, and boll

growth was increased after plants were topped (Yang et al., 2012). Other experiments

have shown an increase in dry weight, plant height, number of branches, and pods and

seed yield per plant in okra (Abelmoschus esculentus) after topping and spraying

gibberellic acid (Marie et al., 2007). It is well documented that topping and control of

sucker growth are required to achieve acceptable yields and higher quality tobacco leaf

(Douglass et al., 1985; Link et al., 1982; Goins et al., 1993; McKee, 1995; Sheets et al.,

1994). In addition to the benefits on yield, quality, and increased alkaloid production, it

has been suggested that topping also reduces the potential for wind damage, increases

fertilizer use efficiency and drought tolerance, as well as provides a reduction in the

population of insects such as aphids and budworms (Bailey et al., 2017; Fisher et al.,

3

2018). Therefore, topping and control of subsequent sucker growth should be managed in

order to avoid detrimental effects on tobacco yield and leaf quality.

In a topping timing study using flue-cured tobacco, it was shown that the highest

yields in hand suckered treatments were observed when tobacco was topped in the button

or early flower stages, with delays resulting in a yield penalty of around 28 kg ha-1 day-1

(Marshall and Seltmann, 1964). Another study found no significant differences in burley

tobacco yield and value when topped at early bloom or mid-bloom stages (Seltmann et

al., 1969). However, the number of leaves left on the plant after topping has been shown

to be positively related to yield (King, 1986), but value has been shown to have a

negative relationship with number of leaves left on the plant (Collins and Hawks, 1993).

Topping burley tobacco at ten to twenty-five percent bloom with an optimum leaf

number of 22-24 leaves has been shown to provide the best yield, leaf quality, and a

better opportunity for an actual tip grade (Bailey et al., 2017).

Topping, which wounds the plant, triggers wound-activated responses in gene

expression and metabolism to activate defense mechanisms (León et al. 2001). The

phytohormone, jasmonic acid (JA), is well known as a regulator in the wound-signaling

pathway (León et al., 2001). Phytohormones, primarily auxin (IAA) and cytokinin (CK),

are known to be involved with the initiation of axillary bud growth (Müller and Leyser,

2011). Wang et al. (2018) performed comparative transcriptomic analyses to find

differentially expressed genes (DEGs) in untopped and topped tobacco plants. They

found that many of the DEGs are involved in starch and sucrose metabolism,

glycolysis/gluconeogenesis, pyruvate metabolism, and plant hormone signal transduction,

along with other processes. The previously-mentioned processes (starch and sucrose

4

metabolism) are believed to contribute significantly to the enlargement of axillary bud

growth (Wang et al. 2018). Another study found that DEGs in flue-cured tobacco roots

after topping are mostly related to secondary metabolism, hormone metabolism,

signaling/transcription, stress/defense, protein metabolism and carbon metabolism (Qi et

al., 2012).

A number of transcription factors (TFs) belonging to R2R3 MYB

(Myeloblastosis), basic helix-loop-helix (bHLH), GRAS (GAI, gibberellic acid

insensitive-RGA, repressor of GAI-Scarecrow), NAC (NAM-no apical meristem, ATAF-

Arabidopsis transcription activator factor, CUC- cup-shaped cotyledon),

homeodomain/leucine zipper (HD/ZIP), and TCP (Teosinte branched1-Cycloidea-

Proliferating cell nuclear antigen factor) families have been identified and characterized

for their roles in meristem development in Arabidopsis, tomato, pepper, and rice (Wang

et al., 2018; Janssen et al., 2014). After investigating the phenotypic and genetic

interactions of mutations in the REVOLUTA (REV) gene, it was found that REV is

required for lateral meristem and floral meristem initiation and encodes a HD/ZIP TF in

Arabidopsis (Otsuga et al., 2001). Schmitz et al. (2001) identified two genes, BLIND and

TOROSA belonging to R2R3 MYBs that control lateral meristem (axillary bud) initiation

in tomato. Müller et al. (2006) found three R2R3 MYB genes in Arabidopsis, which

were homologous to the tomato Blind gene and were designated as REGULATORS OF

AXILLARY MERISTEMS (RAX). RAX control axillary bud formation at a very early

step of initiation in Arabidopsis. BLIND ortholog was also found to reduce axillary

meristem initiation in pepper plants (Jeifetz et al., 2011). The GRAS family TF, lateral

suppressor (LAS) has also been shown to play a role in meristem development in

5

Arabidopsis and tomato (Greb et al. 2003; Schumacher et al. 1999). Double mutant

analyses in tomato and Arabidopsis revealed that LAS and MYB TFs control axillary

meristem formation through separate pathways (Schmitz et al., 2002; Müller et al., 2006).

In tomato, blind mutants did not initiate lateral meristem during shoot and inflorescence

development (Schmitz et al. 2002). The lateral suppressor (ls) mutant almost blocked all

lateral meristem development during vegetative development (Schumacher et al., 1999);

however, during reproductive development, the LS gene is not required for axillary

meristem formation (Greb et al., 2003). The TCP family TF, BRANCHED1, is known to

be involved in axillary meristem development in plants. Axillary buds in Arabidopsis

express only a single BRC1 gene compared to two BRC1-like genes in other Solanaceae

species such as tomato (Martin-Trillo et al., 2011). Martin-Trillo et al. (2011) suggested

that interplay between these two dimerizing transcription factors might result in a more

complex regulation of axillary bud growth patterns in plants like tomato, as two divergent

BRC1-like genes are co-expressed. Li et al. (2003) characterized MONOCULM 1

(MOC1), which is an important gene for rice tillering and encodes a protein highly

homologous to the tomato LAS. MOC1 is a key regulator of tillering and regulates

expression of several important genes involved with axillary bud development, namely,

OSH1. OSH1 is a rice orthologue of the maize TB1 that is expressed in axillary buds and

regulates axillary bud outgrowth (Li et al., 2003).

Each leaf axil of mature tobacco plants can potentially produce three suckers, but

it has been noted that only two suckers develop under normal commercial production

(Seltmann and Kim, 1964). Increased root growth in response to manual topping and

hand suckering has been shown to result in higher potential for the tobacco plant to

6

absorb water and nutrients as well as an increased ability to synthesize nicotine (Collins

and Hawks, 1993). In agreement with the previous statement, Woltz (1955) showed that

topping and suckering flue-cured tobacco resulted in better yield and quality and not-

topped plants resulted in substantial decreases in nicotine and sugar content. Tso (1990)

concluded that topping increased nicotine content and resulted in a net gain in total

alkaloid content. It has also been documented that tobacco that is untopped and grown in

higher plant populations produce less than 1.5% nicotine (Papenfus, 1987).

Tobacco alkaloids are an important component of leaf quality and provide

tobacco consumers a physiological stimulus that makes consumption of tobacco products

pleasurable (Bush, 1999). The major carcinogens found in tobacco are tobacco-specific

nitrosamines (TSNA), which are formed from tobacco alkaloids and are produced

primarily during curing. The amount of specific alkaloid precursor influences the amount

of TSNA accumulation and the most prevalent TSNA in burley tobacco is N-

nitrosonornicotine (NNN), which is converted from nornicotine (Jack et al, 2017).

Control of Axillary Bud Growth

In the broadest sense, three types of chemicals can be used for chemical inhibition

of axillary bud growth. These include contact, local systemic, and systemic suckercides

(Bailey et al., 2017). Tobacco growers relied on intensive labor to remove suckers by

hand prior to chemical sucker control development (Meyer et al., 1987). Therefore, it is

no surprise that chemical sucker control methods were readily adopted by tobacco

growers.

7

Contact suckercides are not absorbed, nor translocated by the plant and effective

control of suckers requires placement of the chemicals directly on the leaf axil. Local

systemic suckercides are absorbed in the leaf axil area to inhibit cell division. Systemic

suckercides, unlike contact and local systemic suckercides, do not need to directly contact

the suckers as they are absorbed by the leaves and translocated to the leaf axils, where

cell division is inhibited. Maleic hydrazide (1,2-dihydro–3,6,-pyridazinedione) is the only

true systemic suckercide that is used in tobacco production (Bailey et al. 2017).

Contact suckercides, also known as long chain fatty acids or alcohols (FA), were

developed in the early 1960s and destroy differentiating plant cells (Tso 1990; Tso and

Chu, 1977). There were many vegetable oils, saturated fatty acids, unsaturated fatty acids

and their analogues evaluated for inhibition of axillary bud development in tobacco (Tso

1990). Tso (1964) found that alkyl esters of C8 – C12 fatty acids were able to inhibit the

growth of suckers without detrimentally affecting the leaf when applied after topping

tobacco. Within the C8 – C12 fatty acids, the C10 and C11 methyl esters were the most

effective but the C11 compound showed an increased amount of phytotoxicity (Steffens et

al., 1967). These products are applied as sprays and as stalk-run down methods to contact

and kill immature actively-growing suckers, however, suckers dormant at the time of

application are not controlled so multiple applications of suckercides are required (Tso

1990). After an application of an emulsified fatty acid ester onto a plant, the ester is not

translocated and is restricted to the general area of application. Therefore, it was

suggested that growth of meristematic tissues are inhibited because of selective

penetration into rapidly dividing cells (Steffens et al., 1967). Wheeler et al. (1991)

studied the mode of action of fatty alcohols and found that the plasma membrane is

8

broken down after application, followed by dehydration of the cell, which leads to cell

death. Cathey et al. (1966) reported that lower alkyl esters of the C8 – C12 fatty acids and

the C8 – C10 fatty alcohols in aqueous emulsions were able to selectively kill the terminal

meristems without damaging axillary meristems, leaves, or stem tissues in herbaceous,

semi-woody, and woody plants. Steffens and Cathey (1969) found that terminal buds or

axillary buds of topped tobacco plants were controlled with use of emulsions containing

2:1 ratios of alcohol and surfactant or 3:1 ester to surfactant ratio. Other fatty alcohol

chain lengths, namely C9, C10, and C11, have also been reported to be highly active and

selective on inhibiting axillary and terminal bud growth of tobacco when using the proper

type and amount of surfactant as these fatty alcohols are nonselective in the absence of

surfactants (Steffens et al., 1967). Cathey et al. (1966) observed that the first visible

plant response occurs within 15 minutes after application of fatty acid esters or alcohols.

Comparable to other agricultural chemicals, there are concerns of fatty alcohol residue

levels on the treated tobacco leaves. However, studies have found that residue levels of

fatty alcohols were not detected 26 days after treatment (Tso 1990).

Local systemic suckercides need to be applied similarly to contacts so that the

solution will contact every leaf axil. Products that are local systemic belong to the

dintroanaline family and butralin and flumetralin are the major active ingredients (Bailey

et al., 2017). Singh et al. (2015) identified 179 common DEGs between tobacco plants

that were topped or topped and treated with local systemic or a contact suckercide. DEGs

related to wounding, phytohormone metabolism, and secondary metabolite biosynthesis

were upregulated after topping and downregulated after suckercide treatment. This study

also found that the application of a local systemic suckercide affected the expression of

9

auxin and cytokinin signaling pathways, which are likely involved with axillary bud

formation (Singh et al. 2015).

Maleic hydrazide (MH) was first synthesized by Curtis and Fosterburg (1895)

from maleic anhydride and hydrazine (Hoffman and Parips, 1964; Meyer et al., 1987).

However, it was not known that it had an effect on plant growth until the late 1940’s

when Schoene and Hoffman reported this effect (Schoene and Hoffman, 1949). Currier

and Crafts (1950) suggested that MH could be used as a selective herbicide. In 1951,

Peterson investigated the ability to control suckers in tobacco using maleic hydrazide and

found that MH provided excellent sucker control with no significant effects on yield,

quality, or burning properties of cured leaves (Peterson, 1952).

Maleic hydrazide has provided excellent sucker control and equivalent cured leaf

yield as opposed to hand suckering without adversely influencing leaf quality (Chaplin,

1967). Maleic hydrazide has also been shown to reduce total alkaloid levels compared to

a hand-suckered control (Cui et al. 1995). Cui et al. (1995) found that alkaloid content

was reduced by 9-34%, 4-20%, and 5-29% in the top, middle, and bottom stalk positions

after MH treatment, respectively. Treatments applied to burley tobacco that were topped

and then treated with MH provided the highest yield and value per acre (Seltmann et al.

1969). Maleic hydrazide was originally applied as a spray over the upper one-third of the

plant to cover the upper leaves (Marshall and Seltmann, 1964) and must be absorbed by

the leaves to be effective. Absorption was enhanced when MH was applied to rapidly

growing plants under conditions of high humidity (Steffens, 1983; Smith et al., 1959).

Smith et al. (1959) studied different factors including temperature, light, humidity, plant

species, plant turgidity, application rate, and formulation on absorption of MH. All

10

previously mentioned factors could impact the absorption of MH and efficacy with the

exception of light. However, relative humidity had the greatest effect as a three- to five-

fold increase in absorption was observed when relative humidity increased from 50% to

100% (Smith et al, 1959). There was only a moderate effect of temperature, possibly due

to changes in relative humidity through changes in temperature (Meyer et al., 1987). The

rate of MH uptake was decreased as the plant turgidity decreased (Smith et al., 1959),

therefore, spraying MH in the morning and evening hours of the day compared to during

the afternoon may lead to faster absorption (Smith and Stone, 1957; Meyer et al., 1987).

MH was most effective when applied on a crop growing under good moisture conditions,

more than likely as a result of less difficult penetration of the leaf cuticle during active

plant growth (Collins and Hawks, 1993).

After MH enters the plant, it is readily translocated throughout the plant

vasculature, in both phloem and xylem tissues (Hoffman and Parips, 1964; Steffens,

1983; Zukel, 1963). Similar patterns of distribution after translocation were observed

when C14 MH was applied to the top, middle, or bottom leaves (Smith et al., 1959). Most

of the chemical leaving the treated leaves went to actively growing tissues, i.e. apical and

axillary bud regions (Smith et al., 1959). Frear and Swanson (1978) also observed a

source to sink translocation pattern using foliar absorbed C14 MH. When MH was applied

to tobacco, it inhibited cell division without affecting cell elongation, thus preventing the

growth of newly developing suckers without hindering the growth of more mature leaves

(Collins and Hawks, 1993).

MH applications have been shown to be related to an increase in starch

accumulation, but these changes with respect to increased photosynthesis or decreased

11

translocation have been unclear (Bush and Sims, 1974; Seltmann and Nichols, 1984).

Callaghan and Norman (1956) concluded that a foliar spray of maleic hydrazide in the

cotyledon stage (Swiss chard) or at five to six leaves (tobacco) increased the rate of

photosynthesis. However, later studies showed that there was an increase in sucrose and

starch accumulation as a result of decreased translocation of assimilate and not due to

increased photosynthesis in burley tobacco in response to MH (Crafts-Brandner and

Sutton, 1994).

Chemical Topping

There are no suckercides that are registered or manufactured specifically for

chemical topping of tobacco, but some experiments have evaluated such products for this

purpose (Long et al., 1989; Steffens and McKee, 1969; Steffens et al. 1967; Peek, 1995).

Chemical topping utilizes sucker control products to inhibit the terminal bud and axillary

bud growth without manually removing the top of the tobacco plant. It was found that

fatty alcohols (FA) with chain lengths of C9, C10, and C11 could inhibit the terminal bud if

applied before the flowers were open and terminate suckers after the FA contacted leaf

axils (Steffens et al., 1967). Another study showed that chemically topped tobacco

yielded significantly higher when FA was applied at the button stage compared to

manually topped at the full bloom stage, but yield from chemically topped tobacco was

not significantly different than manually topped and sprayed with FA at the button stage

(Steffens and McKee, 1967). Long et al. (1989) evaluated chemically topping with MH,

flumetralin, FA, and tank mixtures and found that suppression of the terminal and

axillary buds were successful in all treatments, however, MH alone produced

12

significantly less yield due to reduced sucker control. Similarly, Peek (1995) found that a

tank mixture of MH with flumetralin resulted in the highest total yield and MH alone

resulted in the lowest yield of all chemically topped treatments. Other studies have found

chemical topping to be successful if applied at earlier button growth stages but the

earliest button stages also resulted in the largest yield reduction (Peek, 1995). Peek

(1995) found that all chemically topped treatments resulted in reduced yields when

compared to a manually topped check, except when a mammoth-type variety was

chemically topped at higher leaf numbers.

Exploring the Mode of Action of MH

The biochemical processes through which MH affects plant development are still

not completely understood even though the mode of action for MH has been studied since

1949 (Bush and Sims, 1974). It has been shown that maleic hydrazide acts as an

antimitotic agent in axillary bud tissue (Clapp and Seltmann, 1983). In the early 1970s,

there were two different views on how MH worked; those who believed that MH

interacted with nucleic acid precursors and thus ultimately with nucleic acid synthesis,

and those who did not agree (Coupland and Peel, 1971). Coupland and Peel (1971)

showed that for an increase in the concentration of MH, there was a corresponding

increase in the inhibition of uracil uptake. Their data supports the hypothesis that MH can

inhibit uptake of uracil into cells by a competitive process supporting the claim that MH

has a two-fold effect on plant tissues: 1) inhibits uracil uptake into the cell and 2) once

inside the cell, MH can become incorporated into RNA. This could be due to the close

structural resemblance MH has to uracil (Coupland and Peel, 1971; Cradwick, 1975).

13

Collins and Hawks (1993) reported that MH is absorbed by the tobacco plant and

symplastically translocated to active growing points where the mechanism of action is as

a uracil antimetabolite. Weed Science Society of America (WSSA) (2010) also stated that

maleic hydrazide may act as a uracil anti-metabolite but the mechanism of action is not

well understood. A study conducted by Appleton et al. (1981) showed that MH was

incorporated into RNA in yeast cells where it was substituted for cytosine rather than for

uracil. However, some evidence indicates that MH inhibits cell division and subsequent

sucker growth by inhibiting DNA and RNA synthesis (Nooden, 1969; Nooden, 1972;

Zukel, 1963) but does not influence actively growing cells, as they will enlarge and

differentiate (Steffens, 1983). Other theories have suggested that MH reacts with

sulfhydryl groups (Muir and Hansch, 1953) or a carbonyl reagent (Suzuki, 1966),

however Nooden (1973) showed no reactions between MH and sulfhydryl or carbonyl

compounds and discounted these theories. To summarize, there is no widely accepted

proposed mechanism of action for maleic hydrazide in the literature but it is apparent that

cell division is inhibited.

Fate of Maleic Hydrazide in Plants

Chemically, maleic hydrazide is a very stable molecule in and on plants as several

of the degradation and transfer processes for organic chemicals were not effective on MH

(Ponnapalam et al., 1983; Collins and Hawks, 1993; Nooden, 1970). MH was stable

under ultraviolet irradiation and decomposed at 260°C (WSSA, 2010), thus field and

curing conditions associated with these factors are not likely to influence residual

amounts of MH on cured tobacco leaves. In addition to UV and temperature, the vapor

14

pressure of MH is nearly zero, which leads to insignificant amounts of MH lost to

volatilization (Collins and Hawks, 1993). Therefore, there is a higher potential for MH

residues to be present in and on the surface of cured tobacco leaf since MH can become

fixed and is not believed to be highly metabolized (Collins and Hawks, 1993; WSSA

2010). To address high MH residues, a Maximum Residue Limit (MRL) of 80 µg/g was

established in Germany (Weber, 1974; Wittekindt, 1978).

MH as commonly applied is formulated as a potassium salt of MH which

possesses a high water solubility. This has a two-fold implications: higher penetration

efficiency in the plant (Coresta, 2014), and potential for control of suckers and MH

residues to be significantly influenced by rainfall and irrigation (Collins and Hawks,

1993; Seltmann and Sheets, 1987; Fisher et al., 2018).

Seltmann and Sheets (1987) found reduced sucker control with simulated rainfall

amounts of 0.2-2.0 cm within 12 hours of MH application. Leaf samples from plots

exposed to simulated rainfall 24 hours after application had significantly less MH residue

than the control (no simulated rainfall) in one year of their study. Sheets (1978), in

Collins and Hawks (1993), observed in flue-cured tobacco that mid-stalk positions of

tobacco had decreasing MH residues from harvest through four days after harvest. By

day four, after a 2.2-inch rain on day three, there was a 66% reduction in residues from

harvest. Dew may also contribute to a reduction in unbound MH on the leaf surfaces as

there was a 24% reduction in residues on day three when compared to the initial day of

harvest.

After entering the plant, it is believed that MH can exist as unmodified or free

MH, become bound with cell wall components such as lignin (Nooden, 1970), or

15

detoxified through formation of a glycoside (Towers, 1958; Coresta, 2014). The amount

of free MH has been shown to decrease after harvest, curing, and storage, which alludes

to a gradual conversion to a bound form of MH (Coresta, 2014). Nooden (1970) was able

to show that C14 MH was bound to the cell wall fragments using an energy requiring

process. Towers et al. (1958) concluded that there is formation of glycosides of MH

which could serve as a detoxifying mechanism in leaf segments of wheat. In later

studies, it was shown that MH can be metabolized with glucose to form two different

glucoside conjugates, MH-N-β-D-glucoside (Tagawa et al., 1995) or MH-O-β-D-

glucoside (Frear and Swanson, 1978). Tagawa (1995) showed that ten to thirty percent of

the total MH residue found in MH-treated cured tobacco was attributed to MH-N-β-D-

glucoside.

Conclusions and Dissertation Overview

The focus of this research involved evaluating the feasibility of eliminating

manual topping by utilizing chemical topping to top the plant without sacrificing yield,

quality, and other characteristics of tobacco. There were several research objectives in

order to determine the utility of a chemical topping system: 1) determine if burley

tobacco could be chemically topped with currently registered suckercide products while

maintaining control of subsequent sucker growth; 2) compare chemical topping to

manual topping for yield and leaf quality; 3) Identify burley tobacco varieties that are

better suited for chemical topping systems; 4) determine the optimum plant growth stage

at which chemical topping treatments should be applied; and 5) identify genes that are

differentially expressed following suckercide applications.

16

Burley tobacco production is historically and economically important for

Kentucky and much research has been dedicated to agronomic practices that promote and

maintain good agricultural practices. This review has centered on the act of topping

tobacco, formation of axillary buds, controlling axillary buds, chemical topping instead of

manual topping, and maleic hydrazide. This review demonstrates that using currently

registered suckercides to chemically top burley tobacco has the potential to reduce the

cost of labor associated with manual topping. In this dissertation, field experiments were

conducted at two locations in Kentucky for three years to evaluate different suckercide

products and application rates (Chapter 2) and the optimum application timing and

appropriate variety maturity (Chapter 3) for chemical topping of burley tobacco. In

Chapter 4, we studied changes in gene expression through use of RNA-sequencing in

MH-treated chemically topped burley tobacco and further investigate the mechanisms of

how systemic suckercides inhibit apical and axillary shoot formation. A summary of

chemical topping findings from this series of field and lab experiments is in Chapter 5.

17

Chapter Two: The Effect of Suckercide Product and Rate on Chemical Topping of

Burley Tobacco

Abstract

The act of topping tobacco (Nicotiana tabacum L.) involves the removal of the terminal

bud or inflorescence of the tobacco plant. This practice ordinarily is accomplished by

manually removing the top of each tobacco plant in an entire field which is labor intensive

and costly. The major objectives for this research were to determine which labelled

suckercides could effectively chemically top burley tobacco and the effect of suckercide

rate on sucker control, yield, leaf quality, MH residues, and leaf chemistry. To pursue our

objectives, a study was initiated at Murray, Princeton, and Lexington, KY that investigated

the efficacy of suckercide applications using combinations of maleic hydrazide (MH),

butralin, and fatty alcohols (FA). The terminal bud was not well controlled with FA or

butralin alone nor was adequate sucker control or total yield achieved. A significant

reduction in total yield and sucker control was observed when plants were chemically

topped with MH alone compared to manually topped or chemically topped with a tank

mixture of MH and butralin at Princeton only. At the other locations, all chemically topped

plants had similar yield to manually topped plants. Our data suggested that chemical

topping of burley tobacco with a tank mixture of MH and a local systemic can be an

acceptable alternative to manual topping as total yield and leaf quality grade index were

not significantly different at any location. Total tobacco-specific nitrosamine (TSNA)

content and MH residues were not significantly higher than manual topping.

18

Introduction

Kentucky is the leading state for production of type 31 light air-cured burley

tobacco (Nicotiana tabacum L.), accounting for over seventy percent of the estimated

68,000 metric tons produced in the United States (USDA NASS, 2017). The estimated

average yield of burley tobacco produced in Kentucky for 2014, 2015, and 2016 was 2,400,

2,000, and 1,950 kg ha-1, respectively (USDA NASS, 2017). Burley tobacco is

predominately used as a component in the manufacturing of blended cigarettes (Palmer

and Pearce, 1999), along with flue-cured and oriental tobacco.

The intensive labor requirement for producing tobacco coupled with fluctuating

market prices and increased costs for labor and other inputs has led to declining profit

margins for burley growers. Studies on tobacco production have indicated that it takes

150-200 hours of labor to grow one acre of burley tobacco even with advances that have

increased labor efficiency (Snell and Powers, 2013; Duncan and Wilhoit, 2014). Current

challenges within the tobacco industry involve delivering increasingly regulated, reduced-

risk tobacco products to a decreasing number of consumers (Snell, 2017). Maximizing

yields and reducing input costs will be vital in maintaining a profitable tobacco operation

in a changing marketplace. Therefore, research on improving the efficiency of production

is worth investigating since burley tobacco is significant to Kentucky’s agricultural

economy.

One area to focus research effort involves the practice of removing the terminal bud

or inflorescence of the tobacco plant. This practice, commonly known as topping, is

ordinarily accomplished by manually removing the apical meristem of each tobacco plant

in an entire field, which is labor intensive and costly (Swetnam and Walton, 1998).

19

Removal of the terminal bud or inflorescence prevents reproductive development (i.e. seed

head) and results in energy transferred to increased leaf size, weight, nicotine content, and

other chemical constituents (Tso, 1990). Subsequently, topping suppresses apical

dominance in the plant resulting in axillary bud growth, known as suckers (Decker and

Seltmann, 1971). Each leaf axil of a mature tobacco plant can potentially produce three

suckers, but it has been noted that only two suckers develop under normal commercial

production (Seltmann and Kim, 1964). Effective sucker control and yield are positively

correlated (Collins and Hawks, 1993). It is well documented that topping and control of

sucker growth is required to achieve acceptable yields and higher quality leaf (Douglass et

al., 1985; Link et al., 1982; Goins et al., 1993; McKee, 1995; Sheets et al., 1994). In the

broadest sense, there are three types of chemicals that can be used for chemical inhibition

of axillary bud growth. These three types are contact (fatty alcohols), local systemic

(butralin or flumetralin), and systemic (maleic hydrazide) suckercides (Bailey et al., 2017).

Maleic hydrazide has been shown to result in excellent sucker control and

equivalent cured leaf yield, compared to hand suckering, without adversely influencing

leaf quality (Chaplin, 1967). Chemically, MH is a very stable molecule in and on plants as

several of the degradation and transfer processes for organic chemicals are not effective

(Ponnapalam et al., 1983; Collins and Hawks, 1993; Nooden, 1970). MH is stable under

ultraviolet irradiation and decomposes at 260°C (WSSA, 2010), thus field and curing

conditions associated with these factors are not likely to influence residual amounts of MH

on cured tobacco leaves. In addition to UV and temperature, the vapor pressure of MH is

nearly zero, which leads to insignificant amounts of MH lost to volatilization (Collins and

Hawks, 1993). Therefore, there is a higher potential for MH residues to be present in and

20

on the surface of cured tobacco leaf since MH can become fixed and is not believed to be

highly metabolized (Collins and Hawks, 1993; WSSA 2010). However, MH is formulated

as a potassium salt of MH which possesses a high water solubility and has a two-fold

implication: higher penetration efficiency in plant (CORESTA, 2014), and control of

suckers and MH residues can be significantly influenced by rainfall and irrigation (Collins

and Hawks, 1993; Seltmann and Sheets, 1987; Fisher et al., 2018). Nonetheless, higher

chemical residues can be explained by the chemical properties of MH molecules and use

patterns by tobacco producers (Collins and Hawks, 1993).

Increased root growth in response to manual topping and hand suckering has been

shown to increase the potential for the tobacco plant to absorb water and nutrients as well

as an increased ability to synthesize nicotine (Collins and Hawks, 1993). Woltz (1955)

showed that topping and suckering flue-cured tobacco resulted in better yield and quality

and that untopped plants had substantially lower nicotine and sugar content. Tso (1990)

concluded that topping increases nicotine content and results in a net gain in total alkaloid

content. Cui et al. (1995) found a reduction in total alkaloid levels when MH was applied

compared to a hand suckered control. Long et al. (1989) found that chemically topped

plants had a reduced percentage of total alkaloids compared to manually topped tobacco

plants. It has been shown that applications of MH decreased lamina tobacco-specific

nitrosamine (TSNA) content due to altering the precursor-TSNA relationship (Cui et al.,

1994). TSNAs are nitrogenous compounds that are formed only from tobacco alkaloids

and are detectable in the tobacco leaf and in the particulate phase of tobacco smoke. There

are four major TSNAs: N-nitrosonornicotine (NNN), 4-(methylnitrosamino)-1-(3-

pyridyl)-1-butanone (NNK), N-nitrosoanatabine (NAT), and N-nitrosoanabasine (NAB)

21

(Brunnemann et al. 1983, Fisher et al. 1990, Hecht et al. 1998, Hoffman et al. 1994, Hecht

and Hoffman, 1988).

Suckercides that are currently registered were not intended specifically for

chemically topping tobacco, but some experiments have evaluated use of such chemicals

for this purpose (Long et al., 1989; Steffens and McKee, 1969; Steffens et al. 1967; Peek,

1995). Fatty alcohols (FA) with chain lengths of C9, C10, and C11 could inhibit the terminal

bud if applied before the flowers were open and terminate suckers after the FA contacted

leaf axils (Steffens et al., 1967). Another study showed that chemically topped tobacco

yielded significantly higher when FA was applied at the button stage compared to manually

topped at the full bloom stage but not significantly different than manually topped and

sprayed with FA at the button stage (Steffens and McKee, 1967). Long et al. (1989)

evaluated chemical topping with MH, flumetralin, FA, and tank mixtures and found that

suppression of the terminal and axillary buds were successful in all treatments, however,

MH alone produced significantly less yield due to reduced sucker control. Peek (1995)

found that a tank mixture of MH with flumetralin resulted in the highest total yield and

MH alone resulted in the lowest yield of all chemically topped treatments. Chemically

topping with a tank mixture of MH and flumetralin on photoperiod-sensitive cultivars of

flue-cured tobacco resulted no differences in yield compared to manually topped and

sprayed (Long et al., 1989). Long et al. (1989) found that split treatments of a half rate of

MH or one application of a full rate of MH sprayed without manually topping resulted in

reduced yield compared to other treatments due to poor sucker control.

The primary objective of this research was to determine if burley tobacco could be

chemically topped while simultaneously controlling axillary bud growth (suckers) using

22

currently registered rates of suckercide products without detrimentally impacting yield and

leaf quality.

Materials and Methods

Field experiments were conducted in 2016 and 2017 at the Agricultural Experiment

Station Spindletop Farm near Lexington, KY and the University of Kentucky Research and

Education Center near Princeton, KY. In 2015, this study was conducted at the Spindletop

Farm and the West Farm of Murray State University near Murray, KY. Plants of late-

maturing burley tobacco (‘KT 210’ or ‘KT 215’ depending on location) were produced in

a greenhouse float system according to current University of Kentucky recommendations

(Pearce et al., 2017). Tobacco plants were transplanted to the field in late May/early June

in all years and locations of these experiments. All field production practices, other than

topping, followed University Extension guidelines (Pearce et al., 2017). Prior to harvest,

sucker control data and plant measurements were collected from the center two rows of

each four row plot.

The experimental design was a randomized complete block with four replications.

Suckercides were applied based on product labels with a CO2-pressurized sprayer

calibrated to deliver 468 L ha-1 through an over-the-row three-nozzle row-1 configuration

using solid cone spray tips (TG3 - TG5- TG3). Treatments included maleic hydrazide

(Royal® MH-30, 0.18 kg L -1, Arysta LifeSciences), butralin (Butralin, 0.36 kg L-1, Arysta

LifeSciences), and a fatty alcohol (Off-Shoot-T, 0.31 kg octanol + 0.41 kg decanol + 0.002

kg dodecanol per L, Arysta LifeSciences). All treatments, suckercide application rates,

and dates are listed in Table 2.1. There were six chemically topped treatments including

applications of MH alone at 2.24 (Full MH) or 1.68 (Reduced MH) kg a.i. ha-1, a tank

23

mixture of MH and butralin at 2.24 + 0.56 (Full Mix) or 1.68 + 0.56 (Reduced Mix) kg ha-

1, respectively. A local systemic (butralin) or contact (FA) alone at 1.12 kg ha-1 or at 10%

v/v was also included. There was also a manually topped and not sprayed (Untreated

Control or UTC) and a manually topped and sprayed (Grower Standard or G.S.) treatment

with the full mixture of MH and butralin. Chemically topped treatments were applied at

the pre-bud (10% button) stage and manually topped treatments were imposed at the 10%

bloom stage. Button percentage was calculated by dividing the total number of plants in

the two center rows of each plot by the number of plants with a visible terminal bud

between the apical leaves, or growth stage 51 (Coresta, 2009). Bloom percentage was

calculated by dividing the total number of plants in the two center rows of each plot by the

number of plants with at least one flower open, or growth stage 60 (Coresta, 2009). All

sucker control data were collected within 7 days prior to tobacco harvest and are shown in

fresh weight of suckers (grams). All treatment application dates are provided in Table 2.1.

Thirty tobacco plants from the center two rows in each plot were stalk harvested 3

– 4 weeks after manual topping, placed on sticks, and cured in traditional air-curing barns.

After curing, tobacco leaves were removed from the stalk, sorted into four stalk positions

including flyings (lower stalk), lug (lower mid-stalk), leaf (upper mid-stalk), and tip (upper

stalk), and weighed to calculate yield per hectare. MH residue analyses on cured leaf from

lower (flyings and lug) and upper (leaf and tip) stalk positions were performed by Global

Laboratory Services, Wilson, NC. In 2016 and 2017, a United States Department of

Agriculture (USDA) grader evaluated cured leaf to USDA standards for type 31 light air-

cured burley tobacco and grades were assigned an index value between 1 and 100

(Bowman et al., 1989). Grade index data are a weighted average of grade across stalk

24

positions based on the grade received for each stalk position, and the percent contribution

of that stalk position to total yield. TSNA samples consisted of 20 leaves, collected from

the 4th leaf position from the top of 20 plants in each plot. Samples were then air-dried,

ground to 1 mm, and sent to the University of Kentucky Tobacco Analytical Laboratory

located at the Kentucky Tobacco Research and Development Center for TSNA analysis

following the method described by Morgan et al. 2004. TSNAs are presented as total

TSNA in micrograms per gram, which is the sum of all individual TSNAs (NNN, NAT,

NAB, NNK). All data were subjected to analysis of variance (ANOVA) with the general

linear model procedure (proc GLM), and means were separated using the LS-means

multiple comparison procedure at P = 0.10 using SAS 9.4 (SAS Institute Inc., Cary, N.C.).

Results and Discussion

Data for sucker control effectiveness, plant height, tobacco yield, quality grade

index, MH residue, and total TSNA are presented by year and location as there were

significant environment by treatment interactions.

Sucker Control.

There was a significant treatment effect in each site-year on sucker control. In

2015, there was a significant reduction in sucker control for treatments that did not include

MH. Butralin and FA used alone resulted in significantly less sucker control in Murray,

and Lexington (Table 2.2). Treatments that included MH (G.S., Full MH, Reduced MH,

Full Mix, and Reduced Mix) ranged from 87 to 100% control. Butralin and FA alone

treatments were discontinued for 2016 and 2017 as a result of inadequate sucker control

and the inability to chemically top the apical meristem observed in 2015.

25

Sucker growth (grams of fresh weight) ranged from 0 – 60 grams in treated plots in

2016 with the UTC significantly highest at each location. There was a significant reduction

in control when MH was used alone as compared to the G.S. and mix treatments Princeton.

In Lexington however, only the reduced MH treatment resulted in significantly less sucker

control (94%) when compared to all other treated plots. There were no significant

differences in sucker control between the G.S. and full or reduced mix treatments at either

location in 2016.

The range of sucker control effectiveness in treated plots was 99 to 100% in 2017

at Lexington. Therefore, the addition of butralin in the treatment did not improve the

control of axillary bud growth. There was a statistically significant reduction with the

reduced MH only treatment at Lexington (5 grams) but this difference is likely not

biologically relevant as most MH treated plots controlled all sucker growth. Treatments at

Princeton in 2017 followed a similar trend as 2016 with MH alone treatments resulting in

reduced sucker control. There was a benefit of using the full rate (119 grams) when

compared to the reduced rate (173) of MH, however, only the full and reduced mix

treatments provided equivalent sucker control to the G.S. (94 to 100% control).

Plant Height.

Investigating the total length of the tobacco plant to be harvested and cured was of

interest to determine if there would be limitations with the stalk harvesting and curing as a

result of the chemical topping system compared to traditional manual topping. There was

a significant treatment effect on plant height in all years and locations (Table 2.3). There

was variability in plant height across all environments and treatments ranging from 121 cm

to 251 cm. However, all plant heights above 200 cm came from plots with little to no

26

sucker control in the butralin alone, FA alone, and the UTC treatments in 2015 at each

location. Tobacco in chemically topped treatments that included MH was significantly

shorter than the G.S. at both locations in 2015, except for the Reduced Mix treatment at

Lexington. There was a total range of 12 cm in plant height at Princeton in 2016 across all

treatments. Chemically topped treatments that included MH resulted in significantly

reduced plant height when compared to the G.S. at Lexington in 2016. The total range in

plant height between the G.S. and chemically topped treatments for Princeton in 2017 was

11 cm. However, chemically topped treatments at Lexington resulted in significantly taller

tobacco when compared to the G.S. To summarize, differences in plant height between

treatments were observed. However, other than treatments without MH, these differences

did not result in difficulties in the process of harvesting, handling, and curing.

Total Yield.

There was a significant treatment effect on total yield in each year and location

combination except in 2016 at Lexington (Table 2.4). Total yield ranged from 1697 to

2252 kg ha-1 at Murray in 2015, with no significant differences in total yield between the

G.S. and chemically topped treatments that included MH. Butralin and FA alone were not

different from the UTC and resulted in a significant reduction in total yield compared to

the G.S. and chemically topped treatments that included MH. These reductions in yield

likely resulted from reduced sucker control (Table 2.2). Total yield at Lexington in 2015

ranged from 1817 to 2244 kg ha-1 (Table 2.4). Butralin and FA alone treatments produced

significantly lower total yield and MH treated plots grouped with the higher yielding

treatments. There were no significant differences between the G.S. and all chemically

topped treatments that included MH. The higher yielding treatments also had a

27

corresponding increase in sucker control effectiveness. The butralin and FA alone

treatments were discontinued after the 2015 season as these treatments were not successful

in chemically topping the plant, controlling axillary bud growth, and producing yields that

were comparable to the G.S.

Total yield ranged from 1884 to 2647 kg ha-1 at Princeton in 2016 (Table 2.4). The

G.S., Full Mix, and Reduced Mix treatments resulted in equivalent total yield at Princeton

in 2016 and 2017, however, the MH alone (Full MH and Reduced MH) treatments resulted

in significantly reduced total yield. The reduction in total yield in the Full MH and

Reduced MH treatments at Princeton were accompanied by a significant reduction in

sucker control effectiveness (Table 2.2). The addition of butralin (Full Mix and Reduced

Mix) provided significantly better sucker control and higher total yield. There were no

significant differences in total yield at Lexington in 2016 (P = 0.6447), however, there was

a narrow range of 3320 to 3397 kg ha-1 in treated plots. Therefore, chemically topped

treatments did not result in a significant decrease in yield as compared to the G.S. This

result can be attributed to a high degree of sucker control at Lexington in 2016 (Table 2.2).

There were no statistically significant differences between the G.S. and any chemically

topped treatment at Lexington in 2017, and sucker control was 99 – 100% across all treated

plots.

The Full and Reduced Mix treatments were most consistent and were not

significantly different from the G.S. in any year-location combination for total yield and

sucker control effectiveness (Tables 2 and 4). Chemical topping with MH alone (Full or

Reduced MH) did provide yields that were comparable to the G.S. and tank mix treatments

28

with butralin in four of the six environments that were tested. Butralin or FA alone was

not suitable for chemical topping as yields and sucker control were reduced.

Quality Grade Index.

There were no significant effects of chemically topped treatments (P = 0.3463) on

quality grade index at Murray in 2015 with a range of 38 to 48, as shown in Table 2.5.

Federal quality grade index data were not collected at Lexington in 2015. In 2016, there

was a significant treatment effect in Lexington (P = 0.0107) on quality grade index but this

was not observed at Princeton (Table 2.5). There were no significant differences between

the G.S. and chemically topped treatments with a range of 70 to 73 with the UTC resulting

in significantly higher quality grade index than all other treatments. There were no

significant differences at Princeton (P=0.1884) or Lexington (P=0.6712) in 2017.

Maleic hydrazide residues.

MH residue samples for G.S., Full Mix, and Reduced Mix treatments were

collected in all years and locations of this experiment. Within all years and locations, MH

residues were higher in the upper leaf positions than the lower leaf positions except

Lexington, 2015 and Princeton, 2017 (Table 2.6). There was no consistent reduction in

MH residues due to the application of a reduced rate of MH, as the Full Mix contained only

25% more product and did not always produce higher MH residues. Generally,

precipitation occurring after topping and prior to harvest provided some explanation for

differing MH residues in the different environments. The highest rainfall during this period

(10.39 cm) resulted in the lowest residues at Princeton in 2016. Higher MH residues were

observed in Murray, 2015 and Lexington, 2016 and lower amounts of rainfall after topping

and prior to harvest.

29

At Murray in 2015, the G.S. treatment had numerically higher MH residues (64

ppm) than the Full and Reduced Mix treatments (33 and 59 ppm, respectively), although

this was not statistically significant. The Reduced Mix treatment (19 ppm) had

significantly less MH residues than the G.S. (49 ppm) with the Full Mix (32 ppm) treatment

not significantly different from either at Lexington in 2015 (Table 2.6). In 2016 at

Princeton, the G.S. treatment resulted in significantly higher MH residues compared to the

Full and Reduced Mix treatments (P = 0.0233). Overall, MH residues at Princeton in 2016

were lower than all other location and year combinations likely due to heavy rainfall (10.39

cm) after topping through harvest. There were no significant differences at Lexington in

2016, however, the G.S. had numerically higher MH residues than chemically topped

treatments. Unexpectedly, the Full Mix chemically topped treatment resulted in

significantly lower MH residues compared to the Reduced Mix and G.S. at Princeton in

2017. Although not significant, the G.S. had numerically less MH residues compared to

chemically topped treatments at Lexington in 2017. This is likely due to a rainfall event

that occurred within three to six hours after application. The decision was made not to

reapply this treatment as it may have influenced results; however, sucker control and yield

were not negatively affected (Tables 2 and 4). Theoretically, chemical topping may result

in lower MH residues due to the timing of application as chemical topping applications are

typically made about seven days prior to when growers would normally apply MH

following manual topping. Assuming both are harvested at the same time, the increased

time between application and harvest would allow more time for precipitation and

degradation to reduce MH residue levels. There was no clear evidence of a reduction in

30

MH residues with chemical topping in this study, but MH residues were not higher

compared to the current grower standard for sucker control.

Tobacco-Specific Nitrosamines.

There were no significant differences in TSNA between manually or chemically

topped treatments in 2016 (p=0.8444) or 2017 (p=0.2046) at Princeton as shown in Table

2.7. However, there was a significant treatment effect on total TSNA at Lexington in 2016

(p=0.0019) and 2017 (p=0.0702). The UTC had significantly higher TSNAs than all other

treatments in 2016. Chemically topped treatments (Full Mix and Reduced Mix) resulted

in significantly lower TSNAs than the UTC and the G.S. There was a topping effect and

MH application effect on TSNA. Topping without spraying MH (UTC) resulted in

significantly higher total TSNA compared to manually topping and spraying MH (G.S.).

A similar significant trend was observed at Lexington in 2017, however, there were no

significant differences between the G.S. and UTC. Cui et. al. (1994) suggested that

applying MH reduced TSNA in air-cured burley tobacco due to MH altering the precursor-

TSNA relationship. Tso (1990) concluded that topping increases nicotine content and

results in a net gain in total alkaloid content; therefore, tobacco plants that are not manually

topped should be expected to have less alkaloids and therefore less precursor to TSNA

formation. This is likely due to a combination of increased root growth leading to an

increase in nicotine biosynthesis and upregulated plant defenses due to wound signaling

pathways. This may help explain reduced total TSNA in chemically topped treatments

(Full Mix and Reduced Mix) at Lexington in 2016, as nicotine content (Table 2.7) was

significantly less in chemically topped treatments (p<.0001). Chemically topped

treatments resulted in lower nicotine content at Princeton in 2016 and in both years at

31

Lexington (Table 2.7). Another possible explanation is the timing of MH application

between the G.S. and the chemically topped treatments as the chemically topped MH

treatments were applied seven days prior to the G.S. (Table 2.1), thus altering the timing

of the precursor relationship. Significant reductions in TSNA were only observed in

Lexington, however, numerical trends were observed in Princeton.

Cumulative Distribution Function for Cost Savings

Chemical topping burley tobacco was found to be a suitable alternative to the

traditional manual topping as sucker control, total yield, and leaf quality grade index was

not significantly different in all environments tested. A stochastic simulation model was

developed to evaluate the potential savings from the use of chemical versus manual

topping. The stochastic variables in the model are the number of man-hours required for

manual topping (Min=3, Mean=5.5, Max=10), amount of time required to spray (Min=0.4,

Mean=0.5, Max=0.6), hourly wage (Min=8, Mean=10, Max 12.5), yield (kg ha-1)

(Min=1905, Mean=2242, Max=3138) and the average price per kilogram (Min=2.71,

Mean=3.95, Max=4.41). Minimal research has been conducted on hours to manually top

versus spraying a hectare of tobacco and its impact on yield and quality, which impacts

price. A GRKS distribution was utilized based on parameters using the preceding

minimum, mean, and maximum values in variables. The GRKS1 distribution is an

augmented triangle distribution and is used in situations when minimal information is

available (Richardson et al., 2006). The critical difference between manual and chemical

topping is labor cost savings potential (Figure 2.1). The foundation for the simulation is

1 The Gray, Richardson, Klose, and Schumann (GRKS) distribution is similar to a

triangular distribution and was developed to simulate random variables when insufficient

historical data is available (Richardson, 2016).

32

the 2016 burley tobacco budget used to calculate the cost reduction of chemical topping as

a function of the reduced labor requirement for topping (Snell et al., 2016). Cost saving ($

ha-1) is based on the return over variable cost as opposed to return over total costs. Based

on the assumptions of this simulation, an average of $134.45 ha-1 was saved when chemical

topping was used if topping required only five man-hours in a manual topping system. The

range of cost saving is $28.81 to $288.49 ha-1 based on 500 iterations, with an iteration

representing a possible outcome given the assumptions, under the assumptions with the

simulation. Another simulation was performed assuming that manual topping required ten

man-hours in a manual topping system. An estimated average of $259.23 ha-1 was saved

when chemical topping was used to replace the labor associated with topping. The range

of cost saving would be $142.21 to $438.11 ha-1 if ten man-hours were required to manually

top based off 500 iterations under the assumptions with the simulation.

Conclusion

Chemical topping of burley tobacco at 10% button stage with a tank mixture of MH

and a local systemic suckercide was a suitable alternative to manual topping as sucker

control, total yield, and leaf quality grade index were not different in all years and locations

of this study. Application of a local systemic or fatty alcohol alone did not inhibit the

terminal bud nor control subsequent sucker growth resulting in a reduction in total yield.

MH residues for chemically topped tobacco were not consistently higher than residues

from manually topped and sprayed tobacco. Total TSNA was not increased due to

chemically topped treatments, and at Lexington there was a reduction in total TSNA

compared to manually topping. Future work should further investigate these total TSNA

reductions that were observed. Chemical topping has the potential to reduce labor input

33

and production costs without negatively impacting the yield, quality or chemistry of burley

tobacco.

34

Table 2.1. Suckercide application rate and date for manual and chemical topping treatments.

Treatment Suckercides Application Rate Manually

Topped Treatment Applied

kg a.i. ha-1 Yes/No

Spindletop Princeton

2015 2016 2017 2015* 2016 2017

UTCa - - Yes 7/27 8/9 7/28 8/28 8/14 7/31

G.S. MH + B 2.24 + 0.56 Yes 7/27 8/9 7/28 8/28 8/14 7/31

Full MH MH 2.24 No 7/20 8/2 7/20 8/20 8/8 7/26

Reduced MH MH 1.68 No 7/20 8/2 7/20 8/20 8/8 7/26

Full Mix MH + B 2.24 + 0.56 No 7/20 8/2 7/20 8/20 8/8 7/26

Reduced Mix MH + B 1.68 + 0.56 No 7/20 8/2 7/20 8/20 8/8 7/26

Butralin B 1.12 No 7/20 - - 8/20 - -

FA C8-C10-C12 10% v/v No 7/20 - - 8/20 - - a UTC = Untreated control ; G.S. = Grower Standard ; MH = maleic hydrazide; FA = Fatty Alcohol; B = butralin.

* 2015 location was at Murray, KY

35

Table 2.2. Total weight of suckers per plant prior to harvest for manual and chemical topping treatmentsa.

2015c 2016 2017

Murray Lexington Princeton Lexington Princeton Lexington

Treatmentb ----------------------------------------------- g -----------------------------------------------

UTC 269 d 454 d 324 c 336 c 385 d 676 c

G.S. 2 a 2 a 0 a 0 a 0 a 0 a

Full MH 5 a 30 a 50 b 5 a 119 b 0 a

Reduced MH 25 ab 43 a 60 b 19 b 173 c 5 b

Full Mix 0 a 26 a 12 a 0 a 2 a 0 a

Reduced Mix 1 a 57 a 6 a 1 a 22 a 0 a

Butralin 96 bc 338 c - - - -

FA 121 c 257 b - - - -

p-value <.0001 <.0001 <.0001 <.0001 <.0001 <.0001 a Total weight of suckers per plant calculated from a sample of 10 plants per plot. bUTC = Untreated control ; G.S. = Grower Standard ; MH = maleic hydrazide; FA = Fatty Alcohol; B =

butralin. c Means within a column followed by the same letter are not significantly different according to Fisher’s

Protected LSD at P = 0.10.

36

Table 2.3. Plant height following manual topping and chemical topping treatments.

2015b 2016 2017

Murray Lexington Princeton Lexington Princeton Lexington

Treatmenta -------------------------------------------- cm --------------------------------------------

UTC 187 cd 209 b 184 c 195 a 196 a 161 a

G.S. 190 c 169 c 178 d 185 b 142 b 121 d

Full MH 169 e 151 e 185 bc 168 c 143 b 148 bc

Reduced MH 179 d 159 d 189 ab 171 c 142 b 149 b

Full Mix 167 e 161 d 190 a 168 c 132 c 150 b

Reduced Mix 171 e 168 c 189 abc 166 c 135 c 143 c

Butralin 242 b 219 a - - - -

FA 251 a 218 a - - - -

p-value <.0001 <.0001 <.0001 <.0001 <.0001 <.0001 a UTC = Untreated control ; G.S. = Grower Standard ; MH = maleic hydrazide; FA = Fatty Alcohol; B =

butralin.

b Means within a column followed by the same letter are not significantly different according to Fisher’s

Protected LSD at P = 0.10.

37

Table 2.4. Effect of manual or chemical topping treatments on burley tobacco yield.

2015b 2016 2017

Murray Lexington Princeton Lexington Princeton Lexington

Treatmenta -------------------------------------------- kg ha-1 --------------------------------------------

UTC 1720 b 1872 ab 1884 b 3098 2122 b 2005 b

G.S. 2121 a 2155 ab 2627 a 3371 2725 a 2598 a

Full MH 2166 a 2229 ab 2074 b 3338 2297 b 2779 a

Reduced MH 2233 a 2141 ab 2127 b 3397 2112 b 2705 a

Full Mix 2252 a 2157 ab 2647 a 3356 2690 a 2828 a

Reduced Mix 2148 a 2244 a 2611 a 3320 2614 a 2576 a

Butralin 1697 b 1817 b - - - -

FA 1719 b 1817 b - - - -

p-value 0.0026 0.0371 0.0030 0.6447 <.0001 <.0001 a UTC = Untreated control ; G.S. = Grower Standard ; MH = maleic hydrazide; FA = Fatty Alcohol; B =

butralin. b Means within a column followed by the same letter are not significantly different according to Fisher’s

Protected LSD at P = 0.10.

38

Table 2.5. Effect of manual or chemical topping treatments on quality grade index for type 31 burley tobaccoa.

2015c 2016 2017

Murray Lexington Princeton Lexington Princeton Lexington

Treatmentb -------------------------------------- Quality Grade Index --------------------------------------

UTC 42 - 60 77 a 64 62

G.S. 38 - 61 71 b 57 65

Full MH 43 - 61 70 b 67 62

Reduced MH 47 - 62 71 b 62 58

Full Mix 39 - 61 73 b 63 66

Reduced Mix 48 - 62 73 b 68 70

Butralin 45 - - - - -

FA 41 - - - - -

p-value 0.3463 - 0.1306 0.0107 0.1884 0.6712 aQuality grade index is a numerical representation of Federal quality grade index received for tobacco and is a

weighted average of grade index for all stalk positions following Bowman et al. 1989. bUTC = Untreated control ; G.S. = Grower Standard ; MH = maleic hydrazide; FA = Fatty Alcohol; B =

butralin. cMeans within a column followed by the same letter are not significantly different according to Fisher’s

Protected LSD at P = 0.10.

39

Table 2.6. Maleic hydrazide residues as affected by manual or chemical topping, upper and lower stalk positions, and precipitation.

2015a 2016 2017

Murray Lexington Princeton Lexington Princeton Lexington

Treatmentb ------------------------------------------------------- ppm ------------------------------------------------------

G.S. 64 49 a 15 a 62 41 a 29

Full Mix 33 32 ab 10 b 54 10 b 50

Reduced Mix 59 19 b 11 b 51 36 a 44

p-value 0.1886 0.0944 0.0233 0.7038 0.0231 0.1168

Position

Upper 78 A 38 13 A 85 A 35 53 A

Lower 26 B 26 10 B 27 B 24 29 B

p-value 0.0011 0.1692 <.0001 <.0001 0.2279 0.0078

Precipitationc -------------------------------------------------------- cm --------------------------------------------------------

2.92 6.17 10.39 4.47 3.12 7.49 a Means within a column followed by the same uppercase or lowercase letter are not significantly different according to Fisher’s

Protected LSD at P = 0.10. b Full MH and Reduced MH were excluded from residue analysis to make better comparisons to the G.S. c Total rainfall from topping through harvest.

40

Table 2.7. Tobacco-specific nitrosamines and nicotine content from differing

manual or chemical topping treatments.

2016 2017

Princeton Lexington Princeton Lexington

Total TSNAa

Treatmentb ----------------------- µg g-1 ------------------------

UTC 4.72 3.14 a 1.38 1.34 ab

G.S. 5.04 2.22 b 1.24 1.60 a

Full Mix 4.23 1.01 c 0.87 0.59 b

Reduced Mix 4.63 1.15 c 0.85 0.57 b

p-value 0.8444 0.0019 0.2046 0.0702

Nicotine

-------------------------- % --------------------------

UTC 3.05 c 6.01 a 4.76 6.03 a

G.S. 4.65 a 5.34 a 4.18 5.55 a

Full Mix 4.02 b 2.23 c 4.43 3.58 b

Reduced Mix 3.56 b 3.00 b 4.41 3.29 b

p-value 0.0002 <.0001 0.8773 <.0001 a Means within a column and variable followed by the same letter are not

significantly different according to Fisher’s Protected LSD at P = 0.10. b Full MH and Reduced MH were excluded from leaf chemistry analyses to make

better comparisons to the G.S.

41

Cumulative Distribution Function for Manual and Chemical Topping Burley Tobacco

Dollars saved ($ ha-1

)

0 100 200 300 400 500

Pro

ba

bili

ty

0.0

0.2

0.4

0.6

0.8

1.0

5 Hours to Manually Top

10 Hours to Manually Top

Figure 2.1. Cumulative Distribution Function for Manual and Chemical Topping Burley Tobacco. The lines represent the cost

saved per hectare with the assumption that 5 man-hours (blue) or 10 man-hours (red) was required for manual topping.

42

Chapter 3: The Effect of Suckercide Application Timing and Variety Maturity on

Chemical Topping of Burley Tobacco

Abstract

Experiments were initiated in 2015 to evaluate the efficacy of chemical topping for burley

tobacco. The major objectives for this study were to determine the optimum timing of

suckercide application and appropriate variety maturity for effective chemical topping.

Burley tobacco varieties TN90 (medium maturity), KT210 and KT215 (late maturity) were

chemically topped at the 10% button, 50% button, and 10% bloom growth stages. The

10% button and 50% button application timings were best suited for chemical topping

practices. Treatments that targeted the 10% bloom stage did not completely halt

inflorescence growth, however all application timings resulted in excellent sucker control.

Both medium and late-maturing burley varieties proved to be acceptable for chemical

topping methods, however, timing the suckercide application may be less difficult with

later maturing varieties. Chemically topped treatments generally resulted in shorter,

narrower tip leaves than manually topped treatments. There were no significant differences

in total yield when comparing tobacco that was manually topped at 10% bloom to

chemically topped at 10% button, 50% button, or 10% bloom across all environments in

TN90. In four out of six environments, total yield was not significantly different between

manual topping and any chemically topped application timing in the late-maturing burley

varieties; however, at least one chemically topped application timing was equivalent to

manually topped tobacco in all environments.

43

Introduction

Topping, the removal of the terminal bud or inflorescence of the tobacco (Nicotiana

tabacum L.) plant, is ordinarily accomplished by manually removing the top of each

tobacco plant in an entire field, which is labor intensive and costly. Removal of the

terminal bud or inflorescence prevents reproductive development (i.e. seed head) and

results in energy transferred to increased leaf size, weight, nicotine, and other chemical

constituents (Tso, 1990). Topping eliminates apical dominance in the plant resulting in

axillary bud growth, known as suckers (Decker and Seltmann, 1971). It has been shown

that controlling sucker growth and yield are positively correlated (Collins and Hawks,

1993).

Topping burley tobacco at ten to twenty-five percent bloom with an optimum leaf

number of 22-24 leaves has been shown to provide the best yield, leaf quality, and a better

opportunity for a true tip grade (Bailey et al., 2017). Higher yields were observed when

flue-cured tobacco was topped in the button or early flower stages in hand-suckered

treatments with a yield penalty of around 28 kg-1 day-1 when topping was delayed beyond

this point (Marshall and Seltmann, 1964). Other studies have found no significant

differences in burley tobacco yield and value when topped at early bloom or mid-bloom

stages (Seltmann et al., 1969). However, the number of leaves left on the plant after

topping was shown to be positively related to yield (King, 1986), but value has been shown

to have a negative relationship with number of leaves left on the plant (Collins and Hawks,

1993). A chemical topping study applying a tank mixture of maleic hydrazide (MH) and

flumetralin when the 20th leaf expanded to 15 cm on photoperiod-sensitive cultivars of

flue-cured tobacco found no differences in yield compared to manually topped and sprayed

44

tobacco (Long et al., 1989). Peek (1995) found that chemical topping at the 25% button

stage was the most effective timing of application but resulted in the largest yield reduction.

The associated yield reduction was attributed to reduced leaf size, specifically in the upper

stalk positions (Peek, 1995). Long et al. (1989) found that chemically topped plants

generally resulted in taller plants with shorter, narrower top leaves. The primary objectives

of this research was to determine the optimum stage of apical bud growth to target that

could chemically top the plant while simultaneously controlling axillary bud growth

(suckers) using currently registered suckercide products in medium and late maturing

burley tobacco varieties.

Materials and Methods

Field experiments were conducted in 2016 and 2017 at the Agricultural Experiment

Station Spindletop Farm near Lexington, KY and the University of Kentucky Research and

Education Center near Princeton, KY. In 2015, this study was conducted at the Spindletop

Farm and the West Farm of Murray State University near Murray, KY. Transplants of

burley tobacco varieties ‘KT 210 and KT 215’ (late maturity) and ‘TN90’ (medium

maturity) were grown in a greenhouse float system according to current University of

Kentucky recommendations (Pearce et al. 2017). Tobacco plants were transplanted to the

field in late May/early June in all years and locations of these experiments. All field

production practices, other than topping, followed recommendations based on the

University Extension guidelines (Pearce et al., 2017).

The experimental design was a randomized complete block with treatments

replicated four times. Suckercides were applied with a CO2-pressurized sprayer calibrated

to 468 L ha-1 with a directed three-nozzle row-1 configuration (TG3-TG5-TG3). Maleic

45

hydrazide (Royal MH-30®, 0.18 kg a.i. L-1, Arysta LifeScience) tank mixed with butralin

(Butralin®, 0.36 kg a.i. L-1, Arysta LifeScience) was used as the suckercide application.

Chemical topping treatments were applied at either the 10% button, 50% button, or 10%

bloom stages. There was also a manually topped and not sprayed (Untreated Control or

UTC) and a manually topped and sprayed (Grower Standard G.S.) imposed at the 10%

bloom stage (Figure 3.1). Button percentage was calculated by dividing the total number

of plants in the two center rows of each plot by the number of plants with a visible terminal

bud between the apical leaves, or growth stage 51 (Coresta Guide #7, 2009). Bloom

percentage was with the total number of plants in the two center rows of each plot with at

least one flower open, or growth stage 60 (Coresta Guide #7, 2009). All sucker control

data were collected within 7 days before tobacco harvest and are shown in percent control

of fresh weight of suckers compared to fresh weight of sucker in the manually topped

untreated control that did not receive suckercide treatment.

Thirty tobacco plants from the center two rows in each plot were stalk harvested

three-four weeks after manual topping, placed on sticks, and cured in traditional air-curing

barns. Prior to harvest, sucker control data and plant height measurements were collected

from the center two rows of each four-row plot. After curing, tobacco leaves were removed

from the stalk, sorted into 4 stalk positions including flyings (lower stalk), lug (lower mid-

stalk), leaf (upper mid-stalk), and tip (upper stalk), and weighed to calculate yield per

hectare. A sample of 25 leaves from the tip grade of each plot was measured to determine

leaf length and leaf width. A United States Department of Agriculture (USDA) grader

evaluated cured leaf to USDA standards for type 31 light air-cured burley tobacco and

grades were assigned an index value between 1 and 100. Grade index data are a weighted

46

average of grade across stalk positions based on the grade received for each stalk position,

and the percent contribution of that stalk position to total yield (Bowman et al., 1989). All

data were subjected to analysis of variance (ANOVA) with the general linear model

procedure (proc GLM), and means were separated using the least-square means multiple

comparison procedure at P = 0.10 using SAS 9.4 (SAS Institute Inc., Cary, N.C.).

Results and Discussion

Data for sucker control effectiveness, plant height, tip leaf stalk position length,

tobacco yield, and quality grade index are presented by year, location, and variety maturity

as there were significant environment by treatment interactions.

Sucker Control.

There was a significant application timing effect on percent sucker control in each

environment as shown in Table 3.2. Overall, sucker control ranged from 89 to 100%

control in treated plots across all environments. In 2015 at Murray, there was a significant

reduction in sucker control when suckercides were applied at the 10% bloom stage in the

late maturity group compared to all other timings, however, this difference was only one

percent. There were no significant differences between application timings in the medium

maturity TN 90 at Murray. In 2015 at Lexington, there was a significant reduction in sucker

control when applications were made at the 10% bloom stage in TN 90 but this difference

was only five percent in comparison with the G.S. There was a significant three percent

reduction in sucker control in the 10% button timing as compared to the G.S. in the late

maturing KT 210.

The range of sucker control effectiveness across application timings in treated plots

for medium and late maturing varieties was 91 to 100 % and 89 to 100%, respectively in

47

2016 at Princeton. There were no significant differences in the medium-maturing TN 90

between the G.S. and any application timing at Lexington in 2016 as excellent sucker

control was observed in all treated plots. There was a significant four percent reduction in

sucker control when late-maturing varieties were chemically topped at the 10% bloom

application timing. In 2017, there was a significant two percent reduction in sucker control

in the 50% button application timing when compared to the G.S. at Princeton in the

medium-maturing TN 90. There were no significant differences between the G.S. and any

chemical topping application timing in the late-maturing varieties. In 2017 at Lexington,

the 10% and 50% button chemical topping application timings resulted in significantly

higher sucker control than the G.S. and the 10% bloom stage in the medium maturing TN

90. The 50% button application timing resulted in a significant nine percent reduction in

sucker control compared to all other treated plots.

In summary, excellent sucker control was achieved in all chemical topping

application timings. Peek (1995) observed reduced sucker control when suckercides were

applied at later maturity stages. Chemical topping at 10% bloom resulted in around 10%

flower spikes present at harvest as the blooms were not manually removed. Therefore, we

concluded that the 10% or 50% button application timings were better suited for chemical

topping of burley tobacco.

Plant Height.

Investigating the height of the tobacco plants to be harvested and cured was of

interest to determine if there would be harvest difficulties encountered when using

chemical topping compared to traditional manual topping. Plant height was measured

while plants were still in the field and was determined by measuring from the ground to

48

the uppermost plant part. There was a significant timing effect on plant height in all years

and locations (Figure 3.3). There was variability in plant height across all environments,

maturity, and application timings ranging from 123 to 231 cm. The UTC had significantly

higher plant height compared to all other application timings for the medium maturity TN

90, which was due to no sucker control applied after topping. Within the medium maturity

TN 90, the 10% bloom application timing resulted in significantly lower plant height than

the UTC but significantly higher plant height compared to the G.S. and 10% or 50% button

timings within each environment. There was a 1 to 13 cm difference in plant heights across

all environments when comparing the chemical topping application timings at 10% button

and 50% button to the G.S. within the medium maturity TN 90. Therefore, 10% and 50%

button application timings appeared to be more suitable target timings for chemical topping

when comparing plant height for the medium maturity variety used in these experiments.

Unlike in the medium maturity variety, the UTC did not always result in

significantly higher plant height in all years and locations for the late maturing variety.

Either the UTC or 10% bloom application timing had significantly higher plant height

compared to all other application timings and the G.S within each environment (Figure 3.3)

for the late maturing varieties used in these experiments. In three of the six environments

within the late-maturing varieties (Murray, 2015; Lexington, 2016; and Princeton, 2017),

there were no significant differences between 10% button application timing and the G.S.

The 10% button application timing resulted in significantly lower plant height at

Lexington, 2015 and Princeton, 2016 but significantly higher plant height in Lexington,

2017 compared to the G.S. Within the late-maturing varieties, the 50% button application

timing resulted in significantly higher plant height compared to the G.S. and 10% button

49

in all environments with the exception of 2016 at Lexington where there were no

significant differences between either timing. There were no undesirable plant heights that

caused problems in harvesting or curing except no sucker control within the UTC and the

existence of blooms within the 10% bloom application timing. The UTC was trimmed

immediately prior to harvest to meet the size requirements of the curing facility; however,

the plots were not suckered. Therefore, we concluded that the 10% or 50% button

application timings should be targeted.

Leaf Dimensions.

Leaf dimension data were collected from a 25-leaf sample of cured-leaf from the

tip stalk position. There was a significant application timing effect on leaf length in each

environment except Princeton in 2017 as shown in Figure 3.4. The range of tip leaf length

for medium maturity was 32 – 54 cm across all environments and treatments. The G.S.

resulted in significantly longer tip leaves than any chemically topped application timing

within the medium maturity variety in four of five environments where tip leaf length was

measured. In the late-maturing varieties, either the UTC or the G.S. resulted in

significantly longer tip leaves when comparing all treatments within each environment.

Chemical topping in the late maturity variety resulted in significantly shorter tip leaf length

at Lexington in all years of this study. However, only the 10% button application timing

resulted in significantly shorter tip leaves at Princeton in 2016 with the 50% button and

10% bloom timings not significantly different than the G.S. Significant differences in 2017

at Princeton were likely not biologically relevant as the total range in tip leaf length was

only two cm when comparing all treatments excluding the UTC. The total difference

50

between the 10% button, 50% button, and 10% bloom application timings within each

environment and maturity ranged from one to three cm.

There was a significant application timing effect on tip leaf width within each

environment (Figure 3.5). The range of tip leaf width for medium maturity was 13 – 24

cm across all environments and treatments. Within the medium maturity TN 90, the G.S.

had significantly wider tip leaves compared to all chemically topped application timings

except 10% bloom at Lexington in 2016. The range in tip leaf width for the late-maturing

varieties was 15 – 28 cm across all environments and treatments. Within the late-maturing

varieties, the G.S. had significantly wider leaves than all chemically topped application

timings with the exception of 50% button at Princeton in 2016. The 10% button application

timing was grouped with the significantly narrowest leaf in all environments for each

maturity, except for late maturing varieties at Lexington in 2016.

Generally, chemically topped plants resulted in shorter, narrower leaves in the tip

stalk position compared to treatments that were manually topped, which is comparable to

other previous results (Long et al. 1989; Peek, 1995). It would be expected that tip leaf

length in chemically topped burley tobacco would be equal to or less than manually topped.

Thus, the marketable cured tip leaf stalk position would be expected to have a higher

likelihood to meet the leaf length requirement for tip grade in chemically topped burley

tobacco.

Total Yield.

There was a significant application timing effect on total yield in each year and

location combination except in the late maturing KT 215 at Murray in 2015 (Figure 3.6).

As expected, the UTC resulted in the lowest total yield within each environment, maturity,

51

and application timing as there was no sucker control applied to these plots. There were

no significant differences between the G.S. and chemically topped application timings in

the medium maturity TN 90 at either location in any year. Within chemically topped

treatments, the 10% bloom application timing resulted in significantly higher total yield

compared to chemically topped at 10% button timing at Murray in 2015 (p=0.0040).

Chemically topped at the 50% button application timing resulted in significantly higher

total yield compared to the 10% bloom timing at Lexington in 2015 within the medium

maturity (p=0.0026). Each location in 2016 and 2017 for the medium maturity TN 90

followed the same trend with the G.S. not significantly different than any chemically

topped application timing, which is similar to sucker control effectiveness data.

Within the late maturing varieties, there were no significant differences between

the G.S., 10% button, 50% button, and 10% bloom at Lexington in 2015 and 2017 or

Princeton in 2016. The G.S. resulted in significantly higher total yield than the 50% button

and 10% bloom application timings at Lexington in 2016 but was not different than the

10% button (p=0.0206). The 10% button application timing at Princeton in 2017 had

significantly lower total yield compared to the G.S. and 10% bloom application timing

(p=0.0059). Sucker control effectiveness data does not exclusively explain differences in

total yield for the late maturing varieties, as sucker control across all treated plots ranged

from 89 – 100%, especially considering the excellent sucker control with all treatments at

Princeton in 2017 (Figure 3.2). It should be noted that later maturing/flowering varieties

might be better suited for adopting chemical topping methods, as the transition between

reproductive growth stages is slower than in earlier maturing varieties. To summarize,

there were no significant differences in total yield when comparing the G.S. to tobacco that

52

was chemically topped at 10% button, 50% button, and 10% bloom across all environments

in the medium maturity TN 90. With the exception of two environments, total yield was

not significantly different between the G.S. and any chemically topped application timing

in the later maturing varieties; however, at least one chemical topping timing was

equivalent to the G.S. in all environments.

Quality Grade Index.

There was no significant effect of treatment across all environments and maturities

on quality grade index (Figure 3.7). Quality grade index data were not collected at

Lexington in 2015. There was a difference of 11 grade index points between all treatments

within the medium maturity TN 90 at Murray; however, there was only two grade index

points difference between the G.S. and all chemically topped application timings for

quality grade index. Within TN 90, there was a difference of 3 and 13 grade index points

across all treatments in 2016 and 3 and 9 grade index points in 2017 at Princeton and

Lexington, respectively. There was a difference of 12 grade index points across all

treatments within the late maturing KT 215 at Murray. Within the late maturing varieties,

there was a difference of 5 and 6 grade index points across all treatments in 2016 and 2 and

10 grade index points in 2017 at Princeton and Lexington, respectively. Therefore, our

data suggested that no application timing detrimentally influences quality grade index as

there were no significant differences across manually or chemically topped treatments.

Conclusion

Chemical topping burley tobacco at 10% button (pre-bud) to 50% button (early-

bud) is ideal as application of suckercides at 10% bloom did not completely halt the

development of reproductive growth. Most chemically topped application timings

53

included in these experiments provided similar sucker control, total yield, and leaf quality

compared to manually topping. Chemically topped treatments also appeared to have shorter

tip leaves which may contribute to an increased amount of marketable tip grades compared

to manually topping. Although there were no outstanding differences in yield and quality

between the medium and late maturing varieties used in this experiment, later maturing

varieties tended to yield higher and may be better suited for chemical topping due to less

rapid change from vegetative to reproductive growth, which would result in a wider

window for making chemical topping applications at the most appropriate timings.

54

Table 3.1. Suckercide application date for manual topping and chemical topping application timings.

Maturity Treatment Timing Manually Topped Treatment Applied

Yes/No

Spindletop Princeton

2015 2016 2017 2015b 2016 2017

Medium

UTCa 10% Bloom Yes 7/24 7/29 7/29 8/20 8/8

G.S. 10% Bloom Yes 7/24 7/29 7/29 8/20 8/8

10% Button 10% Button No 7/20 7/26 7/20 8/2

50% Button 50% Button No 7/20 7/26 7/25 8/5

10% Bloom 10% Bloom No 7/27 7/29 7/27 8/20 8/8

Late

UTC 10% Bloom Yes 7/27 8/9 7/28 8/28 8/14 7/31

G.S. 10% Bloom Yes 7/27 8/9 7/29 8/28 8/14 7/31

10% Button 10% Button No 7/20 8/1 7/25 8/20 8/8 7/26

50% Button 50% Button No 7/24 8/1 7/27 8/24 8/11 7/28

10% Bloom 10% Bloom No 7/27 8/9 7/29 8/28 8/14 7/31 aUTC = Untreated control; G.S. = Grower Standard b2015 location was at Murray, KY.

55

Table 3.2. Sucker control effectiveness as percent of the control for manual and chemical topping application timings for

medium and late maturing varieties.

2015a 2016 2017

Murray Lexington Princeton Lexington Princeton Lexington

Maturity Timing ------------------------------------------------%------------------------------------------------

Medium

UTCb 0 B 0 C 0 B 0 B 0 C 0 C

G.S. 100 A 100 A 97 A 100 A 100 A 96 B

10% Button 100 A 97 AB 100 A 100 A 99 AB 100 A

50% Button 100 A 97 AB 91 A 100 A 98 B 100 A

10% Bloom 99 A 95 B 96 A 100 A 99 AB 95 B

p-value <.0001 <.0001 <.0001 <.0001 <.0001 <.0001

Late

UTC 0 c 0 c 0 b 0 c 0 b 0 c

G.S. 100 a 100 a 97 a 100 a 100 a 100 a

10% Button 100 a 97 b 100 a 98 ab 100 a 100 a

50% Button 100 a 100 a 89 a 98 ab 100 a 91 b

10% Bloom 99 b 100 a 94 a 96 b 100 a 100 a

p-value <.0001 <.0001 <.0001 <.0001 <.0001 <.0001 aMeans within a column followed by the same uppercase or lowercase letter are not significantly different

according to Fisher’s Protected LSD at P = 0.10. bUTC = Untreated control; G.S. = Grower Standard.

56

Table 3.3. Plant height following manual topping and chemical topping application timings for medium and late maturing

varieties.

2015a 2016 2017

Murray Lexington Princeton Lexington Princeton Lexington

Maturity Timing ------------------------------------------------cm------------------------------------------------

Medium

UTCb 231 A 199 A 216 A 187 A 169 A 164 A

G.S. 150 D 160 C 166 C 144 E 135 C 134 D

10% Button 137 E 164 C 153 D 149 D 134 C 123 E

50% Button 162 C 164 C 166 C 157 C 137 C 144 C

10% Bloom 185 B 176 B 188 B 166 B 154 B 159 B

p-value <.0001 <.0001 <.0001 <.0001 <.0001 <.0001

Late

UTC 198 c 191 a 177 c 192 b 181 a 165 a

G.S. 179 d 161 c 187 b 177 c 140 d 123 d

10% Button 178 d 144 d 167 d 176 c 139 d 134 c

50% Button 207 b 176 b 201 a 175 c 149 c 154 b

10% Bloom 220 a 180 b 206 a 202 a 156 b 139 c

p-value <.0001 <.0001 <.0001 <.0001 <.0001 <.0001 aMeans within a column followed by the same uppercase or lowercase letter are not significantly different

according to Fisher’s Protected LSD at P = 0.10. bUTC = Untreated control; G.S. = Grower Standard.

57

Table 3.4. Leaf length for tip stalk position following manual topping and chemical topping application timings for medium

and late maturing varieties.

2015a 2016 2017

Murrayc Lexington Princeton Lexington Princeton Lexington

Maturity Timing ------------------------------------------------cm------------------------------------------------

Medium

UTCb - 52 A 42 A 48 A 34 37 B

G.S. - 54 A 41 A 43 B 34 41 A

10% Button - 44 C 39 B 39 C 35 34 C

50% Button - 47 B 40 B 39 C 32 33 C

10% Bloom - 45 BC 39 B 40 C 32 33 C

p-value - <.0001 <.0001 <.0001 0.6979 <.0001

Late

UTC - 56 a 51 a 48 b 40 a 40 b

G.S. - 51 b 48 b 50 a 37 b 45 a

10% Button - 45 c 44 c 44 c 36 bc 36 c

50% Button - 44 c 47 b 43 d 35 c 36 cd

10% Bloom - 43 c 47 b 41 d 35 c 35 d

p-value - <.0001 <.0001 <.0001 <.0001 <.0001 aMeans within a column followed by the same uppercase or lowercase letter are not significantly different

according to Fisher’s Protected LSD at P = 0.10. bUTC = Untreated control; G.S. = Grower Standard. cLeaf length data not collected at Murray in 2015.

58

Table 3.5. Leaf width for tip stalk position following manual topping and chemical topping application timings for medium

and late maturing varieties.

2015a 2016 2017

Murray Lexington Princeton Lexington Princeton Lexington

Maturity Timing ------------------------------------------------cm-------------------------------------------------

Medium

UTCb - 22.7 A 22.4 A 23.0 A 14.8 B 15.0 B

G.S. - 23.5 A 21.3 B 18.9 B 17.1 A 16.7 A

10% Button - 17.4 C 18.2 D 15.7 C 12.7 C 13.5 C

50% Button - 19.1 B 19.4 C 16.1 C 12.9 BC 13.6 C

10% Bloom - 20.0 B 19.8 C 17.8 BC 14.7 B 13.9 C

p-value - <.0001 <.0001 <.0001 <.0001 <.0001

Late

UTC - 24.2 a 28.2 a 22.5 b 18.7 a 19.8 a

G.S. - 22.9 a 26.7 b 25.8 a 17.4 b 20.1 a

10% Button - 18.1 b 22.8 d 18.3 d 15.4 c 15.2 b

50% Button - 18.9 b 26.2 b 17.3 e 15.7 c 15.3 b

10% Bloom - 18.7 b 25.1 c 19.3 c 16.3 c 15.1 b

p-value - <.0001 <.0001 <.0001 <.0001 0.0016 aMeans within a column followed by the same uppercase or lowercase letter are not significantly different

according to Fisher’s Protected LSD at P = 0.10. bUTC = Untreated control; G.S. = Grower Standard. cLeaf width data not collected at Murray in 2015.

59

Table 3.6. Total yield following manual topping and chemical topping application timings for medium and late maturing

varieties.

2015a 2016 2017

Murray Lexington Princeton Lexington Princeton Lexington

Maturity Timing ----------------------------------------------kg ha-1 ----------------------------------------------

Medium

UTCb 1340 C 1803 B 1788 B 2291 B 1965 B 2155 B

G.S. 2068 AB 2094 AB 2692 A 2751 A 2614 A 2456 A

10% Button 1810 B 2122 AB 2566 A 2799 A 2475 A 2513 A

50% Button 2149 AB 2326 A 2580 A 2796 A 2516 A 2579 A

10% Bloom 2246 A 1896 B 2555 A 2680 A 2676 A 2494 A

p-value 0.0040 0.0026 0.0019 0.0003 0.0417 0.0246

Late

UTC 1688 2033 b 1737 b 2552 c 2223 c 2413 c

G.S. 2154 2190 ab 2318 a 3439 a 2878 a 2924 ab

10% Button 2447 2263 ab 2492 a 3057 ab 2516 b 3090 a

50% Button 2244 2561 a 2145 a 2849 bc 2683 ab 2728 bc

10% Bloom 2378 2235 ab 2187 a 2976 b 2826 a 2720 bc

p-value 0.4036 0.0032 0.0182 0.0206 0.0059 0.0070 aMeans within a column followed by the same uppercase or lowercase letter are not significantly different

according to Fisher’s Protected LSD at P = 0.10. bUTC = Untreated control; G.S. = Grower Standard.

60

Table 3.7. Quality grade index following manual topping and chemical topping application timings for medium and late

maturing varieties.

2015a 2016 2017

Murray Lexington Princeton Lexington Princeton Lexington

Maturity Timing ---------------------------------------------- 0-100 ----------------------------------------------

Medium

UTCb 40 - 65 66 64 75

G.S. 51 - 65 73 64 74

10% Button 49 - 68 67 67 68

50% Button 49 - 65 57 67 70

10% Bloom 49 - 67 60 66 66

p-value 0.0543 - 0.7255 0.4727 0.6089 0.7615

Late

UTC 41 - 60 74 63 60

G.S. 41 - 58 73 64 69

10% Button 49 - 60 68 64 66

50% Button 53 - 60 68 63 69

10% Bloom 47 - 63 69 62 70

p-value 0.1317 - 0.1124 0.7864 0.9968 0.5473 aMeans within a column followed by the same uppercase or lowercase letter are not significantly different

according to Fisher’s Protected LSD at P = 0.10. bUTC = Untreated control; G.S. = Grower Standard.

61

Chapter 4: Global Transcriptomic Changes in Chemically Topped Burley Tobacco

Abstract

Information on the influence of suckercides on tobacco gene expression and their

molecular mechanism of action is limited. Therefore, the primary objectives of this

experiment were to study global changes in gene expression in apical (ApB) and axillary

buds (AxB) of “chemically topped” (untopped plants treated with MH) burley tobacco

using RNA-sequencing (RNA-seq) and to propose a possible molecular mechanism of

action of MH on sucker control. Sequencing of RNA libraries from ApB and AxB of

control and MH-treated tobacco generated a total of 450 million (M) clean reads and

more than 75% of the total reads were mapped to reference tobacco genome. Analysis

of the RNA-seq libraries revealed that compared with the control, chemical topping (CT)

significantly altered gene expression in ApB and AxB; 573 (132 upregulated, 441

downregulated) and 2,632 (2,174 upregulated, 458 down-regulated) genes were found to

be differentially expressed in chemically topped ApB and AxB, respectively. Gene

ontology (GO) enrichment analysis was performed on differentially expressed genes

(DEGs) both in ApB and AxB. In MH-treated ApB, upregulated genes were enriched

for phosphorelay signal transduction, leaf proximal/distal pattern formation and

regulation of timing of transition from vegetative to reproductive phase whereas GO

terms related to meristem maintenance, cytokinin metabolism, cell wall synthesis,

photosynthesis and DNA metabolism were enriched in downregulated genes. In MH-

treated AxB, GO terms related to defense response and oxylipin metabolism were enriched

in upregulated genes whereas GO terms related to cell cycle and DNA metabolism,

cytokinin metabolism were enriched in downregulated genes. Genes encoding proteins

62

essential for cell division control and DNA replication were downregulated. Expression

of a number of transcription factor genes known to play crucial roles in apical and axillary

meristem development were downregulated in ApB and AxB after MH treatment. MH

negatively affects the expression of a number of MADS-box family TFs in ApB known to

determine floral organ identity in plants. In addition, MH-treatment induces defense and

secondary metabolism genes in axillary buds. TFs belonging WRKY, AP2/ERF and NAC

families were mostly affected in MH-treated AxB. Furthermore, genes related to

biosynthesis and signaling of a number of phytohormones including CK, JA, ethylene (ET),

abscisic acid (ABA), and gibberellic acid (GA), were affected by MH-treatment in ApB

and AxB of tobacco. In summary, MH profoundly influenced gene expression in ApB

and AxB of tobacco. The number of DEGs were higher in AxB compared to ApB. In

both ApB and AxB the expression of genes related to phytohormones, meristem

development, cell division, DNA repair and recombination were affected following MH

treatment, which likely leads to the inhibition of apical and axillary shoot growth.

Collectively, RNA-seq analysis provides insights into the possible molecular

mechanism of action of MH on apical and axillary buds of tobacco.

Introduction

Removal of the terminal bud or inflorescence of the tobacco (Nicotiana tabacum

L.) plants, commonly known as topping, is usually accomplished by manually removing

the top of each tobacco plant in an entire field, which is labor intensive and costly. There

are other crops that also benefit from topping. It has been shown in cotton (Gossypium

spp.) that a higher number of plants retain cotton bolls and show increased boll growth

after topping (Yang et al., 2012). Topping of okra (Abelmoschus esculentus) has been

63

shown to result in an increase in seed yield per plant (Marie et al., 2007). Removal of the

terminal bud or inflorescence in tobacco prevents reproductive development (i.e. seed

head) and results in energy transferred to increased leaf size, weight, nicotine, and other

chemical constituents (Tso, 1990). Topping also eliminates apical dominance in the plant

resulting in axillary bud growth, known as suckers (Decker and Seltmann, 1971). Topping,

which wounds the plant, triggers wound-responsive gene expression and metabolism to

activate defense mechanisms (León et al. 2001). The phytohormone, jasmonic acid (JA)

and its methyl esters, methyl jasmonate (MeJA), are well known elicitors of the wound-

signaling pathway in plants (León et al., 2001). Wang et al. (2018) performed comparative

transcriptomic analyses to find differentially expressed genes (DEGs) in untopped and

topped tobacco plants. They found that many of the DEGs are involved in starch and

sucrose metabolism, glycolysis/ gluconeogenesis, pyruvate metabolism, and plant

hormone signal transduction, along with other processes. The starch and sucrose

metabolism processes are believed to contribute significantly to the enlargement of axillary

bud growth (Wang et al. 2018). Another study found that DEGs in flue-cured tobacco

roots after topping are mostly related to secondary metabolism, hormone metabolism,

signaling/transcription, stress/defense, protein metabolism and carbon metabolism (Qi et

al., 2012).

Phytohormones, primarily auxin (IAA) and cytokinin (CK), are known to be

involved with the initiation of axillary bud growth in plants (Müller and Leyser, 2011). A

number of transcription factors (TFs) belonging to R2R3 MYB, basic helix-loop-helix

(bHLH), GRAS (GAI, gibberellic acid insensitive-RGA, repressor of GAI-Scarecrow),

NAC (NAM, no apical meristem-ATAF, Arabidopsis transcription activator factor-CUC,

64

cup-shaped cotyledon), homeodomain/leucine zipper (HD/ZIP), and TCP(Teosinte

branched1-Cycloidea-Proliferating cell nuclear antigen factor) families have been

identified and characterized for their roles in meristem development in Arabidopsis,

tomato, pepper, and rice. After investigating the phenotypic and genetic interactions of

mutations in the REVOLUTA (REV) gene, it was found that REV is required for lateral

meristem and floral meristem initiation and encodes a HD/ZIP TF in Arabidopsis (Otsuga

et al., 2001). Schmitz et al. (2001) identified two genes, BLIND and TOROSA belonging to

R2R3 MYBs that control lateral meristem (axillary bud) initiation in tomato. Müller et al.

(2006) found three R2R3 MYB genes in Arabidopsis, which were homologous to the

tomato Blind gene and were designated as REGULATORS OF AXILLARY MERISTEMS

(RAX). RAX control axillary bud formation at a very early step of initiation in Arabidopsis.

A BLIND ortholog was also found to reduce axillary meristem initiation in pepper plants

(Jeifetz et al., 2011). The GRAS family TF, lateral suppressor (LAS) has also been shown

to play role meristem development in Arabidopsis and tomato (Greb et al. 2003;

Schumacher et al. 1999). Double mutant analyses in tomato and Arabidopsis revealed that

LAS and MYB TFs control axillary meristem formation through separate pathways

(Schmitz et al., 2002; Müller et al., 2006). In tomato, blind mutants did not initiate lateral

meristems during shoot and inflorescence development (Schmitz et al. 2002). The lateral

suppressor (ls) mutant blocked almost all lateral meristem development during vegetative

development (Schumacher et al. 1999); however, during reproductive development the LS

gene is not required for axillary meristem formation (Greb et al., 2003). The TCP family

TF, BRANCHED1, is known to be involved in axillary meristem development in plants.

Axillary buds in Arabidopsis express only a single BRC1 gene compared to two BRC1-like

65

genes in other Solanaceae species such as tomato (Martin-Trillo et al., 2011). Martin-Trillo

et al. (2011) suggested that interplay between these two dimerizing transcription factors

might result in a more complex regulation of axillary bud growth patterns in plants like

tomato, as two divergent BRC1-like genes are co-expressed. Li et al. (2003) characterized

MONOCULM 1 (MOC1), which is an important gene for rice tillering and encodes a

protein highly homologous to the tomato LAS. MOC1 is a key regulator of tillering and

regulates expression of several important genes involved with axillary bud development,

namely, OSH1. OSH1 is a rice orthologue of the maize TB1 that is expressed in axillary

buds and regulates axillary bud outgrowth (Li et al., 2003).

Sucker growth control and yield are positively correlated (Collins and Hawks,

1993). Studies dealing with tobacco production have indicated that it takes 150-200 hours

of labor to grow one acre of burley tobacco even with advances that have come with

increased labor efficiency (Snell and Powers, 2013; Duncan and Wilhoit, 2014).

Therefore, non-traditional methods of topping are of interest to eliminate the need for

manual topping to reduce the labor requirement. There are three major types of chemicals

(suckercides) that are typically used for chemical inhibition of axillary bud growth. These

three types consist of contact, local systemic, and systemic suckercides (Bailey et al.,

2017). Contact suckercides are not absorbed, nor translocated by the plant and effective

control of suckers requires placement of the chemicals directly on the leaf axil (Bailey et

al., 2017). Local systemic suckercides are absorbed in the leaf axil area to inhibit cell

division (Bailey et al., 2017). Singh et al. (2015) identified 179 common DEGs between

tobacco plants that were topped, and treated after topping with a local systemic or contact

suckercide. DEGs related to wounding, phytohormone metabolism, and secondary

66

metabolite biosynthesis were upregulated after topping and downregulated after suckercide

treatment. This study also found that the application of a local systemic suckercide affected

the expression of auxin and cytokinin signaling pathways, which are likely involved with

axillary bud formation.

Systemic suckercides, unlike contact and local systemic suckercides, do not need

to be in direct contact with the suckers as they are absorbed by the leaves and translocated

to the leaf axils, where they inhibit cell division (Bailey et al., 2017). Maleic hydrazide

(MH, 1,2-dihydro – 3,6,-pyridazinedione) is the only true systemic suckercide that is used

in tobacco production (Bailey et al. 2017). MH is readily translocated throughout the plant

vasculature, in both phloem and xylem tissues (Hoffman and Parips, 1964; Steffens, 1983;

Zukel, 1963) and inhibits cell division without affecting cell elongation, thus preventing

the growth of newly developing suckers without hindering the growth of more mature

leaves (Collins and Hawks, 1993). The molecular mechanism through which MH affects

bud growth is still not completely understood even though the mode of action for MH has

been studied since 1949 (Bush and Sims, 1974). It has been shown that MH acts as an

antimitotic agent in axillary bud tissue (Clapp and Seltmann, 1983). In the early 1970’s,

there were two different views on how MH works; those who believed that MH interacts

with nucleic acid precursors and thus ultimately with nucleic acid synthesis, and those who

did not agree. Coupland and Peel (1971) showed that for an increase in the concentration

of MH, there is a corresponding increase in the inhibition of uracil uptake. Their data

supports the hypothesis that MH can inhibit uptake of uracil into cells by a competitive

process eluding to the claim that MH has a two-fold effect on plant tissues: 1) inhibits

uracil uptake into the cell and 2) once inside the cell, MH can become incorporated into

67

RNA. This could be due to the close structural resemblance of MH with uracil (Coupland

and Peel, 1971; Cradwick, 1975). Collins and Hawks (1993) reported that MH is absorbed

by the tobacco plant and symplastically translocated to active growing points where the

mechanism of action is a uracil antimetabolite. A study conducted by Appleton et al.

(1981) showed that MH was incorporated into RNA in yeast cells where it was substituted

for cytosine rather than for uracil. However, some evidence indicates that MH inhibits cell

division and subsequent sucker growth by inhibiting DNA and RNA synthesis (Nooden,

1969; Nooden, 1972; Zukel, 1963) but does not affect actively growing cells, as they

enlarge and differentiate (Steffens, 1983). Other theories have suggested that MH reacts

with sulfhydryl groups (Muir and Hansch, 1953) or a carbonyl reagent (Suzuki, 1966);

however, Nooden (1973) showed no reactions between MH and sulfhydryl or carbonyl

compounds and discounted these theories. To summarize, there is no widely accepted

mechanism of action for MH in the literature since the mid-20th century but it is apparent

that cell division is inhibited after MH application. Technology, such as RNA-sequencing,

can be used to address this knowledge gap.

There are no suckercides that are registered or intended specifically for chemical

topping of tobacco but some experiments have evaluated products for this purpose (Long

et al., 1989; Steffens and McKee, 1969; Steffens et al. 1967; Peek, 1995). Field

experiments conducted at the University of Kentucky found that chemically topping burley

tobacco can be achieved with use of MH. Manually topping followed by an application of

suckercides are common agronomic practices in the production of burley tobacco. A few

studies have investigated DEGs prior to and after topping, as well as after suckercide

treatment in tobacco (Wang et al., 2018, Singh et al. 2015). However, the impact of

68

chemical topping on gene expression has not been investigated. The primary objective of

this experiment was to study global changes in gene expression through use of RNA-

sequencing in MH-treated chemically topped burley tobacco and to expand the knowledge

on how systemic suckercides can inhibit apical and axillary shoot formation.

Materials and Methods

Tissues and RNA Sequencing

Plants of a late-maturing burley tobacco variety were produced in 2015 at the

Agricultural Experiment Station Spindletop Farm near Lexington, KY. Tobacco plants

were transplanted to the field in late May/early June and all field production practices, other

than topping, were standard based on the University Extension guidelines (Pearce et al.,

2017). The experimental design was a randomized complete block with treatments

replicated four times. Maleic hydrazide (Royal MH-30, 180 g liter-1, Arysta LifeSciences),

a systemic suckercide, was applied with a CO2-pressurized sprayer calibrated to 468 L ha-

1 with a directed three-nozzle row-1 configuration (TG3 - TG5 - TG3). Chemically topped

and manually topped treatments were applied at the pre-bud (10% button) stage. Button

percentage was calculated by dividing the total number of plants in the two center rows of

each plot by the number of plants with a visible terminal bud between the apical leaves, or

growth stage 51 (Coresta Guide #7, 2009). The axillary (AxB) and apical (ApB) meristems

were collected 24 h after treatment from the control (not-topped, not-sprayed), chemically-

topped (not-topped, sprayed with MH), topped without sprayed, and the grower standard

(topped and sprayed with MH) and were frozen immediately in liquid nitrogen and stored

at −80 °C until RNA extraction.

69

Total RNA was isolated from 100 mg of AxB and ApB tissues using the RNeasy

Plant Mini Kit (Qiagen, Chatsworth, USA) following manufacturer’s instructions. A

NanoDrop ND-1000 spectrophotometer (NanoDrop Technologies, Wilmington, DE, USA)

was used to determine RNA quantity. RNA quality was determined using Agilent 2100

Bioanalyzer (Agilent Technologies, Palo Alto, CA, USA) and only samples with RNA

integrity number (RIN) above eight were used for library preparation. Two micrograms

from each RNA sample was sent to the Sequencing and Genomic Technologies Shared

Resource facility at Duke University for RNA-Seq library preparation and sequencing.

Data processing and gene expression quantification

Raw Illumina sequence reads were processed as described previously (Singh et al.,

2015). In summary, raw Illumina sequence reads were filtered for low-quality reads using

the prinseq-lite-0.20.426 (Schmieder and Edwards, 2011). The preprocessed reads were

assessed for quality control with systemPipeR (Backman and Girke, 2016). Read mapping

was performed by Bowtie2 (Langmead and Salzberg, 2012) using the reference sequence

downloaded from Solgenomics Network database (Bombarely et al., 2011; Sierro et al,

2014). Finally, differential gene expression analysis was carried out using the DESeq2

Bioconductor package in R (Love et al, 2014). Differentially expressed genes (DEGs) were

identified using the following two criteria: (i) log2 fold-change ≥ 1 and (ii) false discovery

rate (FDR) p-value correction of ≤ 0.05. The heatmap was constructed using the

ComplexHeatmap (Gu et al, 2013) function in R through the Bioconductor package (Team,

2013).

70

Functional annotation and gene ontology (GO) analysis

Functional annotation of DEGs was performed with eggNOG 4.5 (Huerta-Cepas et

al, 2016) and InterPro (Mitchell et al, 2015) databases. GO analysis of enriched functional

categories was performed using BiNGO (version 2.44) (Maere et al., 2005) and visualized

in Cytoscape (Shannon et al., 2003). The hypergeometric test with Benjamini &

Hochberg’s false discovery rate (FDR) correction was used to calculate overrepresented

GO categories among differentially expressed genes, using a P-value<0.05. Results from

the gene list analyzed using BiNGO were summarized with REViGO by removing

redundant GO terms (Supek et al., 2011). For pathway analysis, a MapMan mapping file

was specifically generated for the tobacco genes by the Mercator tool, which bins all genes

according to hierarchical ontologies after searching a variety of databases and, finally,

MapMan v.3.5.1 was used to visualize DEGs on different pathways (Thimm et al., 2004).

Quantitative RT-PCR

Gene specific primers for the six candidate genes were designed using Primer3

software3 (Untergasser et al, 2012). RNA isolated from control, topped, MH-treated

samples were reverse-transcribed using the Superscript III Reverse Transcriptase

(Invitrogen, USA), following the manufacturer’s instructions. Quantitative PCR was

performed as described by (Pattanaik et al., 2010). All PCR reactions were performed in

triplicate and repeated two times. The comparative cycle threshold (Ct) method (bulletin

no. 2; Applied Biosystems, http://www.appliedbiosystems.com) was used to measure

transcript levels. In addition to tobacco œ-tubulin (GenBank accession number AJ421411),

tobacco elongation factor-1œ (GenBank accession number D63396) was also used as a

reference gene.

71

Results and Discussion

RNA-seq analysis of MH-treated apical and axillary buds

RNA-seq analysis was performed to study the influence of MH on gene

expression in apical (ApB) and axillary buds (AxB) of chemically topped tobacco plants

and to propose a molecular mechanism of action of MH on sucker control. A total of

12 samples comprised of two different tissues (ApB and AxB) and two different

treatments (control and MH) were used for library preparation and sequencing using the

Illumina HiSeq2500 system (Table 4.1). Sequencing of RNA libraries from ApB and

AxB of tobacco generated a total of 450 million (M) clean reads (Table 4.1). Each

biological sample (control and MH-treated) was represented by an average of more than

100 M reads (Table 4.1). On average, more than 75% of the total reads from control and

MH-treated libraries were successfully mapped to the reference tobacco genome

sequence (Table 4.1).

For our analysis, we considered a transcript as ‘detected’ if the Fragments Per

Kilobase of gene per Million reads mapped (FPKM) value was ≥1. Total number of

transcripts varied from approximately 44,000 to 47,000 in all analyzed samples which were

further divided into three categories, low (1-5 FPKM), moderate (5-20 FPKM), or high

(>20 FPKM), based on transcript abundance. Both control and MH-treated samples had

similar distribution of low, moderate, and highly expressed mRNAs (Figure 4.2A). We

selected 10,000 mRNAs which are most abundant and show distinct accumulation pattern

between the two tissue and treatments for further analysis (Figure 4.2B).

72

MH-treatment significantly alters gene expression both in apical and axillary buds

Compared with the control, chemical topping (CT; untopped plants treated with

MH) significantly altered gene expression in ApB and AxB; 573 (132 upregulated, 441

downregulated) and 2,632 (2,174 upregulated, 458 down-regulated) genes were found to

be differentially expressed in chemically topped ApB and AxB, respectively (Figure

4.3A). A total of 87 genes were commonly affected by MH-treatment in both ApB and

AxB (Figure 4.3B). Among the 87 common genes, 8 were upregulated whereas 18 were

downregulated. The other 61 genes showed contra-regulation (opposite regulation) in ApB

and AxB. These commonly affected genes were enriched for meristem development and

cytokinin (CK) metabolism. The member of KNOX family and BTB-POZ domain

transcription factors were downregulated in both ApB and AxB by MH-treatment. Among

the contra-regulated genes, genes related to secondary metabolism and defense, such as

terpene synthase, pathogenesis-related (PR) genes, chitinase and hormone biosynthesis

(jasmonate and ethylene biosynthetic genes), were upregulated in MH treated AxB but

downregulated in ApB. Genes related to translation (such as member of ribosomal

protein L22p/L17e family) were upregulated in MH treated ApB only.

Gene ontology (GO) enrichment analysis highlights influence of MH on different

developmental and metabolic pathways in ApB and AxB

To gain further insight into the implications of MH-treatment, we performed GO

enrichment analysis on genes that were upregulated and downregulated both in ApB and

AxB. In MH-treated ApB, upregulated genes were enriched for phosphorelay signal

transduction system (GO:0000160), leaf proximal/distal pattern formation

(GO:0010589), regulation of timing of transition from vegetative to reproductive phase

73

(GO:0048510) and vernalization (GO:0010048) while GO terms related to meristem

maintenance, cytokinin metabolism, cell wall synthesis, photosynthesis and DNA

metabolism were enriched in downregulated genes (Figure 4.4A). In MH-treated AxB,

GO terms related to defense response and oxylipin metabolism were enriched in

upregulated genes whereas GO terms related to cell cycle and DNA metabolism, cytokinin

metabolism were enriched in downregulated genes (Figure 4.4B). Genes from several

protein families such as mini-chromosome maintenance (MCM2/3/5) family protein,

origin recognition complex (ORC) proteins family and cell division control protein which

are essential for initiation of DNA replication were downregulated (Shultz et al., 2007).

Genes with known function in DNA repair and recombination such as Replication Protein

A 1B, RAD21.2, ARABIDOPSIS HOMOLOG OF YEAST CDT1 A, BREAST CANCER

ASSOCIATED RING 1 (BARD1) and RECQ helicase l1 were also suppressed by MH

treatment (Singh et al., 2010). The cytokinin metabolism is affected in both ApB and AxB.

Collectively, these findings suggest that MH suppresses the DNA repair and recombination

machinery which, leads to inaccurate DNA replication and cell cycle arrest. Moreover, MH

inhibits apical and axillary shoot development possibly by affecting cytokinin metabolic

processes in tobacco.

MH perturbs expression of transcription factor genes involved in meristem

maintenance and development

Transcription factors (TFs) belonging to the R2R3MYBs, bHLH, GRAS, HD/ZIP,

KNOX and BTB/POZ families are known to play crucial role in meristem maintenance

and development in plants. The R2R3 MYB, BLIND/RAX, bHLH TF ROX, GRAS family

TF LAS and HD/ZIP TF REV are positive regulators of axillary meristem development in

74

Arabidopsis, tomato and pepper. Our transcriptome analysis revealed that expression of

these TFs were downregulated in ApB and AxB after MH treatment (Figure 4.5). The

KNOX genes comprise a small family of TALE homeobox TFs that are found in all plant

species and can be divided into two major subclasses (Gao et al., 2015). Class I KNOX

genes are most similar to maize knotted1 (kn1) gene and are predominantly expressed in

the shoot apical meristem (SAM) (Hake et al., 2004; Gao et al., 2015), whereas Class II

KNOX genes show diverse expression patterns (Gao et al., 2015). KNOX ARABIDOPSIS

THALIANA MEINOX (KNATM) genes are relatively new members of the KNOX

family that encodes a MEINOX domain but not a homeodomain. In Arabidopsis, KNATM

is expressed in proximal-lateral domains of organ primordia and at the boundary of mature

organs and is involved in leaf proximal-distal patterning (Magnani and Hake, 2008). We

identified 19 members of the KNOX family in tobacco and phylogenetic analysis revealed

three major clades, the KNOX I, KNOX II and KNATM, as described previously (Gao et

al., 2015) (Figure 4.6A). MH-treatment repressed the expression of most of the members

of class I KNOX genes in ApB and AxB (Figure 4.6B), whereas members of KNOX II

subfamily were not significantly affected. In Arabidopsis, the Class I KNOX gene SHOOT

MERISTEMLESS (STM) has been shown to play key role in shoot and floral meristem

maintenance (Endrizzi et al., 1996). In addition, STM is shown to activate CK biosynthesis

genes and, consequently, CK accumulation (Yanai et al., 2005). Therefore, it is reasonable

to hypothesize that MH affects CK accumulation in ApB and AxB by repressing the class

I KNOX genes. Unlike typical KNOX family members, KNATM encodes a MEINOX

domain without homeodomain and interacts with TALE-class homeodomain proteins to

modulate their activities (Magnani and Hake, 2008). We identified two KNATM family

75

members in tobacco and both copies of KNATM were upregulated in ApB but not in AxB

(Figure 4.6B).

In MH-treated ApB, expression of a number of MADS-box family TFs were

downregulated (Figure 4.7A). The members of the MADS-box family are well known for

their function in flower and fruit development (Ng and Yanofsky, 2001; Theißen et al.,

2016). MADS-box family genes regulate both flowering time and vegetative to

reproductive phase transition (Borner et al., 2000; Putterill et al., 2004). The vegetative to

reproductive phase transition in plants is accurately controlled by environmental conditions

and endogenous developmental cues. In Arabidopsis, flowering has been proposed to be

regulated by four genetic pathways, photoperiod, autonomous, vernalization, and

gibberellin induced pathways (Boss et al., 2004; Bäurle and Dean, 2006). The current

ABCDE model for flower development proposes that floral organ identity is specified by

five classes of homeotic genes, A (APETALA1, AP1), B (PISTILATA, PI), C

(AGAMOUS, AG), D (SEEDSTICK/AGAMOUS-LIKE11,STK/AGL11) and E

(SEPALLATAs, SEPs) (Rijpkema et al., 2010). Different combinations of these homeotic

genes determine the identities of the floral organs: sepals (A + E), petals (A + B + E),

stamens (B + C + E), carpels (C + E), and ovules (D + E). In Arabidopsis, most of the

members of class A, B, C, D, E belong to the MADS-box TF family. Since a large number

of MADS-box genes were differentially expressed in MH-treated ApB, we looked into the

expression of the homologs of well characterized MADS- box family members in our

transcriptome. SEPALLATA1 (SEP1), SEP2 and SEP3, and SEP4 are required to specify

petals, stamens, and carpels (Ditta et al., 2004). We found that expression of several

homologs of SEPs were repressed by MH application. In addition, expression of genes

76

required for floral organ identity such as PISTILLATA, AGAMOUS, AP1, and AP3 were

also repressed by MH treatment. In Arabidopsis, AGAMOUS-like 22 (AGL22) regulates

flowering time by negatively regulating the expression of the floral integrator, FT, via

direct binding to the CArG motifs in the FT promoter region (Lee et al., 2007). AGL22

was induced in ApB by MH-treatment (Hartmann et al., 2000). A MADS-box TF, AGL6,

which is reported to be a positive regulator of axillary meristem formation and flowering

is also repressed by application of MH (Koo et al., 2010; Huang et al., 2012). In

Arabidopsis, the BTB/POZ domain TFs BLADE ON PETIOLE 1 (BOP1) and BOP2, act

redundantly to control leaf and floral patterning by modulating the meristematic activity.

BOP2 is highly expressed in young floral meristem (Xu et al., 2010). Expression of BOP2

was repressed by MH-treatment. Barley homolog (Cul4) of Arabidopsis BOP2 has also

been shown to express in axil and leaf boundary regions to positively control axillary bud

(Tavakol et al., 2015). Downregulation of multiple MADS-box and BTB/POZ family

genes after MH application was consistent with previous results that MH treatment delays

or inhibits the flower initiation in several plants including tobacco (Naylor, 1950; Klein

and Leopold, 1953).

MH-treatment induces defense and secondary metabolism genes in axillary buds

We found that TFs belonging to WRKY, AP2/ERF and NAC families were mostly

affected in MH-treated AxB (Figure 4.7B). WRKYs are well studied plant-specific TFs

which are involved in diverse biotic and abiotic stress responses as well as in

developmental/physiological processes (Phukan et al., 2016). Expression of several

WRKYs TF genes including WRKY2, WRKY6, WRKY7, WRKY11, WRKY23, WRKY28,

WRKY33, WRKY38, WRKY40, WRKY41, WRKY45, WRKY50, WRKY51, WRKY53,

77

WRKY70 were found to be upregulated in AxB in response to MH treatment. Previous

studies suggests that in Arabidopsis, WRKY50 and WRKY51 act as positive regulators of

SA-mediated signaling and negative regulators of JA signaling (Gao et al., 2011) whereas

WRKY28 and WRKY70 are involved in both SA and JA-signaling pathways in plants (Li

et al., 2004; Chen et al., 2013). WRKY33 is a key regulator of camalexine biosynthesis

and is required for resistance to necrotrophic fungal pathogens in Arabidopsis (Zheng et

al., 2006; Liu et al., 2016). Notably, WRKY13, which is known to activate lignin

biosynthesis-related genes (Li et al., 2015) and repress flowering (Li et al., 2016) in

Arabidopsis, was upregulated by MH treatment in AxB.

MH-treatment affects expression of phytohormone biosynthesis and signaling genes

in apical and axillary buds

Phytohormones play a crucial role in ApB and AxB development (Yang and Jiao,

2016). We identified phytohormone metabolism and signaling related genes in tobacco as

described previously (Prasad et al., 2016). Genes related to biosynthesis and signaling of a

number of phytohormones including CK, SA, JA, ethylene (ET), abscisic acid (ABA), and

gibberellic acid (GA), were affected by MH-treatment in ApB and AxB of tobacco (Tables

4.2-4.3). RNA-seq analysis revealed that compared with the control, 20 and 57 genes

related to phytohormone metabolism and signaling were significantly differentially

expressed in CT-ApB and CT-AxB, respectively. In addition, seven differentially

expressed genes were common to CT-ApB and CT-AxB. These seven genes belong to ET,

JA and CK metabolism, which are downregulated in AxB but upregulated in ApB. Among

seven common genes, 4 are homologs of Arabidopsis ACO4, key gene in ET biosynthetic

pathway. Two (i.e. AOS) are related to JA biosynthesis. One gene belongs to UDP-

78

glycosyltransferase superfamily and was also common between MH-treated ApB and AxB.

UDP-glycosyltransferase superfamily mediate the transfer of glycosyl residues from

activated nucleotide sugars to acceptor molecules (such as hormone) and thus regulate the

homeostasis (Ross et al., 2001). CT-AxB specific genes are related to SA, JA, GA and ET

metabolism (Table 4.3). Interestingly, genes related to ET biosynthesis including 1-

amino-cyclopropane-1-carboxylate (ACC) synthase and ACC oxidase were upregulated by

MH-treatment in AxB. ET induces the expression of ET signaling pathway genes such as

members of ET insensitive (EIN) family. In CT-AxB dataset, EIN homologs of

Arabidopsis, EIN3, was upregulated. ET is known to inhibit cell division, DNA synthesis,

and growth of AxB.

GAs play fundamental roles in plant growth and development. Three classes of

enzyme, i.e. terpene synthases (TPSs), CYP450s and GA oxidases (GAoxs), are required

for the biosynthesis of bioactive GAs from geranylgeranyl diphosphate (GGDP), and the

pathway can be divided into two main steps. The early steps are catalyzed by a series of

genes encoding enzymes such as ENT-COPALYL DIPHOSPHATE SYNTHASE (CPS),

ENT-KAURENE SYNTHASE (KS), ENT-KAURENE OXIDASE (KO), and

ENTKAURENOIC (KAO). The enzymes catalyzing later steps, such as GA2 oxidase

(GA2ox), GA20 oxidase (GA20ox), and GA3 oxidase (GA3ox), belong to the 2OG-Fe (II)

oxygenase superfamily and are encoded by different gene families (Hedden and Phillips,

2000). The genes involved in the later steps of GA biosynthesis are differentially regulated

by developmental and environmental cues and play crucial but antagonistic roles in the

accumulation of bioactive GA levels. For instance, upregulation of GA20ox and GA3ox

increase the GA level whereas higher expression of GA2ox decreases the GA level

79

(Schomburg et al., 2003; Lo et al., 2008). GA is involved in AxB development in different

plants including tomato, rice and aspen (Lo et al., 2008; Martínez-Bello et al., 2015; Rinne

et al., 2016). In rice, GA negatively regulates expression of two TFs, homeobox 1 and

TEOSINTE BRANCHED1 (TB1), which control meristem initiation and AxB outgrowth,

respectively, and inhibits tillering (Lo et al., 2008). Two homologs of Arabidopsis GA2ox

were found to be upregulated in response to MH-treatment in AxB in our dataset (Table

4.3), which likely lowered the concentration of GA and inhibited AxB development.

MapMan visualization highlights the influence of MH-treatment on different

plant metabolic pathways

Pathway-based analysis was performed to associate biological functions with the

genes differentially expressed in response to MH treatment. We used a comprehensive

tool, the MapMan, to visualize the pathways affected by MH-treatment in ApB and AxB

tissues in tobacco. We overlaid the log2 fold change of DEGs to identify and visualize

affected pathways. The number of genes in AxB affected by MH treatment were

significantly higher compared to ApB, indicating a broader impact of MH on AxB.

Genes related to defense such as secondary metabolites, proteolysis, pathogenesis

related genes, and heat shock protein were downregulated in MH-treated ApB (Figure

4.8). However, unlike ApB, genes related to defense pathway and hormone biosynthesis

were upregulated in AxB by MH treatment (Figure 4.9). In AxB, several genes in the

JA biosynthesis pathway such as lipoxygenase and allene oxidase were upregulated in

response to MH treatment. Genes related to auxin homeostasis (IAA-amino acid

hydrolase and GH3 family), ethylene biosynthesis and signaling (ethylene responsive

factor1 (ERF1), ERF2, ERF5, ERF4, 1-aminocyclopropane-1-carboxylate oxidases and

80

1-aminocyclopropane-1-carboxylate synthase) were also induced by MH treatment in

AxB. Plant glutathione S-transferases (GSTs) comprise a large, complex gene family in

plants. For instance, there are 25 GST genes in Glycine max, 42 in Zea mays, and 47 in

Arabidopsis thaliana. Plant GST gene family are divided by sequence similarity into

three categories (I, II, and III) or, alternatively, into six classes (tau, phi, zeta, theta,

lambda, and DHAR), with the tau and phi classes being prevalent. GSTs catalyze the

conjugation of toxic xenobiotics and oxidatively produced compounds to reduced

glutathione, which further facilitates their metabolism, sequestration, or removal

(Dalton et al., 2009). Expression of GSTs are also known to be induced by auxin and

ethylene in plants including tobacco (Van der Zaal et al., 1991; Itzhaki and Woodson,

1993; Droog et al., 1995; van der Kop et al., 1996). Unlike ApB, several homologs of

auxin-responsive GSTs were found to be induced by MH treatment in AxB (Figure 4.9).

Pathogenesis-related (PR) proteins play numerous roles in plant development and defense.

The PR proteins are highly conserved proteins and have been classified into 17 classes

based on their amino acid sequence, serological relationship, and biological activities (Van

Loon and Van Strien, 1999). PR proteins are involved in plant immune responses (Stintzi

et al., 1993) and enhance plants tolerance to both biotic and abiotic stresses (Wu et al.,

2016). For instance, overexpression of PR proteins, such as PR-1, PR-5, or PR-10, in plants

enhances tolerance to a number of pathogens such as Rhizoctonia solani, Phytophthora

nicotianae, Ralstonia solanacearum, and Pseudomonas syringae (Datta et al., 1999;

Sarowar et al., 2005). Those PR proteins have also been reported to have multiple roles in

adaption to abiotic stresses such as salt and heavy metal tolerance (Sarowar et al., 2005; de

las Mercedes Dana et al., 2006). MH treatment was found to activate the expression of

81

several PR genes and secondary metabolite related genes in AxB (Figure 4.9). Taken

together, these results indicate that ApB and AxB respond differently to MH treatment

(Figures 4.8-4.9).

Quantitative RT-PCR analysis of selected DEGs validates the RNA-seq data

To validate the RNA-seq results, expression of six differentially expressed genes

were analyzed by qRT-PCR (Figure 4.10). A list of primers used to conduct RT-qPCR

analysis is shown in Table 4.4. These genes encode the KNOX (KNOX1 and KNOX12)

genes, AGL (AGL6 and AGL20) genes, 1-aminocyclopropane-1-carboxylate oxidase 4

(ACO4) and SALICYCLIC ACID CARBOXYL METHYLTRANFERASE (SAMT). The

qRT-PCR results complemented the RNA-seq data, confirming the reliability and

accuracy of our RNA-seq in this study.

In summary, MH has profound influence on gene expression in ApB and AxB of

tobacco. The number of differentially expressed genes were higher in AxB compared to

ApB. In both ApB and AxB, the expression of genes related to a number of

phytohormones, meristem development, cell division, DNA repair and recombination

were affected following MH treatment, which likely leads to the inhibition of apical and

axillary shoot growth. In addition, MH elicits defense responses in plants by inducing

the expression genes involved in oxylipin biosynthesis, secondary metabolism and

defense-related genes. In addition, MH-treatment induces the expression of a number of

GSTs, which are possibly involved in detoxification processes. Collectively, our RNA-

seq analysis reveals a possible molecular mechanism of action of MH on apical and

axillary buds of tobacco (Figure 4.11).

82

Table 4.1. Summary of sequencing and read mapping in different RNA-seq libraries.

Treatment Symbol Experiment Total raw

reads

Total clean

reads

Total raw

reads per

biological

sample

Total clean reads

mapped to

reference

transcriptome

-------------- million -------------- ---- % ----

Control-

axillary

bud

C-AxB

1 42 36

109

76.47

2 44 39 75.9

3 40 34 73.96

Control-

apical bud C-ApB

1 51 31

118

81.41

2 48 42 78.9

3 52 45 79.06

Chemically

topped

-axillary

bud

CT-

AxB

1 46 40

111

76.06

2 38 32 76.26

3 45 39 75.8

Chemically

topped

-apical bud

CT-

ApB

1 36 31

112

80.38

2 49 43 79.83

3 43 37 79.58

83

Table 4.2. Expression of different phytohormone biosynthesis and signaling genes in MH-treated apical

buds of tobacco.

Gene Hormone Function

Best

Arabidopsis

match

Gene

symbol

ApB

(Log2FC)

AxB

(Log2FC)

Nitab4.5_0007480g0030.1 ABA signaling AT4G34000 ABF3 -1.09 0.85

Nitab4.5_0001924g0060.1 ABA biosynthesis AT3G14440 NCED3 -1.65 0.76

Nitab4.5_0000525g0140.1 Auxin signaling AT3G23050 IAA7 1.64 -0.30

Nitab4.5_0004203g0040.1 Auxin signaling AT4G14550 IAA14 2.03 -0.01

Nitab4.5_0007788g0010.1 Auxin biosynthesis AT4G28720 YUC8 -1.31 -0.86

Nitab4.5_0001119g0070.1 Auxin signaling AT2G14960 GH3.1 -1.58 0.35

Nitab4.5_0002208g0100.1 CK signaling AT3G57040 ARR9 1.03 -0.61

Nitab4.5_0000026g0340.1 CK signaling AT5G62920 ARR6 1.41 -0.27

Nitab4.5_0002818g0060.1 CK biosynthesis AT2G36780 -1.23 3.11

Nitab4.5_0000130g0140.1 ET biosynthesis AT1G05010 EFE -1.41 2.05

Nitab4.5_0004330g0020.1 ET biosynthesis AT1G05010 EFE -1.57 2.98

Nitab4.5_0000130g0130.1 ET biosynthesis AT1G05010 EFE -1.48 2.11

Nitab4.5_0004330g0030.1 ET biosynthesis AT1G05010 EFE -1.09 2.28

Nitab4.5_0004821g0040.1 ET biosynthesis AT1G12010 -1.27 -0.38

Nitab4.5_0003887g0030.1 GA biosynthesis AT1G05160 CYP88A3 -1.05 0.02

Nitab4.5_0008507g0010.1 JA biosynthesis AT5G42650 AOS -1.31 1.30

Nitab4.5_0003281g0080.1 JA biosynthesis AT5G42650 AOS -1.23 2.24

Nitab4.5_0000571g0010.1 SA signaling AT2G41370 BOP2 -1.23 -0.50

Nitab4.5_0003771g0010.1 SA signaling AT2G14580 ATPRB1 -1.57 0.94

Nitab4.5_0001066g0090.1 SA biosynthesis AT3G11480 BSMT1 -2.30 0.43

84

Table 4.3. Expression of different phytohormone biosynthesis and signaling genes in MH-treated axillary buds

of tobacco.

Gene Hormone Function

Best

Arabidopsis

match

Gene

symbol

ApB

(Log2FC)

AxB

(Log2FC)

Nitab4.5_0009622g0010.1 ABA signaling AT2G29380 HAI3 0.00 1.12

Nitab4.5_0000463g0070.1 ABA signaling AT2G29380 HAI3 0.14 1.11

Nitab4.5_0006647g0010.1 ABA biosynthesis AT3G29250 0.32 1.45

Nitab4.5_0000635g0100.1 Auxin signaling AT4G03400 DFL2 0.32 1.06

Nitab4.5_0003159g0030.1 Auxin signaling AT3G15540 IAA19 -0.92 -1.31

Nitab4.5_0003885g0020.1 Auxin signaling AT3G15540 IAA19 -0.48 -1.38

Nitab4.5_0007761g0010.1 Auxin signaling AT2G21210 0.06 -1.34

Nitab4.5_0006273g0010.1 Auxin biosynthesis AT5G05260 CYP79A2 -0.05 1.91

Nitab4.5_0000604g0030.1 Auxin signaling AT2G14960 GH3.1 -0.90 4.24

Nitab4.5_0004933g0020.1 Auxin biosynthesis AT5G56660 ILL2 -0.24 2.15

Nitab4.5_0000996g0050.1 CK signaling AT1G27320 AHK3 -0.60 1.14

Nitab4.5_0014466g0010.1 CK biosynthesis AT2G36760 UGT73C2 -0.40 2.82

Nitab4.5_0006222g0020.1 CK biosynthesis AT1G22380 UGT85A3 0.04 1.35

Nitab4.5_0000601g0080.1 CK biosynthesis AT2G36780 -0.87 2.51

Nitab4.5_0002818g0060.1 CK biosynthesis AT2G36780 -1.23 3.11

Nitab4.5_0000600g0080.1 ET signaling AT4G17500 ERF-1 -0.07 1.99

Nitab4.5_0000130g0140.1 ET biosynthesis AT1G05010 EFE -1.41 2.05

Nitab4.5_0004330g0020.1 ET biosynthesis AT1G05010 EFE -1.57 2.98

Nitab4.5_0000130g0130.1 ET biosynthesis AT1G05010 EFE -1.48 2.11

Nitab4.5_0004330g0030.1 ET biosynthesis AT1G05010 EFE -1.09 2.28

Nitab4.5_0000915g0150.1 ET biosynthesis AT1G01480 ACS2 0.36 1.46

Nitab4.5_0009635g0010.1 ET biosynthesis AT1G05010 EFE -0.41 2.06

Nitab4.5_0002687g0110.1 ET signaling AT3G20770 EIN3 0.24 1.02

Nitab4.5_0002236g0020.1 ET signaling AT4G17500 ERF-1 -0.16 2.33

Nitab4.5_0007571g0020.1 ET signaling AT3G23240 ERF1 -0.25 1.82

Nitab4.5_0002211g0030.1 ET signaling AT4G17500 ERF-1 -0.13 2.27

Nitab4.5_0003058g0050.1 GA signaling AT3G63010 GID1B -0.84 1.86

Nitab4.5_0011064g0010.1 GA signaling AT3G63010 GID1B -0.52 1.43

85

Nitab4.5_0012276g0010.1 GA biosynthesis AT1G30040 GA2OX2 -0.23 1.49

Nitab4.5_0008288g0010.1 GA biosynthesis AT1G30040 GA2OX2 -0.22 2.96

Nitab4.5_0000343g0360.1 JA signaling AT1G19180 JAZ1 -0.03 1.25

Nitab4.5_0004234g0080.1 JA signaling AT1G19180 TIFY10A -0.13 2.34

Nitab4.5_0001799g0060.1 JA biosynthesis AT1G76690 OPR2 0.21 2.19

Nitab4.5_0007594g0020.1 JA signaling AT2G46370 JAR1 0.01 2.04

Nitab4.5_0000305g0060.1 JA signaling AT2G46370 JAR1 -0.70 2.40

Nitab4.5_0000073g0270.1 JA signaling AT1G19180 JAZ1 0.22 2.32

Nitab4.5_0002898g0030.1 JA biosynthesis AT1G17420 LOX3 -0.02 2.22

Nitab4.5_0002262g0110.1 JA signaling AT2G46370 JAR1 -0.26 1.01

Nitab4.5_0006391g0020.1 JA biosynthesis AT1G17420 LOX3 0.28 1.63

Nitab4.5_0000240g0150.1 JA biosynthesis AT1G67560 LOX6 0.11 1.02

Nitab4.5_0000110g0020.1 JA biosynthesis AT5G42650 AOS 0.18 2.10

Nitab4.5_0008239g0010.1 JA biosynthesis AT5G42650 AOS -0.02 1.91

Nitab4.5_0009125g0010.1 JA signaling AT2G46370 JAR1 -0.02 1.07

Nitab4.5_0001546g0010.1 JA signaling AT3G17860 JAZ3 0.20 1.06

Nitab4.5_0008507g0010.1 JA biosynthesis AT5G42650 AOS -1.31 1.30

Nitab4.5_0003281g0080.1 JA biosynthesis AT5G42650 AOS -1.23 2.24

Nitab4.5_0002889g0090.1 JA biosynthesis AT1G19640 JMT -0.11 1.38

Nitab4.5_0002574g0030.1 SA signaling AT5G45110 NPR3 0.00 1.09

Nitab4.5_0004861g0040.1 SA signaling AT4G33720 -0.78 1.31

Nitab4.5_0005030g0030.1 SA signaling AT5G45110 NPR3 0.05 1.16

Nitab4.5_0003642g0050.1 SA signaling AT1G68640 PAN 1.03 -1.14

Nitab4.5_0006853g0020.1 SA signaling AT5G06839 TGA10 -0.40 2.04

Nitab4.5_0012788g0010.1 SA signaling AT3G12250 TGA6 0.08 1.37

Nitab4.5_0002920g0050.1 SA biosynthesis AT3G11480 BSMT1 -0.40 5.88

Nitab4.5_0009504g0020.1 SA biosynthesis AT5G04370 NAMT1 0.08 1.60

Nitab4.5_0000198g0050.1 SA biosynthesis AT1G68040 0.46 1.37

Nitab4.5_0003904g0010.1 SA biosynthesis AT3G11480 BSMT1 -0.46 7.28

86

Table 4.4. Primers used to conduct RT-qPCR analysis.

Gene name Forward primer Reverse primer

------------------------- (5' - 3') --------------------------

AGL6 AGAGGTACCAACGTTGTTGC TCACCAAGCAAGTGCCTTTG

AGL20 CTTCTCAAAGCGCCGGAATG AGTTGGAGCTAGCGAAATCG

ACO4 CCAGCAAAGGTCTTGAAGCTG ATGGCGCAAGAAGAAAGTGC

SAMT ATTGCGGACTTAGGTTGCTC ATTCCGGCGACTGTTTTTGG

KNOX1 AGGAAGCAAGGCAACAACTG ATTCAGCAAGTGCCAGCTTC

KNOX12 TGCAAGAAACAGGTCTGCAG ACGTCGATGGATTGCTATGC

87

Figure 4.1. Schematic diagram of the experimental design for chemical topping and gene expression analysis. C-ApB,

control apical bud; CT-ApB, chemically topped apical bud; C-AxB, control axillary bud; CT-AxB, chemically topped axillary

bud.

88

Figure 4.2. Overview of RNA sequencing analysis (A) Distribution of FPKM normalized transcripts across the four treatments.

(B) Clustered heat-map of top 10,000 highly abundant mRNAs. C-ApB, control apical bud; CT-ApB, chemically topped-apical

bud; C-AxB, control axillary bud; CT-AxB, chemically topped-axillary bud.

89

Figure 4.3. Differentially expressed genes in maleic hydrazide (MH)-treated apical and axillary buds. (A) Number of

upregulated (Red) and downregulated (Blue) genes. (B) Venn diagram depicting the overlap of differentially expressed genes

(DEGs) between MH-treated apical and axillary bud. CT-ApB, chemically topped-apical bud; CT-AxB, chemically topped-

axillary bud.

90

91

Figure 4.4. Gene Ontology (GO) analyses of DEGs in maleic hydrazide (MH)-treated apical and axillary buds. GO analysis

of DEGs in apical (A) and axillary bud (B). Upregulated terms are colored in ‘red’ while downregulated terms are in ‘blue’.

Each circle represents one GO term. Circle size represents the number of genes in each GO category while color represents the

significance level.

92

Figure 4.5. Effect of maleic hydrazide on expression of key transcription factors involved in apical and axillary bud

development. LAS, Lateral suppressor; RAX, Regulator of axillary meristem; ROX, Regulator of axillary meristem formation;

REV, Revoluta; CT-ApB, chemically topped-apical bud; CT-AxB, chemically topped-axillary bud.

93

Figure 4.6. Phylogenetic and gene expression analysis of KNOX gene family from tobacco. (A) A neighbor-joining

phylogenetic tree of members of KNOX gene family from Arabidopsis thaliana and Nicotiana tabacum (tobacco) was

constructed using ClustalX and MEGA7.0 software with 1000 bootstraps. Nodes belong to A. thaliana are represented by ‘blue’

circles while ‘red’ circles represent the genes from tobacco. (B) Heat map showing the FKPM values KNOX genes obtained by

RNA-seq analysis. Rows are probes and columns are samples. The differential expression of each class of KNOX genes is

annotated in the right bar. NSC, not significantly changed.

94

Figure 4.7. Differentially expressed transcription factor (TF) genes in chemically topped axillary and apical buds. The X-

axis represents the names of differentially expressed TF families and Y-axis indicates the number of transcription factors. (A)

apical bud (B) axillary bud.

95

Figure 4.8: MapMan visualization of differential gene expression in chemically topped apical bud compared with

control. Each dot denotes a gene. ‘Blue’ color indicates downregulation while ‘red’ upregulation. The log2 fold changes of

significantly differentially expressed genes were imported and visualized in MapMan for the chemically topped apical bud

sample with regard to pathogen/pest attack.

96

Figure 4.9: MapMan visualization of differential gene expression in chemically topped axillary bud compared with

control. Each dot denotes a gene. ‘Blue’ color indicates downregulation while ‘red’ upregulation. The log2 fold changes of

significantly differentially expressed genes were imported and visualized in MapMan for the chemically topped axillary bud

sample with regard to pathogen/pest attack.

97

Figure 4.10. Validation of RNA-seq results using quantitative real-time PCR (qRT-PCR). Six differentially expressed genes

were selected for qRT-PCR. Tobacco tubulin was used an internal control for normalization. Data represents mean±SD of three

biological replicates.

98

Figure 4.11. A model depicting the effects of maleic hydrazide on different developmental and metabolic processes in

apical (ApB) and axillary (AxB) buds of tobacco. Solid arrows represent positive regulation; solid T-bars represents negative

regulation. Dashed arrow or T-bars represent possible regulation through combined effects of up- or down-regulated genes.

99

Conclusion

This research has shown that chemical topping burley tobacco at 10% button stage

with a tank mixture of MH and a local systemic suckercide was a suitable alternative to

manual topping as sucker control, total yield, and leaf quality grade index were not

different between manually topped and chemically topped tobacco (Chapter 2 and 3).

However, applications of a local systemic or fatty alcohol alone did not inhibit the terminal

bud or control sucker growth, resulting in reduced yield. Chemical topping with MH alone

did not provide adequate sucker control and equivalent yields when compared to manual

topping in all years and locations of these studies. MH residues for chemically topped

tobacco were not consistently different from residues from manually topped and sprayed

tobacco, and often were observed to be lower within an environment. Total TSNA was not

increased due to chemically topped treatments, and at Lexington there was a significant

reduction in total TSNA compared to manually topping, a similar result was also shown in

nicotine content. Future work should further investigate these total TSNA and nicotine

content reductions that were observed. Chemical topping has the potential to reduce labor

input and production costs without negatively impacting the yield, quality or chemistry of

burley tobacco (Chapter 2).

Applications of MH plus Butralin at 10% button (pre-bud) to 50% button (early-

bud) was found to be an ideal application timing for applying suckercides to chemically

top burley tobacco as applications at 10% bloom did not completely halt the development

of reproductive growth (Chapter 3). Most chemically topped application timings included

in these experiments provided similar sucker control, total yield, and leaf quality compared

to manually topping. Chemically topped treatments also appeared to have shorter tip leaves

which may contribute to an increased amount of marketable tip grades compared to

100

manually topping. Later maturing varieties may be better suited for chemical topping due

to less rapid change from vegetative to reproductive growth, which would result in a wider

window for making chemical topping applications at the most appropriate timings.

MH has a profound influence on gene expression in apical and axillary buds of

tobacco (Chapter 4). The number of differentially expressed genes were higher in

axillary buds compared to apical buds. Expression of genes related to a number of

phytohormones, meristem development, cell division, DNA repair and recombination

were affected following MH treatment in both apical and axillary buds, which likely

leads to the inhibition of apical and axillary shoot growth. In addition, MH elicits

defense responses in plants by inducing the expression genes involved in oxylipin

biosynthesis, secondary metabolism and defense-related genes. Collectively, RNA-

sequencing analysis may have revealed a possible molecular mechanism of action of

MH on apical and axillary buds of tobacco.

Chemical topping is a viable labor saving alternative to manual topping without

negatively affecting the yield, quality, or chemistry of burley tobacco.

101

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Mitchell D. Richmond

Vita

EDUCATION

University of Kentucky 2014-2018

PhD Integrated Plant and Soil Science emphasis in Crop Science

University of Kentucky 2012-2014 M.S. Crop Science

Thesis: Evaluation of Correlation between Within-Barn Curing Environment and

TSNA Accumulation in Dark Air-Cured Tobacco.

Morehead State University 2008-2012 B.S. Agriculture, Education, Communication and Leadership

West Carter High School, Olive Hill, Kentucky 2004-2008

RESEARCH/WORK EXPERIENCE

Team Leader, Canadian Tobacco Research Foundation 2018

Graduate Research Assistant PhD Integrated Plant and Soil Science 2014-2018

Graduate Research Assistant M.S. Crop Science 2012–2014

Work Study, Derrickson Agricultural Complex, Morehead, Kentucky 2009–2012

Work Study, IKON Document Services, Morehead, Kentucky 2008-2009

Assistant Farm Manager/Labor, Paradise Valley Farms, Grayson, Kentucky2005–2012

TEACHING, ADVISING, and MENTORSHIP

Guest lecturer, Senior Seminar, Morehead State University 2017

Guest lecturer, Integrated Weed Management, University of Kentucky 2017

Guest lecturer, Senior Seminar, Morehead State University 2016

Teaching Assistant, University of Kentucky, Integrated Weed Management 2016

Guest lecturer, Senior Seminar, Morehead State University 2015

Integrated Plant and Soil Sciences Graduate Student Association 2012-2018

Agriculture Education Student Teaching Practicum, West Carter High School 2012

125

SERVICE AND RECOGNITION

Gerald O. Mott Meritorious Graduate Student in Crop Science, University of

Kentucky 2018

Graduate Senator-at-Large, Student Government, University of Kentucky 2017

President, UK IPSS GSA 2017

Summer Senate, Student Government Association, University of Kentucky 2017

Executive Board, Graduate Senator-at-Large, Graduate Student Congress 2017

Plant and Soil Science Department Head Search Committee, University of Kentucky 2017

Certified Non-Commercial Pesticide Applicator, Category 10 R&D 2017

Department of Plant and Soil Science, Seminar Committee 2016

Plant and Soil Science Graduate Student Spotlight 2016

National FFA Convention Judge, Agriscience Fair-Plant Systems 2015

Gamma Sigma Delta, the Honor Society of Agriculture. 2013-2015

Carter County Fair Board 2005-2018

Science Fair Judge for Fayette County Schools 2014

UK Graduate Student Weed Science Team 2013

Morehead State University Outstanding Agricultural Sciences Student 2012

Agriculture Ambassadors, Senior Coordinator 2009-2012

Morehead State University FFA Collegiate Chapter, President 2008-2012

Morehead State University Deans List 2008-2012

Delta Tau Alpha honorary fraternity, Morehead State University 2010-2012

West Carter FFA Alumni, Treasurer 2009-2011

American FFA Degree 2011

Appleseed Scholarship 2009-2011

Farm Credit Services Scholarship 2010

Center for Regional Engagement, Gateway Homeless Shelter Grant, $1,500 2010

Kentucky State FFA Officer Candidate 2008

West Carter FFA, President 2004-2008

GRANTS/FELLOWSHIPS

Altria Client Services Graduate Fellowship Recipient. 18,000/year, tuition, and health

insurance 2017-2018

Altria Client Services Graduate Fellowship Recipient. 18,000/year, tuition, and health

insurance 2016-2017

Jeffrey’s Fellowship Recipient. 15,000/year, tuition, and health insurance. 2015-2016

CORESTA Study Grant Recipient. 15,000/year. Internationally competitive. 2012-2014

PUBLICATIONS

126

Refereed Journal Articles

Richmond, M. D., R.C. Pearce, B.M. Goff, and W.A. Bailey. 2017. Analysis of

Variability in Curing Conditions and Tobacco-Specific Nitrosamines within Barns of

Dark Air-Cured Tobacco. Tobacco Science, 54:1-9.

Richmond, M. D., R.C. Pearce, and W.A. Bailey. 2016. Dark Fire-Cured Tobacco

Response to Potassium Rate and Application Method. Tobacco Science, 53:12-15.

Bailey, W. A., T.W. Lax, R.A. Hill, and M.D. Richmond. 2013. Evaluation of Herbicide

Systems for Dark Fire-Cured Tobacco. Tobacco Science, 50:34-38.

Thesis

Richmond, M. D. 2014. Evaluation of Correlation between Within-Barn Curing

Environment and TSNA Accumulation in Dark Air-Cured Tobacco.

Abstracts Published in Proceedings

Richmond, M.D., W. A. Bailey, and R. C. Pearce. Chemically Topping Burley Tobacco.

Paper 25. Tobacco Workers Conference. January 2018. Myrtle Beach, SC, USA.

Bailey, W.A., J.C. Rodgers, A.B. Keeney, and M.D. Richmond. Evaluation of Dark

Tobacco Transplanting Intervals Following 2,4-D and Saflufenacil Herbicide

Applications. Paper 21. Tobacco Workers Conference. January 2018. Myrtle Beach, SC,

USA.

Richmond, M.D. R.C. Pearce, and W.A. Bailey. Chemically Topping Burley Tobacco.

Paper AP14. Proceedings. 2017 CORESTA Agro-Phyto Meeting. Santa Cruz do Sul,

Brazil.

Bailey, W.A., J.C. Rodgers, A.B. Keeney, and M.D. Richmond. Evaluation of Dark

Tobacco Transplanting Intervals Following 2,4-D and Saflufenacil Herbicide

Applications. Paper AP43. Proceedings. 2017 CORESTA Agro-Phyto Meeting. Santa

Cruz do Sul, Brazil.

Richmond, M.D., W. A. Bailey, and R. C. Pearce. Chemical Topping Burley Tobacco.

Paper 22. Tobacco Workers Conference. January 2016. Nashville, TN, USA.

Hill, R.A., M.D. Richmond, J.C. Rodgers, and W.A. Bailey. Dark Fire-Cured Tobacco

Response to Potassium Application Rate and Timing. Paper 5. Tobacco Workers

Conference. January 2016. Nashville, TN, USA.

Richmond, M.D., W.A. Bailey, and R.C. Pearce. 2014. Evaluation of Correlation

between Within-Barn Curing Environment and TSNA Accumulation in Dark Air-Cured

127

Tobacco. Paper AP03. Proceedings. 2014 CORESTA Agro-Phyto Meeting. Quebec

City, Canada.

Richmond, M.D., W. A. Bailey, and R. C. Pearce. Preliminary Evaluation of Correlation

between within-barn curing environment and TSNA Accumulation in Dark Air-cured

Tobacco. Paper 10. Tobacco Workers Conference. January 2014. Pinehurst, NC, USA

Hill, R.A., W.A. Bailey, J.C. Rodgers, and M.D. Richmond. Dark Tobacco Response to

Potassium Rate and Application Timing. Paper 56. Tobacco Workers Conference.

January 2014. Pinehurst, NC, USA

Bailey, W.A., J.C. Rodgers, R.A. Hill, and M.D. Richmond. Preliminary Investigation of

Common Pesticide Residues in Dark Air-Cured and Dark Fire-Cured Tobacco. Paper 13.

Tobacco Workers Conference. January 2014. Pinehurst, NC, USA

M. D. Richmond, W. A. Bailey, and R. C. Pearce. Preliminary Evaluation of Correlation

between within-barn curing environment and TSNA Accumulation in Dark Air-cured

Tobacco. Paper AP27. Proceedings. CORESTA Agro/Phyto. October 2013. Brufa di

Torgiano, Italy

Bailey, W.A., J.C. Rodgers, R.A. Hill, and M.D. Richmond. Preliminary evaluation of

cured leaf residues of common pesticides used in dark air-cured and dark fire-cured

tobacco. Paper AP06. Proceedings, 2013 CORESTA Agro-Phyto Meeting. October 2013.

Brufa di Torgiano, Italy

Popular Press

Tobacco Research Roundup. Tobacco Farm Quarterly, Summer Lookout. 2016. pg. 12-

13.

Interviewed by Tobacco Farm Quarterly magazine on September 7, 2012 describing

importance of tobacco research, current research project, and future tobacco research.

Reviewer Service

Assisted in review of articles for Tobacco Science journal.

Assisted in review of one article for American Society of Agricultural and Biological

Engineers journal.

MEETINGS AND PRESENTATIONS

Moderator, First Annual IPSS 3-Minute-Thesis 2018

128

Moderator and presenter, IPSS Graduate Student Symposium 2017

University of Kentucky Tobacco Field Day, Presenter 2017

Twilight Tour Field Day, Murray State University, Presenter 2017

Tobacco, Beef, and More Field Day, Presenter, University of Tennessee 2017

Twilight Tour Field Day, Murray State University, Presenter 2016

University of Kentucky Tobacco Field Day, Presenter 2016

University of Kentucky Research and Education Center Field Day, Presenter 2016

Woodford County Field Day, Presenter 2016

Moderator and presenter, IPSS Graduate Student Symposium 2016

Twilight Tour Field Day, Murray State University, Presenter 2015

University of Kentucky Research and Education Center Field Day, Tour Guide 2015

Evaluation of Curing Environment and TSNA Accumulation Within Barns in Dark Air-

cured Tobacco, Graduate Student Day at the Capitol. Frankfort, Kentucky 2015

TSNA Subgroup meeting, CORESTA 2014

Residue Field Trial Task Force meeting, CORESTA 2014

Moderator, Integrated Plant and Soil Science Graduate Student Symposium 2014

Evaluation of Curing Environment and TSNA Accumulation Within Barns in Dark Air-

cured Tobacco. IPSS Mini-Symposium. 2014

Progress reports and final report on Analysis of Variability in Curing Conditions and

TSNA

within Barns of Dark Air-Cured Tobacco. CORESTA Scientific Commission. 2012-2014

Tobacco Specific Nitrosamine (TSNA) Content in Dark Air-Cured Tobacco can be

Correlated to Variability in Curing Conditions. IPSS Mini-symposium. 2013

University of Kentucky Research and Education Center Field Day, Tour Guide 2013